Open access peer-reviewed chapter

From Genes to Biomarkers: Understanding the Biology of Malaria Gametocytes and Their Detection

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Daniel Kepple, Anthony Ford, Ebony Little, Gabrielle Kolesar, Beka R. Abagero, Ashley N. Blackwell, Swarnapali De Silva Indrasekara, Delenasaw Yewhalaw and Eugenia Lo

Submitted: 29 June 2021 Reviewed: 09 July 2021 Published: 30 August 2021

DOI: 10.5772/intechopen.99364

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Genetic Polymorphisms - New Insights

Edited by Mahmut Çalışkan

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Each year, approximately 230 million malaria cases and 400,00 malaria deaths are reported worldwide. Malaria is a life-threatening disease caused by Plasmodium parasites that are transmitted from one individual to another through the bites of infected female Anopheles mosquitoes. Malaria parasites replicate asexually in the human host, and, in each replication cycle, a portion of the asexual stages develops into sexual gametocytes that permit transmission. The proportion of infections that carries gametocytes and the infectivity of gametocytes are indicators of human-to-mosquito transmission potential. In P. falciparum, gametocytes appear 10–14 days after infection, whereas in P. vivax gametocytes appear simultaneously with asexual schizonts. Such difference in development not only increases the length of time that an individual is infectious, but also increases the likelihood of transmission before treatment. The conversion from asexual parasites to gametocytes is also highly variable between infections. Differences in age, host immune response, parasite genetic composition, density of red blood cells, presence of co-infecting parasite strains, and antimalarial drug use could affect gametocytes production. In P. vivax, the unique ability to produce hypnozoites, a dormant liver stage of the parasite, may allow gametocytes to be produced periodically from relapse and contribute to transmission. In this chapter, we will provide an overview of the biology of Plasmodium gametocytes, existing tools for gametocyte detection, and features of gametocyte genes. The biological insights and genetic findings are essential to developing better detection biomarkers and effective strategies to reduce transmission in malaria-endemic countries.


  • Plasmodium
  • gametocyte
  • epidemiology
  • biomarkers
  • transmission

1. Introduction

1.1 Malaria epidemiology

Malaria is a mosquito-borne disease. In humans, malaria is primarily caused by six different Plasmodium parasite species including P. falciparum, P. vivax, P. ovale, P. malariae, P. knowlesi, and P. simium [1]. To date, there is an estimated 229 million cases of malaria worldwide each year resulting in approximately 409,000 deaths [2]. This has resulted in an estimated direct cost (treatments, premature death, and loss of household income) of $12 billion USD per year, with lost economic growth being many times greater [3]. The deadliest of these is P. falciparum, primarily located in the African continent [4], due to the high parasitemia levels and rapid growth [4]. P. falciparum utilizes several host receptors, including Band 3 and Glycophorin A, B, and C for red blood cell invasion [5], which contributes to high parasite loads in infected patients and spread across the ethnically diverse continent. Contrary to P. falciparum, P vivax primarily uses the Duffy antigen receptor found on the surface of reticulocytes for invasion, leading to lower parasitemia levels. P. vivax is globally widespread in Asia, South America, and northern and eastern parts of Africa [6]. This parasite can cause relapse from weeks to months after the initial infection [6].

Most P. ovale cases were reported in the tropics of western Africa [7] and is considerably less prevalent and possibly less fatal in comparison to P. falciparum and P. vivax [8]. P. ovale has two different subspecies: P. ovale curtisi and P. ovale wallikeri, both of which may produce dormant stage hypnozoites similarly to P. vivax [9, 10]. P. ovale has been shown to be transmitted across international borders from Africa to China [11] as well as to Korea [11, 12], in countries that were considered malaria free [12]. P. malariae is mostly reported in sub-Saharan Africa and southeast Asia. Similar to P. ovale, P. malariae is less common and less fatal. In sub-Saharan Africa, infections of P. malariae are often found to also contain P. falciparum [13]. Though less common, P. knowlesi, a primate parasite infecting long-tailed and pig-tailed macaques [14], has recently been reported to infect humans in Southeast Asia [15, 16] and is mainly transmitted by Anopheles leucosphyrus and A. hackeri [16, 17, 18]. P. knowlesi utilizes two gene families: DBL and RBP, to bind the Duffy antigen/chemokine receptor (DARC) to invade human erythrocytes [19]. Last but not least, P. simium is a common primate parasite that has recently been detected through molecular assays in humans from Atlantic Forest region of southeastern Brazil [20]. P. simium remains largely uncharacterized due to its novelty and rarity in humans, but is thought to have similar invasion mechanisms to P. vivax due to genetic and morphological similarities [20]. The presence of these less-common malarial species emphasizes the need for better detection tools and control measures especially in countries approaching the elimination phase.

1.2 Development and transmission of Plasmodium

The lifecycle of Plasmodium is divided into two stages: the human asexual reproduction stage (also known as the erythrocytic stage) and the mosquito sexual reproduction stage (known as the sporogonic cycle) [20], both of which are necessary for population growth and genetic diversification [19]. The parasite first enters the human body in the form of sporozoites through the infected salivary gland of an Anopheles mosquito during a blood meal. Within minutes, sporozoites infect the liver and begin asexual replication to form schizonts (a process called exoerythrocytic schizogony) within a few days to a few weeks [20]. Once matured, schizonts bust to release hundreds of merozoites into the blood stream [20]. P. falciparum, P. ovale, P. malariae, and P. simium infect mature erythrocytes [2, 19, 21] while P. vivax and P. knowlesi infect younger reticulocytes [2, 19, 22, 23, 24] to feast on hemoglobin and further reproduce asexually [2, 19]. At this stage, the intracellular Plasmodium is considered as a ring stage parasite because the young parasite takes up hemoglobin in a single, large vacuole that looks like a ring. Once the hemoglobin is engulfed, the parasite then begins feeding and hemoglobin is acquired by endocytosis of erythrocyte cytoplasm within cytostomes, known as the trophozoite stage. After the hemoglobin is consumed, the trophozoite may either undergo schizogony to asexually reproduce and start the erythrocytic cycle with new merozoites or mature into a macrogametocyte (egg) or microgametocyte (sperm). Because human body temperature is considerably warm for Plasmodium sexual reproduction [25], both forms of gametocytes are taken up by an Anopheles mosquito where sexual reproduction occurs within the gut of the mosquito producing zygotes. Once matured, the zygote will become mobile and elongated, developing into ookinete to infect the midgut wall of the mosquito [26] and develop further into oocyst. Oocysts asexually divide and eventually rupture into sporozoites that travel to the salivary gland of the mosquito to infect a new human host [2, 19].

