Open access peer-reviewed chapter

Genotyping for Plasmodium spp.: Diagnosis and Monitoring of Antimalarial Drug Resistance

Written By

Jean Bernard Lekana-Douki and Larson Boundenga

Submitted: October 10th, 2017 Reviewed: March 19th, 2018 Published: November 5th, 2018

DOI: 10.5772/intechopen.76601

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Abstract

Malaria is one the world’s most widespread lethal diseases. Plasmodium falciparum, P. vivax, P. ovale, P. malariae and P. knowlesi induce human pathology. These species could be differentially diagnosed using the genotyping of cytochrome b, Pfdhfr and RNA 18S. The persistence of P. falciparum, the most lethal parasite, is mainly due to antimalarial drug resistance. Indeed, a few years after the start of the ambitious malaria eradication program in 1960, chloroquine resistance emerged in Asia and spread widely in all the endemic areas. It was associated with genotypes in P. falciparum chloroquine resistance transporter (CVIET, SVMNT, CVMNT, CVIDT, SVIET and CVMET). The use of new drugs such as sulfadoxine-pyrimethamine (SP) leads quickly to SP-resistant parasites associated with genotypes on P. falciparum DiHydroFolate reductase (I51-R59-N108-I164) and P. falciparum DiHydroPteroate synthetase (436-437-580-613). Recently, the delay of parasite clearance has been described with artemisinine (the most efficacious antimalarial drug). This resistance was associated with the K13 propeller genotype. Since malaria species and antimalarial drug resistance markers could be characterized using nucleic acid sequences, genotyping is needed for malarial monitoring of species distribution and antimalarial drug resistance.

Keywords

  • Plasmodium parasites
  • drug resistance
  • diagnostic
  • genotyping

1. Introduction

Malaria remains a major public health problem. More than 40% of the world’s population (3.3 billion) live malarial endemic areas in varying degrees. Despite tremendous efforts in the fight, and though this strategy or plan resulted in significantly decreasing the burden in the last 20 years, Malaria still is persistent in nearly 91 countries (Figure 1). In 2016, the overall incidence was 216 million cases among these and 445,000 deaths were recorded [1]. Africa continues to account for 90% of malaria burden. African children under 5 years of age are the most affected. This infectious disease is due to the invasion of Plasmodium spp. Currently, the five species are known to infect and to induce significantly malaria in humans are Plasmodium ovale, Plasmodium malariae, Plasmodium knowlesi, Plasmodium vivax and Plasmodium falciparum.

Figure 1.

Countries endemic for malaria in 2000 and 2016. From WHO [1].

P. falciparum is the most virulent species as it is responsible for more than 90% of malaria deaths [2, 3]. Its specific biology with antigenic variation, sequestration of infected blood cells and interactions with host cells leads to severe malaria [4, 5]. Also, it is the most recent human infection with limited adaptation in the host [6]. P. knowlesi, which is a specific Asian monkey parasite (Macaca genus), was recently transferred to humans, causing high mortality in the south of Asia [7, 8, 9]. One of its differences from other species is the time of its life cycle which is 24 h whereas 40–48 h for other human Plasmodium spp. P. vivax is the most prevalent in Asia and South America. Due to the Duffy-negative statue of Black people, it is the rarest in Black Africa. However, recent studies reported the presence of P. vivax in Blacks with Duffy-negative from some countries of central Africa such as Equatorial Guinea, Congo Republic and so on [10, 11]. P. malariae is less prevalent in Asia, while it is most common in sub-Saharan Africa and southwest Pacific [12]. It often finds minor prevalence compared to P. falciparum. This parasite is thought to be a zoonotic infection because is genetically close to P. brasilianum which infects monkeys of South America [12, 13]. P. ovale is prevalent in Sub-Saharan Africa, South-East Asia, India, Papua New Guinea, Timor and Indonesia [14]. It is the less-prevalent human malarial parasite. However, in most places where P. ovale is observed, it is relatively uncommon and its prevalence rarely exceeds 5% [12].

Decisions concerning malaria treatment depend on the identification of the species causing the disease. Traditionally, this diagnosis was based on the microscopic detection of Plasmodium parasites in Giemsa-stained blood slides. In recent decades, antigen detection assays and molecular detection assays were introduced as alternatives to microscopy [15]. These approaches were very useful; however, they are not very reliable. Indeed, the morphological features and life history traits of a parasite species can vary from one host species to another [16, 17]. For antigen detection assays are mainly aimed at the identification of P. falciparum. Indeed [18], only a few assays are able to identify infections caused by other human malaria parasites [15, 19]. However, the development of molecular tools for the identification of species in diagnosis and genotyping permits a better reading of plasmodial diversity circulating among the human population and allows best highlighting the phenomenon of resistance of Plasmodium than microscopic tools.

Plasmodium species like several other genera have specific genetic markers such as 18S rRNA, ITS, cytochrome b and so on, used in studying speciation. These markers play an important role in molecular analysis of genotyping and monitoring antimalarial drug resistance. The persistence of malaria burden is partly due to the emerging and widespread nature of the antimalarial drug resistance. Indeed, in the 1950s, WHO launched the malaria eradication program, with chloroquine and dichloro-diphenyl-trichloroethan (DDT). But in the early 1960s, chloroquine resistance emerged [20, 21]. Antimalarial drug resistance is defined as the parasite’s ability to survive after the absorption of drug doses is greater than patient-tolerated doses. This chapter described the methods of genotyping the malaria for species diagnosis that helps to monitor drug resistance.

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2. Genotyping for Plasmodium spp. diagnosis

The genotyping of Plasmodium spp. infections allows for the characterization of distinct species and subpopulations present in hosts. The genotyping techniques presented in the following allow for the characterization of different Plasmodium species by sizing the polymerase chain reaction (PCR) product of the polymorphic marker gene merozoite surface protein.

