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Advances in the dissection of Anopheles–Plasmodium interactions

  • Sally A. Saab,

    Affiliation W. Harry Feinstone Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland, United States America

    ⨯
  • Victor Cardoso-Jaime,

    Affiliation W. Harry Feinstone Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland, United States America

    ⨯
  • Mary Kefi,

    Affiliation W. Harry Feinstone Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland, United States America

    ⨯
  • George Dimopoulos

    gdimopo1@jhu.edu, gdimopou@jhsph.edu

    Affiliation W. Harry Feinstone Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, Baltimore, Maryland, United States America

    ⨯

Abstract

Malaria is a life-threatening mosquito-borne disease caused by the Plasmodium parasite, responsible for more than half a million deaths annually and principally involving children. The successful transmission of malaria by Anopheles mosquitoes relies on complex successive interactions between the parasite and various mosquito organs, host factors, and restriction factors. This review summarizes our current understanding of the mechanisms regulating Plasmodium infection of the mosquito vector at successive plasmodial developmental stages and highlights potential transmission-blocking targets and strategies.

Citation: Saab SA, Cardoso-Jaime V, Kefi M, Dimopoulos G (2025) Advances in the dissection of Anopheles–Plasmodium interactions. PLoS Pathog 21(3): e1012965. https://doi.org/10.1371/journal.ppat.1012965

Editor: Eric Calvo, National Institutes of Health, UNITED STATES OF AMERICA

Published: March 31, 2025

Copyright: © 2025 Saab et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work has been funded by the NIH / NIAID grants (R01AI170692 and R01AI158615 to GD) and Bloomberg Philanthropies (to GD). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

1. Introduction

Mosquitoes are vectors for a number of human pathogens that cause severe disease, such as malaria, Zika, chikungunya, dengue, and yellow fever [1]. According to the most recent World Health Organization (WHO) report, malaria caused an estimated 249 million cases and 608,000 deaths worldwide in 2022, representing an increase of 5 million cases over 2021 numbers [2]. Six species of the Plasmodium protozoan parasite have been shown to cause human malaria: Plasmodium falciparum, P. vivax, P. ovale, P. malariae, P. knowlesi, and P. cynomologi [3]. In Africa, P. falciparum is the most commonly fatal species and is mainly transmitted by adult female members of the Anopheles gambiae species complex [4]. This complex comprises eight morphologically identical sibling species that vary in their geographical prevalence [5–7]. Apart from the invertebrate mosquito vector, the life cycle of Plasmodium also involves the vertebrate human host; however, all of the previous attempts that have focused on the human host have failed to produce a vaccine that can effectively eradicate malaria [8]. Despite the progression to advanced-clinical trials, most available vaccines cannot be considered sufficient as a stand-alone measure to worldwide malaria eradication [9]. Moreover, emerging parasite resistance against the anti-malarial drugs jeopardizes their efficiency [10, 11]. For instance, the emerging Plasmodium resistance to the fast-acting antimalarial drug artemisinin has led to a significant reduction in the drug’s efficacy, resulting in lower treatment success rates and prolonged illness [12]. Thus, control efforts have also been focused on transmission-blocking strategies that are designed to prevent the parasite from infecting the mosquito vector. One promising strategy is based on generating genetically engineered mosquitoes resistant to Plasmodium infection [13], a strategy that requires extensive knowledge of both mosquito and parasite biology. Here, we review our current understanding of Anopheles–Plasmodium interactions and potential manipulations to block the mosquito’s vectorial capacity.

2. Plasmodium development in the lumen of the mosquito gut

2.1. Gametogenesis

The sexual reproductive phase of the Plasmodium parasite begins when the mosquito ingests, along with mosquito saliva, an infected gametocyte-containing blood meal from a vertebrate host. Once the ingested gametocytes reach the mosquito midgut, they are exposed to the mosquito-derived byproduct of the tryptophan degradation, xanthurenic acid (XA), along with a temperature drop and pH elevation, and they respond by differentiating into mature gametes in a process known as gametogenesis [14–16]. Gametogenesis starts with the rounding up of the parasites within the ingested erythrocytes, preceding the “inside-out” egress of the parasites into the mosquito gut. This egress involves a sequential membrane rupture that starts with the rupture of the inner parasitophorous vacuole membrane and is followed by the rupture of the external erythrocyte membrane [17,18].

Gametogenesis differs between male and female gametocytes. Recent transcriptomic studies have reported that the histone variant H3.3 protein, which is enriched in female gametocytes, is responsible for the repression of the male-specific genes to favor female properties [19]. Male gametogenesis, which is known as exflagellation, involves three rapid consecutive DNA replication cycles that give rise to eight motile microgametes within 15 min after activation [18,20]; in contrast, female gametogenesis results in a single non-motile female macrogamete that is ready for fertilization [21].

A well-orchestrated molecular mechanism occurs in Plasmodium gametocytes to initiate gametogenesis. XA-mediated boosting of guanylyl cyclase (GC) activity increases the level of the second messenger 3′−5′-cyclic guanosine monophosphate (cGMP) in the gametocyte [22]. Despite the presence of two membrane GC proteins (GCα and GCβ) in P. falciparum [23], only GCα seems to be involved in cGMP production during gametogenesis, since disrupting GCβ does not prevent XA-stimulated gamete formation [24]. Recent studies in Plasmodium yoelii using CRISPR/Cas9 genome editing have shown that this XA-stimulated cGMP production is mediated by a membrane protein that spans multiple membranes. Known as gametogenesis essential protein 1 (GEP1), this protein interacts with GCα, enhancing its activity [25]. Another membrane protein known as G-protein-coupled receptor 180 (GPR180) promotes this cGMP elevation in P. berghei [26]. Protein kinase G (PKG) responds to the elevated levels of cGMP by phosphorylating a multipass membrane protein, recently named important for Ca2+ mobilization 1 (ICM1), which stimulates the mobilization of stored calcium into the cytosol [27–29]. This increase in cytosolic Ca2+ activates several calcium-dependent protein kinases (CDPK), which in turn translate this signal into various cellular responses required for gametogenesis. For instance, CDPK1 in P. falciparum is critical for both male and female gametogenesis. In the absence of CDPK1, female gametocytes are incapable of rounding up post-activation, as opposed to male gametocytes, which can round up but do not engage in exflagellation [30]. On the other hand, CDPK2 is only essential during male exflagellation [31]. Another major CDPK in male gametogenesis is CDPK4, which was recently shown to be responsible for essential processes, including DNA replication, mRNA translation, and cell motility [32].

In addition to CDPKs, other families of protein kinases are also implicated in Plasmodium male gametogenesis. The male-specific mitogen-activated protein kinase 2 (MAP-2) in P. falciparum has been reported to be essential for proper axonemal beating [33]. Recently, cyclin-dependent kinase-related kinases (CRKs) have also been found to be critical for male exflagellation, with deletion of CRK5 in P. falciparum resulting in defective male gametogenesis [34].

2.2. Gamete fertilization

After their egress from the ingested erythrocytes, Plasmodium male and female gametes fuse to form a diploid zygote within the first hour after intake of a blood meal [20,35]. Several surface proteins present on both male and female gametes are involved in this process (Fig 1). Two gamete proteins, P48/45 and P230, of the six-cysteine protein family form a complex on the surface of the gamete that is required for male fertility and subsequent fusion with the female macrogamete in P. falciparum [36,37]. Another surface six-cysteine protein, known as P47, is solely expressed on the female macrogamete surface and is necessary for female fertility in P. berghei but not P. falciparum [38,39]. Moreover, deleting the highly conserved hapless 2 protein (HAP2, also known as generative cell-specific 1) in P. berghei male microgametes does not prevent the male and female gametes from adhering to each other, but it inhibits their fusion [40]. In addition to HAP2, HAP2 paralog (HAP2p) is essential for the fertilization of P. falciparum gametes. Deleting either of these two membrane fusogens, which are located throughout the flagella of the male microgametes, will prevent gamete fertilization [41]. In addition, the histone chaperone protein FACT-L (named after the large subunit of the human protein, facilitates chromatin transcription [FACT]) plays a crucial role in male gamete fertility and subsequent gamete fusion, but the mechanism by which this nuclear protein affects fertilization is still not clear [42]. Recent evidence underlines the requirement for additional membrane proteins, Pb115 and Pbs54, for the attachment of male and female gametes in P. berghei [43,44].