All six human malaria parasites require gametocytes to infect female Anopheles mosquito to reproduce sexually and continue development into sporozoites before infecting a new human host [27]. This has led to complications in the control of malaria transmission and gametocyte detection [28]. Furthermore, P. vivax and P. ovale’s unique ability to form dormant stage hypnozoites complicates parasite clearance and can reintroduce old parasite strains into transmission reservoirs [219]. Gametocytes are critical for malaria transmission and possible immune evasion in both the human and mosquito hosts [29, 30], emphasizing the importance of accurate detection. This chapter aims to provide a systemic review that highlights the complexity of Plasmodium gametocytes at the biological and genetic levels and current methods used to track and detect Plasmodium gametocytes. We further examine the capability of biomarkers used across Plasmodium species and provide new candidate biomarkers to further enhance detection protocols. Lastly, we examine the immunogenicity of gametocytes in both humans and mosquitoes.


2. Gametocytogenesis

2.1 Gametocyte commitment and development

Gametocytogenesis is the commitment of a Plasmodium parasite to produce male and female gametocytes through mitotic division during the trophozoite stage that involves multiple epigenetic and transcriptional regulations [31]. Changes in temperature, pH, and host age help stimulate gametogenesis, the emergence of the gametes in the mosquito mid-gut. A drop in temperature from 38°C to 20-26°C, the exposure to gametocyte activating factors and/or mosquito exflagellation factors, as well as a rise in pH can trigger gametogenesis [32, 33, 34]. Some merozoites, for reasons not fully understood, differentiate into the sexual forms of the parasites, the gametocytes. Upon ingestion by the mosquito, the decline in pH, a drop in temperature, and other mosquito derived factors such as xanthurenic acid can together activate gametocytes to transform within 5–10 minutes to male (microgamete) and female gametes (macrogamete) within the mosquito midgut. When gametocytes are taken up during a mosquito’s blood meal, a number of factors including temperature, oxygen and carbon dioxide concentration, pH and exflagellation factor contribute to the maturation of gametocytes [35]. Some species, such as P. ovale in humans and P. yoelii in mice, can develop into morphologically distinct male and female gametocytes directly from hepatic merozoites for further transmission [7]. Gametocyte commitment is largely based on stress factors including high parasitemia, anemia, drug treatments, and host immune responses [36, 37, 38, 39]. Although no clinical symptoms are experienced during gametocytogenesis, this developmental stage is critical for sexual replication in the mosquitoes and subsequent infection of a new human host in the form of sporozoites. There is considerable variation in the development time among the different human Plasmodium species, ranging from 7 to 10 days after the initial establishment of asexual parasites for P. falciparum [40] and 7–15 days for P. vivax [41]. It is yet unclear about the time for gametocyte development in P. knowlesi, P. malariae, P. ovale, and P. simium. During this prolonged maturation period, gametocytes undergo five morphologically distinct stages (I-V) [42]. The immature gametocytes developed during stages I-IV sequester mostly in host tissues, particularly in bone marrow and spleen [43]. At stage V during maturation, gametocytes become more deformable and return in the blood circulation for uptake by a new mosquito host [44]. For proper sporogonic development to occur, both male and female gametes must be taken up during a blood meal as merozoites are incapable at forming both male and female gametocytes from the same schizont [45]. Most regulation is performed through RNA binding proteins, such as acetylation lowers binding efficiency 4 (ALBA4), that increases exflagellation events in an unknown manner [46, 47].

To date, most of our knowledge on gametocyte commitment is derived from P. falciparum due to a lack of viable culturing methods for P. vivax [48] and low prevalence of other Plasmodium species. Molecular processes including epigenetic regulation and histone posttranslational modifications play vital roles in gametocyte commitment [49, 50, 51]. A well-studied example of histone regulation in P. falciparum is H3K9me3, which is normally restricted to multigene families in subtelomeric regions [52], and recruits heterochromatin protein 1 (HP1) [53, 54]. Both proteins are strongly associated with the AP2-G locus, the primary regulator for gametocytogenesis [55, 56]. Previous studies have shown that a majority of asexual parasites have AP2-G silenced [47] and this gene silencing may be induced by histone deacetylase (Hda2) [57]. Depletion of HP1 through GDV1, another gene that regulates gametocyte production, was shown to increase schizont development into gametocytes by 50% [47, 58]. Further, the presence of Stabilization ligand Shield 1 (Shld1) may also increase transcription of AP2-G by stabilizing the protein complex for transcription and enhance gametocyte production. Although the molecular mechanisms of gametocytogenesis are well established in P. falciparum, there are currently no sensitive and reliable methods available in clinical settings for front-line detection of in vivo gametocytes due to strain polymorphisms, limiting gametocyte densities, and variations in the timing of gametocyte production and development.

2.2 Polymorphisms of gametocyte genes

Recently, advances in next generation sequencing provide new insights in our understanding of genetic variation and gene expression across different stages of Plasmodium species [59]. The mosquito immune system is a significant barrier for some Plasmodium isolates to infect the mosquito hosts. For example, in P. falciparum, highly expressed Pfs47 protein allows the parasites to become “invisible” and escape the immune system of the mosquito. This protein is expressed on the surface of female gametocytes and ookinetes. The African isolates of P. falciparum that are known to express Pfs47 have been shown to escape the Anopheles gambiae immune system by suppressing the Jun-N-terminal kinase (JNK) signaling and avoiding the induction of epithelial nitration [59]. The Pfs47 protein gene is highly polymorphic implying that the mosquito immune system may be the driving force for diversity, and that parasites with compatible Pfs47 haplotypes and/or escape immune evasion are preferably selected [60].

