Plasmodium Genomics and Genetics: New Insights into Malaria Pathogenesis, Drug Resistance, Epidemiology, and Evolution

Protozoan Plasmodium parasites are the causative agents of malaria, a deadly disease that continues to afflict hundreds of millions of people every year. Infections with malaria parasites can be asymptomatic, with mild or severe symptoms, or fatal, depending on many factors such as parasite virulence and host immune status. Malaria can be treated with various drugs, with artemisinin-based combination therapies (ACTs) being the first-line choice.

SUMMARY

Protozoan Plasmodium parasites are the causative agents of malaria, a deadly disease that continues to afflict hundreds of millions of people every year. Infections with malaria parasites can be asymptomatic, with mild or severe symptoms, or fatal, depending on many factors such as parasite virulence and host immune status. Malaria can be treated with various drugs, with artemisinin-based combination therapies (ACTs) being the first-line choice. Recent advances in genetics and genomics of malaria parasites have contributed greatly to our understanding of parasite population dynamics, transmission, drug responses, and pathogenesis. However, knowledge gaps in parasite biology and host-parasite interactions still remain. Parasites resistant to multiple antimalarial drugs have emerged, while advanced clinical trials have shown partial efficacy for one available vaccine. Here we discuss genetic and genomic studies of Plasmodium biology, host-parasite interactions, population structures, mosquito infectivity, antigenic variation, and targets for treatment and immunization. Knowledge from these studies will advance our understanding of malaria pathogenesis, epidemiology, and evolution and will support work to discover and develop new medicines and vaccines.

INTRODUCTION

Malaria is a deadly disease caused by Plasmodium species, a large and diverse taxonomic group that includes parasites of birds, reptiles, rodents, monkeys, apes, and humans. Hundreds of millions of people are infected annually across tropical and subtropical regions, largely Africa, South and Central America, India, Southeast Asia, and Oceania (1–3). In addition to studies of the major species infecting humans (Plasmodium falciparum, Plasmodium vivax, Plasmodium malariae, Plasmodium ovale, and Plasmodium knowlesi), malaria research employs models including infections of nonhuman primates (e.g., Plasmodium reichenowi [infecting chimpanzees] and Plasmodium cynomolgi and P. knowlesi [infecting Macaca fascicularis]), rodents (e.g., Plasmodium berghei, Plasmodium chabaudi, and Plasmodium yoelii [infecting Mus musculus]), and birds (e.g., Plasmodium gallinaceum [infecting Gallus gallus domesticus]) (4–10).

Malaria parasites are transmitted by mosquitoes. The sporozoite forms inoculated by mosquito feeding produce blood stages that cause a wide range of clinical outcomes, ranging from no symptom at all to severe malaria and death. The onset of malaria often manifests by constitutional symptoms similar to those of “the flu,” such as fever, chills, headache, dizziness, back pain, myalgia, joint and bone pains, cough, chest pain, weakness, prostration, nausea, vomiting, and diarrhea. Progression to severe malaria happens in a small fraction of patients, with life-threatening or outright lethal developments of coma (cerebral malaria), pulmonary edema, acute renal failure, severe anemia, acidosis, hypoglycemia, and/or bleeding (11). Interestingly, many adults in regions of endemicity in African countries often carry parasites in the bloodstream, including gametocytes that are infectious to mosquitoes, but do not have clinical symptoms and are thus considered “asymptomatic.” However, some adults can develop recurrent episodes of symptomatic parasitemia, chronic anemia, maternal morbidity, coinfection with invasive bacterial disease, and cognitive impairment (12).

Sporozoites inoculated into the skin by mosquito feeding transit through the dermis and enter the bloodstream, where they are carried to the liver (13). The parasites then enter hepatocytes and complete a cycle of schizogony, growing and dividing into exoerythrocytic merozoites. In P. vivax and P. ovale infections, some parasites become dormant forms, called “hypnozoites,” after invasion of their host hepatocytes (14), where they can remain quiescent for months or years before entering schizogony (15, 16); P. falciparum, P. malariae, and P. knowlesi parasites do not produce such hypnozoites. After schizogony, merozoites are released from the hepatocytes into bloodstream, where they invade erythrocytes. There they start new cycles of schizogony, with each cycle progressing from morphologically characteristic ring-stage forms to trophozoite and then schizont stages. Mature schizonts have a segmented appearance from mitotic divisions that produce many daughter merozoites, each for invasion of a new erythrocyte upon its release from the consumed and destroyed host cell. The duration of each erythrocytic cycle is approximately 48 h for P. falciparum, P. vivax, and P. ovale, 24 h for P. knowlesi, and 72 h for P. malariae.

After several erythrocytic cycles and under likely stimulations from host immune responses, fever, and even antimalarial therapy, some of the ring forms develop into male and female gametocytes that are infective to mosquitoes (schizogenic replication is asexual). When mosquitoes take blood meals from the infected patient, male and female gametes differentiated from gametocytes fertilize in the mosquito midgut to produce zygotes that develop into motile ookinetes. Ookinetes penetrate through the mosquito midgut wall and develop into oocysts. Approximately 2 weeks later (depending on the parasite species), thousands of sporozoites result from schizogony in each oocyst. The mature sporozoites then migrate via hemolymph circulation and penetrate the mosquito salivary gland, where they await injection into another person when the mosquito takes a blood meal again (17).

Various drugs are used for malaria prevention and cure. These include quinine (QN), chloroquine (CQ), amodiaquine, piperaquine (PPQ), mefloquine (MQ), lumefantrine (LUM), pyrimethamine (PYR), proguanil, sulfadoxine, atovaquone, primaquine, and artemisinin and its derivatives (ART) (18). Artemisinin-based combination therapies (ACTs) are now recommended worldwide for treatment of P. falciparum infections (19). Primaquine and the recently approved tafenoquine are the only antimalarials available against liver-stage parasites and hypnozoites (20, 21). Unfortunately, resistance (or treatment failure) has been reported from nearly all malarious regions (22). Efforts are being made to discover new antimalarial drugs and to understand the molecular mechanisms of drug resistances (23).

THE PLASMODIUM GENOMES

Genome Sequences and Characteristics of Important Plasmodium Species

Since completion of the first draft sequence of the P. falciparum 3D7 genome in 2002 (5), genomic research on malaria parasites has advanced rapidly in step with next-generation sequencing (NGS) technologies and reduction in costs (24). In addition to the sequences of important species that infect humans (4, 6, 8, 25–27), public databases now provide the genome information of primate, rodent, and avian parasites, including those of widely used disease models such as P. berghei, P. chabaudi, P. yoelii, P. reichenowi, P. cynomolgi, P. relictum, and P. gallinaceum (Table 1) (7, 28–33). The evolutionary relationships of Plasmodium spp. have been extensively investigated using various DNA sequences, with clustering of parasite species largely depending on their host origins (34–36). Data from large numbers of laboratory lines and field isolates of P. falciparum and P. vivax have been deposited from studies of genome diversity, parasite evolution, population genetics, and drug resistances (37–44), leading to generation of an updated reference P. falciparum genome with greatly improved annotation (45). More recently, single-cell sequencing techniques have begun to yield exciting information for mixed infections, genetic recombination, and parasite differentiation (46–48).

TABLE 1.

