- Open Access
Size and sequence polymorphisms in the glutamate-rich protein gene of the human malaria parasite Plasmodium falciparum in Thailand
© The Author(s). 2018
- Received: 30 September 2017
- Accepted: 8 January 2018
- Published: 22 January 2018
The glutamate-rich protein (GLURP) of the malaria parasite Plasmodium falciparum is a key surface antigen that serves as a component of a clinical vaccine. Moreover, the GLURP gene is also employed routinely as a genetic marker for malarial genotyping in epidemiological studies. While extensive size polymorphisms in GLURP are well recorded, the extent of the sequence diversity of this gene is rarely investigated. The present study aimed to explore the genetic diversity of GLURP in natural populations of P. falciparum.
The polymorphic C-terminal repetitive R2 region of GLURP sequences from 65 P. falciparum isolates in Thailand were generated and combined with the data from 103 worldwide isolates to generate a GLURP database. The collection was comprised of 168 alleles, encoding 105 unique GLURP subtypes, characterized by 18 types of amino acid repeat units (AAU). Of these, 28 GLURP subtypes, formed by 10 AAU types, were detected in P. falciparum in Thailand. Among them, 19 GLURP subtypes and 2 AAU types are described for the first time in the Thai parasite population. The AAU sequences were highly conserved, which is likely due to negative selection. Standard Fst analysis revealed the shared distributions of GLURP types among the P. falciparum populations, providing evidence of gene flow among the different demographic populations.
Sequence diversity causing size variations in GLURP in Thai P. falciparum populations were detected, and caused by non-synonymous substitutions in repeat units and some insertion/deletion of aspartic acid or glutamic acid codons between repeat units. The P. falciparum population structure based on GLURP showed promising implications for the development of GLURP-based vaccines and for monitoring vaccine efficacy.
- DNA sequence
- Genetic variation
- Plasmodium falciparum
- Glutamate-rich protein
The rapid and accurate identification of malaria parasite species is critical for clinical malaria diagnosis and subsequent treatment. A microscopic examination of the blood stage malaria on blood films is the most commonly used method in the laboratory and clinics. This technique is simple, low-cost and easy to perform, but it has many limitations, such as the requirement of high parasitemia levels and skills for diagnostic interpretation. More importantly, microscopic examination alone cannot reveal the multiplicity of an infection, which is the feature that occurs in malaria-endemic regions. Molecular tools, such as the polymerase chain reaction (PCR), may be considered as a suitable alternative. A number of PCR-based diagnostic assays have been developed and increasingly adopted for clinical malaria diagnostics [1, 2]. For identification of malaria species, house-keeping genes have been chosen as molecular targets for developing PCR assays, including the small subunit ribosomal RNA gene in the nuclear genome as well as the cytochrome c oxidase subunit 1 gene in the mitochondrial genome [3–6]. Additionally, PCR-based assays can be performed using polymorphic loci, including amplified fragment length polymorphism, microsatellites and antigen-coding genes, facilitating the identification of the parasite strains, the identification of monoclonal vs polyclonal infections, and the differentiation between relapse vs re-infections [7–11]. One of the polymorphic antigen-encoding genes often employed as a key genetic marker for strain identification of the human malaria parasite P. falciparum is the glutamate-rich protein (GLURP) gene.
The glutamate-rich protein (GLURP), a 220 kDa antigen expressed throughout the life-cycle of the malaria parasites in a mammalian host [12, 13], is localized to the Plasmodium falciparum Pfs38 protein complex on the merozoite surface, and the protein contains a module of six cysteine residues forming three intramolecular disulphide bonds. The Pfs38 complex binds to host erythrocytes directly via glycophorin A as a receptor . The GLURP protein consists of the three defined regions of an N-terminal non-repetitive region (R0), a central repetitive sequence (R1) and a C-terminal repetitive region (R2). The R0 region is predicted to be hydrophobic and may act as a signal peptide, which is a feature of secretory proteins .
Evidence that supports the role of GLURP as an immune target comes from several studies. First, immunizing mice with a vaccinia virus-expressed GLURP of P. falciparum induced a humoral immune response . To further map the immunodominant region of GLURP, four purified polypeptides, including the full-length protein and the R0, R1 and R2 regions, were produced in Escherichia coli and then tested for reactivity with sera from adults living in endemic areas in Liberia. The antibody reactivity against the R2 and R0 regions was observed in nearly 100% of the serum samples, suggesting a role for the R2 and R0 regions as the immune target . Subsequent epidemiological studies performed in endemic areas in Africa and South America consistently showed that malaria-infected patients had high levels of antibodies that react specifically with GLURP, and the levels of anti-GLURP antibodies against the R0 and R2 regions were positively associated with a reduced risk of clinical malaria [17–21]. Anti-GLURP antibodies against the R0 region act in synchronization with monocytes or mediate opsonic phagocytosis, conferring protection against febrile malaria [22, 23]. Together, these studies demonstrate that GLURP R0 and R2 may participate in eliciting protective immunity in humans, thereby giving further impetus for the development of malaria vaccines targeting GLURP.