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Fig 1. The Plasmodium life cycle in the mosquito vector.

Following an infected blood meal, the ingested Plasmodium gametocytes develop into male and female gametes that can fuse together to form a zygote. The zygote then differentiates into a motile ookinete and invades the midgut epithelium after traversing the peritrophic matrix separating the ingested blood meal from the epithelial cells. Right after it reaches the extracellular space between the epithelium and the basal lamina, the ookinete develops into an early oocyst, which in turn undergoes sporogony to give rise to a mature oocyst holding thousands of sporozoites. Following rupture of the oocyst, the released sporozoites invade the salivary glands and are ready for transmission to a new host. Proteins mediating the various phases of the life cycle are mentioned in the panels below the corresponding developmental stages. Created in BioRender. Saab, S. (2025) https://BioRender.com/p65s417.

https://doi.org/10.1371/journal.ppat.1012965.g001

Most of the previously mentioned Plasmodium surface proteins have been studied for their transmission-blocking vaccine (TBV) potential, and continuous efforts have been made to identify various domains in P230, P48/45, P47, and HAP2 that could be used for inducing transmission-reducing antibodies in the host [45–50].

2.3. Development of the zygote into a motile ookinete

Following gamete fusion, the diploid zygote undergoes DNA replication, resulting in a tetraploid phase that persists up to the formation of the haploid sporozoites within the oocysts. This process has been shown to be regulated by the NIMA-related kinases Nek-2 and Nek-4, which in turn are essential for ookinete differentiation [51].

At 18–30 h after a blood meal, the zygote differentiates into a motile ookinete stage, which invades the midgut epithelium of the mosquito after traversing the peritrophic matrix, the semipermeable chitinous layer separating the blood bolus from the midgut epithelial cells (Fig 1) [52–54]. Studies of the dynamics of zygote-ookinete development in An. coluzzii mosquitoes by live immunofluorescence microscopy have demonstrated that P. falciparum passes through several intermediate stages before becoming a mature ookinete. Formation of mature ookinetes peaks at 23 h following the ingestion of infected blood and persists in the gut up to 36 hpost-infection [55].

Recent single-cell RNA-seq data have linked these various stages to their gene expression profiles, with the essential genes for DNA replication and metabolism detected in early zygotes, and those required for host invasion identified in later ookinete stages [56]. Also, studies in P. berghei have shown that only the maternally inherited alleles are active in the zygote-ookinete stages; the paternally inherited genes are expressed later on, during midgut invasion [57]. Such gene expression profiles are translated into several cell structure modifications, including the establishment of a special organelle known as the inner membrane complex (IMC), located beneath the Plasmodium cell membrane, as well as the cell polarization that occurs during zygote-to-ookinete differentiation [58].

While the transition between the various Plasmodium developmental stages is strongly coordinated by various apicomplexan Apetala-2 transcription factors [59–61], translational regulation has also been reported to play an essential role in repressing the expression of certain mRNAs [62]. Recently, Hirai et al. have identified an RNA binding protein in P. berghei known as Pb103, which has the two zinc finger domains required for zygote differentiation, most probably through translational repression [63]. In addition to translational regulation, post-translational modifications are also crucial for zygote differentiation. Studies using P. yoelii have pointed to several pellicle proteins, including IMC subcompartment proteins 1 and 3 (ISP1 and ISP3), IMC‐residing palmitoyl‐S‐acyl‐transferase (PAT) DHHC2, and the cytoskeletal microtubule β-tubulin, being involved in the proper arrangement of the cytoskeletal subpellicular microtubules through a process that requires palmitoylation to promote zygote elongation [64]. Thus, inhibiting palmitoylation might be a promising way to prevent Plasmodium development [65].

2.4. Anti-plasmodial immune responses in the lumen of the mosquito midgut

Upon the ingestion of an infected blood meal, the mosquito gut microbiota, mainly composed of gram-negative bacteria [66,67], proliferate and play a protective role against the Plasmodium parasite by immune priming and the secretion of factors that can inhibit parasite development. Hence, elimination of the microbiota in antibiotic-treated An. gambiae mosquitoes leads to a significant increase in P. falciparum oocyst numbers, thus rendering the mosquitoes more susceptible to the parasite [66]. Accordingly, an Enterobacter species isolated from Anopheles arabiensis in Zambia can directly kill Plasmodium through the production of reactive oxygen species (ROS) [68]. Furthermore, the Chromobacterium species, which was initially isolated from Aedes aegypti mosquitoes in Panama, exerts an anti-Plasmodium effect by secreting romidepsin, a histone deacetylase inhibitor, which inhibits Plasmodium [69,70]. In addition, Serratia marcescens bacteria isolated from mosquito midguts also secrete uncharacterized metabolites that can directly inhibit the Plasmodium ookinete stage, independent of the mosquito vector [71]. Recently, certain strains of Serratia ureilytica, isolated from wild An. sinensis in Tengchong, China, have been shown to produce an anti-plasmodial lipase that can target the early gametocyte stage of the parasite in the mosquito midgut, thus preventing ookinete formation [72].

Apart from their direct effects, the midgut microbiota can also indirectly protect a host against Plasmodium by triggering the expression of the mosquito immune genes. After a blood meal, the gut microbiota proliferates and releases immune elicitors that can activate the immune deficiency (Imd) pathway, which targets the P. falciparum ookinete stage through multiple effectors [73–76]. The activation of the Imd signaling pathway results in the nuclear translocation of the NF-κB transcription factor Rel2, triggering the expression of several immune genes, including anti-Plasmodium factors and AMPs [74]. Defensins, cecropins, attacin, and gambicin are the four different classes of AMPs identified in the An. gambiae genome [77]. These AMPs target a wide range of pathogens, including Plasmodium, gram-positive and gram-negative bacteria, fungi, and yeasts [74]. The Imd pathway acts in concert with the Toll signaling pathway, contributing to the expression of defensin 1, cecropin 1, and gambicin [78]. While activating the Imd pathway abolishes P. falciparum infection in three different Anopheles species, it has no effect on P. berghei, highlighting the specificity of various immune pathways for various pathogens [75]. Furthermore, the antiparasitic role of the Imd signaling pathway has been further emphasized in genetically modified An. stephensi overexpressing Rel2 in both the midgut and the fat bodies, which renders the mosquitoes resistant to P. falciparum [79].

Recently, DNA methylation was shown to play a crucial role in regulating the midgut anti-plasmodial immune responses driving the differential susceptibility of various Anopheles albimanus phenotypes to P. berghei [80]. Interestingly, the transgenic depletion of several microRNAs in An. gambiae using microRNA sponges modulates the expression of the midgut immune genes, making the mosquitoes more resistant to parasitic infection [81].

3. Plasmodium ookinete invasion of the mosquito midgut epithelium

3.1. Ookinete invasion of the mosquito gut

The ookinete first crosses the chitin-containing peritrophic matrix by releasing chitinase and then invades the epithelial cells through a complex process that requires several interactions between the parasite and the mosquito factors [82] (Fig 1). The epithelial cell invasion is mediated in part by enolase on the ookinete surface, which interacts with the ingested plasminogen and the epithelial cells, allowing midgut invasion to occur [83]. Other ookinete-surface proteins also facilitate the traversal of the midgut epithelium. For example, P25/28, P47, and Plasmodium infection of the mosquito midgut screen 43 (PIMMS43) help the ookinete evade the mosquito’s immune responses. At the same time, PIMMS2 is essential for midgut invasion [84–87].

Apart from the proteins expressed on its surface, mature ookinetes also secrete other micronemal proteins that mediate midgut invasion. These proteins include the secreted ookinete adhesive protein (SOAP), circumsporozoite- and TRAP-related protein (CTRP), cell-traversal protein for ookinete and sporozoites (CelTOS), Plasmodium perforin-like protein 3-5 (PPLP3-5), chitinase, and GPI-anchored micronemal antigen (GAMA) [88–95]. In addition, the A domain of CTRP is crucial for the ookinete’s gliding motility, which in turn makes possible the penetration of the mosquito midgut [96].