Summary of genome sequence statistics for important human and animal Plasmodium species^a

Host	Parasite species or strain	No. of PIRb genes	Genome size (Mb)	GC content (%)	No. of scaffolds	No. of predicted genesc	No. of predicted proteinsc	Data release date (mo/day/yr)	Data update date (mo/day/yr)	Reference(s)
Humans	P. falciparum 3D7	189	23.2	19.3	14	5,712	5,460	10/2/02	3/31/16	27, 29
	P. vivax	1,212	29.1	39.7	374	6,830	6,677	10/6/16	10/6/16	8, 285
	P. ovale	>2,100	33.5	29.4	779	6,986	6,228	8/3/16	9/15/16	33
	P. ovale subsp. wallikeri	1,375	33.5	28.9	1,914	8,582	8,421	6/16/16	6/16/16	8
	P. ovale subsp. curtisi	1,949	33.5	28.4	4,025	7,280	7,162	6/16/16	6/16/16	8
	P. malariae	255	33.6	24.7	63	6,709	6,573	9/23/16	9/23/16	8
	P. knowlesi	71	24.4	38.7	28	5,483	5,323	5/15/17	5/15/17	6, 8
	P. inui San Antonio		27.4	42.4	323	5,879	5,832	1/31/14	4/31/14	https://plasmodb.org/plasmo/app/record/organism/NCBITAXON_1237626
Chimpanzees	P. reichenowi	351	24.0	19.3	261	5,909	5,741	3/25/18	3/25/18	29
Old World monkeys	P. cynomolgi B	265	26.2	40.4	1,663	5,776	5,716	5/31/12	9/16/15	31
	P. coatneyi Hackeri	771	27.7	39.7	14	5,575	5,516	7/6/16	2/2/17	https://plasmodb.org/plasmo/app/record/organism/TMPTX_pcoaHackeri
Rodents	P. yoelii 17X	980	22.8	21.1	154	6,257	6,091	8/27/14	10/26/17	28, 286
	P. berghei ANKA	217	18.5	22.1	100	5,245	4,928	8/27/14	10/24/17	28, 286
	P. chabaudi AS	201	18.9	23.6	39	5,364	5,217	8/27/14	5/13/16	28, 286
	P. vinckei		18.2	23.4	49	5,009	4,954	6/16/14	7/30/14	287, 288
Birds	P. relictum SGS1	4	22.6	18.4	498	5,306	5,138	11/17/16	1/9/17	30
	P. gallinaceum 8A	20	23.8	17.8	152	5,439	5,280		1/9/17	30, 289, 290

^aAll information was obtained through PubMed searches. More sequences from different strains of each species may be available.

^bPIR, Plasmodium interspersed repeat.

^cIncludes updated information from PlasmoDB.

Except for a short diploid phase after fertilization in the mosquito midgut, Plasmodium parasites are haploid throughout their life cycle. The genomes of different species are roughly 2 to 3 times larger than that of brewer’s yeast (Saccharomyces cerevisiae), ranging from 20 to 35 megabases (Mb) and containing 14 chromosomes, a circular plastid genome of ∼35 kb, and multiple copies of a 6-kb mitochondrial DNA (Table 1). Comparison of genomes from different species showed that homologous genes are often found in syntenic blocks arranged in different orders among different chromosomes (49, 50). The adenine-thymine (AT) contents of Plasmodium spp. can also be very different: e.g., ∼80% AT in P. falciparum, P. reichenowi, and P. gallinaceum; ∼75% AT in rodent malaria parasites; and ∼60% AT in P. vivax, P. knowlesi, and P. cynomolgi (Table 1). AT content is often higher in introns and intergenic noncoding regions than in protein-coding exons, with an average of 80.6% AT for the whole P. falciparum genome versus 86.5% for noncoding sequences (27). The high AT content of P. falciparum reflects large numbers of low-complexity regions, simple sequence repeats, and microsatellites, as well as a highly skewed codon usage bias (51–54). Polymorphisms of AT-rich repeats provide abundant markers for linkage mapping of drug resistance genes (55–58) and for tracing the evolution and structure of parasite populations (59–61).

Gene Families Playing Important Roles in Parasite Development, Virulence, and Transmission

Malaria parasite genomes carry multigene families that serve important roles in parasite interactions with their hosts, including antigenic variation, signaling, protein trafficking, and adhesion (27, 62, 63). Several polymorphic families are found among species of the Laverania subgenus (P. falciparum [infecting humans], P. reichenowi, P. gaboni, and P. billcollinsi [infecting chimpanzees], and P. praefalciparum, P. adleri, and P. blacklocki [infecting gorillas]) (64). Among the gene families, the genes encoding P. falciparum erythrocyte membrane protein 1 (PfEMP1) (65, 66) have been studied most extensively. Each individual P. falciparum parasite carries a unique set of 50 to 150 copies of the var genes in its genome, where switches of gene expression produce antigenic variation (67–69). PfEMP1 plays an important role in the pathogenesis of clinical developments such as in cerebral and placental malaria, in which it mediates the cytoadherence of infected red blood cells (iRBCs; infected erythrocytes) in the deep tissues (70–72). Different PfEMP1 molecules bind to various host molecules, including α2-macroglobulin, CD36, chondroitin sulfate A (CSA), complement 1q, CR1, E-selectins and P-selectins, endothelial protein C receptor (EPCR), heparan sulfate, ICAM1, IgM, IgG, PECAM1, thrombospondin (TSP), and VCAM1 (62). Such binding leads to activation of various host inflammatory responses. Hemoglobinopathies, including the hemoglobin C and hemoglobin S trait conditions, interfere with PfEMP1 display in knob structures of the iRBCs. This poor display of PfEMP1 on the host cell surface offers protection against malaria by reducing the cytoadherence and activation of inflammatory processes that promote the development of severe disease (73–75).

A second group of genes receiving recent attention is the large Plasmodium interspersed repeat (pir) multigene family (63). Members of the pir family are named differently by parasite species: yir in P. yoelii, bir in P. berghei, vir in P. vivax, and so on. Several P. falciparum gene families (stevor, rif, and PfMC-2TM) are classified with pir by their similar gene structures (63), which characteristically include a short first exon, a long second exon, and a third exon encoding a transmembrane domain. In a recent study, the pir genes from P. chabaudi (cir) were shown to be expressed in different cellular locations, within and on the surface of iRBCs, and in merozoites (76). Additionally, a subset of recombinant CIR (P. chabaudi) proteins bound to mouse red blood cells (RBCs), suggesting a role for CIR in rosette formation and/or invasion. In another study, sequences of the P. berghei fam-a, fam-b, and bir multigene families were compared with those of P. yoelii and P. chabaudi; expressions of mRNA and selected proteins were analyzed (77). The majority of fluorescently tagged proteins were transported into the iRBC cytoplasm and into the parasitophorous vacuole of the liver stage, suggesting potential functions in parasite development and/or in manipulating the host immune response. Interestingly, Fam-A proteins carrying a steroidogenic acute regulatory-related lipid transfer (START) domain were found to transfer phosphatidylcholine in vitro, suggesting that these proteins transport host phosphatidylcholine for parasite membrane synthesis. Using the P. chabaudi AS parasite and C57BL/6 mice, Brugat et al. (78) showed that chronic infections were characterized by expression of distinctive clusters of cir genes, independent of adaptive immunity, and that the initial composition of parasite population dictated chronicity and virulence of infection. These observations suggest involvement of some cir genes in regulating the establishment of chronic infections and virulence. In an earlier study, vector transmission was shown to influence subsequent P. chabaudi asexual blood-stage infection (79). Transmission through the mosquito was also associated with a reduction in the severity of the resulting infection and altered the mammalian immune response as evidenced by decreased levels of circulating proinflammatory chemokines and cytokines during the acute stage (79). Overall, the expression of the cir multigene family was increased following mosquito transmission, as opposed to the hierarchical expression pattern observed in non-vector-transmitted parasites (80). Malaria parasites devote large portions of their genomes to gene families that ensure evasion of host immune defenses and protection of molecular processes essential to infection. These families emphasize the importance of research on their roles in parasite-host interactions and virulence, despite the difficulties inherent to their investigation.