The innate immune response in the mosquito vector is mediated in most part by the hemocytes, which eliminate pathogens such as bacteria, fungi, and protozoa. Anopheles mosquitoes are known to have a complement C3-like protein called thioester-containing proteins (TEP). TEP of An. gambiae (AgTEP1) has been shown to initiate immune defense against P. berghei [61]. TEP1 facilitates the interaction between the parasite and hemocytes with late encapsulation that kill the parasite. Knockdown of the TEP1 gene renders genetically selected refractory Anopheles strain susceptible to infection and increases the infectivity rates [61]. However, P. falciparum has been shown to bypass the TEP-based defense mechanism using its 6-cystein protein P47-like [61]. In P. berghei, the P47-like protein is important for female gamete fertility, whereas in P. falciparum, it promotes the gametocyte-to-ookinete development and protects the ookinete from complement-dependent lysis [61]. Furthermore, infection of An. gambiae mosquitoes by ookinetes of P. berghei has been shown to module the mosquito’s immune system by up-regulating expression of the peptide defensin and a putative gram-negative bacteria-binding protein and a TNF-α factor-like transcription factor (LL3) [61].

2.3 Expression of gametocyte genes

During the different stages of gametocyte development, various genes express differentially. For instance, Pfs16 expresses the earliest and highest in stage II gametocytes, the alpha-tubulin II gene shows maximum expression levels in both stage II and III gametocytes, Pfs230 expresses in stage III gametocytes, PfsMR5 in stage IV gametocytes, and Pfs28 in stage V gametocytes [62]. Such gene expression pattern is directly related to the activities involved in each specific stage. Pfs16 is known as a marker for detecting sexually committed ring stages parasites and it likely plays a key role in gametocyte maturation given its continual expression during the entire gametocyte maturation process [63]. Recent immunofluorescences assay identified a small population of schizonts that expressed the Pfs16 gene. Pfs16 mRNA increases in the asexual cycle before schizonts develop into stage I gametocytes, though translation is delayed until the onset of stage I gametocytogenesis [62]. This finding suggests that both Pfs16 transcription and translation may begin prior to invasion of a committed merozoite and development into a stage I gametocyte.

The female gametocyte specific gene Pfs25, which expresses solely in the mosquito, is a glycosylphosphatidylinositol-linked protein expressed on the surface of ookinetes [64]. This gene is used for the detection of stage V female mature gametocytes as well as in the quantification of gametocytes from field studies [63]. Because Pfs25 is solely expressed inside the mosquito hosts with limited immune selective pressure, sequence variation between isolates is relatively low [65]. Antibodies against Pfs25 have been shown to reduce oocyst production in in vitro membrane feeding experiments. Both high sequence conservation and antibody response make Pfs25 a leading target for transmission-blocking vaccine design. Apart from Pfs25, another family of gametocyte protein with cysteine-rich domains includes Pfs230 and Pfs48/45 that have also been targets of transmission-blocking vaccine development [66, 67].

Pfs230 expresses in both male and female gametocytes with a prodomain that is processed during gametocytogenesis and mediates red blood cell binding, specifically during oocyst development [68]. The Pfs230 protein appears on the surface of gametes as a complex with Pfs48/45 (a glycosylphosphatidylinositol (GPI) anchored protein [69]) and appears to be critical for gamete fusion. The double domains in Pfs230 reveal a structure resembling the surface antigen 1 (SAG1) protein with a double beta-sandwich structure found in Toxoplasma gondii, another apicomplexan parasite. The complete gene encoding the Pfs230 protein from different isolates of P. falciparum showed that 27 nonsynonymous polymorphic sites [70]. Among them, the amino acids at eight polymorphic sites map to positions that point their side chains toward the surface of the protein [70]. Five of those eight sites map to a confined region on the same side of the beta-sandwich of the protein structure. Even though the distribution of the amino acids are over much of the length of the domain IV sequence, the polymorphic sites appear to outline, although loosely, a contiguous surface region of the model [70]. This structural feature may provide relevant sites with respect to the interaction of male and female gametes [70].

In both male and female gametocytes, Pfs230 is expressed without a known membrane anchor and appears as a complex with the Pfs48/45 proteins on the surface of gametes, though this colocalization is not required. Pfs230 has been observed on the surface of live macrogametes in the absence of Pfs48/45 [71]. Pfs48/45 is a cysteine rich surface protein that is vital in male gamete fertility. Sequence analyses of Pfs48/45 revealed that polymorphisms are rare for residues involved at the binding interface [72]. Furthermore, unlike other pre-erythrocytic blood stage antigens Pfs48/45 is less polymorphic. The number of synonymous substitutions per synonymous site exceeds the number of non-synonymous substitutions per non-synonymous site [73].

Recent clustering analyses indicated that there is a clear distinction in the expression levels between male and female gametocyte genes. Male and female gametocytes have been shown to be differentiated by their gene expression levels as early as stage III. The female gametes are generally separated by stage, indicating differentiating gene expression levels with respect to mid and late-stage female gametocytes [74]. For male gametocytes, PF3D7_1325200, a putative lactate dehydrogenase gene, is highly expressed [74]. Another male gametocyte gene PF3D7_1311100 is a putative meiosis-specific nuclear structure protein 1. This gene is essential for normal assembly of the sperm flagella in mice, suggesting that it may have a role in the male gamete development and exflagellation. Other genes including Pf3D7_1114000, Pf3D7_1122900, Pfg14–748, HAP2, and MAPK2 are found to be associated with male gametocytes, though both Pf3D7_1114000 and Pf3D7_1122900 are also expressed in a few female gametocytes [74].