Recently, another member of the perforin-like proteins, PPLP4, was also reported to have a role in traversing the epithelia [97]. Along with the parasite proteins, peritrophic matrix-associated fibrinogen-related protein 1 (FREP1), which is released by the epithelial cells, can directly bind to ookinetes, enabling them to penetrate both the peritrophic matrix and the epithelium [98]. The fibrinogen-like domain of FREP1, which is responsible for its direct interaction with Plasmodium gametocytes and ookinetes, is highly conserved between the various Anopheles species and is an effective TBV candidate [99,100].

Recently, Plasmodium α-tubulin-1, expressed on the apical end of the ookinetes, was shown to interact with FREP1 to direct the ookinetes toward the peritrophic matrix, thus facilitating parasite invasion [101]. Blocking the direct interaction between FREP1 and the parasite through anti-fibrinogen-like domain or anti-α-tubulin-1 antibodies significantly reduces P. falciparum infection in An. gambiae mosquitoes, making the FREP1-tubulin interaction a robust transmission-blocking target [99,101]. Interestingly, An. gambiae alanyl aminopeptidase N1 (AgAPN1), which is expressed on midgut epithelial cells, can also play an essential role in facilitating the invasion by P. berghei and P. falciparum ookinetes [102]. Despite this significant research, more work is still needed to elucidate more fully the interactions between AgAPN1 and the parasite; however, determining the crystal structure of AgAPN1 has been of great help in working out how to target this enzyme with domain-specific antibodies that are promising for transmission-blocking [103,104].

Ookinetes have been suggested to penetrate the midgut epithelium through both intercellular and intracellular routes by using their ability to move by gliding. Use of either of these invasion routes was recently shown to be a mosquito species-specific process [105]. Studies with P. yoelii have linked this gliding motility to the IMC sub-compartment protein 1 [ISP1]-mediated polarization of guanylate cyclase β from the cytoplasm to the “ookinete extrados site”, a site posterior to the apical structure in mature ookinetes, thereby activating the cGMP-dependent PKG signaling pathway that is responsible for the gliding motility [106].

While in the ookinete stage, the parasite passes through a major population bottleneck, making this stage a target for future antimalarial transmission-blocking interventions [107]. Surface or secreted ookinete proteins that are involved in motility and invasion are robust candidates for TBVs [108,109]. In addition to vaccines, transmission-blocking strategies also include the generation of transgenic mosquitoes engineered to block the parasite and prevent its invasion [110]. For instance, transgenic An. stephensi expressing exogenous single-chain antibodies (scFv) that act against chitinase or Pf25, along with the sporozoite circumsporozoite protein (CSP), can successfully clear P. falciparum sporozoites from mosquitoes [111]. Also, CRISPR/Cas9-mediated FREP1 knockout in An. gambiae suppresses both human and rodent parasite infection [112].

3.2. Immune responses within the epithelial cells

Despite its ability to digest and penetrate the peritrophic matrix, the parasite remains protected in part from epithelium-elicited immune responses, thanks to cross-linkage of the mucus layer proteins by immunomodulatory peroxidase and the intestinal dual oxidase [113]. Another redox system mediates a more robust anti-plasmodial immune response, known as midgut-epithelial cell nitration, that can further modify the ookinete and make it a target for the complement system [114].

Nitration is a two-step reaction involving the synthesis of nitric oxide (NO), followed by a peroxidase reaction [115]. The synthesis of NO after Plasmodium infection requires the upregulation of nitric oxide synthase (NOS) downstream of the JAK-STAT signaling pathway. NOS plays a critical role in reducing Plasmodium oocyst survival [116]. Also, a heme peroxidase (HPX2)/NADPH oxidase 5 (NOX5) system, induced by the Jun-N-terminal kinase (JNK) signaling pathway, is responsible for mediating epithelial nitration and subsequent ookinete surface modification. These reactions mark the ookinete for targeting by the mosquito’s complement-like system through recruitment of the complement-like thioester-containing protein 1 (TEP1) as the ookinetes reach the extracellular space between the epithelium and the basal lamina and become exposed to the hemolymph [114,117]. Certain Pfs47 variants specific for the African Plasmodium strains can suppress midgut epithelial nitration in An. gambiae mosquitoes through binding to a certain Pfs47 midgut receptor (P47Rec), thereby enabling evasion of the mosquito’s immune response [118–120]. Recent studies addressing the mechanistic function of P47Rec have revealed the involvement of heat shock protein 70 cognate 3 (Hsc70-3) in inhibiting the caspase-mediated apoptosis of the invaded midgut cells and therefore disrupting epithelial nitration [121].

3.3. Hemolymph immune responses against the ookinete stage

After bypassing the local immune responses induced in the midgut, ookinetes become exposed to systemic immune responses on the basal side of the epithelium that are triggered in the fat body and hemocytes [122]. For instance, abdominal tissues such as the fat body can respond to the diffusion of NO and hydrogen peroxide from the mosquito midgut to the hemolymph by releasing anti-plasmodial AMPs and other factors [123]. Hemocytes can also be activated, after the nitration of infected epithelial cells, to release vesicles containing unknown factors that promote the binding of TEP1 to the ookinete [124], targeting the parasite for elimination through either lysis or melanization (Fig 2) [125,126].

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Fig 2. Mosquito immune responses against the early ookinete stage.

Plasmodium ookinete midgut invasion induces nitration, modifying the parasite surface for TEP1 recruitment. TEP1 is secreted into the hemolymph in its full form (TEP1-F). Upon activation, TEP1-F is cleaved by an unknown factor into the active cut form (TEP1cut), which is stabilized in the hemolymph by binding to the APL1C/LRIM1 heterodimer. Recognition of the ookinete results in the deposition and accumulation of TEP1 cut on the parasite surface which is promoted by SPCLIP1 due to the cleavage of more TEP1-F by an unknown TEP1 convertase. TEP1-marked ookinetes are then targeted for lysis or, in rare cases, melanization, which mainly occurs in certain refractory backgrounds (left panel). Melanization requires a cascade of catalytic and noncatalytic CLIP serine proteases that regulate the cleavage of PPO zymogen into active PO, the key protein in melanin synthesis. Altogether, thus far, TEP1 seems to be the factor farthest upstream, followed by the core cSPH module (SPCLIP1-CLIPA8-CLIPA28) that acts upstream of the catalytic CLIPC9. Downstream of the CLIPC9 comes CLIPBs, with CLIPB4 and CLIPB17 seeming to be the farthest upstream; the cascade then bifurcates into two different branches: one converging on CLIPB8 and the other on CLIPB10. Dashed arrows indicate that the steps require further characterization (right panel). Created in BioRender. Saab, S. (2025) https://BioRender.com/r16n889.

https://doi.org/10.1371/journal.ppat.1012965.g002

Although TEP1 expression was first attributed to hemocytes, recent studies have suggested that other organs, such as the fat body, are also involved in this process [127,128]. After the full-length TEP1-F is secreted, a portion of it is cleaved into TEP1cut to become activated [129]. TEP1cut is then stabilized in the hemolymph by a disulfide-linked heterodimer made up of two leucine-rich repeat proteins (LRRs), leucine-rich repeat immune protein 1 (LRIM1) and Anopheles Plasmodium-responsive leucine-rich repeat 1 (APL1C) (Fig 2). Interestingly, a knockdown of LRIM1/APL1C not only results in a higher number of oocysts in the midgut but also in a nonspecific deposition of TEP1 cut on mosquito tissues, suggesting the participation of LRIM1/APL1C in avoiding an autoimmune response [130–133]. APL1 is encoded by a family of three genes, APL1A, APL1B, and APL1C. APL1A is induced by the Imd pathway and is P. falciparum-specific; in contrast, APL1C is induced by the Toll pathway and is P. berghei-specific [134,135]. APL1C acts against both extracellular ookinetes and circulating sporozoites of rodent Plasmodium, but not human [136].

TEP1 function is further promoted by the non-catalytic CLIP serine protease (cSPH) known as SPCLIP1, which mediates the accumulation of TEP1cut on the microbial surface, thereby triggering lysis or melanization [137]. Another cSPH, CLIPA2, has also been shown to negatively regulate TEP1 activity by preventing its cleavage into TEP1cut [138]. It is well established that the complement-like system plays a crucial role in parasite elimination; however, the process by which the TEP1/LRIM/APL1C complex induces pathogen lysis has not been fully dissected and warrants further investigation.