Another polymorphic gene family worth mentioning is a group of 14 genes encoding proteins with six cysteines (6-Cys) (81). These proteins often localize on the parasite surface interacting with host proteins, are likely under various selection pressures such as immunity, and are promising vaccine candidates (82–85). These genes are expressed at different parasite developmental stages: pf230, pf48/45, pf230p, pf47, and pfPSOP12 are expressed in P. falciparum sexual stages; pf52, pf36, pfLISP2, and pfB9 are expressed in the preerythrocytic stages; and pf12, pf12p, pf41, pf38, and pf92 are expressed in asexual erythrocytic stages (81). The proteins have diverse functions. Pf48/45, Pf230, and Pf47 share unique disulfide-bonded structures and have been shown to play an essential role in parasite fertilization (86). Both Pf48/45 and Pf47 were shown to have high inbreeding coefficients, suggesting that these two molecules may play a significant role in mating interaction (87). Indeed, the pf47 gene was shown to play a critical role in the evasion of the Anopheles gambiae innate immune response, and replacement of the pf47 haplotypes in a P. falciparum isolate changed its compatibility to a different mosquito species (88, 89). PfP52 and PfP36 mediate sporozoite invasion of hepatocytes (90, 91). The proteins expressed in asexual stages are generally polymorphic and/or under selection, suggesting that they could be targets of the host immune response; however, their functions in parasite development remain largely unknown (81).

GENOME POLYMORPHISMS AND LINKAGE ANALYSIS

Early Work on Restriction Fragment Length Polymorphism

The polymorphic nature of Plasmodium genomes was recognized before large-scale genome sequencing was available. Early studies of the smaller P. falciparum chromosomes by pulsed-field gel electrophoresis (PFGE) showed size polymorphisms involving tens to hundreds of kilobases (92–94). Successful separation of the larger chromosomes by PFGE subsequently defined the nuclear content of 14 chromosomes (95) and, by analysis with rare-cutting restriction enzymes, demonstrated that the chromosome structure in P. falciparum is largely conserved in central regions but extensively polymorphic in both length and sequence near the telomeres (96–98). Much of this subtelomeric variation was explained by recombination within blocks of repetitive sequences and families of genes (99, 100). At a time when whole-genome sequencing was not available, restriction fragment length polymorphism (RFLP) analysis was employed to map the linkage groups of chromosomes in different parasite strains (101). These advances supported the localization of genes in P. falciparum crosses governing the heritable traits of PYR and CQ resistance (CQR) (102, 103).

Simple Sequence Repeats (Microsatellites)

The frequency of simple sequence repeats (microsatellites) in P. falciparum is estimated to be approximately one polymorphic microsatellite per kb DNA, a high rate that may involve the AT-rich nature of the genome (52, 53). This abundance of microsatellites facilitated the generation of high-density genetic linkage maps (53, 104) and expedited PCR-based mapping of the CQR locus in P. falciparum (57). Additionally, the rich wealth of genome variations supported development of rapid genotyping methods that have greatly facilitated parasite characterization and studies of P. falciparum epidemiology, population structure, and transmission (105, 106). Hundreds of microsatellites have also been identified from the P. yoelii genome and have been employed to type various strains and map genes contributing to virulence (104, 107, 108). Microsatellites seems to be less frequent in other Plasmodium species that have genomes with lower AT contents (104, 107–109).

Single Nucleotide Polymorphisms

Advances in DNA sequencing technologies have also facilitated the identification of large numbers of single nucleotide polymorphisms (SNPs). Early work compared amplified DNA segments from 204 P. falciparum genes on chromosome 3 (110). A total of 403 polymorphic sites (238 SNPs and 165 microsatellites) were identified from five parasite clones, establishing a chromosome-wide map with one polymorphic marker per 2.3 kb. Similarly, 191 SNPs and 44 size polymorphisms were identified from five P. vivax isolates after amplification and sequencing a DNA segment of ∼100 kb of DNA syntenic to a segment of P. falciparum chromosome 3 (109). Later, three independent laboratories comprehensively sequenced additional P. falciparum isolates, establishing high-density SNP maps for the parasite (111–113). Using these SNP maps, signatures of selection and drug-selective sweeps were identified from field parasite populations.

Copy Number Variation

Another important type of genomic variation in Plasmodium is copy number variation (CNV). CNVs have been found to affect important traits of drug resistance (114–119), erythrocyte invasion (120, 121), cytoadherence (122), and transcriptional regulation (123, 124). Increased gene copy numbers and levels of expression of P. falciparum multiple-drug resistance gene 1 (pfmdr1) were associated with decreased susceptibility to MQ and halofantrine (114). Amplification of the P. falciparum GTP-cyclohydrolase I (pfgch1) gene, associated with resistance to antifolate drugs, has also been described (44, 117, 125). Genomic breakpoints are frequently located in AT-rich regions or near homopolymeric tracks of poly(A) or poly(T) nucleotides (126, 127). Gene amplification is often unstable and can revert to single copy without pressure. However, DNA duplications flanked by distant A/T tracks could be a first step for desirable evolutionary events under selection pressure (127). Duplication between chromosome subtelomeric regions followed by sequence divergence was a likely mechanism for the generation of two histidine-rich genes of P. falciparum, PfHRP-II and PfHRP-III (95, 128).

GENETICS, GENOMICS, AND DRUG RESISTANCE

Candidate Gene Association Studies

Genetics and genomics information has been widely applied to identify genes contributing to various parasite traits, particularly those of drug resistance. Such studies have included strategies of linkage mapping using genetic crosses, genome-wide association studies (GWAS) of parasite field populations, and in vitro selection followed by gene expression analysis and/or genome-wide sequence surveys for mutations. Candidate genes may also be suggested based on known determinants of biological processes in different species. For example, some ATP-binding cassette (ABC) transporters are known to confer drug resistance in cancer treatment by efflux of drugs out of cancer cells. One of the ABC transporters in the P. falciparum parasite, P. falciparum multiple-drug resistance 1 (PfMDR1, the product of the pfmdr1 gene), was thought to confer CQR by pumping the drug out of the parasite food vacuole (129, 130). An association between CQR and several alleles of pfmdr1 was reported (131); however, analysis of a genetic cross between the CQ-sensitive (CQS) HB3 and the CQR Dd2 parasites found a lack of linkage between the CQR phenotype and inheritance of pfmdr1 (102) (see “Genetic Crosses and Quantitative Trait Analysis of Parasite and Host Phenotypes,” below). Interestingly, pfmdr1 amplification was associated with higher half-maximal inhibitory concentrations (IC₅₀) for MQ in P. falciparum, P. vivax, and P. chabaudi (114, 132, 133); in other work, parasites with amplified pfmdr1 genes became more sensitive to CQ (134). A large number of studies using candidate gene association have been published; space is not available to discuss and cite all of the work here (for a recent review, see reference 22).

Genetic Crosses and Quantitative Trait Analysis of Parasite and Host Phenotypes

Experimental crosses of malaria parasites provide a powerful tool for the identification of genes underlying parasite traits (Fig. 1). To this end, four P. falciparum crosses have been performed through chimpanzee hosts and additional crosses through humanized mice (102, 135–138). These crosses have been employed to study the determinants for a variety of important phenotypes, including drug resistance, parasite invasion of RBCs, nutrient transport, infectivity to mosquitoes, hemoglobin catabolism, and gene expression (56, 88, 123, 136, 139–144). The first P. falciparum cross was generated using HB3 and 3D7 parasites and was used to map a phenotype of PYR resistance to the parasite dihydrofolate reductase-thymidylate synthase gene (pfdhfr-ts) (103, 135). The second genetic cross was performed using the CQR Dd2 and CQS HB3 parasites and linked CQR to a determinant on P. falciparum chromosome 7 (57, 102); the gene for the P. falciparum CQ resistance transporter (pfcrt) was identified after further fine-mapping and parasite transformation experiments (145). The third cross, between 7G8 and GB4 P. falciparum clones, identified a novel erythrocyte-binding protein, PfRH5, that mediates species-specific erythrocyte invasion and virulence of P. falciparum infection to Aotus nancymaae monkeys (136). PfRH5 was independently shown to bind human basigin, a key erythrocyte receptor used by P. falciparum merozoites to invade RBCs (146), and is a promising vaccine candidate against parasite invasion of RBC (147). The HB3 × Dd2 and 7G8 × GB4 crosses also supported discovery of the clag3 genes, which are responsible for channel-mediated nutrient uptake by iRBC (141), pfs47 alleles, which mediate evasion of the mosquito immune system (88), and modulation of parasite responses to amodiaquine and CQ by pfcrt and pfmdr1 (148). The fourth P. falciparum cross was performed between Cambodian 803 and Ghanaian GB4 parasites to investigate ART response phenotypes in vitro and in vivo and to evaluate phenotypes attributed to the PfK13 C580Y Kelch propeller mutation (137). In monkeys receiving three daily doses of intravenous artesunate (AS) after infection with the progenies, recrudescences were not more frequent in propeller mutant than in wild-type PfK13 infections, nor were clearances of the mutant parasites substantially slower in vivo, even though a greater in vitro ring-stage survival rate was linked to the PfK13 C580Y mutation in progenies of the cross after an ART pulse. Additional laboratory crosses of human malaria parasites are being performed through human-liver chimeric mice now that research use of chimpanzees is restricted (138).