2.1. PCR using the specificity of 18S RNA

rRNA is one of the ribosome components. Among rRNA, the 18S rRNA gene is the most frequently cited marker for malaria detection. It is composed of highly conserved regions which can be targeted for a qualitative detection of Plasmodium spp. and of variable zones allowing species identification. Early in the 1990s, Snounou and colleagues reported high-sensitivity methods for the detection of plasmodium species using nested PCR [22]. This method was based on the conversion of 18S rRNA among human plasmodium and the specificity of 18S rRNA from each parasite. So, the method includes the first PCR with primers named rPLU1/5 and rPLU3/4 that match with human plasmodium. This step is followed by the amplification of the product of primary PCR separately with the four species-specific primer pairs: rFAL1 and rFAL2, rVIV1 and rVIV2, rMAL1 and rMAL2 and rOVA1 and rOVA2 to identify the species P. falciparum, P. vivax, P. malariae and P. ovale, respectively. The primary PCR with rPLU1/5 and rPLU3/4 gives a 1100 bp band in a 2% agarose gel electrophoresis (Figure 2), in the presence of Plasmodium.

Figure 2.

Specificity and sensitivity of the PCR detection assay. (a) Nested PCR amplification for the demonstration of the specificity of the primers employed. Control genomic DNA from P. falciparum (F), P. vivax (V), P. malariae (M), P. ovale (O) and human blood (H); (b) nested PCR assay for the detection of in vitro cultured P. falciparum ring-stage parasites; (c) nested PCR amplification using diluted control DNAs; (d) product of amplification of diluted control DNAs, using the PCR assay (Source: Snounou et al. [22]).

The nested PCR rFAL1 and rFAL2 generate a 205 bp in presence of the P. falciparum parasite. The nested PCR rVIV1 and rVIV2 generate a 120 bp in presence of the P. vivax parasite. The nested PCR rMAL1 and rMAL2 generate a 144 bp in presence of the P. malaria parasite. The nested PCR rOVA1 and rOVA2 generate an 800 bp in presence of the P. ovale parasite [22, 23].

2.2. RT-PCR NASBA 18S rRNA

Nucleic acid sequence-based amplification (NASBA) is a method in molecular biology, which is used to amplify RNA sequences. This novel approach of genotyping, based on the amplification of nucleic acid sequence (real-time QT-NASBA), was developed by Compton [24]. Immediately after its discovery, the NASBA method was used for the rapid diagnosis and quantification of HIV-1 in patients [25]. Some years later, Schooner et al. [26] developed a real-time quantitative nucleic acid sequence-based amplification (real time QT-NASBA) for the detection of Plasmodium falciparum 18S rRNA with a sensitivity of 10–50 parasites/ml [26, 27]. Thus, NASBA method that uses primers and probes were selected on the basis of the sequence of the small subunit 18S rRNA gene [26, 28], to characterize or identify the different human Plasmodium species [18]. For NASBA, Schooner et al. have defined primers Plas-1F (59-TCAGATACCGTCGTAATCTTA-39) and Plas-2R T7 (59-AATTCTAATACGACTCACTATAGGGAGAGAACTTTCTCGCTTGCGCGAA-39) which were used [26]. The RNAs from P. falciparum, P. malariae, P. vivax and P. ovale are specifics and they should all be amplified by the NASBA isolates with these primers. Therefore, detection is done by capture probe (59-ACCATAAACTATG CCGACTAGG-39) which is bound to magnetic beads. Finally, samples are hybridized separately to ruthenium-labeled WT (59-CCTTATGAGAAATCAAAGTC 39) and Q (59-AATAACTGCACCAGTGTATA-39) detection probes, followed by ECL detection in a NucliSens ECL reader [26, 29]. The NASBA method is very sensitive and specific. It can be used for the detection, identification and quantitative measurement of low parasitaemia of Plasmodium species, that is, the lower detection limit of the assay is 100–1000 molecules in vitro RNA for all human malaria parasites.

2.3. PCR sequencing cytochrome b

Cytochrome b (Cyt-b) is one of the respiratory chain systems present in mitochondria. It is highly conserved in Plasmodium species. But some of its region shows diversities and species specificities. It is also in multiple copies (20–100 copies) in the haploid genome, suggesting the increase of sensitivity. So, the alignment of Cyt-b sequences obtained after amplification by the nested PCR portion of mitochondrial DNA using couples of primers (Table 1) allows distinguishing the Plasmodium species present in the host (Figure 3).

Table 1.

Amplification primers of the cytochrome b gene (Cyt-b). Cyt-b is amplified by nested PCR.

Figure 3.

The tree of relationship of Plasmodium species. This tree was built with a portion of Cyt-b sequences characterized among some patients and other Cyt-b sequences from Genbank (accession numbers AB489194, EU880499, GU723548 and GU723532). The tree was built using maximum likelihood with Cyt-b sequences of 925pb sizes.

The first round of PCR (PCR1) and use of the primers DW2/F and DW4/R produce the fragments of 1253 bp. In the second round of PCR (PCR2), primers Cyt-b1F and Cyt-b2R are used and this generates PCR fragments of 939 bp [30]. This approach was used more for the characterization of malaria parasites in primates (human and non-human) [31, 32] because it allows to identify all species known or unknown circulating in vertebrates [33].

2.4. PCR-RFLP from cytochrome b

Due to the specie specificity and diversity of Cytochrome b, it could be digested with the restriction enzyme Alu I leading to species-specific patterns [34]. In this case, a nested PCR is used. Primers Plas 1 (5′-GAGAATTATGGAGTGGATGGTG-3′) and Plas 2 (5′-GTGGTAATTGACATCCWATCC-3′) are used for primary PCR producing an 816 bp fragment, following by a nested with Plas 3 and Plas 4 primers. The n-PCR produces a 787 bp fragment. The digestion with Alu I gives 159 and 640 bp fragments; 187, 249 and 381 bp fragments; 111, 169, 187, 249 and 270 bp; and 224 and 584 bp fragments for P. falciparum, P. malariae, P. vivax and P. ovale, respectively, on agarose gel electrophoresis (Figure 4).

Figure 4.

Illustration of a digestion cytochrome fragment using Alu enzyme.