In addition to TEP1-mediated lysis, the melanization reaction observed in certain refractory genetic backgrounds makes An. gambiae mosquitoes highly resistant to Plasmodium ookinetes [139–141]. The deposition of melanin on the surface of a pathogen depends on a series of biochemical reactions that require active phenoloxidases (POs), which are released into the hemolymph as prophenoloxidase (PPO) zymogens. PPO cleavage is regulated by a cascade of CLIP domain serine proteases, which represent a large family of five subgroups (A to E) in An. gambiae mosquitoes [142,143]. Previous genetic studies have identified several cSPHs of subfamily A as playing a major nonredundant role in P. berghei melanization. For instance, SPCLIP1 (CLIPA30), CLIPA8, and CLIPA28 form a core cSPH module that positively regulates P. berghei ookinete melanization [137,144,145]. Such cSPHs are activated in an ordered manner downstream of TEP1, with SPCLIP1 being the farthest upstream, followed by CLIPA8 and then CLIPA28 [145]. Two CLIP-domain serine proteases, CLIPA2 and CLIPA14, have been shown to act as negative regulators of the melanization response [138,146]. In addition to cSPHs, several catalytic CLIP serine proteases of subfamilies B and C have also been identified as playing an essential role in the melanization response in An. gambiae mosquitoes. Individually silencing CLIPB4, CLIPB8, CLIPB10, CLIPB14, CLIPB17, or CLIPC9 partially reverses P. berghei melanization in refractory An. gambiae mosquitoes [144,147–149]. Recent in vivo studies have shown redundancy in the antimicrobial function of CLIPBs, which can be partially explained by a bifurcation of the cascade downstream of CLIPB4 and CLIPB17 into two branches, one converging on CLIPB8 and the other on CLIPB10 (Fig 2) [150]. Based on their capacity to cleave Manduca sexta PPOs in vitro, CLIPB4, CLIPB9, and CLIPB10 have been classified as prophenoloxidase-activating proteins (PAPs), which are inhibited by serine protease inhibitor 2 (serpin 2 or SRPN2) [148,151,152]. The core cSPH module acts upstream of the catalytic proteases, despite the cleavage of CLIPA8 by several recombinant CLIPBs in vitro [150], highlighting the complexity of this network.

Conversely, the C-type lectin 4 (CTL4) and MA2 (CTL2) play regulatory roles in protecting the parasite from melanization. For instance, silencing CTL4 in An. gambiae triggers TEP1-mediated melanization of P. berghei ookinetes [140,153,154], and CRISPR/Cas9-based CTL4 knockout An. gambiae mosquitoes show an enhanced ability to melanize P. falciparum ookinetes in a TEP1-independent manner [141]. Injecting recombinant CLT4/CTLMA2 into CTL4-silenced An. gambiae mosquitoes reversed the increase in the PO activity, suggesting a common role for the CLT4/CTLMA2 heterodimer in the melanization response [155]. The mechanisms governing CTL4-mediated protection of the ookinete from melanization and how cSPHs intervene to regulate PPO cleavage are still unclear and require further study.

A recent single-cell transcriptomic study in An. gambiae has demonstrated that the CLIP-domain serine proteases, CTL4/CTLMA2 and LRIM1, which are involved in the regulation of melanization, are highly expressed in the fat body, but PPOs are exclusively produced by hemocytes [156], suggesting that the mechanisms limiting Plasmodium infection also involve a systemic immune response.

Apart from the humoral immune responses, hemocytes are also critical for killing malaria parasites such as P. berghei that infect rodents. Hemocyte depletion in An. gambiae drastically increases the number of P. berghei parasites in the mosquito [127]. In addition to TEP1, hemocytes can also express other factors that either facilitate or limit Plasmodium development [156–160]. For example, a functional assay performed by Lombardo et al. [160] has shown that An. gambiae hemocytes express genes that function as P. berghei agonists, including a putative LRR protein, lipopolysaccharide-induced tumor necrosis factor alpha factor-like 6 (LL6), laminin A homolog, peptidase and trypsin-like domain-containing transmembrane protein, and a vesicular-type ATPase. They also express P. berghei antagonists, including von Willebrand factor-type A domain protein, collagen type IV protein, hexamerin 2 beta homolog, fibrinogen-related FBN8, and a CLIPB [161]. Smith and colleagues have reported similar findings in their proteomic analysis of An. gambiae, which revealed that phagocytic hemocytes play a dual role in P. falciparum infections by expressing anti-plasmodial immune effectors such as TEP1, defensin1, HPX2, and two components of the Ras family, the small GTPases Ras-related and Ras-homolog family member A, as well as Plasmodium protective factors such as lysozyme c-1 (LYSC1), scavenger-receptors SCRASP1 and SCRBQ2, LRIM15, and pretaporter (Prtp) [162]. Interestingly, several members of the fibrinogen-related proteins (FREP or FBN) that are mainly abundant in the hemolymph, including FBN8, FBN9, FBN30, FBN39, act as Plasmodium antagonists [161,163–165].

4. Plasmodium development in the hemolymph and hemolymph-triggered immune responses

4.1. Oocyst development and sporogony

After reaching the extracellular space separating the midgut epithelial cells from the basal lamina, the ookinete rounds up and develops into an oocyst, which in turn undergoes extensive mitotic divisions known as sporogony [166]. The mechanism triggering the developmental progression from ookinete to oocyst is still unclear; however, several mosquito and Plasmodium genes have been shown to be involved (Fig 1). Recently, three novel ookinete-expressed genes, PIMMS01, PIMMS57, and PIMMS22, were found to be critically involved in the development of the oocyst from the ookinete stage [167]. Moreover, the oocyst capsule also includes several mosquito-derived proteins, including laminin, matrix metalloprotease 1 (MMP1), and lysozyme c-1 (LYSC1), that assist the oocyst in hiding from the mosquito’s immune responses, thus favoring oocyst development [168]. Other parasite-derived proteins in the oocyst capsule, including the P. berghei oocyst capsule protein 380 (PbCap380), oocyst capsule-associated protein 93 (PbCap93), and the ookinete surface and oocyst capsule protein (OSCP), are essential for oocyst development and maintenance, and knocking out the genes encoding such proteins results in a reduced oocyst number [169–171].

Interestingly, a proteomic study has revealed unique, temporally regulated signatures in early, mid, and late oocysts in the rodent parasite [172]. For instance, expression of CSP as an oocyst capsule component after the onset of sporogony in rodent parasites is necessary for its escape from the melanization immune response in An. stephensi mosquitoes [173]. Sporogony eventually produces oocysts containing thousands of sporozoites by 14 days after the intake of a blood meal [168]. Recently, adipokinetic hormone signaling has been shown to be involved in facilitating P. falciparum sporogony [174]. Live fluorescence imaging of oocysts undergoing sporogony has revealed that sporozoite release is preceded by capsule thinning and small opening formation, thereby facilitating the sporozoites’ egress [175]. Although several proteins, including parasite-derived CSP and oocyst rupture protein (ORP) 1 and 2 [176], have been reported to be required for the oocyst rupture, the molecular mechanism driving this capsule thinning and the consequent sporozoite excystation remain poorly understood. Residence in the mosquito vector in limited numbers for about 2 weeks, along with their immobility, render the oocyst stage an appropriate transgenic target for blocking parasite transmission, but little is yet known about anti-oocyst effectors.

4.2. Immune responses against oocysts

Since they reside between the midgut epithelial cells and the basal lamina for about 2 weeks, oocysts become extensively exposed to the immune factors that are involved in ookinete killing. However, oocysts seem to be the most resilient parasite stage in the mosquito [168,176]. Recently, oocysts of P. yoelii were shown to evade An. stephensi immune responses through the circumsporozoite protein (CSP). CSP disruption makes the parasite visible to the immune system through midgut nitration and hemocyte recruitment, with subsequent activation of the TOLL signaling pathway and oocyst TEP1-dependent melanization (Fig 3) [173]. Interestingly, in An. gambiae, PPO2, PPO3, and PPO9 limit the survival of P. berghei oocysts without triggering melanization, hinting at alternative functions that are not related to the classical roles of POs [127]. Recently, the oocyst stage has been shown to also be susceptible to the melanization immune response, in which co-silencing CLIPA2 and CLIPA14, the negative regulators of melanization, triggers the melanization of both P. berghei and P. falciparum oocysts [177]. Despite the fact that only late-stage P. berghei oocysts were melanized, this was not the case for P. falciparum, in which both early and late oocysts were found to be melanized [177].