FIG 1.

Diagram illustrating the principle of a malaria genetic cross of rodent malaria parasites. A cross starts with an intravenous or intraperitoneal injection of blood samples containing gametocytes from two parasite strains (in this case, Plasmodium yoelii subsp. nigeriensis N67 and P. yoelii subsp. yoelii 17XNL, that have different growth characteristics and virulence in mice). Mice infected with mixtures of gametocytes are anesthetized and fed to Anopheles stephensi mosquitoes. Approximately 15 to 17 days after feeding, the infected mosquitoes with salivary gland sporozoites are allowed to feed on new mice. Daily blood smears are made to monitor parasitemia. The resulting parasites are cloned through limiting dilution by injecting a single parasite into a mouse or are frozen in liquid nitrogen for future studies. The parasite mixtures can be also used for linkage group selection (LGS) after applying selection pressure such as drugs.

Genetic crosses of rodent malaria parasites have also been performed to investigate various phenotypes such as drug resistance, disease severity, or parasite development (104, 149–155). To optimize recombination in these crosses, the parasitemias of the parents are first tested in different ratios for gametocytemias that give maximum cross-fertilization (156). For characterization of inherited traits and linkage relationships, progeny clones may be obtained by injecting a large number of mice with inocula containing individual parasites (usually obtained by limiting dilution), a process that is more difficult and expensive than the cloning of P. falciparum progenies from in vitro cultures. Analysis of the crosses is sometimes possible by an alternative strategy of linkage group selection (LGS) (132, 149–152, 157–159) (Fig. 2). In LGS, samples of uncloned progeny populations are subjected to a specific pressure that selects an inherited trait (151). The survivors of selection and the original mixtures of unselected progenies are then compared in genome-wide searches for evidence of reduced marker diversity (“selection valleys”) in a chromosome region. This strategy has been used to map the genes conferring resistances to PYR (dihydrofolate reductase or pcdhfr) and ART (a gene encoding a deubiquitinating enzyme) and strain-specific immunity in P. chabaudi (150–152). LGS avoids the labor-intensive processes of parasite cloning and evaluations of their individual phenotypes.

FIG 2.

Diagram illustrating the principle of genetic recombination and linkage group selection. (A) Genetic recombination between parental lines (purple and blue) results in progenies harboring various combinations of the parental chromosomal segments. The bars represent chromosomal segments with a putative resistance marker from parent 1 (red dot) distributed among the progenies. Black arrows at the bottom denote positions of polymorphic genetic markers between the two parents. (B) Ratios of resistant alleles from parent 1, showing approximately 50% of allele ratios of the parents before selection. (C) After selection, only the parasites carrying the resistant allele (red dot) survive. (D) A plot of the ratios of resistant alleles shows increased frequency (to 100%) at one locus of the chromosome segment, suggesting at least one genetic determinant contributing to parasite survival or resistance to selection pressure in the locus. Fine-mapping with additional recombinant progenies and genetic markers may identify the gene(s) conferring the resistance.

Genetic crosses of rodent malaria parasites are a valuable resource for studies of phenotypes that are affected by host environment and the interplay of infections from different parasite strains. A number of P. yoelii crosses have been performed to map genes involved in host-parasite interaction, the host immune response, parasite growth, and virulence (104, 153, 160–162). In a recent example, a genome-wide quantitative trait locus (QTL) scan of 43 progenies from a P. yoelii cross linked two major loci on chromosomes 1 and 7 to phenotypes of parasite growth and host mortality, respectively (153). Fine-mapping of the chromosome 7 locus identified a gene encoding a HECT-like E3 ubiquitin ligase (pyheul), which was confirmed by allelic exchange experiments and modification of gene expression to modulate both parasite growth and host mortality.

Progenies of Plasmodium crosses in mice are also a powerful resource for the identification of host genes that respond to parasite infections. More than 1,000 host genes were significantly linked to many parasite genetic loci after transspecies expression QTL (ts-eQTL) analysis (161). In one example, the host transcriptome responses to infections of 24 progenies from a P. yoelii genetic cross were obtained as phenotypes (by microarray analysis) and were analyzed against the parasite genotypes of hundreds of parasite microsatellites. Genome-wide patterns of logarithm-of-odds (LOD) scores (GPLS) were used to cluster host genes that likely function in related pathways of host responses, identifying a number of regulators of the type I interferon (IFN-I) response (Fig. 3). Functional assays have confirmed the roles of individual candidate regulators in IFN-I responses (163).

FIG 3.

Plots of host genes based on genome-wide patterns of LOD scores. This figure is based on data published in reference 161. In the study, mice were individually infected with 24 progenies from a genetic cross of Plasmodium yoelii subsp. yoelii 17XNL and Plasmodium yoelii subsp. nigeriensis N67 parasites. mRNA samples from infected spleens were extracted at day 4 postinfection and were hybridized to a microarray representing ∼19,100 unique mouse genes. Transspecies expression quantitative trait locus (ts-eQTL) analysis was used to analyze genome-wide transcription data from the infections against 479 microsatellite (MS) markers typed on the corresponding progenies. This analysis provided a genome-wide pattern of LOD scores (GPLS) for each host gene. Genes with expression levels linked to at least one MS marker (out of 479) with a LOD score of ≥2.0 were chosen, and their GPLSs were clustered based on pattern similarity. This figure shows a group of genes with similar GPLSs including a major peak of LOD score on one end of the parasite chromosome 13. Genes in an individual cluster are often related through roles in the same host response pathways; examples shown in this figure include type I interferon (IFN-I)-stimulated genes (isg) or genes that likely regulate IFN-I responses (Oas1a, Oas1g, Adar, Mx2, Irf7, S1pr5, or Oasl1). The names of the genes in the cluster are indicated under the plot.

Genome-Wide Association Studies for Genes Contributing to Parasite Drug Responses

The development of a relatively high-density linkage map with hundreds of microsatellites (53) made it possible to conduct GWAS using parasite isolates adapted to in vitro culture. In an early study, 87 culture-adapted worldwide P. falciparum isolates were typed with 342 highly polymorphic microsatellite markers (59). Extensive linkage disequilibrium (LD) of markers near pfcrt with decay of LD suggested that strong CQ directional selective sweeps originated from multiple CQR founder events (59). Publication of the first draft of the P. falciparum genome sequence (27) provided a further resource for the identification and characterization of large numbers of SNPs and microsatellites. Sequencing of additional P. falciparum parasites and their use in GWAS identified a large number of SNPs and expanded the availability of these high-density polymorphic markers (111, 112, 164). In one of these GWAS, the responses to seven antimalarial drugs in 189 culture-adapted P. falciparum parasites were analyzed with a custom-built array containing approximately 3,000 SNPs, averaging ∼1 SNP per 7 kb (165). In addition to identifying candidate genes significantly associated with parasite responses to various drugs, the results of the study provided information about parasite population structure, recombination rate, and loci under recent positive selection. Another GWAS, performed with a higher-density array of over 17,000 SNPs (∼0.7 SNP/kb) and 57 culture-adapted P. falciparum parasites, suggested an association of the Duffy binding-like merozoite surface protein 2 gene (pfdblmsp2) (gene ID, PF3D7_1036300 or PF10_355) with parasite resistance to halofantrine, MQ, and LUM (166); genetic knockout of this gene was subsequently found to increase parasite sensitivity to these three drugs (167). In a GWAS of 22 antimalarial drug responses among 35 Kenyan P. falciparum isolates (168), signals were detected from previously unreported genes as well as genes known to be involved in drug resistance (pfdhfr, pfdhps, pfmdr1, pfnhe, and pfcrt).