2.5. PCR-DHFR

Figure 5 shows the equation of transformation of dihydrofolate to tetrahydrofolate. The dihdrofolate reductase is one of the important malaria proteins involved in the plasmodium folate synthesis. This gene is coded in chromosome 4 and is highly conserved between distantly related species, like plasmodium species. Its linker region revealed significantly a higher sequence diversity than the relatively conserved enzymatic diversity. Different species of Plasmodium are characterized by a unique linker sequence. So, it has been used to identify human plasmodium species [35]. Nonsynonymous mutations on DHFR are associated with pyrimethanime resistance. Using primers ATGGARSAMSTYTSMGABGTWTTYGA and AAATATTGRTAYTCTGGRTG for primary PCR gives a 1000 bp fragment. The nested PCR species specific with primers shown in Table 2 allow to amplify specifically 160, 177, 144, 231/237/243 and 134 bp fragments for P. falciparum, P. malariae, P. vivax, P. ovale and P. knowlesi.

Figure 5.

The equation of the DHF-THF reaction. Dihydrofolate reductase is an enzyme that catalyzes the NADPH-dependent reduction of dihydrofolate to tetrahydrofolate. This reaction is essential for the de novo synthesis of purines and certain amino acids. This enzyme is essential for rapid growth and is the target for the action of the important antimalarial drugs pyrimethamine and proguanil.

PrimerSpeciesSequence (5′–3′)Annealing (°C)No. of cyclesProduct (bp)
Pla-DHFR-F
Pla-TS-R
Plasmodium sp.ATGGARSAMSTYTSMGABGTWTTYGA
AAATATTGRTAYTCTGGRTG
50301000
Pla-DHFR-NF
Pla-TS-R
Plasmodium sp.AAATGYTTYATYATWGGDGG
AAATATTGRTAYTCTGGRTG
5535509–587
PF-Lin-F
PF-Lin-R
P. falciparumAAAAGGAGAAGAAAAAAATAA
AAAATAAACAAAATCATC
5035160
PM-Lin-F
PM-Lin-R
P. malariaeGACCCAAGAATCCCTCCC
CCCATGAAGTTATATTCC
5035177
PV-Lin-F
PV-Lin-R
P. vivaxCGGGAGCACTGCGGACAGCG
CACGGGCACGCGGCGGGGC
5535144
PO-Lin-F
PO-Lin-R
P. ovaleGACACACAAAATGATGGGGA
ATTGTCCTTTCCTTGACTCG
5535231, 237 or 243
PK-Lin-F
PK-Lin-R
P. knowlesiCGATGGATATGGATAGTGG
CGCGGGAGAGCATTTCCTC
5835134

Table 2.

Primer sequences and the PCR condition for detection of Plasmodium spp. that infect humans.

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3. Genotyping for the monitoring of antimalarial drug resistance

3.1. Antimalarial drug resistance

Antimalarial drug resistance is the ability of P. falciparum to survive after absorption of the drug at concentrations greater than concentrations tolerated by the patient. Drug resistance arises rarely because it is the result of some non-lethal mutations, but it spreads relatively quickly. The clinical failure treatment is the lasted event in the long way of changes in parasites genes. Antimalarial drug resistance markers are genes associated with antimalarial drug resistance. Among them, the most characterized are Pfcrt (P. falciparum chloroquine resistance transporter), Pfmdr1 (P. falciparum multidrug resistance 1), Pfdhfr (P. falciparum dihydrofolate reductase), Pfdhps (P. falciparum dihydropteroate synthase), Pfmrp (P. falciparum multidrug resistance protein), Pfatpase 6 (P. falciparum atpase 6) and PfK13 (P. falciparum kelch 13).

3.2. The genotyping of markers associated with antimalarial drug resistance

3.2.1. P. falciparum chloroquine resistance transporter (Pfcrt)

Pfcrt is an ATP binding cassette (ABC) protein, able to fixe and hydrolyze ATP. The general structure of the ABC transporter contains cytosolic nucleotide binding domains (NBDs), a nucleotide hydrolysis site and transmembrane segments. These transporters extrude effective drugs from the digestive vacuole and function as an efflux pump leading to the decrease of intracellular accumulation of drugs in the parasite. The genetic cross-experiment between chloroquine-sensitive and chloroquine-resistance strains allowed the identification of P. falciparum chloroquine-resistance transporter (Pfcrt) in 2000 [20, 36]. It is a 45 kDa protein coding in chromosome 7, containing 10 predicted transmembrane domains located on the membrane of the digestive vacuole (Figure 6).

Figure 6.

The predicted structure and representative haplotypes of P. falciparum chloroquine-resistance transporter. PfCRT is predicted to have 10 transmembrane domains, with its N and C terminals located on the cytoplasmic side of the digestive vacuole membrane. Mutations identified in the Pfcrt cDNA sequences from CQR lines (black circles), the crucial K76T mutation common to all CQR strains (red) and the S163R mutation identified in the amantadine- and halofantrine-resistant parasites (yellow circle) are indicated (this figure was pulled in [40]).

Several mutations have been identified in this transporter. The main mutation T76 allows for the abolition of accumulation of the drug chloroquine in the digestive vacuole. The association of mutations in codons 72, 73, 74, 75 and 76 defined different haplotypes. These haplotypes show a spatio-temporal specificity. CVMNK is the wild-type haplotype that is found in chloroquine-sensitive parasites. In Africa, the most prevalent chloroquine-resistance haplotype is the CVIET. It was also found with less prevalence in South America and in Southeast of Asia. Another Pfcrt resistance haplotype, named South American haplotype, is the SVMNT. That is rarely found in Africa and Asia, has relatively little fitness cost and was associated with the emergence of amodiaquine resistance too [37, 38]. The other main Pfcrt-resistance haplotypes were CVMNT, CVMET, SVIET and CVIDT. CVMNT is most prevalent in South America and in Asia but rarely found in Africa [39, 40]. CVMET is the rarest haplotype found in Asia whereas SVIET is the South American haplotype. CVIDT is the specific haplotype from Asia. Haplotypes CVIET and SVMNT were also associated with the plasmodial resistance against amodiaquine and lumefantrine [41]. In our recent study from Gabon and Congo we found new haplotypes. But the involvement of these in antimalarial resistance is needed yet (unpublished yet). The withdrawal of chloroquine has led to the decrease of the T76 genotype. Furthermore, recent data showed that the use of artemisinine-based combination therapies, mainly artemether-lumefantrine, selected the wild-type K76 genotype. Other polymorphisms in Pfcrt were described including A144F, I194T, A220S, Q271E and N326S [40].