3. Genetic markers for detection and prevention

3.1 Conventional genes used for gametocyte detection

Previous study has indicated that approximately 10% of P. falciparum and 60% of P. vivax infections had concurrent detectable low-density gametocytemia [75]. Gametocyte densities are typically lower than asexual parasite densities (Figure 1A) and directly associated with total asexual parasite densities (Figure 1B). Given the relatively low gametocyte densities, it is conceivable that many of these infections are submicroscopic and remain undetected in communities where malaria occurs, contributing to continuous transmission.

Figure 1.

(A) Comparison of asexual parasitemia and gametocytemia among P. vivax infections from Ethiopia. (B) Significant correlation was observed between asexual parasitemia and gametocytemia.

Molecular tests for diminutive amounts of gametocytes use reverse-transcription polymerase chain reaction (RT-PCR) to amplify RNA transcripts of gametocyte-specifically expressed genes. Compared to DNA-based assay, qRT-PCR of targeted RNA transcripts revealed higher sensitivity in detecting gametocytes of considerably low densities. For example, there are more than 106 copies of 18S rRNA transcript per cell but only 5 copies of the 18 s rRNA gene per genome [76]. The production of high transcript copies in parasite cells allow for greater detection limits. For P. falciparum, several targeted genes such as Pfs25, Pfg136, Pfg84, and Pfg17 are specific to female gametocytes. The lack of genes specific to male gametocytes could underestimate the total gametocytes in an infection and thus its transmission potential. Moreover, genes often show wide variations in detection limit based on the level of RNA transcript production, ranging from 100 gametocytes per microliter of blood by Pfg84, 70.7 gametocytes/μL by Pfg136, 25.3 gametocytes/μL by Pfs25 to as low as 10 gametocytes/μL by Pfg17 [77]. Pfg17 is considered as the most sensitive biomarker so far because it can detect the lowest number of gametocytes. For example, in malaria-endemic areas of Ghana, Pfg17 offers higher sensitivity than Pfs25 in detecting P. falciparum gametocytes in clinical samples collected from children and adults [61]. Of the 80 children, 47 were tested negative and 33 were positive for asexual blood stage P. falciparum. Nine out of the 47 (19%) negative children were detected positive for P. falciparum gametocytes by Pfg17. Among 30 adults aged from 32 to 60 years, 21 were (70%) tested positive for P. falciparum gametocytes by Pfg17, but only 7 (23.3%) were positive by Pfs25 [61].

For P. vivax, Pvs 25 and Pvs16 that are specific to female gametocytes are two conventional gene markers for gametocyte detection [78]. One gametocyte roughly corresponds to four Pvs25 transcripts per cell [79], and Pvs25 can detect from a mean of 0.34 gametocytes per μL blood from P. vivax patients in Papua New Guinea [78] to a mean of 2 gametocytes per μL blood in patients from Ethiopia [75]. Such low gametocyte densities make them extremely difficult to be detected by microscopy. The number of Pvs25 gene transcript copes detected by qRT-PCR directly correlates with the number of mature gametocytes as well as the overall parasite densities [41, 80] and was shown a nearly normal distribution with a mean of 1.2 × 107 copies/μL (ranging from 1.1 to 4.8 × 108 copies/μL) blood among 42 symptomatic P. vivax patients from northwestern Brazil [81]. Prior studies showed that age is tightly associated with gametocytemia. A lower proportion of infections with gametocytes was found with increasing age [79, 82]. Gametocytes are generally detected in ~20% of the infections among adults [41], but at much higher proportions in children under the age of 12 [82, 83]. Yet, gametocytemia in adults is up to 20-fold higher than in children [84, 85]. In areas with low levels of transmission, a large proportion of infections that are undetected by microscopy could be reservoir for parasites with high infectious gametocytes [86]. In Ethiopia, symptomatic P. vivax infections are nearly four times more infectious than asymptomatic ones [87]. Other factors such as host immune response, parasite strains, red blood cell density, antimalarial drug use, and relapse can also affect gametocyte production [79, 88, 89]. For P. malariae and P. ovale, there is yet no gametocyte assays due to little success in finding Pfs25 or Pvs25 orthologues in these species.

3.2 Novel gene candidates to improve detection sensitivity

Recently, the male-specific gene pfs13 that offers the lowest female to male expression ratio was identified as a new male gametocyte biomarker used in qRT-PCR assays from field isolates [90]. Other male-specific markers including PF3d7_1311100, PF3D7_1325200, and Pfg14–748 were also shown with abundant stage V male gametocytes in infected samples that made them as sensitive biomarker candidates [74], although their specific functions are unclear. Furthermore, CCp1, CCp3, and P25 were identified as better biomarkers in differentiating female from male gametocytes [74, 91]. These genes are highly expressed in stages III to V female gametocytes. Though another gene NEK4 is female-specific, it is not sufficiently expressed in the transcriptome until stages IV and V, suggesting that it could be a good late-stage female biomarker candidate. PF3d7_1107800, a putative AP2 transcription factor, also strongly correlates as a female specific gametocyte gene similar to NEK4. PF3d7_1107800 is highly expressed only during stages IV and V, and thus, could potentially a biomarker for late-stage female gametocyte detection [74]. Besides, female gametocyte marker gene CCp4 was also identified as a new target for gametocyte detection [92]. The design of intron-spanning primers of these novel genes allows for the amplification of mRNA only without a DNA digestion step [92].

A recent study of 26 P. vivax samples from Cambodian patients indicated that the expression profile of 21 predicted gametocyte genes were clustered in two distinct groups [93]. One group includes Pvs25, ULG8, gametocyte developmental protein 1, guanylate kinase, HMGB1, and five CPW-WPC proteins that associate with intracellular trafficking and histone remodeling in the female gametocytes. The other group includes Pvs47, Pvs48/45. Hap2, the gamete egress and sporozoite traversal protein, s16, and three CPW-WPC proteins that associate with microtubular development in the male gametocytes. It remains to be determined if these male and female gametocyte genes show higher expression than the conventional marker Pvs25, offer high detectability of the total potentially transmitting gametocyte densities (both male and female gametocytes), and allow for robust gametocyte sex-ratio estimates in field studies given their stability under suboptimal storage conditions [92].