Accordingly, P. falciparum GLURP has currently been employed in vaccine development. One example is the GMZ2 vaccine, which combines the immunodominant GLURP R0 region and MSP-3 of P. falciparum. The vaccine is designed to mimick naturally acquired anti-malarial immunity . In pre-clinical studies, the vaccine appeared to be safe and induced strong protective immune responses against the malaria parasites in simian models [25, 26]. In Phase 1 clinical trials, GMZ2 was safe, well tolerated and immunogenic in both children and adults [27–29]. Serological analyses of the antibody levels showed high levels of specific and functional antibodies in vaccinated subjects, with the capacity to control the parasite multiplication [30, 31]. Following these trials, a Phase 2 clinical trial was conducted in participants of 1 to 5 years of age in four African countries, and the GMZ2 vaccine was confirmed to be well tolerated, immunogenic and capable of reducing the malaria incidence, but the efficacy was only moderate . It is likely that the efficacy of the vaccine was compromised by the parasite antigenic diversity . Therefore, it is necessary to investigate the extent of genetic polymorphism of any vaccine candidate for malaria parasites in the natural populations where the vaccine is to be used.
The GLURP gene of P. falciparum is located on chromosome 10 and encodes a long polypeptide of 1271 amino acids (data of strain F32 from Tanzania) [12, 13]. The repetitive sequences of the R2 region of GLURP are highly conserved, but the number of repeat amino acid units (AAUs) varies among P. falciparum isolates, resulting in a size polymorphism. Hence, GLURP is routinely employed in PCR assays as a genetic tool for parasite identification and studying the multiplicity of malaria infections [34–39]. While the size polymorphism of the GLURP R2 region in P. falciparum populations is well reported, surprisingly, there is limited information on the sequence variation that underlines the polymorphisms in GLURP. To date, most of the available GLURP R2 region sequences (and their translated GLURP amino acid sequences) deposited at the NCBI nucleotide database are derived from P. falciparum isolates in India [34, 35], with a few unpublished sequences from P. falciparum from Iran, Indonesia and China. However, there are no reports of GLURP sequences in P. falciparum populations in Thailand. To address this deficiency, this study aimed to describe the detailed molecular variation in the GLURP R2 region. We generated GLURP sequences from 65 parasite isolates from five representative malaria hotspots in Thailand and combined them with the GLURP sequence data from P. falciparum isolates worldwide, thereby generating a global catalogue of GLURP and, after in silico translation, GLURP. This collection of GLURP sequences can be used to investigate the basis of molecular variations and the signature of natural selection as well as to infer the population structure of P. falciparum.
Amplification and DNA sequencing of the GLURP R2 region
The DNA templates for the PCR amplification of GLURP were extracted from 72 P. falciparum samples collected from five localities (Mae Hong Son, Kanchanaburi, Ranong, Trat and Ubon Ratchatani) in Thailand, as previously described , and shown in Additional file 1: Figure S1. The species of malaria parasites were confirmed by microscopic examinations of the thin blood film and by genotyping with the genetic markers of merozoite surface protein-1 (msp-1) and msp-3, as previously reported [40, 41]. For amplification of the R2 region of the GLURP gene, the nOF-primer (forward: 5′-CAA CCA AAT CCA CAA GAA CCA GTT GA-3′) and the R2-primer (reverse: 5′-TCT GGT TTA GTG GAT TCA CCT TCA GAT-3′) primer pair were used to yield sequences that corresponded to nucleotide positions 2215–2240 and 3441–3467 of the full-length GLURP gene of P. falciparum strain F32 (GenBank: M59706.1) . The total PCR reaction volume (50 μl) contained 200 μM of dNTPs, 2 mM of MgCl2, 0.3 μM of each primer, 200–300 ng of parasite DNA and 2.5 units of Taq polymerase enzyme in 1X Taq PCR buffer (Biotechrabbit, Hennigsdorf, Germany). The PCR conditions started with 95 °C for 5 min, followed by 40 cycles of 95 °C for 30 s, 65 °C for 30 s and 72 °C for 90 s, and then followed by a final 72 °C for 5 min. The PCR products were analysed by standard agarose gel electrophoresis in a 2% (w/v) agarose-TBE gel. The quality of the PCR products was determined under UV transillumination after exposing the gels to ethidium bromide for DNA staining. All the positive PCR products produced a single band and were submitted for commercial automatic DNA sequencing using the Big Dye Terminator System (Applied Biosystems, California, USA) on an ABI3730XL DNA analyser. DNA sequencing was performed twice for each PCR fragment. The sequencing primers included the above PCR primer pair and the additional Int-3 primer (5′-GAA GTG GCT CAT CCA GAA ATT GTT GAA-3′). A total of 65 sequences were unambiguous and included in the present analysis.