For the genotype haplotype 72–76 of Pfcrt, PCR sequencing and PCR-RFLP are used. That is based on nested PCR with several primers (Table 3). The RFLP pattern indicates a specific genotype for each codon.

Genes, codonsPrimer namesPrimersT°CRestriction enzymeSizes (bp)*
Pfcrt, C72SCRT72MSTTTATATTTTAAGTATTATTTATTTAAGTGGA55Mbo I55 + 38
76-D2CAAAACTATAGTTACCAATTTTG
Pfcrt, M74ICRT745MSTAAGTATTATTTATTTAAGTGTATGTGTCAT55Nla III53 + 31
76-D2CAAAACTATAGTTACCAATTTTG
Pfcrt, N75MCRT745MSTAAGTATTATTTATTTAAGTGTATGTGTCAT50BspHI53 + 31
76-D2CAAAACTATAGTTACCAATTTTG
Pfcrt, 76Pfcrt-76AGCGCGCGCATGGCTCACGTTTAGGTGGAG55Apo I136 + 56
Pfcrt-76BGGGCCCGGCGGATGTTACAAAACTATAGTTACC

Table 3.

Sequences of primer sets and restriction enzymes used to characterize polymorphisms.

Sizes* indicate the sizes of fragments generated after restriction enzyme digestions. T°C= hybridization temperature during PCR program.

3.2.2. Plasmodium falciparum multidrug resistance 1 (Pfmdr1)

P. falciparum multidrug resistance 1 is a homologous of the bacterial multidrug resistance 1 protein. It is one of the main antimalarial drug resistance markers. Like Pfcrt, Pfmdr1 is also an ABC transporter, located in the membrane of a digestive vacuole, coded in chromosome 5 by the 4758 bp gene. Its structure shows two homologous parts containing six transmembrane domains plus a nucleotide binding domain (Figure 7).

Figure 7.

Genetic polymorphisms in P. falciparum multidrug resistance-1. PfMDR1 has two homologous halves, each with six predicted transmembrane domains and a nucleotide-binding pocket. The nucleotide-binding domains (NBD1 and NBD2; orange boxes) are each formed by large cytoplasmic domains. Polymorphic amino acids found in the K1 allele (N86Y) and the 7G8 allele (Y184F, S1034C, N1042D and D1246Y) are indicated. The D1246Y mutation is located in the predicted NBD2 (this figure was pulled, from Ref. [40]).

Some isolates exhibit multicopies of Pfmdr1. Polymorphisms in codons 82, 184, 1034, 1042 and 1246 (N86Y, Y184F, S1034C, N1042D and D1246Y) were associated with antimalarial resistance against mefloquine, lumefantrine, artemether, halofantrine, quinine and chloroquine [36, 42]. Copy number and polymorphisms of the pfmdr1 gene have been investigated as molecular markers of mefloquine resistance. With the treatment of artemether-lumefantrine, a selection of N86 genotype was reported [43]. That was confirmed in areas where this ACT was implemented [44]. In same way, it was shown that ACT led to the selection of haplotypes NFD, NYD in codons 86, 184, 1246.

To the genotype Pfmdr1, followed primers and restriction enzymes are used for PCR-RFLP or for PCR followed by sequencing gene. Digestion of PCR products gives fragments of 126 and 165 bp for mutant 86Y whereas wild type N86 is not digested by restriction enzyme Afl III. In codon 1246, wild-type D1246 is digested by restriction enzyme Bgl II, giving fragments of 113 and 90 bp, whereas the mutant is not digested (Table 4). For codon 184, the genotype is assessed using the PCR followed by sequencing.

Genes, CodonsPrimer namesPrimersT°CRestriction enzymeSizes (bp)*
Pfmdr1, N86Ymdr86D1TTTACCGTTTAAATGTTTACCTGC45Afl III126 + 165
mdr86D2CCATCTTGATAAAAAACACTTCTT
Pfmdr1, D1246Ymdr1246D1AATGTAAATGAATTTTCAAACC45Bgl II113 + 90
mdr1246D2CATCTTCTCTTCCAAATTTGATA

Table 4.

Sequences of primer sets and restriction enzymes used to characterize Pfmdr1 polymorphisms.

Sizes* indicate the sizes of fragments generated after restriction enzyme digestions. T°C = hybridization temperature during PCR program.

3.2.3. Plasmodium falciparum dihydrofolate reductase (PfDHFR)

P. falciparum dihydrofolate reductase is mainly involved in the synthesis of the thymidine base. This enzyme is the target of pyrimethamine which blocked the synthesis of DNA and lead to the death of the parasite. After implementation of this drug, isolates with resistance against it were described. This parasite carried the mutations in the Pfdhfr gene. Among these, the mutation of serine to asparagine in codon 108 is the main resistance mutation. Additional mutations in codons 16, 51, 59 and 164 contribute to the increase of the resistance level of the parasite against pyrimethamine. The double mutant 108N + 51I and 108N + 59R increased the IC50 of parasites from 2 to 16 times, compared to the simple mutant N108 [45, 46]. In this way the triple mutant 51I + 59R + 108N or 59R + 108N + 164R shows the highest resistance level compared to the double mutant. Quadruple mutants exhibit a higher level of resistance compared to the triple mutant [47]. In Africa, the AIRNI haplotype is the most prevalent. PCR-RFLP was developed to genotype Pfdhr codons associated with pyrimethamine resistance. For these, primers and restriction enzyme are contained in Table 5..