Genetic Analysis of P. falciparum Responses to Artemisinin Derivatives and Artemisinin-Based Combination Therapy

Artemisinin and its derivatives, collectively termed ART, are now the critical components of first-line antimalarial therapies recommended worldwide (169). In these therapies, the ART component serves as a powerful, short-acting drug for rapid parasitemia reduction, and a second, longer-acting partner drug is provided to eliminate parasites that can survive ART and cause recrudescence. The requirement for a partner drug in ART treatment was recognized in early clinical studies by Li et al. (170), who demonstrated that 3 days of ART monotherapy was followed within 28 days by recrudescence in more than 40% of P. falciparum-infected patients. The importance of ACTs for definitive cures was further highlighted by findings of frequent recrudescence after 3 to 7 days of ART treatment alone, including results from a trial in Vietnam where patients received artemisinin doses of 10 mg/kg on day 1 followed by 5 mg/kg/day on days 2 to 7 (170–172).

The failures of ART monotherapy to cure P. falciparum infections were recognized by Li et al. (170) as a phenotype of RI resistance, defined by the World Health Organization (WHO) as microscopic clearance followed by recrudescence within 28 days (173). This category is one of the four WHO classifications of drug response (173): (i) S, drug sensitive by clearance of asexual parasitemia within 7 days of the first day of treatment without recrudescence; (ii) RI, resistance characterized by microscopic clearance of asexual parasitemia, followed by recrudescence within 28 days; (iii) RII, resistance defined by reduction of asexual parasitemia but no clearance observed by microscopy; and (iv) RIII, high-level resistance in which there is no substantial reduction of asexual parasitemia. Resistances to CQ, sulfadoxine-PYR (SP), and certain other drugs have been reported at all levels of RI, RII, and RIII. Infections that fail CQ or SP treatment yield isolates that have half-maximum inhibitory concentrations (IC₅₀s) that are significantly increased over those of drug-sensitive isolates in conventional 72-h dose-response in vitro assays. IC₅₀ increases of 10× to 1,000× are linked to mutations in the pfcrt and pfdhfr-ts genes, respectively (145, 174). These mutations are useful molecular markers for drug resistance in population surveys (175).

ACT failures in Southeast Asia have raised concerns about increasing ART or partner drug resistance or both (176–178). In these areas of Southeast Asia, however, parasite isolates were not found to have substantially higher ART IC₅₀ values than elsewhere (165, 179–182). Explorations for ART “resistance” therefore turned to alternative phenotypes such as delayed parasite clearance (DPC) or clearance half-life (t_1/2) (179, 183); however, in patients treated with monotherapy, these measures were not associated with increased recrudescence or fever clearance time (180, 184–187). The t_1/2s of erythrocyte ring stages were instead associated with improved survival of these forms (but not later trophozoite and schizont stages) after a short pulse of high-concentration ART in vitro (ring survival assay [RSA]) (188).

Using data from a number of field studies, GWAS have been conducted to identify genes associated with these various phenotypes. An early study examined dihydroartemisinin (DHA) IC₅₀ values from culture-adapted parasites to search for SNPs associated with ART responses (165); however, the small differences between IC₅₀ values precluded their definitive associations with candidate resistant genes. Cheeseman et al. (189) analyzed 6,969 SNPs in 91 parasites collected from Cambodia, Thailand, and Laos and identified 33 genome regions under strong selection; further study of 715 isolates from Thailand with SNPs using microsatellites revealed a signature of selective sweep on chromosome 13 that was significantly associated with DPC. Takala-Harrison et al. (190) used a P. falciparum array of 8,079 SNPs to survey 342 parasite samples from artemisinin-treated patients in Bangladesh, in northwestern Thailand near the Myanmar border, and at two sites in western Cambodia. One SNP on chromosome 10, two on chromosome 13, and one on chromosome 14 were significantly associated with DPC, of which the two SNPs on chromosome 13 were near those identified by Cheeseman et al. (189). Whole-genome sequencing of clinical isolates from Cambodia and an in vitro line selected for ART resistance then identified a gene with Kelch propeller domain mutations (pfk13; gene ID, PF3D7_1343700 or PF13_0238) from this chromosome segment. Correlation of these mutations with in vivo parasite clearance t_1/2s and RSA survivals provided evidence for selection of pfk13 alleles under ACT pressure in Cambodia (182). More recently, a large multicenter GWAS of parasite samples from 15 locations in Southeast Asia found that at least 20 mutations in the pfk13 gene were associated with a low parasite clearance rate after treatment with ART (191). Multiple additional candidate gene association studies have been recently reviewed (192).

In view of the frequent recrudescences recognized ever since the original clinical studies of ART (170), the absence of ART IC₅₀ signals from isolates after treatment failure, and the studies showing no significant association of clinical recrudescences with DPC or clearance t_1/2s after 3 days of monotherapy (180, 184–187), the nature of ART “resistance” and whether clinical levels of resistance to the drug are now substantially greater have been controversial (183, 193–197). The debate is compounded by observations of greater gametocytogenesis with pfk13 mutant strains (198, 199). Because mature gametocytes are poorly cleared by ART, the mutant parasites may have better survival rates than those with wild-type pfk13, and P. falciparum transmission to mosquitoes may be protected as a survival mechanism. Additionally, parasites from infections clearing more slowly after treatment (i.e., those with a t_1/2 of >5 h) (200) may have IC₅₀ values as low as or lower than those for parasites from shorter clearance infections (t_1/2s of <3 h). Clearance t_1/2s can be affected by host genetic background and factors including immunity, fever, and anemia as well as parasitemia, and differences of 2-fold or less may be difficult to assess in infected patients (196, 201).

RSA is an in vitro parasite survival assay that was developed based on the observation that the ring stage of some parasite strains could survive better under a short period of ART treatment (202, 203). For P. falciparum, this surrogate measure of ART resistance has been set at a threshold ring-stage survival rate of ≥1% after exposure to 700 nM DHA for 6 h (181, 188). This measure ignores the full life cycle, during which ring-stage parasites mature through trophozoite and schizont stages that show little or no K13-mediated difference in survival after the DHA pulse, which is approximately 1% regardless of their pfk13 status (188). In the continuing presence of drug, a ring-stage parasite that survives 6 h of exposure will be subsequently killed as it becomes a more mature trophozoite stage, so that RSA cannot reliably reflect or predict failure to cure infection.

In a recently reported cross of P. falciparum parasites (803 × GB4), parasite clearances and recrudescence in Aotus monkeys treated with three daily doses of intravenous AS were not significantly associated with the inheritance of different parental pfk13 alleles (137), although the C580Y codon in pfk13 was linked to increased RSA values. Moreover, a C580 wild-type clone, produced by allelic exchange of a C580Y mutant-type progeny, yielded a monkey infection that recrudesced every time after 13 individual 3-day artesunate treatments over a period of 500 days. These results reinforce the fact that ART-treated P. falciparum infections frequently recrudesce with or without K13 mutations and highlight the need, as emphasized by Li et al. (170), for effective ACT partner drugs to completely cure parasites that can persist after exposure to ART. Unfortunately, ACT treatment failures from ineffective partner drugs have been increasingly reported from western Cambodia. For example, DHA plus PPQ, an ACT with a historically good track record, now fails frequently in regions of Cambodia (204, 205). PPQ resistance from new mutations in pfcrt is a major contributor to these failures (206), necessitating switches to alternative ACTs such as MQ plus AS (200, 207). Recent findings of plasmepsin gene amplifications, modified pfmdr1 copy numbers, and selection of different pfmdr1 genotypes (208–210) in Cambodian parasites have led to suggestions that rotations among different ACTs or triple ACT drug combinations may help reduce difficulties with drug resistance (211, 212). Multicopy plasmepsin 2 was proposed as a surrogate molecular marker to track PPQ resistance; however, the roles of plasmepsin 2 require further investigation relative to the H97Y, F145I, M343L, or G353V mutation of PfCRT in parasite resistance to PPQ (206). Readers who are interested in additional information on antimalarial drug development and mechanisms of drug resistance can consult the several excellent reviews on these subjects (22, 23, 213–218).