Retrieval of GLURP sequences from the NCBI database
A total of 89 nucleotide sequences of the R2 region of the GLURP gene from India (n = 81), Indonesia (n = 3), Iran (n = 2) and China (n = 1) and the laboratory strains 3D7 and F32 were retrieved from the NCBI nucleotide database (https://www.ncbi.nlm.nih.gov/nuccore) (see Additional file 2: Table S1). In addition, a BLASTN search was performed using the nucleotide sequence of the P. falciparum reference strain 3D7  to search against 36 whole-genome shotgun contigs of P. falciparum laboratory strains and wild isolates that are deposited in the NCBI database. The whole-genome shotgun contigs with the complete GLURP R2 region were identified in the sequences from P. falciparum strains HB3, Dd2, FCC-2/Hainan and VS/1, which were submitted by the Broad Institute (Massachusetts, USA), and a further 10 sequences were from P. falciparum isolates from Mali (Africa), which were submitted by the Institute for Genome Sciences (Maryland, USA). Taken together, all the GLURP R2 region sequences from the 103 P. falciparum isolates worldwide and those from the 65 P. falciparum Thai isolates in Thailand formed a global database comprised of 168 alleles of the GLURP gene.
Sequence analysis and population genetics analysis
The GLURP nucleotide sequences were aligned and then manually adjusted, where necessary, using the BioEdit program . Sequence alignment was performed using ClustalW software . The pairwise interpopulation indices or the Wright Fst indices were calculated using the Arlequin software . Other molecular indexes, including the nucleotide diversity (π), haplotype index (hd), and neutrality tests (Tajima’s D test, Fu and Li’s D* test and Fu and Li’s F* test) and linkage disequilibrium, were calculated using the DnaSP software [46, 47].
Analysis of the AAU composition and GLURP subtypes
Theoretical (in silico) translations of the GLURP nucleotide sequences were performed using BioEdit software , and the resulting GLURP amino acid sequences were exported into a text format. The published amino acid repeat units (AAU) of blocks 1–15 in the R2 region were used to search against the collection of GLURP sequences in order to identify any novel AAU types [34, 35]. A total of 10 AAU types were previously identified between blocks 1–15 of the GLURP R2 region, including one AAU type (*) in block 1, two AAU types (I and II) in block 2, one AAU type (1) in block 3 and 10 AAU types (a, b, c, d, e, f, g, h, i and j) in blocks 4–15 [34, 35]. The arrangements of the AAUs were manually inspected to identify the unique AAU arrangement (GLURP subtype). When novel subtypes were identified, the sequence was BLASTP searched in the NCBI database for confirmation. The total number of AAU types in each block and the parasite isolates with a unique AAU arrangement were recorded. The percentage of AAU types was calculated by dividing the number of a particular AAU type by the total number of AAU types in one block. The diversity of infection (DOI) was calculated by dividing the number of different parasite genotypes that were detected with the total number of parasites that were genotyped .
Nucleotide diversity and neutrality test
Summary of statistics for the GLURP R2 region in P. falciparum populations and tests for departure from neutrality
hd ± SD
π ± SD
Fu and Li’s D*
Fu and Li’s F*
0.833 ± 0.038
0.00611 ± 0.00058
-0.53408 (P > 0.10)a
0.80029 (P > 0.10)
0.38828 (P > 0.10)
0.944 ± 0.016
0.00377 ± 0.00103
-0.29581 (P > 0.10)
-0.88322 (P > 0.10)
-0.79358 (P > 0.10)
1 ± 0.272
0.00102 ± 0.00064
1 ± 0.050
0.00159 ± 0.00079
1 ± 0.045
0.00915 ± 0.00219
-0.70120 (P > 0.10)
-0.73515 (P > 0.10)
-0.81915 (P > 0.10)
1 ± 0.126
0.01170 ± 0.00439
-0.29817 (P > 0.10)
-0.29817 (P > 0.10)
-0.31445 (P > 0.10)
0.757 ± 0.031
0.00777 ± 0.00066
-0.13448 (P > 0.10)
-2.05874 (P > 0.10)
-0.21350 (P > 0.10)
Composition of the AAUs in the GLURP R2 region
The R2 region of GLURP contained 6–15 blocks of AAUs, which could be classified into 18 types. Sequence alignments of the AAU in each block revealed the conserved polymorphic sites and specific distributions of the 18 AAU types in the R2 region (Additional file 3: Table S2).
Blocks 1 and 3 contained single AAU types (* and 1), which were identical in all the P. falciparum isolates. Block 2 was characterized by three AAU types (I, II and III), where type I was the most common type, representing 96% of the block 2 sequence. The AAU types I and II, previously reported in P. falciparum populations in India [34, 35], were also detected in the P. falciparum populations in Thailand, while AAU type III was novel and was detected in the whole-genome shotgun sequences of P. falciparum isolate 309 from Mali (Additional file 2: Table S1). The other 13 AAU types were located between blocks 4–15. Of these, three AAU types (a, b and c), were common in all the P. falciparum isolates. In contrast, the other ten AAU types (d, e, f, g, h, i, j, k, l and m) were relatively less common, and their distributions were reserved in specific blocks. Of these, six AAU types (d, e, f, g, h and i) were previously reported in P. falciparum populations in India [34, 35], while three others (i, j and l) were detected in the whole-genome shotgun sequences of the P. falciparum populations in Mali (Additional file 2: Table S1).
Analysis of the GLURP R2 region in the Thai isolates of P. falciparum identified a total of 13 AAU types, including one AAU type in block 1 (*), two AAU types in block 2 (I and II), one AAU type in block 3 (1) and six AAU types (a, b, c, k, l and m) between blocks 4–14. Of these, two AAU types (k and m) were described for the first time in the P. falciparum populations in Thailand. Specifically, AAU type k was located in block 5 in the P. falciparum isolates RN130 and RN133, while AAU type m was detected in block 12 in the P. falciparum isolates RN72 and RN122. These parasite isolates were all from the same population in Ranong Province.