Table 5.

Primer sets and RFLP conditions to genotypes PfDHFR and PfDHPS.

3.2.4. Plasmodium falciparum dihydropteroate synthase (PfDHPS)

P. falciparum dihydropteroate synthase (Pfdhps) is one of the enzymes involved in the line of thymidine synthesis. It transforms pteridine to dihydropteroate in the presence of pABA. It is coded by the gene located in chromosome 8 of P. falciparum. The implementation of sulfadoxine to treat malaria led to P. falciparum isolates’ resistance against this drug. Genotype analysis reported the association of Pfdhps mutations with sulfadoxine resistance [48]. So, mutations S436A, A437G, K540E, G581A and A613S were reported [49]. The mutation A437G increased the IC50 of isolates by five times [50]. Isolates with triple mutations S436A + A437G + K540E and S436A + A437G + A613S showed the level of sulfadoxine resistance around 9–24 times higher than the single mutant.

Triple mutant I51R59N108 in Pfdhfr and double mutants G437E540 and G437S581S in Pfdhps increase the risk of failure when treated with sulfadoxine-pyrimethamine [51]. The drug resistance against sulfadoxine and pyrimethamine could be monitored by genotyping Pfdhfr and Pfdhps according to the PCR-RFLP conditions shown in Table 5.

3.2.5. Plasmodium falciparum ATPase 6 (PfATPASE6)

The sarcoplasmic/endoplasmic reticulum Ca2+-ATPase, ortholog of P. falciparum (PfSERCA or PfATPase6), is an active Ca2+ protein transporter. This is a part of P-type ATPases enzymes that transport ions across biological membranes with the energy provided by the ATP hydrolysis. Several P-type ATPases, that were reported in P. falciparum and Pfserca, correspond to type 6. The gene coding this protein is located in chromosome 1 and contains 3687 bp. The protein allows the trafficking of calcium though the sarcoplasmic-endoplasmic membrane. Several polymorphisms such as S769N, Y243H, K431E, G110A, A2694T or E623A were reported in PfATPase 6.

PfATPase 6 has been associated with antimalarial drug resistance [52]. The most important paper on this protein was the report of its involvement in artemisinin resistance [53]. But, the role of PfATPase 6 in artemisinin resistance was strongly disputed. S769 N was associated with artemether resistance in French Guinea [53]. However, several articles investigating PfATAPase 6 in artemisinin drug resistance failed to confirm this point [54, 55, 56]. Genotyping using PCR followed by sequencing is usually used to monitor antimalarial drug resistance. Primers used are previously described by Tahar et al. [57].

3.2.5.1. P. falciparum multidrug resistance protein (PfMRP)

P. falciparum multidrug resistance protein is another marker of antimalarial drug resistance. It belongs to the C subfamily of ABC transporters containing two NBDs, two membrane-spanning domains and six transmembrane domains. Pfmrp can transport glutathione, glucuronate, as well as glucuronide and sulfate-conjugated compounds that increased susceptibility to several antimalarial drugs like chloroquine, quinine or artemisinin [58]. Pfmrp1 and Pfmrp2 are expressed in the plasma membrane in all asexual stages of the parasite. Their expression is upregulated by mefloquine and chloroquine. Pfmrp1 is a 215 kDa protein coding on chromosome 1. Mutations on codons H191Y and S437A were associated with quinoline resistance [59]. But no association was found between these polymorphisms and resistance against pyronaridine [60]. The hypothetic mechanism of involving of Pfmrp on antimalarial drug resistance is the pump efflux mechanism associated with the extrusion of glutathione. The genotyping of Pfmrp 1 and Pfmrp2 is achieved using PCR followed by sequencing. The primers used for amplification and sequencing are pfmrp-501F 5′-TTT CAA AGT ATT CAG TGG GT-3′ and pfmrp-1409R 5’-GGC ATA ATA ATT GAT GTA AA-3′.

3.2.5.2. P. falciparum Kelch 13 (PfK13)

Chinese populations used Artemisia annua to treat malaria for a long time. Over the years, artemisinin was extracted from this plant. That is a sesquiterpene lactone. Now, artemisinin is one of the best antimalarial drugs. So, WHO recommends the use of this drug in association with other antimalarial drugs, artemisinin combination therapy (ACT) for the treatment of uncomplicated malaria. Since the 1990s, the use of artemisinin was highly intensified in Asia. Consequently, the delay of P. falciparum clearance was reported after treatment with artemisinin which translates to artemisinin resistance [56]. These slow clearance rates are associated with enhanced survival rates of ring-stage parasites briefly exposed in vitro to dihydroartemisinin. Recently, a large-scale study identified molecular markers of this resistance: polymorphisms on propeller kelch 13 (K13) [61]. That is a region on chromosome 13 (Figure 8) [62].

Figure 8.

P. falciparum kelch13 (K13) protein. The parasite K13 protein consists of plasmodium-specific sequences, a BTB-POZ domain and six kelch domains that are predicted to form a six-blade propeller. In the structural model, the original M476I mutation discovered by Ariey et al. [61] and six other mutations associated with artemisinin resistance (all details of this figure was pulled of Fairhurst and Dondrop [62]) are shown.

Several investigations on K13 genotyping reported that mutations M476I, Y493H, R539T, I543T, P553L and C580Y conferred a greater artemisinin resistance [63]. Other mutations F446I and A578S were described in PfK13. A578S, widespread in Africa, is not associated with artemisinin resistance. These genotypes are investigated by PCR sequencing.

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4. Conclusions

Due to the nucleotide specificities of each Plasmodium species and the molecular changes associated with antimalarial drug resistance, genotyping is used for Plasmodium species diagnosis and monitoring antimalarial drug resistance. This genotyping could be achieved by specific PCR, PCR-RFLP, PCR-sequencing or RT-PCR of some molecular markers.