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Fig 3. Mosquito late-phase immune responses against the oocyst stage.

The main cellular immune response known to be triggered against the oocyst stage involves hemocyte differentiation driven by the action of the LITAF-Like 3 and STAT pathway. Hemocytes are the unique suppliers of PPOs, and PPO2, PPO3, and PPO9 limit oocyst survival without triggering melanin deposition (left panel). However, the oocyst stage can evade most of the hemolymph-mediated immune responses by exploiting its capsule components, such as laminin, collagen, LYSC1, MMP1, and CSP (right panel). Created in BioRender. Saab, S. (2025) https://BioRender.com/q60q481.

https://doi.org/10.1371/journal.ppat.1012965.g003

Other studies have indicated that the STAT pathway and LITAF-like 3 limit early P. berghei and P. falciparum oocyst development through processes involving midgut and carcass nitration along with hemocyte differentiation (Fig 3) [116,178]. Interestingly, a unique stem-cell mediated response downstream of the JAK-STAT pathway has been shown to be responsible for eliminating oocysts [179]. Taken together, these studies suggest the involvement of systemic immune responses in limiting oocyst development through hemocyte participation, not only as effectors but also as signaling mediators. Nevertheless, further research is required to understand the communication mechanisms that exist between hemocytes and other mosquito organs.

During their development in the abluminal portion of the gut, oocysts make use of their extracellular cell wall (capsule) to evade the mosquito’s immune system [175]. The oocysts consume mosquito resources when undergoing sporogony, which involves a drastic increase in the size of the oocysts [180] along with the enlargement of the midgut basal lamina that envelops the oocysts. Interestingly, it has been reported that multiple blood feedings enhance oocyst growth and shorten the period of oocyst maturation [180–182]. Nevertheless, a recent study has shown that subsequent blood feedings reduce the oocyst number of rodent, but not human, Plasmodium parasites. This effect occurs because of the rupture of the basal lamina following blood meal-induced epithelium distention, which promotes direct exposure of the P. berghei oocysts to the mosquito’s complement-like system, an immune response that P. falciparum can evade [181].

Most of the previously mentioned mosquito-derived components of the oocyst capsule that favor oocyst maturation, such as laminin, collagen, LYSC1, and MMP1 [182–186], are highly expressed in hemocytes and can negatively regulate the complement-like system and melanization (Fig 3). Overall, these effects suggest that under certain conditions, hemocytes may be involved in oocyst protection. In addition, mosquito lipophorins, which are ingested by oocysts [187], can also enhance the survival of P. berghei oocysts [188].

In recent decades, several studies have revealed a plethora of mechanisms underlying the hemocyte-mediated immune response, with a main focus on their anti-plasmodial functions; however, their agonistic functions have received less attention. Furthermore, most of these studies have been performed using the rodent malaria parasite P. berghei, but it is well established that the mosquito’s immune system responds differently to P. berghei and the human malaria parasite P. falciparum [122,189,190]. To gain a better understanding of the role of hemocytes in Plasmodium infection, future studies should consider the compatibility of the experimental models (i.e., the Plasmodium and mosquito species used). Furthermore, to develop effective strategies for combating malaria by using genetically modified mosquitoes (GMOs), further studies are needed to focus on abolishing the mechanisms governing Plasmodium’s evasion of immunity.

4.3. Sporozoite invasion of the salivary gland

After their release into the hemolymph, the circulating sporozoites passively reach the basal lamina of the salivary glands (Fig 1) [191]. Salivary glands consist of three lobes attached to a common duct and are surrounded by epithelial cells arranged in a single layer [192]. Infecting this tissue provides the parasite access to the host, but it also represents a physical barrier to transmission [193]. Prior to the entry of the sporozoites into the salivary gland cavity, recognition and epithelial cell-crossing processes occur. These processes involve specific Anopheles receptors and other factors responsible for the initial attachment and invasion, as well as parasite proteins, including the surface proteins CSP, thrombospondin-related anonymous protein (TRAP), and membrane antigen/erythrocyte binding-like protein (MAEBL) [194].

CSP-mediated recognition involves salivary gland glycans and proteins such as CSP-binding protein (CSPBP). TRAP interacts with mosquito Saglin and salivary gland surface protein 1 (SGS1) and mediates directional migration [176]. Recent evidence shows that the P. berghei sporozoite protein, claudin-like apicomplexan microneme protein (CLAMP), is an essential participant in salivary gland traversal and sporozoite gliding motility, with the underlying mechanism involving the shedding of TRAP [195]. Also, the mosquito-encoded epithelial serine protease (ESP), expressed on the basal side of the epithelial cells, is essential for invasion of the sporozoite salivary gland [196]. Furthermore, RNAi-mediated silencing of a salivary gland-transcribed transmembrane glucose transporter (AGAP007752) results in significantly decreased sporozoite numbers, presumably because of its putative function as a sporozoite receptor [197]. Interestingly, a small peptide known as salivary gland and midgut binding peptide 1 (SM1) has been identified via a phage display peptide library and found to be distinctive in its ability to bind to epithelial receptors in the mosquito midgut and salivary gland, thereby interfering with parasite transmission [198].

4.4. Mosquito immune responses against sporozoites

Sporozoites represent the parasite stage that experiences the greatest exposure to the hemolymph, since these final-stage parasites migrate to the heart and make use of the flow of the hemolymph as a transport medium to reach the salivary glands [199]. During this journey, they are entirely exposed to both humoral and cellular immune factors. As a result, in models involving An. dirus-P. vivax and Ae. aegypti-P. gallinaceum, only about 20% of the sporozoites produced in the oocysts actually reach the salivary glands, and the rest disappear in the hemolymph ([199,200]. Nevertheless, hemocyte-mediated immune responses, including phagocytosis, nodulation, encapsulation, and melanization in An. gambiae, An. albimanus, and Ae. aegypti only partially contribute to the elimination, respectively, of P. berghei, P. vivax, and P. gallinaceum sporozoites; instead, lysis is suggested as the major mechanism of their killing (Fig 4) [199,201–203].

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Fig 4. Mosquito immune responses against the sporozoite stage.

Following the rupture of the mature oocysts, the released sporozoites become directly exposed to all the cellular and humoral immune responses in the hemolymph. Apart from hemocyte-mediated phagocytosis, sporozoites can also be subjected to fat body-secreted AMPs that can play an essential role in sporozoite lysis (right panel). After infection, the high hemolymph flow drives the hemocyte accumulation around the periostial region, promoting direct contact between circulating pathogens and hemocytes, and therefore phagocytosis. Also, AMPs are produced in pericardial cells and may be involved in the lysis of sporozoites that accumulate in the heart (right panel). AMPs, lysozyme-related genes, C-type lectins, serine proteases, leucine-rich immune proteins, and TEP1 are all expressed in the mosquito salivary glands; however, their effect on the sporozoite stage needs to be further investigated (left panel). Created in BioRender. Saab, S. (2025) https://BioRender.com/c22y701.

https://doi.org/10.1371/journal.ppat.1012965.g004

The primary function of the mosquito heart is to pump the hemolymph around the mosquito’s body [204]. However, after infection, hemocytes accumulate around periostial regions, which are sites of high hemolymph flow, facilitating contact between circulating pathogens and hemocytes, and therefore phagocytosis (Fig 4) [201,205]. Interestingly, P. berghei sporozoites also accumulate in the periostial regions, where they undergo fragmentation [201]. Although hemocytes have been suggested to be the primary cell type involved in these processes, other cell types located in periostial regions, such as pericardial cells, could also play a significant role in parasite killing. Immune factors such as lysozyme and cecropin, produced in pericardial cells, could be involved in the lysis of pathogens accumulated in the heart [206–208]. In addition, other factors such as NO, ROS, and lysozymes are produced by pericardial cells and hemocytes, and they are implicated in establishing interactions between these cells [206,208–210]. However, whether these cells are involved directly or whether they cooperate with hemocytes in the elimination of sporozoites, as well as the mechanism(s) involved in this interplay, are largely unknown and warrant further study.