Size polymorphism in the GLURP R2 region
Frequency, subtypes and size of the P. falciparum GLURP polypeptide in global database. Size indicates the number of amino acid residues detected between blocks 4–15
No. of repeat units
C1 to C2
D1 to D4
E1 to E19
F1 to F22
G1 to G14
H1 to H26
I1 to I15
J1 to J2
In addition to the variation in the numbers of repeat units, the size polymorphism of the R2 region of GLURP resulted from the insertion or deletion of negatively charged amino acids, including aspartic acid (D) and glutamic acid (E), between the repeat units. The addition of aspartic acid was more frequently detected between blocks 4–9, while the addition of glutamic acid appeared more frequent between blocks 10–12 (Additional file 4: Table S3). Taken together, these results indicated that variations in the repeat units and the insertions/deletions of aspartic acid and glutamic acid between the repeat units contributed to the size polymorphism in GLURP.
Distribution of the GLURP subtypes and the population structure of P. falciparum
The analysis conducted using the 65 alleles of the GLURP R2 region from P. falciparum populations in Thailand identified 28 GLURP subtypes, containing 8–14 repeat units (Additional file 5: Table S4). The majority (41%) of these Thai P. falciparum isolates inherited GLURP with 11 repeat units. This is contradictory to a previous analysis of GLURP sequences from Indian populations [34, 35], where the GLURP with 13 repeat units was the most common. The DOI analysis of the Thai P. falciparum populations revealed different levels of genetic diversity of GLURP among the Thai subpopulations of P. falciparum, where the DOIs of P. falciparum at Mae Hong Son, Kanchanaburi and Ranong (Western Thailand) were 0.60, 0.76 and 0.56, while those at Ubon Ratchatani and Trat (Eastern Thailand) were both 0.83. The implication then is that the parasite subpopulations near eastern Thailand were more genetically diverse than those near western Thailand.
Of the Thai GLURP subtypes, the F8 and F11 subtypes were the most common, being identified in 13 and nine P. falciparum isolates, respectively, representing 37% of the parasite populations in Thailand (Additional file 5: Table S4). Interestingly, the D3 subtype matched the GLURP R2 sequence of P. falciparum isolate 2/GL from Indonesia (Eisen et al., unpublished data), while eight other subtypes (D4, E9, F8, F11, H1, H10, H22 and I5) corresponded to the GLURP sequences of P. falciparum isolates from India [34, 35]. It is also noteworthy that the F11 subtype was identical to the sequence of the P. falciparum reference strain Dd2. However, no parasite isolates in Thailand or worldwide had GLURP sequences that were similar to those of strains 3D7 and F32, which were employed in the genome sequencing project  and the production of the GLURP-based malaria vaccine, respectively . As a result, the present study identified 19 novel GLURP subtypes in Thailand.
Pairwise F st indices of the GLURP alleles between P. falciparum populations in Thailand. Fst values were low and non-significant in all pairs of P. falciparum populations
(P = 0.40)
(P = 0.23)
(P = 0.81)
(P = 0.80)
(P = 0.68)
(P = 0.32)
(P = 0.11)
(P = 0.16)
(P = 0.10)
(P = 0.14)
The present study reports the genetic structure and diversity of the immunodominant GLURP R2 region in P. falciparum populations in Thailand and worldwide. As of September 2017, 103 sequences of the GLURP gene were available in the NCBI database. Sequence analysis led to the identification of 86 unique GLURP subtypes. In the present study, 65 GLURP sequences were generated from P. falciparum populations in Thailand, and 28 GLURP subtypes were identified, including 19 new subtypes that were identified for the first time. Overall, the combined database of GLURP revealed 105 GLURP subtypes in 168 alleles of the P. falciparum isolates worldwide.
Variations in the repeat unit numbers led to size polymorphisms in the GLURP R2 region. In the present study, GLURP, with 8–14 repeat unit blocks was detected in P. falciparum in Thailand, with 11 repeat unit blocks being the most frequent. This result contrasts with previous analyses of GLURP sequences from P. falciparum in Indian isolates [34, 35], where GLURP genes with 9–15 repeat unit blocks were detected, with 13 repeat units being the most common. Within the whole-genome shotgun sequences of P. falciparum laboratory strains, the shortest GLURP R2 region was found to contain only six repeat units, which was detected in P. falciparum HB3. Overall, the data demonstrated the extensive size variation in the GLURP gene in both laboratory strains and natural isolates of P. falciparum.
Comparison of the amino acid repeat units revealed conserved amino acid substitutions amongst the repeat units. Of the 18 AAU types identified, the majority were located between blocks 4–15, with fewer AAU types in blocks 1–3. Indeed, blocks 1 and 3 had identical sequences in all P. falciparum isolates. The limited amino acid substitutions in the AAU likely reflects the functional constraints of the GLURP protein. A recent study reported that GLURP formed the Pfs38 protein complex on the merozoite surface that binds to erythrocytes via glycophorin A and participates in the parasite invasion of erythrocytes . This is consistent with the three neutrality tests (Tajima’s D, Fu and Li’s D* and Fu and Li’s F* tests) that all gave non-significant values, suggesting that the conserved AAU sequences were maintained under negative purifying selection. Together, these results indicated that the AAU sequences of GLURP are highly conserved, supporting the view that GLURP is an attractive target for vaccine development [17–21].