References

  1. 1. WHO. World Malaria Report 2016. 2017. p. 196. ISBN: 978-92-4-156552-3
  2. 2. Hawley WA, Phillips-Howard PA, ter Kuile FO, Terlouw DJ, Vulule JM, Ombok M, Nahlen BL, Gimnig JE, Kariuki SK, Kolczak MS. Community-wide effects of permethrin-treated bed nets on child mortality and malaria morbidity in western Kenya. The American Journal of Tropical Medicine and Hygiene. 2003;68:121-127
  3. 3. Bull PC, Marsh K. The role of antibodies to Plasmodium falciparum-infected-erythrocyte surface antigens in naturally acquired immunity to malaria. Trends in Microbiology. 2002;10:55-58
  4. 4. Berendt A, Ferguson D, Newbold C. Sequestration in Plasmodium falciparum malaria: Sticky cells and sticky problems. Parasitology Today. 1990;6:247-254
  5. 5. Kyes S, Horrocks P, Newbold C. Antigenic variation at the infected red cell surface in malaria. Annual Review of Microbiology. 2001;55:673-707
  6. 6. Loy DE, Liu W, Li Y, Learn GH, Plenderleith LJ, Sundararaman SA, Sharp PM, Hahn BH. Out of Africa: Origins and evolution of the human malaria parasites Plasmodium falciparum and Plasmodium vivax. International Journal for Parasitology. 2017;47:87-97
  7. 7. Cox-Singh J, Singh B. Knowlesi malaria: Newly emergent and of public health importance? Trends in Parasitology. 2008;24:406-410
  8. 8. Lee KS, Divis PC, Zakaria SK, Matusop A, Julin RA, Conway DJ, Cox-Singh J, Singh B. Plasmodium knowlesi: Reservoir hosts and tracking the emergence in humans and macaques. PLoS Pathogens. 2011;7:e1002015
  9. 9. Cox-Singh J, Davis TM, Lee K-S, Shamsul SS, Matusop A, Ratnam S, Rahman HA, Conway DJ, Singh B. Plasmodium knowlesi malaria in humans is widely distributed and potentially life threatening. Clinical Infectious Diseases. 2008;46:165-171
  10. 10. Ngassa Mbenda HG, Das A. Molecular evidence of Plasmodium vivax mono and mixed malaria parasite infections in Duffy-negative native Cameroonians. PLoS One. 2014;9:e103262
  11. 11. Russo G, Faggioni G, Paganotti GM, Djeunang Dongho GB, Pomponi A, De Santis R, Tebano G, Mbida M, Sanou Sobze M, Vullo V, Rezza G, Lista FR. Molecular evidence of Plasmodium vivax infection in Duffy negative symptomatic individuals from Dschang, West Cameroon. Malaria Journal. 2017;16:74
  12. 12. Mueller I, Zimmerman PA, Reeder JC. Plasmodium malariae and Plasmodium ovale—The ‘bashful’malaria parasites. Trends in Parasitology. 2007;23:278-283
  13. 13. Ayala F, Escalante A, Rich S. Evolution of Plasmodium and the recent origin of the world populations of Plasmodium falciparum. Parassitologia. 1999;41:55-68
  14. 14. Collins WE, Jeffery GM. Plasmodium ovale: Parasite and disease. Clinical Microbiology Reviews. 2005;18:570-581
  15. 15. Makler MT, Palmer CJ, Ager AL. A review of practical techniques for the diagnosis of malaria. Annals of Tropical Medicine and Parasitology. 1998;92:419-433
  16. 16. Manwell RD. Malaria infections by four species of Plasmodium in the duck and chicken, and resulting parasite modifications. American Journal of Hygiene. 1943;38:211-222
  17. 17. Jordan HB. The effect of host constitution on the development of Plasmodium floridense. The Journal of Eukaryotic Microbiology. 1975;22:241-244
  18. 18. Mens PF, Schoone GJ, Kager PA, Schallig HD. Detection and identification of human Plasmodium species with real-time quantitative nucleic acid sequence-based amplification. Malaria Journal. 2006;5:80
  19. 19. Moody A. Rapid diagnostic tests for malaria parasites. Clinical Microbiology Reviews. 2002;15:66-78
  20. 20. Trape JF. The public health impact of chloroquine resistance in Africa. The American Journal of Tropical Medicine and Hygiene. 2001;64:12-17
  21. 21. Payne D. Spread of chloroquine resistance in Plasmodium falciparum. Parasitology Today. 1987;3:241-246
  22. 22. Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, Thaithong S, Brown KN. High sensitivity of detection of human malaria parasites by the use of nested polymerase chain reaction. Molecular and Biochemical Parasitology. 1993;61:315-320
  23. 23. Johnston SP, Pieniazek NJ, Xayavong MV, Slemenda SB, Wilkins PP, da Silva AJ. PCR as a confirmatory technique for laboratory diagnosis of malaria. Journal of Clinical Microbiology. 2006;44:1087-1089
  24. 24. Compton J. Nucleic acid sequence-based amplification. Nature. 1991;350:91-92
  25. 25. Kievits T, van Gemen B, van Strijp D, Schukkink R, Dircks M, Adriaanse H, Malek L, Sooknanan R, Lens P. NASBATM isothermal enzymatic in vitro nucleic acid amplification optimized for the diagnosis of HIV-1 infection. Journal of Virological Methods. 1991;35:273-286
  26. 26. Schoone GJ, Oskam L, Kroon NC, Schallig HD, Omar SA. Detection and quantification of Plasmodium falciparum in blood samples using quantitative nucleic acid sequence-based amplification. Journal of Clinical Microbiology. 2000;38:4072-4075
  27. 27. Schneider P, Schoone G, Schallig H, Verhage D, Telgt D, Eling W, Sauerwein R. Quantification of Plasmodium falciparum gametocytes in differential stages of development by quantitative nucleic acid sequence-based amplification. Molecular and Biochemical Parasitology. 2004;137:35-41