Mapping Genes Using Signatures of Selection and Selective Sweeps

Malaria parasites are under continual evolutionary selection from pressures including host immunity and antimalarial drugs. Because the blood-stage parasites have a haploid genome, signatures of selection may be comparatively easy to identify in the neighborhood (valley or peak) of relevant genes. Such signatures can be followed across regions where malaria is endemic by analyzing the temporal and spatial patterns of genetic markers in parasite populations. Signatures of selective sweeps have thus been reported for CQ, SP, and ART (59, 61, 219–223). For discovery of the selective-sweep signals of reduced genetic diversity or LD decay in gene neighborhoods, microsatellites have been particularly useful because of their relatively high mutation rates (52). Microsatellites were employed in the first demonstration of sweeps from multiple founder events of CQ-resistant parasites, including a major sweep from southeast Asia into Africa (59). In addition to mutations directly responsible for drug resistance, selective sweeps can identify signals of positive versus negative (purifying) selections as well as balancing selection. For example, genome-wide analysis of sequence variation was performed on more than 150 P. falciparum clinical isolates from a region of the Republic of Guinea with high endemicity; signatures of selection by antimalarial drugs and host immunity were detected (224). The citations here have included just a few of many reports from selective studies.

CHEMICAL GENOMICS: SELECTION, SCREENING, AND TARGET IDENTIFICATION

As data and tools to study genome-wide variations grow in availability and high-throughput chemical screening platforms expand, innovative combinations of these powerful approaches will drive new discoveries of gene functions and gene interactions. Analysis of differential responses to small molecules (SMs) and various biologics will unveil targets for exploration in programs to discover new drugs and malaria interventions. For large-scale studies, parasites exposed to libraries of SMs can generate large numbers of mutants with phenotypic variations (Fig. 4). These may involve mutations in direct targets or in other molecules (e.g., transporters) that affect SM susceptibility.

FIG 4.

Chemical genomics approaches to identify interactions of small molecules (SMs) and parasite genes. Malaria parasites, either progenies from genetic crosses or field isolates, can be screened for response (half-maximal inhibitory concentration [IC₅₀]) against libraries of SMs using quantitative high-throughput screening (qHTS). Parasites can also be placed under long-term SM pressure for mutations that may play a role in parasite survival. The parasites are genotyped with a large number of genetic markers or are genome sequenced to detect polymorphism or mutations under SM pressure. Linkage analysis, including LGS, or genome-wide association analysis (GWAS) can be performed to link SM and parasite genes playing roles in the SM responses. Genetic mutations and changes in gene expression can be detected after genome sequencing or microarray/RNA-seq from parasites before and after SM pressure. Candidate genes can be further verified using CRISPR/Cas9-based gene knockout, knock-in, or regulation of gene expression to confirm the potential SM target or genes in SM transport.

Mutations underlying the phenotypic changes such as responses to SMs can be identified using various genetic and genomic approaches, while changes in gene expression can be evaluated by microarray or RNA sequencing (RNA-seq) methods (225). Variations in proteins or metabolite levels can be detected using spectrometry-based methods such as matrix-assisted laser desorption ionization–time-of-flight mass spectrometry (MALDI-TOF MS) or gas chromatography-MS (GC-MS) (226, 227), chromatin modifications can be revealed by chromatin immunoprecipitation (ChIP) with microarray technology (ChIP-chip) or ChIP-sequencing (ChIP-seq) (228), and DNA polymorphisms can be mapped using genetic linkage analysis or GWAS (229–231). Mass spectrometry has been employed to characterize the metabolomic profiles of P. falciparum-infected RBCs (232) or parasite development throughout its 48-h intraerythrocytic developmental cycle (233). In a recent example of the use of this approach, Lewis et al. (144) performed QTL analysis of metabolites as phenotypes and identified elevated levels of hemoglobin-derived peptides in CQR parasites of the HB3 × Dd2 genetic cross.

Differential Chemical Phenotypes, Selection of Mutations with Small Molecules, and Identification of Target Genes

The power of using SMs to detect differences between malaria parasites was demonstrated in a quantitative high-throughput screening (qHTS) study of seven P. falciparum strains exposed to the LOPAC 1280 library (Sigma) (Fig. 4) (229). More than 600 differential chemical phenotypes (DCPs), defined as pairwise IC₅₀ differences of 5-fold or more between parasite lines, were detected. Particular SMs (e.g., dihydroergotamine methanesulfonate, trimethoprim, and triamterene) that produced large differences in DCP values were further tested on 32 progenies from a GB4 × 7G8 cross (136). QTL analysis of the DCPs with the genotypes of 285 microsatellite markers identified a major locus on chromosome 5 (LOD score = 16.4) and a second locus on chromosome 12 (LOD score = 3.0), both of which were linked to the dihydroergotamine methanesulfonate response. A candidate gene on chromosome 5, pfmdr1, was confirmed by allelic substitutions to have an important role in this response (229).

Another powerful genomic approach for identifying drug targets is to subject a parasite to prolonged pressure of drugs or SMs (Fig. 4). Under drug pressure, the parasite may increase expression of genes that can help its survival by changing gene copy number or level of transcription. Nucleotide substitutions (or SNPs) in target genes may also occur, leading to parasites that are more resistant to a drug. Large-scale detection of these changes has been employed to identify a number of mutated genes after selection with SMs (214, 234–237). In one study, CNV of the P. falciparum 1-deoxy-d-xylulose 5-phosphate reductoisomerase gene (pfdxr) was detected using a tiling microarray after in vitro selection with fosmidomycin (235). Studies using ART/DHA-selected P. falciparum parasites, genome sequencing, and transcriptome analysis have also reported changes in CNV, gene expression, and/or nucleotide substitutions in genes such as pfmdr1, pfk13, and pfcoronin and genes in the antioxidant defense network (182, 238, 239). In a recent work, 262 P. falciparum parasites were selected to become resistant to 37 diverse compounds. Subsequent genome sequence analysis revealed 159 gene amplifications and 148 nonsynonymous changes in 83 genes that could be associated with resistance to the compounds (240). In another study, selection of P. falciparum with putative inhibitors of P. falciparum NADH dehydrogenase 2 (PfNDH2) led to mutations in the more likely target: the parasite cytochrome B gene expressed by the mitochondrial genome (237). Consistent with these findings, deletion of the pfndh2 gene has been shown to be dispensable in asexual blood stages, and further, its deletion does not alter the parasite’s susceptibility to multiple mitochondrial electron transport chain (mtETC) inhibitors (241).

Genome-Wide Association Studies of Genes Playing a Role in Parasite Responses to Small Molecules

GWAS is also useful for insights into the responses of P. falciparum isolates to SMs (230). In a qHTS screen of parasite responses to serial dilutions of 2,816 individual compounds, Yuan et al. (230) used 3,354 SNPs and 61 P. falciparum parasite lines to identify 32 highly active compounds and several chromosomal loci significantly associated with 49 DCPs (230). These loci were further confirmed by linkage analysis and the production and testing of parasites with genetically modified candidate genes. The results of this study identified not only SMs that were more effective against parasites expressing wild-type genes but also compounds that had better efficacy against parasites with mutant forms of these genes, suggesting the potential for drug combinations to treat both wild-type and mutant forms of molecules such as PfCRT and PfMDR1.