Another important finding in the present study was that the GLURP R2 region in Thai populations of P. falciparum exhibited extensive sequence variation, with a total of 13 AAU types and 28 GLURP subtypes. The majority of these GLURP subtypes (19 subtypes, 67%) in Thailand were novel. However, the number of GLURP subtypes in P. falciparum in Thailand was lower than that in Indian isolates, where 61 GLURP subtypes were seen [34, 35]. This contrasts to a previous genetic analysis of P. falciparum using msp-3 gene sequences, in which fewer msp-3 haplotypes were detected in the P. falciparum populations in India than in Thailand . Accordingly, the genetic diversity in P. falciparum populations should be evaluated using multiple genetic loci. If genetic markers that exhibit low or limited polymorphisms are employed for genotyping, it is likely that the genetic variation of parasite populations will be underestimated. Because GLURP is highly polymorphic in size and is not under the influence of positive selection, this marker would be suitable for epidemiological and population genetic studies.
The mechanisms of the molecular variation in GLURP can be summarized as follows. First, the emergence of the novel GLURP subtypes was attributed to non-synonymous amino acid substitutions in different repeat units that resulted in the formation of new AAU types. Of the 18 AAU types identified in P. falciparum worldwide, two AAU types (l and m, see Additional file 3: Table S2) are reported here for the first time. In addition, novel GLURP subtypes might have arisen from genetic recombination, resulting in a new assortment of AAU types, as well as by the insertion and deletion of extra amino acids, including aspartic acid (D) and glutamic acid (E), between the repeat units. Taken together, our study revealed the mechanisms contributing to the genetic diversity of the GLURP gene in P. falciparum populations.
Comparison of GLURP subtypes within the Thai P. falciparum populations revealed different levels of genetic diversity among the different P. falciparum populations. From the DOI indices, P. falciparum populations in eastern Thailand were more diverse than those in western Thailand. The low DOI simply reflected that the P. falciparum populations in Mae Hong Son, Kanchanaburi and Ranong had large numbers of individuals with the same GLURP subtypes. The analysis also showed that different subpopulations of P. falciparum in Thailand shared a large number of GLURP subtypes, indicating that gene exchange may occur between the parasites near the eastern and western borders of Thailand. This hypothesis was supported by the Wright’s statistical analysis, where low and non-significant Fst values were found for all the pairs of compared P. falciparum populations. In addition, the Fst analysis of the GLURP subtypes in P. falciparum in India and Thailand also showed a low and non-significant Fst value, indicating similar GLURP alleles between the parasite populations. Together, these results demonstrated that P. falciparum populations in different geographical locations may exhibit different levels of genetic diversity, but gene flow likely exists among the P. falciparum populations in the Greater Mekong Subregion and India.
The sequence analysis and population structure presented here may be useful for developing future GLURP-based vaccines and for monitoring vaccine efficacy. Currently, the GMZ2 vaccine, which incorporates the GLURP R0 region of P. falciparum strain F32 and MSP-3, is being tested in clinical studies in various endemic areas [32, 51]. Although GMZ2 reduced the malaria incidence, its efficacy was still low and so vaccines using a more immunogenic formulation are needed. One proposed strategy is to incorporate the GLURP R2 region of P. falciparum strain Dd2 in a vaccine, since the R2 region also contains B-cell epitopes and might elicit functional protective antibodies , while the most common GLURP subtype (F11) in Thailand and India is identical to that of Dd2. Thus, such a vaccine might exhibit improved efficacy against clinical cases of malaria.
The present study generated a database of GLURP gene sequences, and the in silico translated GLURP sequences, from worldwide populations of P. falciparum. The database revealed not only the sequence diversity but also the distribution of the GLURP subtypes. Because the GLURP gene exhibited extensive DNA size and sequence polymorphism, it might serve as a genetic marker for parasite identification as well as for population and epidemiological studies. The sequence information provides knowledge for a better design of GLURP-based malaria vaccines and reveals the population structure of P. falciparum across Thailand, which is of use for monitoring and controlling this parasite.
We are grateful to Dr Robert Butcher for language editing and proofreading the manuscript. We would like to thank the editor and reviewers for their insightful comments and suggestions on the paper, as these comments led us to an improvement of the work.
This work was supported by grants from the Development and Promotion of Science and the Technology Talents Project (DPST) graduate with First Placement, Royal Thai Government to SP. The sponsor of the study had no role in the study design, data collection, data analysis, data interpretation or writing of the manuscript.
Availability of data and materials
The nucleotide sequence data of the different GLURP genes reported in this paper have been submitted to the GenBank database with accession numbers MG578444–MG578508. The data are also available upon request. Requests for data may be made by contacting SP.
SP and PH provided the conceptual framework for the project, guidance for the interpretation of the data, performed the data analysis and participated in the manuscript preparation, revision and coordination. SP, CT and PS performed the molecular genetic work. SP, NW and MK analysed the data. PH was responsible for the parasite cultivation. All authors read and approved the final manuscript.