  28. 28. Schneider P, Wolters L, Schoone G, Schallig H, Sillekens P, Hermsen R, Sauerwein R. Real-time nucleic acid sequence-based amplification is more convenient than real-time PCR for quantification of Plasmodium falciparum. Journal of Clinical Microbiology. 2005;43:402-405
  29. 29. Van Gemen B, van Beuningen R, Nabbe A, van Strijp D, Jurriaans S, Lens P, Kievits T. A one-tube quantitative HIV-1 RNA NASBA nucleic acid amplification assay using electrochemiluminescent (ECL) labelled probes. Journal of Virological Methods. 1994;49:157-167
  30. 30. Ollomo B, Durand P, Prugnolle F, Douzery E, Arnathau C, Nkoghe D, Leroy E, Renaud FA. New malaria agent in African hominids. PLoS Pathogens. 2009;5:e1000446
  31. 31. Prugnolle F, Durand P, Neel C, Ollomo B, Ayala FJ, Arnathau C, Etienne L, Mpoudi-Ngole E, Nkoghe D, Leroy E, Delaporte E, Peeters M, Renaud F. African great apes are natural hosts of multiple related malaria species, including Plasmodium falciparum. Proceedings of the National Academy of Sciences of the United States of America. 2010;107:1458-1463
  32. 32. Boundenga L, Ollomo B, Rougeron V, Mouele LY, Mve-Ondo B, Delicat-Loembet LM, Moukodoum ND, Okouga AP, Arnathau C, Elguero E, Durand P, Liégeois F, Boué V, Motsch P, Le Flohic G, Ndoungouet A, Paupy C, Ba CT, Renaud F, Prugnolle F. Diversity of malaria parasites in great apes in Gabon. Malaria Journal. 2015;14:111
  33. 33. Boundenga L, Makanga B, Ollomo B, Gilabert A, Rougeron V, Mve-Ondo B, Arnathau C, Durand P, Moukodoum ND, Okouga AP, Delicat-Loembet L, Yacka-Mouele L, Rahola N, Leroy E, Ba CT, Renaud F, Prugnolle F, Paupy C. Haemosporidian parasites of antelopes and other vertebrates from Gabon, Central Africa. PLoS One. 2016;11:e0148958
  34. 34. Steenkeste N, Incardona S, Chy S, Duval L, Ekala M-T, Lim P, Hewitt S, Sochantha T, Socheat D, Rogier C, Mercereau-Puijalon O, Fandeur T, Ariey F. Towards high-throughput molecular detection of Plasmodium. New approaches and molecular markers. Malaria Journal. 2009;8:86
  35. 35. Tanomsing N, Imwong M, Theppabutr S, Pukrittayakamee S, Day NP, White NJ, Snounou G. Accurate and sensitive detection of Plasmodium species in humans by use of the dihydrofolate reductase-thymidylate synthase linker region. Journal of Clinical Microbiology. 2010;48:3735-3737
  36. 36. Basco LK, Ringwald P, Thor R, Doury JC, Le Bras J. Activity in vitro of chloroquine, cycloguanil, and mefloquine against African isolates of Plasmodium falciparum: Presumptive evidence for chemoprophylactic efficacy in central and West Africa. Transactions of the Royal Society of Tropical Medicine and Hygiene. 1995;89:657-658
  37. 37. Beshir K, Sutherland CJ, Merinopoulos I, Durrani N, Leslie T, Rowland M, Hallett RL. Amodiaquine resistance in Plasmodium falciparum malaria in Afghanistan is associated with the pfcrt SVMNT allele at codons 72 to 76. Antimicrobial Agents and Chemotherapy. 2010;54:3714-3716
  38. 38. Warhurst DC, Steele JC, Adagu IS, Craig JC, Cullander C. Hydroxychloroquine is much less active than chloroquine against chloroquine-resistant Plasmodium falciparum, in agreement with its physicochemical properties. The Journal of Antimicrobial Chemotherapy. 2003;52:188-193
  39. 39. Sa JM, Twu O. Protecting the malaria drug arsenal: Halting the rise and spread of amodiaquine resistance by monitoring the PfCRT SVMNT type. Malaria Journal. 2010;9:374
  40. 40. Valderramos SG, Fidock DA. Transporters involved in resistance to antimalarial drugs. Trends in Pharmacological Sciences. 2006;27:594-601
  41. 41. Mwai L, Kiara SM, Abdirahman A, Pole L, Rippert A, Diriye A, Bull P, Marsh K, Borrmann S, Nzila A. In vitro activities of piperaquine, lumefantrine, and dihydroartemisinin in Kenyan Plasmodium falciparum isolates and polymorphisms in pfcrt and pfmdr1. Antimicrobial Agents and Chemotherapy. 2009;53:5069-5073
  42. 42. Bray PG, Ward SA. A comparison of the phenomenology and genetics of multidrug resistance in cancer cells and quinoline resistance in Plasmodium falciparum. Pharmacology & Therapeutics. 1998;77:1-28
  43. 43. Sisowath C, Stromberg J, Martensson A, Msellem M, Obondo C, Bjorkman A, Gil JP. In vivo selection of Plasmodium falciparum pfmdr1 86N coding alleles by artemether-lumefantrine (Coartem). The Journal of Infectious Diseases. 2005;191:1014-1017
  44. 44. Lekana-Douki JB, Dinzouna Boutamba SD, Zatra R, Zang Edou SE, Ekomy H, Bisvigou U, Toure-Ndouo FS. Increased prevalence of the Plasmodium falciparum Pfmdr1 86N genotype among field isolates from Franceville, Gabon after replacement of chloroquine by artemether-lumefantrine and artesunate-mefloquine. Infection, Genetics and Evolution. 2011;11:512-517
  45. 45. Peterson DS, Walliker D, Wellems TE. Evidence that a point mutation in dihydrofolate reductase-thymidylate synthase confers resistance to pyrimethamine in falciparum malaria. Proceedings of the National Academy of Sciences of the United States of America. 1988;85:9114-9118