To date, only a few studies have highlighted the mosquito’s immune responses against sporozoites within the salivary glands. In An. gambiae and An. coluzzii, RT-PCR studies as well as serial analysis of gene expression (SAGE) and RNA-seq studies have revealed the induction of several immune-related genes by P. berghei, including AMPs, TEP1, and CTLMA2 [197,211,212]. A comparative transcriptomic analysis between P. vivax-infected and uninfected An. dirus salivary glands has demonstrated an elevated immune response in the infected glands, with up-regulation of several known AMPs as well as leucine-rich immune proteins and C-type lectins, among others (Fig 4) [213]. Moreover, several lysozyme-related genes, lectins, secreted serine proteases, and other uncharacterized secreted proteins have been identified in Anopheles salivary glands [214]. Remarkably, the transcriptional response at the organ level (salivary glands) after bacterial immune challenge is different from that induced systemically [215]. The studies just mentioned clearly indicate that Anopheles salivary glands not only serve as a physical barrier that has to be bypassed by Plasmodium and other microorganisms but also exert local immune activity that is induced after invasion/challenge. The relative contribution of the local salivary gland immune responses against sporozoites, where they remain up until the infectious bite occurs, has yet to be unveiled.

5. Sporozoite transmission to the host

As mosquitoes probe for blood, the infectious mature sporozoites exit the mosquito vector, accompanied by a mosquito saliva protein cocktail co-injected into the host [216]. Apart from facilitating the blood meal through their immunomodulatory, coagulation, anti-inflammatory, anti-hemostatic, and vasodilatory functions, a number of these sialome factors also affect the sporozoites’ transmission and infectivity in the vertebrate recipients [217,218]. Remarkably, the number of sporozoites injected into the mammalian host is by far smaller than the number of parasites found in the salivary glands [193], yet it correlates with the initial load [219]. Sporozoite infectivity in the host is initially locally affected either directly by mosquito factors that bind to the parasite [220] or indirectly by the initial dermal immune reaction that is prompted in response to the injected parasite, the mosquito proteins, and the mechanical damage caused by the bite [221]. Mosquito GILT, named after its similarity to human gamma interferon-inducible thiol reductase, binds to sporozoites and decreases their motility in mice [220], while sporozoite-associated mosquito saliva protein 1 (SAMSP1) positively affects sporozoite entry into host cells, thereby facilitating sporozoite infectivity [222]. Although the effectiveness of the host’s immune response against mosquito factors in influencing the subsequent parasite infectivity is controversial [223,224], accumulating recent evidence has demonstrated that some of these factors have immunomodulatory activity, with the potential to be harnessed as vaccine targets [216,225,226]. In this context, experiments on mice involving immunization with saliva-derived antiserum have resulted in a reduction in host-parasite infection as a result of decreased vascular permeability, mostly attributed to the secreted salivary gland protein AgTRIO [223]; in contrast, the sporozoite-associated factor (SAP) seems to affect sporozoite infectivity by modulating the host’s inflammatory responses [225]. Also, the saliva microRNA (miRNA) repertoire of hematophagous insects has been proposed to contribute to antiparasitic host responses [227]. Interestingly, the An. coluzzii miRNA patterns are distinct for the saliva and the salivary glands, with some of them being identical to human miRNAs with immune-regulatory roles [227,228]. However, as far as mosquitoes are concerned, the effect of these non-immunogenic factors during rapid blood-meal acquisition remains elusive and favors hypotheses regarding host immune regulation with long-term evolutionary advantages to the host [227]. Finally, the presence of the microbiota in salivary glands and saliva, and the transfer of specific bacteria during blood meals into mammalian hosts highlight additional saliva factors that could potentially influence Plasmodium transmission [229].

6. A possible role for salivary gland proteins in host-to-vector transmission

The mosquito salivary proteins blended into the blood meal have recently been shown to play a crucial role in facilitating parasitic infection of the mosquito gut. For example, the salivary mosquito protein known as Saglin was shown to mediate Plasmodium infection in the mosquito gut; however, the mode of action of this protein is still not known [230]. In addition to Saglin, the Anopheles salivary apyrase has been shown to facilitate fibrinolysis, thus promoting Plasmodium parasite infection through the activation of the tissue plasminogen activator in the blood bolus [231,232]. These studies highlight the potential of such mosquito salivary proteins to be harnessed as malaria transmission-blocking targets, either through knocking out such factors or exploiting salivary gland promoters to drive the expression of antiparasitic effectors. For instance, the use of transgenic Anopheles mosquitoes capable of targeting the fibrinolytic system by expressing human plasminogen activator inhibitor 1 (PAI-1) has been successful in reducing Plasmodium transmission to mosquitoes [233].

7. Mosquito immune memory and adaptive ability

Despite their lack of adaptive immunity, mosquitoes possess an innate immune memory that protects them against a repeated exposure to the same pathogen during a second encounter, a phenomenon known as immune priming [234]. Mosquitoes that were previously infected with P. berghei or P. falciparum have shown a reduction in the number of oocysts during the second exposure to the same parasite [235–238]. The mechanisms of immune memory in mosquitoes are not clear; some mechanisms have been proposed that involve hemocytes and midgut epithelial cell immune responses. One of these mechanisms posits direct contact of the microbiome with the epithelial cells when the ookinetes invade the midgut epithelium, triggering a systemic immune response that involves hemocyte differentiation [237,238]. This response produces an increase in the granulocyte subpopulation, changes in hemocyte morphology, and an overexpression of anti-plasmodial genes, including TEP1 and LRIM1. Thus, the mosquito triggers an enhanced immune response against the parasite during the second encounter [238]. Furthermore, midguts from mosquitoes infected with Plasmodium have shown high rates of DNA synthesis [235,236], and the inhibition of this process results in the abolition of protection during the second encounter with the parasite [236]. It has been suggested that some immune genes could be expanded as a result of endoreplication to increase their expression during a second encounter [236].

Although mosquitoes’ repertoire of pattern recognition receptors (PRRs) is limited, they provide a broad recognition for most pathogens. In addition to classic PRRs such as lectins and others, insects encode an immunoglobulin superfamily member called Down syndrome cell adhesion molecule (DSCAM) [234]. Through alternative splicing, DSCAM can produce more than 30,000 isoforms [239]. In mosquitoes, the splicing of some DSCAM isoforms is favored, depending on the pathogen, and this repertoire correlates with the affinity and activity against each pathogen [240]. Some DSCAM isoforms can discriminate between Plasmodium species and provide species-specific protection; however, it is not clear whether they interact with other effector molecules to kill the parasite [240,241]. Furthermore, it is unclear whether the isoform repertoire is maintained for a sufficient time after the immune challenges to provide immune memory. Innate immune memory in insects is probably the consequence of many mechanisms working together, which need to be deeply explored.

8. Conclusion

A great body of literature pinpoints the critical processes that facilitate Plasmodium invasion of the mosquito vector, yet many questions remain open and require deeper exploration to allow us to understand more thoroughly how various immune responses are coordinated to eliminate the invading parasites. However, the capacity of Plasmodium to overcome the well-orchestrated immune responses by the mosquito necessitates a focus of current research on developing transmission-blocking strategies for reducing mosquito vector competence. Recent efforts have been directed at generating genetically modified mosquitoes that are resistant to Plasmodium [79,112,141,242]. Other recent advances in dissecting the parasite’s complex life cycle in the mosquito vector have helped to identify potential TBV targets. However, to eradicate malaria, such strategies must be combined with an effective malaria vaccine and preventive drugs.

Acknowledgments

We thank Dr Deborah McClellan for editorial assistance.