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Zheng Z, Cheng Z. Advances in molecular diagnosis of malaria. Adv Clin Chem. 2017;80:155–92.View ArticlePubMedGoogle Scholar
- Cordray MS, Richards-Kortum RR. Emerging nucleic acid-based tests for point-of-care detection of malaria. Am J Trop Med Hyg. 2012;87:223–30.View ArticlePubMedPubMed CentralGoogle Scholar
- Imai K, Tarumoto N, Misawa K, Runtuwene LR, Sakai J, Hayashida K, et al. A novel diagnostic method for malaria using loop-mediated isothermal amplification (LAMP) and MinION™ nanopore sequencer. BMC Infect Dis. 2017;17:621.View ArticlePubMedPubMed CentralGoogle Scholar
- Batista-dos-Santos S, Raiol M, Santos S, Cunha MG, Ribeiro-dos-Santos Â. Real-time PCR diagnosis of Plasmodium vivax among blood donors. Malar J. 2012;11:345.View ArticlePubMedPubMed CentralGoogle Scholar
- Pattaradilokrat S, Tiyamanee W, Simpalipan P, Kaewthamasorn M, Saiwichai T, Li J, et al. Molecular detection of the avian malaria parasite Plasmodium gallinaceum in Thailand. Vet Parasitol. 2015;210:1–9.View ArticlePubMedGoogle Scholar
- Snounou G, Viriyakosol S, Jarra W, Thaithong S, Brown KN. Identification of the four human malaria parasite species in field samples by the polymerase chain reaction and detection of a high prevalence of mixed infections. Mol Biochem Parasitol. 1993;58:283–92.View ArticlePubMedGoogle Scholar
- Brockman A, Paul RE, Anderson TJ, Hackford I, Phaiphun L, Looareesuwan S, et al. Application of genetic markers to the identification of recrudescent Plasmodium falciparum infections on the northwestern border of Thailand. Am J Trop Med Hyg. 1999;60:14–21.View ArticlePubMedGoogle Scholar
- Rubio JM, Berzosa PJ, Benito A. Amplified fragment length polymorphism (AFLP) protocol for genotyping the malarial parasite Plasmodium falciparum. Parasitology. 2001;123:331–6.View ArticlePubMedGoogle Scholar
- Nyachieo A, VAN Overmeir C, Laurent T, Dujardin JC, D'Alessandro U. Plasmodium falciparum genotyping by microsatellites as a method to distinguish between recrudescent and new infections. Am J Trop Med Hyg. 2005;73:210–3.PubMedGoogle Scholar
- Pumpaibool T, Arnathau C, Durand P, Kanchanakhan N, Siripoon N, Suegorn A, et al. Genetic diversity and population structure of Plasmodium falciparum in Thailand, a low transmission country. Malar J. 2009;8:155.View ArticlePubMedPubMed CentralGoogle Scholar
- Ranford-Cartwright LC, Taylor J, Umasunthar T, Taylor LH, Babiker HA, Lell B, et al. Molecular analysis of recrudescent parasites in a Plasmodium falciparum drug efficacy trial in Gabon. Trans R Soc Trop Med Hyg. 1997;91:719–24.View ArticlePubMedGoogle Scholar
- Borre MB, Dziegiel M, Høgh B, Petersen E, Rieneck K, Riley E, et al. Primary structure and localization of a conserved immunogenic Plasmodium falciparum glutamate rich protein (GLURP) expressed in both the preerythrocytic and erythrocytic stages of the vertebrate life cycle. Mol Biochem Parasitol. 1991;49:119–31.View ArticlePubMedGoogle Scholar
- Høgh B, Thompson R, Zakiuddin IS, Boudin C, Borre M. Glutamate rich Plasmodium falciparum antigen (GLURP). Parassitologia. 1993;35:47–50.PubMedGoogle Scholar
- Paul G, Deshmukh A, Kaur I, Rathore S, Dabral S, Panda A, et al. A novel Pfs38 protein complex on the surface of Plasmodium falciparum blood-stage merozoites. Malar J. 2017;16:79.View ArticlePubMedPubMed CentralGoogle Scholar
- Theisen M, Cox G, Høgh B, Jepsen S, Vuust J. Immunogenicity of the Plasmodium falciparum glutamate-rich protein expressed by vaccinia virus. Infect Immun. 1994;62:3270–5.PubMedPubMed CentralGoogle Scholar
- Theisen M, Vuust J, Gottschau A, Jepsen S, Høgh B. Antigenicity and immunogenicity of recombinant glutamate-rich protein of Plasmodium falciparum expressed in Escherichia coli. Clin Diagn Lab Immunol. 1995;2:30–4.PubMedPubMed CentralGoogle Scholar
- Dodoo D, Theisen M, Kurtzhals JA, Akanmori BD, Koram KA, Jepsen S, et al. Naturally acquired antibodies to the glutamate-rich protein are associated with protection against Plasmodium falciparum malaria. J Infect Dis. 2000;181:1202–5.View ArticlePubMedGoogle Scholar
- Lusingu JP, Vestergaard LS, Alifrangis M, Mmbando BP, Theisen M, Kitua AY, et al. Cytophilic antibodies to Plasmodium falciparum glutamate rich protein are associated with malaria protection in an area of holoendemic transmission. Malar J. 2005;4:48.View ArticlePubMedPubMed CentralGoogle Scholar