3.3 Treatment and prevention

In P. falciparum infections, gametocytes express surface antibodies against the surface antigens for circumsporozoite protein (PfCSP) 2A10, apical membrane antigen 1 (PfAMA1), and thrombospondin-related adhesive protein (TRAP). In P. vivax infections, Pvs48/45 and PfCLAG9 have been shown to elicit naturally acquired immune responses [65, 66]. Gene 1613 has been identified to be critical for the development and maturation of gametocytes, which could potentially impact the elicitation of humoral immune response and the ability of gametocytes to transmit [94]. These could be critical antigen proteins for vaccine design. As gametocytes mature from one stage to another, the production of antibodies in human hosts will cause gametocytes to develop and mature at a much slower rate than normal in the bone marrow [94, 95, 96]. Combining multiple antigens involved in different stages of gametocyte development and sexual progression can help interfere and diminish transmission [69]. Predicting the structure of the parasite antigen binding domains e.g., in AMA1 and Pfs48/45, and delineating epitopes targeted by the host antibodies could uncover key elements in blocking transmission and provide a benchmark for evaluating vaccine efficacy [70]. Vaccines that block the ability of gametocytes to transmit can reduce infections and lower the transmission potential from mosquitoes to humans in malaria-endemic areas [67]. Future studies should focus on expanding our understanding of gene interactions and their functions as well as disease control by way of transmission blocking vaccine development [71]. Furthermore, uncharacterized chemotypes that possess activity against sexual parasites have also been recently identified to inhibit transmission to mosquito and human contact [68]. Interference by JmjC inhibitor known as ML324 causes expression to subsequently cease methylation activity followed by gametocyte lysis for P. falciparum. Identifying key surface proteins present in multi-parasite life rostrums could potentially be means of a preventative antimalarial against gametocytes [68].


4. Conclusions

To date, gametocytogenesis and gametocyte transmission tracking remain largely uncharacterized due to low prevalence and technological hurdles. As several countries approach the elimination phase for malaria, the need for sensitive and reliable biomarkers for gametocyte detection is more urgent than ever. Microscopy has low detection limit to overcome low parasitemia loads and existing qPCR biomarkers fail to accurately detect both sexes of the parasites. Information on genetic polymorphisms and expression levels of gametocyte biomarkers enable researchers to develop a more sensitive and accurate diagnostic test for Plasmodium gametocytes. Knowledge of gametocyte reservoirs and their interactions with host immune system will help develop effective treatment and preventive strategies to minimize the risk of malaria transmission. We recommend future studies focus on gene interactions and protein functions involved in gametocyte development, as well as polymorphisms in novel gametocyte genes that could be targets for developing better diagnostics or transmission blocking vaccines.



We thank our colleagues at the University of North Carolina at Charlotte and Jimma University for the inspiration and discussion that lead to the conceptualization of this paper. We also thank the reviewers for their insightful comments.


Conflict of interest

The authors declare no conflict of interest.


Author contributions

Conceptualization, D.K., A.F., and E.L.; resources, D.Y. and E.L.; writing— original draft preparation, D.K., A.F., E.L., G.K., D.Y., and E.L.; writing—review and editing, D.K., B.A., A.B., S.D.S.I., D.Y., and E.L.; funding acquisition, E.L. All authors have read and agreed to the published version of the manuscript.



This research was funded by NIH/NIAID, grant number R15 AI138002.