  46. 46. Triglia T, Cowman AF. Primary structure and expression of the dihydropteroate synthetase gene of Plasmodium falciparum. Proceedings of the National Academy of Sciences of the United States of America. 1994;91:7149-7153
  47. 47. Nzila-Mounda A, Mberu EK, Sibley CH, Plowe CV, Winstanley PA, Watkins WM. Kenyan Plasmodium falciparum field isolates: Correlation between pyrimethamine and chlorcycloguanil activity in vitro and point mutations in the dihydrofolate reductase domain. Antimicrobial Agents and Chemotherapy. 1998;42:164-169
  48. 48. Osman ME, Mockenhaupt FP, Bienzle U, Elbashir MI, Giha HA. Field-based evidence for linkage of mutations associated with chloroquine (pfcrt/pfmdr1) and sulfadoxine–pyrimethamine (pfdhfr/pfdhps) resistance and for the fitness cost of multiple mutations in P. falciparum. Infection, Genetics and Evolution. 2007;7:52-59
  49. 49. Triglia T, Menting JG, Wilson C, Cowman AF. Mutations in dihydropteroate synthase are responsible for sulfone and sulfonamide resistance in Plasmodium falciparum. Proceedings of the National Academy of Sciences of the United States of America. 1997;94:13944-13949
  50. 50. Triglia T, Wang P, Sims PF, Hyde JE, Cowman AF. Allelic exchange at the endogenous genomic locus in Plasmodium falciparum proves the role of dihydropteroate synthase in sulfadoxine-resistant malaria. The EMBO Journal. 1998;17:3807-3815
  51. 51. Picot S, Bienvenu AL, Konate S, Sissoko S, Barry A, Diarra E, Bamba K, Djimde A, Doumbo OK. Safety of epoietin beta-quinine drug combination in children with cerebral malaria in Mali. Malaria Journal. 2009;8:169
  52. 52. Eckstein-Ludwig U, Webb R, Van Goethem I, East J, Lee A, Kimura M, O’neill P, Bray P, Ward S, Krishna S. Artemisinins target the SERCA of Plasmodium falciparum. Nature. 2003;424:957-961
  53. 53. Jambou R, Legrand E, Niang M, Khim N, Lim P, Volney B, Ekala MT, Bouchier C, Esterre P, Fandeur T, Mercereau-Puijalon O. Resistance of Plasmodium falciparum field isolates to in-vitro artemether and point mutations of the SERCA-type PfATPase6. Lancet. 2005;366:1960-1963
  54. 54. Fairhurst RM, Nayyar GM, Breman JG, Hallett R, Vennerstrom JL, Duong S, Ringwald P, Wellems TE, Plowe CV, Dondorp AM. Artemisinin-resistant malaria: Research challenges, opportunities, and public health implications. The American Journal of Tropical Medicine and Hygiene. 2012;87:231-241
  55. 55. Kiaco K, Teixeira J, Machado M, do Rosario V, Lopes D. Evaluation of artemether-lumefantrine efficacy in the treatment of uncomplicated malaria and its association with pfmdr1, pfatpase6 and K13-propeller polymorphisms in Luanda, Angola. Malaria Journal. 2015;14:504
  56. 56. Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, Lwin KM, Ariey F, Hanpithakpong W, Lee SJ, Ringwald P, Silamut K, Imwong M, Chotivanich K, Lim P, Herdman T, An SS, Yeung S, Singhasivanon P, Day NP, Lindegardh N, Socheat D, White NJ. Artemisinin resistance in Plasmodium falciparum malaria. The New England Journal of Medicine. 2009;361:455-467
  57. 57. Tahar R, Ringwald P, Basco LK. Molecular epidemiology of malaria in Cameroon. XXVIII. In vitro activity of dihydroartemisinin against clinical isolates of Plasmodium falciparum and sequence analysis of the P. falciparum ATPase 6 gene. The American Journal of Tropical Medicine and Hygiene. 2009;81:13-18
  58. 58. Sanchez CP, Dave A, Stein WD, Lanzer M. Transporters as mediators of drug resistance in Plasmodium falciparum. International Journal for Parasitology. 2010;40:1109-1118
  59. 59. Pirahmadi S, Zakeri S, Afsharpad M, Djadid ND. Mutation analysis in pfmdr1 and pfmrp1 as potential candidate genes for artemisinin resistance in Plasmodium falciparum clinical isolates 4years after implementation of artemisinin combination therapy in Iran. Infection, Genetics and Evolution. 2013;14:327-334
  60. 60. Pradines B, Briolant S, Henry M, Oeuvray C, Baret E, Amalvict R, Didillon E, Rogier C. Absence of association between pyronaridine in vitro responses and polymorphisms in genes involved in quinoline resistance in Plasmodium falciparum. Malaria Journal. 2010;9:339
  61. 61. Ariey F, Witkowski B, Amaratunga C, Beghain J, Langlois AC, Khim N, Kim S, Duru V, Bouchier C, Ma L, Lim P, Leang R, Duong S, Sreng S, Suon S, Chuor CM, Bout DM, Ménard S, Rogers WO, Genton B, Fandeur T, Miotto O, Ringwald P, Le Bras J, Berry A, Barale JC, Fairhurst RM, Benoit-Vical F, Mercereau-Puijalon O, Ménard D. A molecular marker of artemisinin-resistant Plasmodium falciparum malaria. Nature. 2014;505:50-55
  62. 62. Fairhurst RM, Dondorp AM. Artemisinin-resistant Plasmodium falciparum malaria. Microbiology Spectrum. 2016;4:EI10-0013-2016
  63. 63. Straimer J, Gnädig NF, Witkowski B, Amaratunga C, Duru V, Ramadani AP, Dacheux M, Khim N, Zhang L, Lam S, Gregory PD, Urnov FD, Mercereau-Puijalon O, Benoit-Vical F, Fairhurst RM, Ménard D, Fidock DA. Drug resistance. K13-propeller mutations confer artemisinin resistance in Plasmodium falciparum clinical isolates. Science. 2015;347:428-431

Written By

Jean Bernard Lekana-Douki and Larson Boundenga

Submitted: October 10th, 2017 Reviewed: March 19th, 2018 Published: November 5th, 2018