VACCINE TARGET IDENTIFICATION, MOLECULAR EVOLUTION, AND EPIDEMIOLOGY

In addition to its usefulness for insights into drug resistance and important parasite traits, information on the genome diversity and variation of malaria parasites is valuable for research on vaccine development, disease epidemiology, and parasite evolution. A large amount of work has been done in these subject areas. Here we draw attention to just some examples.

Genome Surveys for Selection Signatures of Vaccine Targets

Vaccine candidates such as PfMSP1, PfAMA1, PfCSP, or PfEMP1 have highly polymorphic regions where these molecules are under host immune pressure (242, 243). Vaccines developed from these polymorphic antigens may be confounded by allelic restriction of the host immune response, and parasites expressing alternative epitopes can escape host immune clearance. A vaccine that does not provide complete protection may select parasite variants having elevated virulence and thereby be “imperfect” (244). Based on the observation of a polymorphic nature of immune targets, potential antigens can be identified through genome-wide analysis of genetic diversity (111). Population genetic studies may uncover signatures of balancing selection in the polymorphisms of genes encoding immune targets (245). In a genome-wide survey of P. falciparum parasites isolated from 65 Gambian clinical patients, candidate genes with signatures of balancing selection were identified for studies of immune mechanisms and potential vaccine development (246). The gene with the highest values of Tajima’s D was Pfdblmsp2 (PF3D7_1036300 or PF10_0355), a gene previously associated with parasite responses to halofantrine, MQ, and LUM by GWAS and gene expression analysis (166). Interestingly, indirect immunofluorescence assay (IFA) analysis showed that the PfDBLMSP2 protein was expressed in only a small fraction of mature schizonts (246). In another study, a survey of 3,539 predicted P. falciparum genes from four cloned isolates (Dd2, HB3, D10, and 7G8) for elevated diversity detected 56 highly polymorphic genes (with θ values 2 standard deviations higher than the mean θ value for 1,920 genes). In vitro protein expression and Western blotting using pooled infected human sera confirmed the antigenicity of 11 proteins, including seven previously unknown antigens (111). Using samples collected from patients, Conway et al. (247) measured fixation index (F_ST) values for multiple microsatellite loci and polymorphic sites in block 2 of PfMSP1 and showed that the block 2 was under immune selection or an immune target. These studies demonstrate that survey of genome diversity is an effective approach for identifying genes under immune selection for vaccine candidates.

Applications to Studies of Parasite Molecular Evolution

Genome diversity data support inferences about parasite origin and evolutionary history, including estimates of time to the most recent common ancestor (TMRCA). Genetic polymorphism information can be used to date historical events based on assumptions of a molecular clock and the absence of selection. Under such assumptions, the rate of molecular evolution for the average protein is taken to be approximately constant over time, although estimates of rates can be time dependent and influenced by factors of natural selection, calibration errors, model misspecification, and other artifacts (248). Thus, special care should be taken to collect data on “neutral” genetic sequences for evolutionary analysis. Such care helps to avoid the use of antigen genes that can inflate overall estimates of genome diversity and mutation rate. On the other hand, the effects of antimalarial drugs can reduce genome diversity of parasite populations by removing drug-sensitive parasites and overrepresenting parasites that carry drug resistance genes. A single haploid mutant parasite can multiply, produce male and female gametocytes, infect mosquitoes, and quickly generate a genetically homogeneous population (clonal expansion outbreak). Parasite samples from such a malaria outbreak or from parasite populations massively treated with antimalarial drugs may show greatly reduced genome diversity, potentially affecting an estimate of TMRCA.

Studies of P. falciparum populations and their evolutionary origin have led to different estimated TMCRAs and proposed “malaria’s eve” hypotheses (249). Some studies, finding poor correlation between the strength of LD and nucleotide distances in the circumsporozoite (csp) gene, as well as no synonymous substitutions in 10 genes, have suggested that all extant world populations of the P. falciparum parasite are from an ancestral strain (or a small number of strains) that was subjected to an ancestral sweep a few thousand years ago (250, 251). Additional data from SNPs in 25 introns of 10 housekeeping genes and the 6-kb mitochondrial genome were also consistent with a relatively recent origin of P. falciparum populations (252, 253). However, a much longer TMRCA and large effective population size have also been proposed (109, 110, 254, 255). Recent findings from DNA sequences in the fecal samples of wild-living apes suggest that a bottleneck or eve event likely occurred from an ancient transmission of a great ape parasite to a human in Africa (36, 256). Such an event would be consistent with close genomic relationships between P. falciparum and other Plasmodium species of the Laverania subgenus, which infect gorillas (P. praefalciparum, P. adleri, and P. blacklocki) and chimpanzees (P. reichenowi, P. gaboni, and P. billcollinsi) (257). Indeed, recent analysis suggests that P. falciparum began to emerge in humans from P. praefalciparum around 40,000 to 60,000 years ago, followed by a population bottleneck around 4,000 to 6,000 years ago (64).

Population Structure and Geographic Origin of Parasites

Population structures of malaria parasites have been investigated using polymorphic genes and genome-wide distributions of sequence variations. Using 342 highly polymorphic microsatellite markers from a genetic map, a survey of a worldwide collection of P. falciparum isolates demonstrated geographical and continental clustering (59). Additional insight into the structure of P. falciparum populations was gained from genotypes of ∼3,000 SNPs, which suggested an evolutionary separation of Cambodian parasites from those of Thailand (165). More recently, analysis of genome variation in 825 P. falciparum samples from Asia and Africa identified unique divisions in the parasite population structure in western Cambodia, suggesting three different subpopulations associated with ACT responses (41). Differences among these subpopulations were not associated with regional gene flow or high rates of transmission (258). In other studies, clustering of polymorphic P. falciparum apical membrane antigen 1 (PfAMA1) sequences from 97 parasite clones from around the world and 61 isolates from Mali showed patterns of polymorphisms that were independent of geographic location (259), which is likely due to multiple instances of diversifying selection on the gene (260).

Knowledge of population structure can be useful for studies of epidemiological developments and changes in patterns of disease. Initiatives such as the Malaria Genomic Epidemiology Network (MalariaGEN; https://www.malariagen.net) and the PlasmoDB resource (http://plasmodb.org/plasmo/app/search/dataset/AllDatasets/result) provide genomic sequences, omics data sets, disease information, genetic resources, and analysis capabilities for multiple Plasmodium species and infections from regions where malaria is endemic worldwide. Approximately 3,500 P. falciparum parasite isolates from 23 countries and 228 P. vivax samples from 13 countries have been sequenced by MalariaGEN. Sequence data from the project have been used to investigate drug resistance, parasite population structure, diversity, epidemiology, and transmission (41, 43, 191, 258, 261, 262). A recent study examined genome sequences from 1,492 P falciparum samples from 11 locations across southeast Asia, including 464 samples from western Cambodia, and found that an amino acid substitution within the product of a gene encoding an exonuclease (E415G) and plasmepsin 2/3 amplification are frequently associated with piperaquine resistance and dihydroartemisinin-piperaquine failures in Cambodia (261).

GENOMICS AND APPLICATIONS ON POPULATION DIVERSITY, PARASITE ORIGIN, AND RELAPSE OF P. VIVAX INFECTION

Genome Diversity of P. vivax and Genes under Selection

P. vivax is another malaria parasite of major public health burden and was responsible for ∼7.5 million clinical cases in 2018 (263). P. vivax and other non-P. falciparum human malaria species have attracted more attention recently. A number of reports now provide genetic and genomic analyses on parasite population diversity, signatures of selection, and molecular evolution (25, 33, 39, 49, 262, 264–266). One of the early attempts to characterize P. vivax genome diversity was the sequencing of a 100-kb contiguous chromosome segment from five isolates. The data showed P. vivax to be a genetically diverse species with an abundance of SNPs but a relatively lower level of polymorphic tandem repeats than that in P. falciparum, possibly due to its more balanced guanine-cytosine content (109). Whole-genome sequencing of a Peruvian P. vivax isolate obtained ex vivo without further propagation identified a large number of SNPs and genes under selection, including the homolog of pfcrt (pvcrt), genes encoding reticulocyte-binding proteins, and AMA-1 (265). More recently, genome sequencing of global collections of P. vivax parasite isolates confirmed higher genomic diversity in P. vivax than in the more virulent P. falciparum parasite and revealed signals of recent evolutionary selection on drug resistance genes (39, 262, 267). The higher levels of SNPs also led to estimates of TMRCA that were longer than that of P. falciparum (267–269).