- Nebie I, Diarra A, Ouedraogo A, Soulama I, Bougouma EC, Tiono AB, et al. Humoral responses to Plasmodium falciparum blood-stage antigens and association with incidence of clinical malaria in children living in an area of seasonal malaria transmission in Burkina Faso, West Africa. Infect Immun. 2008;76:759–66.View ArticlePubMedGoogle Scholar
- Adu B, Cherif MK, Bosomprah S, Diarra A, Arthur FK, Dickson EK, et al. Antibody levels against GLURP R2, MSP1 block 2 hybrid and AS202.11 and the risk of malaria in children living in hyperendemic (Burkina Faso) and hypo-endemic (Ghana) areas. Malar J. 2016;15:123.View ArticlePubMedPubMed CentralGoogle Scholar
- Pratt-Riccio LR, Lima-Junior JC, Carvalho LJ, Theisen M, Espíndola-Mendes EC, Santos F, et al. Antibody response profiles induced by Plasmodium falciparum glutamate-rich protein in naturally exposed individuals from a Brazilian area endemic for malaria. Am J Trop Med Hyg. 2005;73:1096–103.PubMedGoogle Scholar
- Oeuvray C, Theisen M, Rogier C, Trape JF, Jepsen S, Druilhe P. Cytophilic immunoglobulin responses to Plasmodium falciparum glutamate-rich protein are correlated with protection against clinical malaria in Dielmo, Senegal. Infect Immun. 2000;68:2617–20.View ArticlePubMedPubMed CentralGoogle Scholar
- Kana IH, Adu B, Tiendrebeogo RW, Singh SK, Dodoo D, Theisen M. Naturally acquired antibodies target the glutamate-rich protein on intact merozoites and predict protection against febrile malaria. J Infect Dis. 2017;215:623–30.View ArticlePubMedGoogle Scholar
- Theisen M, Soe S, Brunstedt K, Follmann F, Bredmose L, Israelsen H, et al. A Plasmodium falciparum GLURP-MSP3 chimeric protein; expression in Lactococcus lactis, immunogenicity and induction of biologically active antibodies. Vaccine. 2004;22:1188–98.View ArticlePubMedGoogle Scholar
- Carvalho LJ, Alves FA, Bianco C Jr, Oliveira SG, Zanini GM, Soe S, et al. Immunization of Saimiri sciureus monkeys with a recombinant hybrid protein derived from the Plasmodium falciparum antigen glutamate-rich protein and merozoite surface protein 3 can induce partial protection with Freund and Montanide ISA720 adjuvants. Clin Diagn Lab Immunol. 2005;12:242–8.PubMedPubMed CentralGoogle Scholar
- Tamborrini M, Stoffel SA, Westerfeld N, Amacker M, Theisen M, Zurbriggen R, et al. Immunogenicity of a virosomally-formulated Plasmodium falciparum GLURP-MSP3 chimeric protein-based malaria vaccine candidate in comparison to adjuvanted formulations. Malar J. 2011;10:359.View ArticlePubMedPubMed CentralGoogle Scholar
- Esen M, Kremsner PG, Schleucher R, Gässler M, Imoukhuede EB, Imbault N, et al. Safety and immunogenicity of GMZ2 - a MSP3-GLURP fusion protein malaria vaccine candidate. Vaccine. 2009;27:6862–8.View ArticlePubMedGoogle Scholar
- Mordmüller B, Szywon K, Greutelaers B, Esen M, Mewono L, Treut C, et al. Safety and immunogenicity of the malaria vaccine candidate GMZ2 in malaria-exposed, adult individuals from Lambaréné, Gabon. Vaccine. 2010;28:6698–703.View ArticlePubMedPubMed CentralGoogle Scholar
- Bélard S, Issifou S, Hounkpatin AB, Schaumburg F, Ngoa UA, Esen M, et al. A randomized controlled phase Ib trial of the malaria vaccine candidate GMZ2 in African children. PLoS One. 2011;6:e22525.View ArticlePubMedPubMed CentralGoogle Scholar
- Jepsen MP, Jogdand PS, Singh SK, Esen M, Christiansen M, Issifou S, et al. The malaria vaccine candidate GMZ2 elicits functional antibodies in individuals from malaria endemic and non-endemic areas. J Infect Dis. 2013;208:479–88.View ArticlePubMedGoogle Scholar
- Mamo H, Esen M, Ajua A, Theisen M, Mordmüller B, Petros B. Humoral immune response to Plasmodium falciparum vaccine candidate GMZ2 and its components in populations naturally exposed to seasonal malaria in Ethiopia. Malar J. 2013;12:51.Google Scholar
- Sirima SB, Mordmüller B, Milligan P, Ngoa UA, Kironde F, Atuguba F, et al. GMZ2 trial study group. A phase 2b randomized, controlled trial of the efficacy of the GMZ2 malaria vaccine in African children. Vaccine. 2016;34:4536–42.View ArticlePubMedGoogle Scholar