  1. 1. Mayxay, M., et al., Mixed-species malaria infections in humans. Trends Parasitol, 2004. 20(5): p. 233-240.
  2. 2. Muhseen, Z.T., et al., Natural products for treatment of Plasmodium falciparum malaria: An integrated computational approach. Comput Biol Med, 2021. 134: p. 104415.
  3. 3. Organization, G.W.H., World malaria report 2020: 20 years of global progress and challenges. 2020.
  4. 4. Snow, R.W., et al., The prevalence of Plasmodium falciparum in sub-Saharan Africa since 1900. Nature, 2017. 550(7677): p. 515-518.
  5. 5. Jaskiewicz, E., et al., Erythrocyte glycophorins as receptors for Plasmodium merozoites. Parasit Vectors, 2019. 12(1): p. 317.
  6. 6. Dayananda, K.K., R.N. Achur, and D.C. Gowda, Epidemiology, drug resistance, and pathophysiology of Plasmodium vivax malaria. J Vector Borne Dis, 2018. 55(1): p. 1-8.
  7. 7. Okafor, C.N. and N.A. Finnigan, Plasmodium Ovale Malaria, in StatPearls. 2021, StatPearls Publishing Copyright © 2021, StatPearls Publishing LLC.: Treasure Island (FL).
  8. 8. Baird, J.K., Purnomo, and S. Masbar, Plasmodium ovale in Indonesia. Southeast Asian J Trop Med Public Health, 1990. 21(4): p. 541-4.
  9. 9. Groger, M., et al., A systematic review of the clinical presentation, treatment and relapse characteristics of human Plasmodium ovale malaria. Malar J, 2017. 16(1): p. 112.
  10. 10. Robinson, L.J., et al., Strategies for understanding and reducing the Plasmodium vivax and Plasmodium ovale hypnozoite reservoir in Papua New Guinean children: a randomised placebo-controlled trial and mathematical model. PLoS Med, 2015. 12(10): p. e1001891.
  11. 11. Zhou, R., et al., Characterization of Plasmodium ovale spp. imported from Africa to Henan Province, China. Sci Rep, 2019. 9(1): p. 2191.
  12. 12. Shin, H.I., et al., Diagnosis and Molecular Analysis on Imported Plasmodium ovale curtisi and P. ovale wallikeri Malaria Cases from West and South Africa during2013-2016. Korean J Parasitol, 2020. 58(1): p. 61-65.
  13. 13. Yman, V., et al., Persistent transmission of Plasmodium malariae and Plasmodium ovale species in an area of declining Plasmodium falciparum transmission in eastern Tanzania. PLoS Negl Trop Dis, 2019. 13(5): p. e0007414.
  14. 14. Singh, B. and C. Daneshvar, Human infections and detection of Plasmodium knowlesi. Clin Microbiol Rev, 2013. 26(2): p. 165-184.
  15. 15. Barber, B.E., et al., World Malaria Report: time to acknowledge Plasmodium knowlesi malaria. Malar J, 2017. 16(1): p. 135.
  16. 16. White, N.J., Plasmodium knowlesi: The Fifth Human Malaria Parasite. Clinical Infectious Diseases, 2008. 46(2): p. 172-173.
  17. 17. Sallum, M.A., E.L. Peyton, and R.C. Wilkerson, Six new species of the Anopheles leucosphyrus group, reinterpretation of An. elegans and vector implications. Med Vet Entomol, 2005. 19(2): p. 158-199.
  18. 18. WHARTON, R.H. and D.E. EYLES, Anopheles hackeri, a vector of Plasmodium knowlesi in Malaya. Science, 1961. 134(3474): p. 279-280.
  19. 19. Milner, D.A., Jr., MalariaPathogenesis. Cold Spring Harb Perspect Med, 2018. 8(1).
  20. 20. Kori, L.D., N. Valecha, and A.R. Anvikar, Insights into the early liver stage biology of Plasmodium. J Vector Borne Dis, 2018. 55(1): p. 9-13.
  21. 21. Sterling, C.R., et al., Erythrocyte membrane alterations induced by Plasmodium simium infection in Saimiri sciureus: relation to Schüffner's dots. J Parasitol, 1975. 61(2): p. 177-188.
  22. 22. Golassa, L., et al., The biology of unconventional invasion of Duffy-negative reticulocytes by Plasmodium vivax and its implication in malaria epidemiology and public health. Malar J, 2020. 19(1): p. 299.
  23. 23. Meyer, E.V., et al., The reticulocyte binding-like proteins of P. knowlesi locate to the micronemes of merozoites and define two new members of this invasion ligand family. Mol Biochem Parasitol, 2009. 165(2): p. 111-121.
  24. 24. Semenya, A.A., et al., Two functional reticulocyte binding-like (RBL) invasion ligands of zoonotic Plasmodium knowlesi exhibit differential adhesion to monkey and human erythrocytes. Malar J, 2012. 11: p. 228.
  25. 25. Rossati, A., et al., Climate, environment and transmission of malaria. Infez Med, 2016. 24(2): p. 93-104.
  26. 26. Beier, J.C., Malaria parasite development in mosquitoes. Annu Rev Entomol, 1998. 43: p. 519-543.
  27. 27. Ngotho, P., et al., Revisiting gametocyte biology in malaria parasites. FEMS Microbiol Rev, 2019. 43(4): p. 401-414.
  28. 28. Basu, S. and P.K. Sahi, Malaria: An Update. Indian J Pediatr, 2017. 84(7): p. 521-528.
  29. 29. Kepple, D., et al., Alternative InvasionMechanismsand Host Immune Response to Plasmodium vivax Malaria: Trends and Future Directions. Microorganisms, 2020. 9(1).
  30. 30. Bansal, G.P. and N. Kumar, Immune Responses in Malaria Transmission. Current Clinical Microbiology Reports, 2018. 5(1): p. 38-44.
  31. 31. Ngwa, C.J., T.F.d.A. Rosa, and G. Pradel, The Biology of Malaria Gametocytes, Current Topics in Malaria, in Current Topics in Malaria, A.J. Rodriguez-Morales, Editor. 2016: IntechOpen.
  32. 32. Saini, E., et al., Photosensitized INA-Labelled protein 1 (PhIL1) is novel component of the inner membrane complex and is required for Plasmodium parasite development. Sci Rep, 2017. 7(1): p. 15577.
  33. 33. Carter, R., et al., Plasmodium falciparum: an abundant stage-specific protein expressed during early gametocyte development. Exp Parasitol, 1989. 69(2): p. 140-149.
  34. 34. Drakeley, C.J., et al., Transmission-reducing immunity is inversely related to age in Plasmodium falciparum gametocyte carriers. Parasite Immunol, 2006. 28(5): p. 185-190.
  35. 35. Sinden, R.E., Infection of mosquitoes with rodent malaria, in The Molecular Biology of Insect Disease Vectors, J.M. Crampton, C.B. Beard, and C. Louis, Editors. 1997, Springer, Dordrecht. p. 67-91.