References

  1. 1. Beerntsen BT, James AA, Christensen BM. Genetics of mosquito vector competence. Microbiol Mol Biol Rev. 2000;64(1):115–37. pmid:10704476
  2. 2. World Malaria Report 2023. World Health Organization. 2023.
    • 3. Rénia L, Goh YS. Malaria parasites: the great escape. Front Immunol. 2016;7:463. pmid:27872623
    • 4. CDC. Anopheles gambiae. Malaria. 2020 Available from: https://www.cdc.gov/malaria/about/biology/#tabs-1-5
      • 5. White GB. Anopheles gambiae complex and disease transmission in Africa. Trans R Soc Trop Med Hyg. 1974;68(4):278–301.
      • 6. Fanello C, Santolamazza F, della Torre A. Simultaneous identification of species and molecular forms of the Anopheles gambiae complex by PCR-RFLP. Med Vet Entomol. 2002;16(4):461–4. pmid:12510902
      • 7. Coetzee M, Hunt RH, Wilkerson R, Torre AD, Coulibaly MB, Besansky NJ. Anopheles coluzzii and Anopheles amharicus, new members of the Anopheles gambiae complex. Zootaxa. 2013;3619(3):.
      • 8. Gosling R, von Seidlein L. The future of the RTS,S/AS01 malaria vaccine: an alternative development plan. PLoS Med. 2016;13(4):e1001994.
      • 9. El-Moamly AA, El-Sweify MA. Malaria vaccines: the 60-year journey of hope and final success-lessons learned and future prospects. Trop Med Health. 2023;51(1):29. pmid:37198702
      • 10. Jensen M, Mehlhorn H. Seventy-five years of Resochin in the fight against malaria. Parasitol Res. 2009;105(3):609–27.
      • 11. Suresh N, Haldar K. Mechanisms of artemisinin resistance in Plasmodium falciparum malaria. Curr Opin Pharmacol. 2018;42:46–54.
      • 12. Hanboonkunupakarn B, Tarning J, Pukrittayakamee S, Chotivanich K. Artemisinin resistance and malaria elimination: where are we now?. Front Pharmacol. 2022;13:876282. pmid:36210819
      • 13. Dong S, Dong Y, Simões ML, Dimopoulos G. Mosquito transgenesis for malaria control. Trends Parasitol. 2022;38(1):54–66. pmid:34483052
      • 14. Billker O, Lindo V, Panico M, Etienne AE, Paxton T, Dell A, et al. Identification of xanthurenic acid as the putative inducer of malaria development in the mosquito. Nature. 1998;392(6673):289–92. pmid:9521324
      • 15. Billker O, Shaw MK, Margos G, Sinden RE. The roles of temperature, pH and mosquito factors as triggers of male and female gametogenesis of Plasmodium berghei in vitro. Parasitology. 1997;115 (Pt 1):1–7. pmid:9280891
      • 16. Garcia GE, Wirtz RA, Barr JR, Woolfitt A, Rosenberg R. Xanthurenic acid induces gametogenesis in Plasmodium, the malaria parasite. J Biol Chem. 1998;273(20):12003–5. pmid:9575140
      • 17. Andreadaki M, Hanssen E, Deligianni E, Claudet C, Wengelnik K, Mollard V. Sequential membrane rupture and vesiculation during Plasmodium berghei gametocyte egress from the red blood cell. Sci Rep. 2018;8(1):3543.
      • 18. Yahiya S, Jordan S, Smith HX, Gaboriau DCA, Famodimu MT, Dahalan FA, et al. Live-cell fluorescence imaging of microgametogenesis in the human malaria parasite Plasmodium falciparum. PLoS Pathog. 2022;18(2):e1010276. pmid:35130301
      • 19. Tateishi YS, Araki T, Kawai S, Koide S, Umeki Y, Imai T, et al. Histone H3.3 variant plays a critical role on zygote-to-oocyst development in malaria parasites. Parasitol Int. 2024;100:102856. pmid:38199522
      • 20. Janse CJ, Van der Klooster PF, Van der Kaay HJ, Van der Ploeg M, Overdulve JP. Rapid repeated DNA replication during microgametogenesis and DNA synthesis in young zygotes of Plasmodium berghei. Trans R Soc Trop Med Hyg. 1986;80(1):154–7. pmid:3088783
      • 21. Sinden RE. Gametocytogenesis of Plasmodium falciparum in vitro: an electron microscopic study. Parasitology. 1982;84(1):1–11. pmid:7038594
      • 22. Muhia DK, Swales CA, Deng W, Kelly JM, Baker DA. The gametocyte-activating factor xanthurenic acid stimulates an increase in membrane-associated guanylyl cyclase activity in the human malaria parasite Plasmodium falciparum. Molecular Microbiology. 2001;42(2):553–60.
      • 23. Carucci DJ, Witney AA, Muhia DK, Warhurst DC, Schaap P, Meima M, et al. Guanylyl cyclase activity associated with putative bifunctional integral membrane proteins in Plasmodium falciparum. J Biol Chem. 2000;275(29):22147–56. pmid:10747978
      • 24. Taylor CJ, McRobert L, Baker DA. Disruption of a Plasmodium falciparum cyclic nucleotide phosphodiesterase gene causes aberrant gametogenesis. Molecular Microbiology. 2008;69(1):110–8.
      • 25. Jiang Y, Wei J, Cui H, Liu C, Zhi Y, Jiang Z. An intracellular membrane protein GEP1 regulates xanthurenic acid induced gametogenesis of malaria parasites. Nature Communications. 2020;11(1):1764.
      • 26. Wang P-P, Jiang X, Zhu L, Zhou D, Hong M, He L, et al. A G-protein-coupled receptor modulates gametogenesis via PKG-Mediated Signaling Cascade in Plasmodium berghei. Microbiol Spectr. 2022;10(2):e0015022. pmid:35404079
      • 27. McRobert L, Taylor CJ, Deng W, Fivelman QL, Cummings RM, Polley SD, et al. Gametogenesis in malaria parasites is mediated by the cGMP-dependent protein kinase. PLoS Biol. 2008;6(6):e139. pmid:18532880
      • 28. Brochet M, Collins MO, Smith TK, Thompson E, Sebastian S, Volkmann K, et al. Phosphoinositide metabolism links cGMP-dependent protein kinase G to essential Ca²⁺ signals at key decision points in the life cycle of malaria parasites. PLoS Biol. 2014;12(3):e1001806. pmid:24594931
      • 29. Balestra AC, Koussis K, Klages N, Howell SA, Flynn HR, Bantscheff M, et al. Ca(2+) signals critical for egress and gametogenesis in malaria parasites depend on a multipass membrane protein that interacts with PKG. Science Advances. 2021;7(13):.
      • 30. Bansal A, Molina-Cruz A, Brzostowski J, Liu P, Luo Y, Gunalan K, et al. PfCDPK1 is critical for malaria parasite gametogenesis and mosquito infection. Proc Natl Acad Sci U S A. 2018;115(4):774–9.
      • 31. Bansal A, Molina-Cruz A, Brzostowski J, Mu J, Miller LH. Plasmodium falciparum calcium-dependent protein kinase 2 is critical for male gametocyte exflagellation but not essential for asexual proliferation. mBio. 2017;8(5).
      • 32. Kumar S, Haile MT, Hoopmann MR, Tran LT, Michaels SA, Morrone SR, et al. Plasmodium falciparum calcium-dependent protein kinase 4 is critical for male gametogenesis and transmission to the mosquito vector. mBio. 2021;12(6):e0257521. pmid:34724830
      • 33. Hitz E, Balestra A, Brochet M, Voss T. PfMAP-2 is essential for male gametogenesis in the malaria parasite Plasmodium falciparum. Sci Rep. 2020;10(1):11930.
      • 34. Kumar S, Gargaro OR, Kappe SHI. Plasmodium falciparum CRK5 Is Critical for Male Gametogenesis and Infection of the Mosquito. mBio. 2022;13(5):e0222722. pmid:36154191
      • 35. Sinden RE, Croll NA. Cytology and kinetics of microgametogenesis and fertilization in Plasmodium yoelii nigeriensis. Parasitology. 1975;70(1):53–65. pmid:1118188
      • 36. Eksi S, Czesny B, van Gemert G-J, Sauerwein RW, Eling W, Williamson KC. Malaria transmission-blocking antigen, Pfs230, mediates human red blood cell binding to exflagellating male parasites and oocyst production. Mol Microbiol. 2006;61(4):991–8. pmid:16879650