Origin of P. vivax Parasites

The P. vivax parasite is genetically closely related to several parasites infecting nonhuman primates, such as P. cynomolgi, P. knowlesi, and P. vivax-like parasites (264, 270, 271). Early phylogenetic analyses using mitochondrial DNA or genes from the plastid and nuclear genome placed P. vivax among the Asian primate malaria parasites (Fig. 5A) (35, 269). The near-perfect genetic identity between P. vivax and P. simium, a New World primate malaria species, also suggested the possibility of transfer between humans and the New World monkeys (272–274). Indeed, studies using the complete mitochondrial genome sequence or genes from the plastid and nuclear genome indicated an Asian origin via a host switch from macaque monkeys (268, 269). On the other hand, a relative protection against P. vivax in western and central Africa by the highly prevalent Duffy-negative blood type suggests that P. vivax might have a long history in Africans (275–277). More recent identification and sequencing of genomes of P. vivax-like parasites support an evolutionary transfer of P. vivax-like parasites from African great apes to humans (Fig. 5B) (36, 264, 278). Additionally, analyses of the apicoplast genome from 18 Plasmodium species have shown that P. vivax is positioned next to the divergence of the African guenon parasite, P. gonderi, before the common ancestor of Asian primate parasites, supporting an African origin (279). Much about the relationships and evolutionary history of P. vivax, P. vivax-like, P. simium, P. cynomolgi, and P. knowlesi parasites remains to be discovered.

FIG 5.

Molecular phylogenetic trees of Plasmodium vivax and other related nonhuman primate parasites. (A) A phylogenetic tree of P. vivax and related parasites based on sequences from two nuclear genes (β-tubulin and cell division cycle 2) and a plastid gene (the elongation factor Tu). The tree is adapted from Fig. 1B of reference 269 with permission of the National Academy of Sciences. (B) A phylogenetic tree of P. vivax and P. vivax-like parasites from nonhuman primates, including apes. The tree is adapted (simplified) from Fig. 2 of reference 278 with permission of Springer Nature/Macmillan Publishers Limited.

P. vivax Relapses versus Recrudescences and Reinfections

An important characteristic of P. vivax infection is malaria relapse arising from dormant liver-stage forms called hypnozoites (16). One of the main challenges in studying relapses in regions of endemicity is the difficulty in distinguishing parasites of relapse from those of reinfection or recrudescence of drug-resistant parasites. Genetic typing and more recently genome sequencing have been employed to distinguish parasites of relapse from reinfection and/or recrudescence of drug-resistant parasites (280–282). A study on Australian soldiers experiencing relapses of P. vivax malaria after exposure in East Timor used the polymorphic regions of three P. vivax genes to demonstrate the predominance of a single allelic type in relapsing parasites. However, the second relapse was often from a different allelic type, indicating a different strain (283). Therefore, multiple relapses in an individual may arise from the activation of hypnozoites of different parasite strains. Another study showed that relapsing parasites are often polyclonal, but with some degrees of relatedness to the parasites in a mixed initial infection (282). Different parasite clones might emerge during the course of the relapsing infections, which is consistent with the study of Australian soldiers. Genome sequencing studies including single-cell analysis of relapsing parasites can provide high resolution with a large number of polymorphic sites; however, adequate studies of the low-level parasite populations characteristic of P. vivax will remain challenging. The recent advances in genomics will certainly contribute greatly to the study and resolution of relapses in P. vivax infection (284).

CONCLUSIONS

Knowledge of genetic variations and genome diversity of malaria parasite populations has greatly improved our understanding of the biology, gene function, drug resistance, population dynamics, transmission, and molecular evolution of malaria parasites. Information gained from these studies is particularly important when ethical considerations, resources limitations, or confounding factors limit direct experimental infections of human hosts. In addition, investigations of disease phenotypes in rodent or other animal models of malaria under controlled laboratory conditions, combined with parasite genetics and genomics, can provide helpful insights into pathogenesis and new information on disease control and management in humans.

Biographies

Xin-zhuan Su, Ph.D., is a senior investigator and the Chief of the Malaria Functional Genomics Section, National Institutes of Health, USA. Dr. Su obtained his bachelor’s and master’s degrees in parasitology from Xiamen University, China, and his Ph.D. in parasitology from the University of Georgia. His research interests include mechanisms of drug resistance, drug screening, genetic mapping, genomics, molecular evolution, population genetics, host-parasite interaction, and molecular signaling of the host immune response. His major contributions to malaria research include mapping of the gene encoding the Plasmodium falciparum chloroquine resistance transporter (pfcrt), discovery of P. falciparum var genes, development of the first P. falciparum linkage map, and detection of worldwide selective sweeps of pfcrt. Dr. Su is a board member for several scientific journals and has received various awards, including the Bailey K. Ashford Award from the American Society of Tropical Medicine and Hygiene and the NIH Director’s Award. He is a Fellow of the American Academy of Microbiology.

Kristin D. Lane, Ph.D., M.S., is a research associate in the Division of Intramural Research, National Institute of Allergy and Infectious Diseases. She holds a Ph.D. in microbiology and immunology with concentrations in molecular biology and microbial pathogenesis from Virginia Commonwealth University and a master’s of science in biology from Old Dominion University. Her interests encompass mechanisms of molecular pathogenesis and drug resistance in the malaria parasite Plasmodium falciparum, transcriptional regulation, molecular piracy, and viral pathogenesis. Dr. Lane’s major contributions to malaria research include drug target identification and resistance mechanisms in P. falciparum, particularly those involving cytochrome B (PfCytB) and the P. falciparum chloroquine resistance transporter (PfCRT).

Lu Xia is a Ph.D. student majoring in medical genetics at the State Key Laboratory of Medical Genetics of the Central South University, China. In 2017, she joined the Laboratory of Malaria and Vector Research, NIAID, as a student trainee supported by the China Scholarship Council. Her research focuses mainly on genetics and bioinformatics of human inherited diseases and Plasmodium genetics and genomics. Her expertise includes genome-wide expression and association analyses to study molecular mechanisms of neurodevelopmental disorders and immune pathways in response to malaria infections. She has published 12 peer-reviewed papers in the fields of malaria research and human genetic diseases.

Juliana M. Sá, Ph.D., received a bachelor’s degree in biological sciences from the Universidade Estadual Paulista, Brazil. She obtained a master’s degree in biochemistry and a Ph.D. in parasitology from the Universidade de São Paulo, Brazil. Dr. Sá pursued postdoctoral training at the National Institute of Allergy and Infectious Diseases (NIAID), USA, where she holds a Staff Scientist position investigating antimalarial resistance. Her interest in genetics of drug resistance results from a passion for basic science and applicability. Dr. Sá has been in the field of malaria drug resistance for 18 years.

Thomas E. Wellems, M.D., Ph.D., is an NIH Distinguished Investigator in the Division of Intramural Research, National Institute of Allergy and Infectious Diseases. His interests embrace the biology, pathogenesis, and drug responses of the various species of malaria parasites. Major reports from his work include the descriptions of the Plasmodium falciparum chloroquine resistance transporter (PfCRT), the P. falciparum var gene family (responsible for antigenic variation and immune evasion), a molecular mechanism for malaria protection by sickle-cell trait and hemoglobin C, the PfHRP-II protein used in malaria rapid diagnostic tests (RDTs), and development of P. falciparum transformation methods. Dr. Wellems is a member of the U.S. National Academy of Sciences and the National Academy of Medicine and a former president of the American Society of Tropical Medicine and Hygiene, and he has served on a number of advisory committees for foundations and public-private partnerships, including the Medicines for Malaria Venture.