  36. 36. Talman, A.M., et al., Influence of chemotherapy on the Plasmodium gametocyte sex ratio of mice and humans. Am J Trop Med Hyg, 2004. 71(6): p. 739-744.
  37. 37. Gautret, P., et al., Enhanced gametocyte formation by Plasmodium chabaudi in immature erythrocytes: pattern of production, sequestration, and infectivity to mosquitoes. J Parasitol, 1996. 82(6): p. 900-6.
  38. 38. Smalley, M.E. and J. Brown, Plasmodium falciparum gametocytogenesis stimulated by lymphocytes and serum from infected Gambian children. Trans R Soc Trop Med Hyg, 1981. 75(2): p. 316-317.
  39. 39. Trager, W. and G.S. Gill, Enhanced gametocyte formation in young erythrocytes by Plasmodium falciparum in vitro. J Protozool, 1992. 39(3): p. 429-432.
  40. 40. Gardiner, D.L. and K.R. Trenholme, Plasmodium falciparum gametocytes: playing hide and seek. Ann Transl Med, 2015. 3(4): p. 45.
  41. 41. Bousema, T. and C. Drakeley, Epidemiology and infectivity of Plasmodium falciparum and Plasmodium vivax gametocytes in relation to malaria control and elimination. Clin Microbiol Rev, 2011. 24(2): p. 377-410.
  42. 42. Hawking, F., M.E. Wilson, and K. Gammage, Evidence for cyclic development and short-lived maturity in the gametocytes of Plasmodium falciparum. Trans R Soc Trop Med Hyg, 1971. 65(5): p. 549-559.
  43. 43. Joice, R., et al., Plasmodium falciparum transmission stages accumulate in the human bone marrow. Sci Transl Med, 2014. 6(244): p. 244re5.
  44. 44. Tibúrcio, M., et al., A switch in infected erythrocyte deformability at the maturation and blood circulation of Plasmodium falciparum transmission stages. Blood, 2012. 119(24): p. e172-e180.
  45. 45. Silvestrini, F., P. Alano, and J.L. Williams, Commitment to the production of male and female gametocytes in the human malaria parasite Plasmodium falciparum. Parasitology, 2000. 121Pt 5: p. 465-471.
  46. 46. Muñoz, E.E., et al., ALBA4 modulates its stage-specific interactions and specific mRNA fates during Plasmodium yoelii growth and transmission. Mol Microbiol, 2017. 106(2): p. 266-284.
  47. 47. Trelle, M.B., et al., Global histone analysis by mass spectrometry reveals a high content of acetylated lysine residues in the malaria parasite Plasmodium falciparum. J Proteome Res, 2009. 8(7): p. 3439-3450.
  48. 48. Gunalan, K., et al., Plasmodium vivax Infections of Duffy-Negative Erythrocytes: Historically Undetected or a Recent Adaptation? Trends in Parasitology, 2018. 34(5): p. 420-429.
  49. 49. Josling, G.A., K.C. Williamson, and M. Llinás, Regulation of Sexual Commitment and Gametocytogenesis in Malaria Parasites. Annu Rev Microbiol, 2018. 72: p. 501-519.
  50. 50. Coetzee, N., et al., Quantitative chromatin proteomics reveals a dynamic histone post-translational modification landscape that defines asexual and sexual Plasmodium falciparum parasites. Sci Rep, 2017. 7(1): p. 607.
  51. 51. Miao, J., et al., The malaria parasite Plasmodium falciparum histones: organization, expression, and acetylation. Gene, 2006. 369: p. 53-65.
  52. 52. Lopez-Rubio, J.J., L. Mancio-Silva, and A. Scherf, Genome-wide analysis of heterochromatin associates clonally variant gene regulation with perinuclear repressive centers in malaria parasites. Cell Host Microbe, 2009. 5(2): p. 179-190.
  53. 53. Flueck, C., et al., Plasmodium falciparum heterochromatin protein 1 marks genomic loci linked to phenotypic variation of exported virulence factors. PLoS Pathog, 2009. 5(9): p. e1000569.
  54. 54. Fraschka, S.A., et al., Comparative Heterochromatin Profiling Reveals Conserved and Unique Epigenome Signatures Linked to Adaptation and Development of Malaria Parasites. Cell Host Microbe, 2018. 23(3): p. 407-420.e8.
  55. 55. Bechtsi, D.P. and A.P. Waters, Genomics and epigenetics of sexual commitment in Plasmodium. International Journal for Parasitology, 2017. 47(7): p. 425-434.
  56. 56. Campbell, T.L., et al., Identification and genome-wide prediction of DNA binding specificities for the ApiAP2 family of regulators from the malaria parasite. PLoS Pathog, 2010. 6(10): p. e1001165.
  57. 57. Coleman, B.I., et al., A Plasmodium falciparum histone deacetylase regulates antigenic variation and gametocyte conversion. Cell Host Microbe, 2014. 16(2): p. 177-186.
  58. 58. Chawla, J., J. Oberstaller, and J.H. Adams, Targeting Gametocytes of the Malaria Parasite Plasmodium falciparum in a Functional Genomics Era: Next Steps. Pathogens, 2021. 10(3).
  59. 59. Ramphul, U.N., et al., Plasmodium falciparum evades mosquito immunity by disrupting JNK-mediated apoptosis of invaded midgut cells. Proc Natl Acad Sci U S A, 2015. 112(5): p. 1273-1280.
  60. 60. Molina-Cruz, A., et al., Plasmodium evasion of mosquito immunity and global malaria transmission: The lock-and-key theory. Proc Natl Acad Sci U S A, 2015. 112(49): p. 15178-15183.
  61. 61. Kengne-Ouafo, J.A., et al., Immune Responses to the Sexual Stages of Plasmodium falciparum Parasites. Front Immunol, 2019. 10: p. 136.
  62. 62. Eksi, S., A. Suri, and K.C. Williamson, Sex- and stage-specific reporter gene expression in Plasmodium falciparum. Mol Biochem Parasitol, 2008. 160(2): p. 148-151.
  63. 63. Jafari-Guemouri, S., et al., Dynamics of Plasmodium falciparum gametocyte carriage in pregnant women under intermittent preventive treatment with sulfadoxine-pyrimethamine in Benin. Malar J, 2018. 17(1): p. 356.
  64. 64. Scally, S.W., et al., Molecular definition of multiple sites of antibody inhibition of malaria transmission-blocking vaccine antigen Pfs25. Nat Commun, 2017. 8(1): p. 1568.
  65. 65. McLeod, B., et al., Potent antibody lineage against malaria transmission elicited by human vaccination with Pfs25. Nat Commun, 2019. 10(1): p. 4328.
  66. 66. Williamson, K.C., Pfs230: from malaria transmission-blocking vaccine candidate toward function. Parasite Immunol, 2003. 25(7): p. 351-359.