- Takala SL, Plowe CV. Genetic diversity and malaria vaccine design, testing and efficacy: preventing and overcoming ‘vaccine resistant malaria’. Parasite Immunol. 2009;31:560–73.View ArticlePubMedPubMed CentralGoogle Scholar
- Patel P, Bharti PK, Bansal D, Raman RK, Mohapatra PK, Sehgal R, et al. Genetic diversity and antibody responses against Plasmodium falciparum vaccine candidate genes from Chhattisgarh, Central India: implication for vaccine development. PLoS One. 2017;12:e0182674.View ArticlePubMedPubMed CentralGoogle Scholar
- Kumar D, Dhiman S, Rabha B, Goswami D, Deka M, Singh L, et al. Genetic polymorphism and amino acid sequence variation in Plasmodium falciparum GLURP R2 repeat region in Assam, India, at an interval of five years. Malar J. 2014;13:450.View ArticlePubMedPubMed CentralGoogle Scholar
- Mohd Abd Razak MR, Sastu UR, Norahmad NA, Abdul-Karim A, Muhammad A, Muniandy PK, et al. Genetic diversity of Plasmodium falciparum populations in malaria declining areas of Sabah, East Malaysia. PLoS One. 2016;11:e0152415.View ArticlePubMedPubMed CentralGoogle Scholar
- Akter J, Thriemer K, Khan WA, Sullivan DJ Jr, Noedl H, Haque R. Genotyping of Plasmodium falciparum using antigenic polymorphic markers and to study anti-malarial drug resistance markers in malaria endemic areas of Bangladesh. Malar J. 2012;11:386.View ArticlePubMedPubMed CentralGoogle Scholar
- Congpuong K, Hoonchaiyapoom T, Inorn K. Plasmodium falciparum genotype diversity in artemisinin derivatives treatment failure patients along the Thai-Myanmar border. Korean J Parasitol. 2014;52:631–7.View ArticlePubMedPubMed CentralGoogle Scholar
- Haddad D, Snounou G, Mattei D, Enamorado IG, Figueroa J, Ståhl S, et al. Limited genetic diversity of Plasmodium falciparum in field isolates from Honduras. Am J Trop Med Hyg. 1999;60:30–4.View ArticlePubMedGoogle Scholar
- Simpalipan P, Pattaradilokrat S, Siripoon N, Seugorn A, Kaewthamasorn M, Butcher RD, et al. Diversity and population structure of Plasmodium falciparum in Thailand based on the spatial and temporal haplotype patterns of the C-terminal 19-kDa domain of merozoite surface protein-1. Malar J. 2014;13:54.View ArticlePubMedPubMed CentralGoogle Scholar
- Pattaradilokrat S, Sawaswong V, Simpalipan P, Kaewthamasorn M, Siripoon N, Harnyuttanakorn P. Genetic diversity of the merozoite surface protein-3 gene in Plasmodium falciparum populations in Thailand. Malar J. 2016;15:517.View ArticlePubMedPubMed CentralGoogle Scholar
- Gardner MJ, Hall N, Fung E, White O, Berriman M, Hyman RW, et al. Genome sequence of the human malaria parasite Plasmodium falciparum. Nature. 2002;419:498–511.View ArticlePubMedGoogle Scholar
- Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucl Acids Symp Series. 1999;41:95–8.Google Scholar
- Sievers F, Higgins DG. Clustal omega. Curr Protoc Bioinformatics. 2014;48:3.13.1–16.View ArticleGoogle Scholar
- Excoffier L, Lischer HE. Arlequin suite ver 3.5: a new series of programs to perform population genetics analyses under Linux and Windows. Mol Ecol Resour. 2010;10:564–7.View ArticlePubMedGoogle Scholar
- Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bioinformatics. 2009;25:1451–2.View ArticlePubMedGoogle Scholar
- Rozas J. DNA sequence polymorphism analysis using DnaSP. Methods Mol Biol. 2009;537:337–50.View ArticlePubMedGoogle Scholar
- A-Elbasit IE, A-Elgadir TM, Elghazali G, Elbashir MI, Giha HA. Genetic fingerprints of parasites causing severe malaria in a setting of low transmission in Sudan. J Mol Microbiol Biotechnol. 2007;13:89–95.View ArticlePubMedGoogle Scholar
- Tajima F. Simple methods for testing the molecular evolutionary clock hypothesis. Genetics. 1993;135:599–607.PubMedPubMed CentralGoogle Scholar
- Fu YX, Li WH. Statistical tests of neutrality of mutations. Genetics. 1993;133:693–709.PubMedPubMed CentralGoogle Scholar
- Theisen M, Adu B, Mordmüller B, Singh S. The GMZ2 malaria vaccine: from concept to efficacy in humans. Expert Rev Vaccines. 2017;16:907–17.View ArticlePubMedGoogle Scholar
- Theisen M, Soe S, Oeuvray C, Thomas AW, Vuust J, Danielsen S, et al. The glutamate-rich protein (GLURP) of Plasmodium falciparum is a target for antibody-dependent monocyte-mediated inhibition of parasite growth in vitro. Infect Immun. 1998;66:11–7.PubMedPubMed CentralGoogle Scholar