  67. 67. Theisen, M., M.M. Jore, and R. Sauerwein, Towards clinical development of a Pfs48/45-based transmission blocking malaria vaccine. Expert Rev Vaccines, 2017. 16(4): p. 329-336.
  68. 68. Eksi, S., et al., Malaria transmission-blocking antigen, Pfs230, mediates human red blood cell binding to exflagellating male parasites and oocyst production. Mol Microbiol, 2006. 61(4): p. 991-998.
  69. 69. Kumar, N., Target antigens of malaria transmission blocking immunity exist as a stable membrane bound complex. Parasite Immunol, 1987. 9(3): p. 321-335.
  70. 70. Gerloff, D.L., et al., Structural models for the protein family characterized by gamete surface protein Pfs230 of Plasmodium falciparum. Proc Natl Acad Sci U S A, 2005. 102(38): p. 13598-13603.
  71. 71. Singh, K., et al., Structure and function of a malaria transmission blocking vaccine targeting Pfs230 and Pfs230-Pfs48/45 proteins. Commun Biol, 2020. 3(1): p. 395.
  72. 72. Kundu, P., et al., Structural delineation of potent transmission-blocking epitope I on malaria antigen Pfs48/45. Nat Commun, 2018. 9(1): p. 4458.
  73. 73. Escalante, A.A., et al., Polymorphism in the gene encoding the Pfs48/45 antigen of Plasmodium falciparum. XI. Asembo Bay Cohort Project. Mol Biochem Parasitol, 2002. 119(1): p. 17-22.
  74. 74. Walzer, K.A., et al., Single-Cell Analysis Reveals Distinct Gene Expression and Heterogeneity in Male and Female Plasmodium falciparum Gametocytes. mSphere, 2018. 3(2).
  75. 75. Tadesse, F.G., et al., The shape of the iceberg: quantification of submicroscopic Plasmodium falciparum and Plasmodium vivax parasitaemia and gametocytaemia in five low endemic settings in Ethiopia. Malar J, 2017. 16(1): p. 99.
  76. 76. Nishimoto, Y., et al., Evolution and phylogeny of the heterogeneous cytosolic SSU rRNA genes in the genus Plasmodium. Mol Phylogenet Evol, 2008. 47(1): p. 45-53.
  77. 77. Essuman, E., et al., A Novel Gametocyte Biomarker for Superior Molecular Detection of the Plasmodium falciparum Infectious Reservoirs. J Infect Dis, 2017. 216(10): p. 1264-1272.
  78. 78. Wampfler, R., et al., Strategies for detection of Plasmodium species gametocytes. PLoS One, 2013. 8(9): p. e76316.
  79. 79. Koepfli, C., et al., Blood-Stage Parasitaemia and Age Determine Plasmodium falciparum and P. vivax Gametocytaemia in Papua New Guinea. PLoS One, 2015. 10(5): p. e0126747.
  80. 80. Bharti, A.R., et al., Experimental infection of the neotropical malaria vector Anopheles darlingi by human patient-derived Plasmodium vivax in the Peruvian Amazon. Am J Trop Med Hyg, 2006. 75(4): p. 610-616.
  81. 81. Lima, N.F., M.S. Bastos, and M.U. Ferreira, Plasmodium vivax: reverse transcriptase real-time PCR for gametocyte detection and quantitation in clinical samples. Exp Parasitol, 2012. 132(3): p. 348-354.
  82. 82. Nacher, M., et al., Seasonal fluctuations in the carriage of Plasmodium vivax gametocytes in Thailand. Ann Trop Med Parasitol, 2004. 98(2): p. 115-120.
  83. 83. Olliaro, P.L., et al., Implications of Plasmodium vivax Biology for Control, Elimination, and Research. Am J Trop Med Hyg, 2016. 95(6 Suppl): p. 4-14.
  84. 84. Dixon, M.W., et al., Sex in Plasmodium: a sign of commitment. Trends Parasitol, 2008. 24(4): p. 168-175.
  85. 85. Reece, S.E., R.S. Ramiro, and D.H. Nussey, Plastic parasites: sophisticated strategies for survival and reproduction? Evol Appl, 2009. 2(1): p. 11-23.
  86. 86. Hofmann, N.E., et al., Assessment of ultra-sensitive malaria diagnosis versus standard molecular diagnostics for malaria elimination: an in-depth molecular community cross-sectional study. Lancet Infect Dis, 2018. 18(10): p. 1108-1116.
  87. 87. Tadesse, F.G., et al., The Relative Contribution of Symptomatic and Asymptomatic Plasmodium vivax and Plasmodium falciparum Infections to the Infectious Reservoir in a Low-Endemic Setting in Ethiopia. Clin Infect Dis, 2018. 66(12): p. 1883-1891.
  88. 88. Baker, D.A., Malaria gametocytogenesis. Mol Biochem Parasitol, 2010. 172(2): p. 57-65.
  89. 89. Drakeley, C., et al., The epidemiology of Plasmodium falciparum gametocytes: weapons of mass dispersion. Trends Parasitol, 2006. 22(9): p. 424-430.
  90. 90. Santolamazza, F., et al., Detection of Plasmodium falciparum male and female gametocytes and determination of parasite sex ratio in human endemic populations by novel, cheap and robust RTqPCR assays. Malar J, 2017. 16(1): p. 468.
  91. 91. Bennink, S., M.J. Kiesow, and G. Pradel, The development of malaria parasites in the mosquito midgut. Cell Microbiol, 2016. 18(7): p. 905-918.
  92. 92. Meerstein-Kessel, L., et al., A multiplex assay for the sensitive detection and quantification of male and female Plasmodium falciparum gametocytes. Malar J, 2018. 17(1): p. 441.
  93. 93. Kim, A., et al., Plasmodium vivax transcriptomes reveal stage-specific chloroquine response and differential regulation of male and female gametocytes. Nat Commun, 2019. 10(1): p. 371.
  94. 94. Julien, J.P. and H. Wardemann, Antibodies against Plasmodium falciparum malaria at the molecular level. Nat Rev Immunol, 2019. 19(12): p. 761-775.
  95. 95. de Jong, R.M., et al., Immunity against sexual stage Plasmodium falciparum and Plasmodium vivax parasites. Immunol Rev, 2020. 293(1): p. 190-215.
  96. 96. Hall, C.E., et al., Mosquito Bite-Induced Controlled Human Malaria Infection with Plasmodium vivax or P. falciparum Generates Immune Responses to Homologous and Heterologous Preerythrocytic and Erythrocytic Antigens. Infect Immun, 2019. 87(3).

Written By

Daniel Kepple, Anthony Ford, Ebony Little, Gabrielle Kolesar, Beka R. Abagero, Ashley N. Blackwell, Swarnapali De Silva Indrasekara, Delenasaw Yewhalaw and Eugenia Lo

Submitted: 29 June 2021 Reviewed: 09 July 2021 Published: 30 August 2021