      • 37. van Dijk MR, Janse CJ, Thompson J, Waters AP, Braks JA, Dodemont HJ, et al. A central role for P48/45 in malaria parasite male gamete fertility. Cell. 2001;104(1):153–64. pmid:11163248
      • 38. van Dijk MR, van Schaijk BCL, Khan SM, van Dooren MW, Ramesar J, Kaczanowski S, et al. Three members of the 6-cys protein family of Plasmodium play a role in gamete fertility. PLoS Pathog. 2010;6(4):e1000853. pmid:20386715
      • 39. van Schaijk BCL, van Dijk MR, van de Vegte-Bolmer M, van Gemert G-J, van Dooren MW, Eksi S, et al. Pfs47, paralog of the male fertility factor Pfs48/45, is a female specific surface protein in Plasmodium falciparum. Mol Biochem Parasitol. 2006;149(2):216–22. pmid:16824624
      • 40. Liu Y, Tewari R, Ning J, Blagborough AM, Garbom S, Pei J, et al. The conserved plant sterility gene HAP2 functions after attachment of fusogenic membranes in Chlamydomonas and Plasmodium gametes. Genes Dev. 2008;22(8):1051–68. pmid:18367645
      • 41. Kumar S, Valansi C, Haile MT, Li X, Flyak K, Dwivedy A, et al. Malaria parasites utilize two essential plasma membrane fusogens for gamete fertilization. Cell Mol Life Sci. 2022;79(11):549. pmid:36241929
      • 42. Laurentino EC, Taylor S, Mair GR, Lasonder E, Bartfai R, Stunnenberg HG, et al. Experimentally controlled downregulation of the histone chaperone FACT in Plasmodium berghei reveals that it is critical to male gamete fertility. Cell Microbiol. 2011;13(12):1956–74. pmid:21899698
      • 43. Liu F, Liu Q, Yu C, Zhao Y, Wu Y, Min H, et al. An MFS-domain protein Pb115 plays a critical role in gamete fertilization of the malaria parasite Plasmodium berghei. Front Microbiol. 2019;10:2193. pmid:31616399
      • 44. Pang W, Bai J, Zhu L, Liu F, Wu Y, Yang F, et al. Functional characterization of a conserved membrane protein, Pbs54, involved in gamete fertilization in Plasmodium berghei. Int J Parasitol. 2024;54(2):99–107. pmid:37774810
      • 45. Tachibana M, Miura K, Takashima E, Morita M, Nagaoka H, Zhou L, et al. Identification of domains within Pfs230 that elicit transmission blocking antibody responses. Vaccine. 2019;37(13):1799–806. pmid:30824357
      • 46. Lee S-M, Wu C-K, Plieskatt JL, Miura K, Hickey JM, King CR. N-terminal Pfs230 domain produced in baculovirus as a biological active transmission-blocking vaccine candidate. Clin Vaccine Immunol. 2017;24(10):e00140-17. pmid:28747311
      • 47. Pourhashem Z, Nourani L, Sani J, Yousefi H, Pirahmadi S, Sabouri M. Evaluation of a new fusion antigen, cd loop and HAP2-GCS1 domain (cd-HAP) of Plasmodium falciparum Generative Cell Specific 1 antigen formulated with various adjuvants, as a transmission blocking vaccine. Malaria J. 2023;22(1).
      • 48. Angrisano F, Sala KA, Da DF, Liu Y, Pei J, Grishin NV, et al. Targeting the conserved fusion loop of HAP2 inhibits the transmission of Plasmodium berghei and falciparum. Cell Rep. 2017;21(10):2868–78. pmid:29212032
      • 49. Yenkoidiok-Douti L, Canepa G, Barletta A, Barillas-Mury C. In vivo characterization of Plasmodium berghei P47 (Pbs47) as a malaria transmission-blocking vaccine target. Front Microbiol. 2020;11:1496.
      • 50. Molina-Cruz A, Barillas-Mury C. Pfs47 as a Malaria Transmission-Blocking Vaccine Target. Am J Trop Med Hyg. 2022.
      • 51. Guttery DS, Zeeshan M, Ferguson DJP, Holder AA, Tewari R. Division and transmission: malaria parasite development in the mosquito. Annu Rev Microbiol. 2022;76:113–34. pmid:35609946
      • 52. Bannister LH, Sinden RE. New knowledge of parasite morphology. Br Med Bull. 1982;38(2):141–5. pmid:7052194
      • 53. Vlachou D, Zimmermann T, Cantera R, Janse CJ, Waters AP, Kafatos FC. Real-time, in vivo analysis of malaria ookinete locomotion and mosquito midgut invasion. Cell Microbiol. 2004;6(7):671–85. pmid:15186403
      • 54. Shao L, Devenport M, Jacobs-Lorena M. The peritrophic matrix of hematophagous insects. Arch Insect Biochem Physiol. 2001;47(2):119–25. pmid:11376458
      • 55. Siciliano G, Costa G, Suárez-Cortés P, Valleriani A, Alano P, Levashina EA. Critical steps of Plasmodium falciparum ookinete maturation. Front Microbiol. 2020;11269. pmid:32256458
      • 56. Mohammed M, Dziedziech A, Sekar V, Ernest M, Alves E Silva TL, Balan B, et al. Single-cell transcriptomics to define Plasmodium falciparum stage transition in the mosquito midgut. Microbiol Spectr. 2023;11(2):e0367122. pmid:36847501
      • 57. Ukegbu CV, Cho J-S, Christophides GK, Vlachou D. Transcriptional silencing and activation of paternal DNA during Plasmodium berghei zygotic development and transformation to oocyst. Cell Microbiol. 2015;17(8):1230–40. pmid:25728487
      • 58. Ouologuem DT, Dara A, Kone A, Ouattara A, Djimde AA. Plasmodium falciparum development from gametocyte to oocyst: insight from functional studies. Microorganisms. 2023;11(8):1966. pmid:37630530
      • 59. Yuda M, Kaneko I, Iwanaga S, Murata Y, Kato T. Female-specific gene regulation in malaria parasites by an AP2-family transcription factor. Mol Microbiol. 2020;113(1):40–51. pmid:31231888
      • 60. Modrzynska K, Pfander C, Chappell L, Yu L, Suarez C, Dundas K, et al. A knockout screen of ApiAP2 genes reveals networks of interacting transcriptional regulators controlling the Plasmodium life cycle. Cell Host Microbe. 2017;21(1):11–22. pmid:28081440
      • 61. Nishi T, Kaneko I, Iwanaga S, Yuda M. Identification of a novel AP2 transcription factor in zygotes with an essential role in Plasmodium ookinete development. PLoS Pathog. 2022;18(8):e1010510. pmid:35947628
      • 62. Lasonder E, Rijpma SR, van Schaijk BCL, Hoeijmakers WAM, Kensche PR, Gresnigt MS, et al. Integrated transcriptomic and proteomic analyses of P. falciparum gametocytes: molecular insight into sex-specific processes and translational repression. Nucleic Acids Res. 2016;44(13):6087–101. pmid:27298255
      • 63. Hirai M, Maeta A, Mori T, Mita T. Pb103 regulates zygote/ookinete development in Plasmodium berghei via double zinc finger domains. Pathogens. 2021;10(12).
      • 64. Wang X, Qian P, Cui H, Yao L, Yuan J. A protein palmitoylation cascade regulates microtubule cytoskeleton integrity in Plasmodium. EMBO J. 2020;39(13):e104168.
      • 65. Yadav P, Ayana R, Garg S, Jain R, Sah R, Joshi N, et al. Plasmodium palmitoylation machinery engineered in E. coli for high-throughput screening of palmitoyl acyl-transferase inhibitors. FEBS Open Bio. 2019;9(2):248-64.
      • 66. Dong Y, Manfredini F, Dimopoulos G. Implication of the mosquito midgut microbiota in the defense against malaria parasites. PLoS Pathog. 2009;5(5):e1000423. pmid:19424427
      • 67. Saab SA, Dohna HZ, Nilsson LKJ, Onorati P, Nakhleh J, Terenius O, et al. The environment and species affect gut bacteria composition in laboratory co-cultured Anopheles gambiae and Aedes albopictus mosquitoes. Sci Rep. 2020;10(1):3352. pmid:32099004
      • 68. Cirimotich CM, Dong Y, Clayton AM, Sandiford SL, Souza-Neto JA, Mulenga M, et al. Natural microbe-mediated refractoriness to Plasmodium infection in Anopheles gambiae. Science. 2011;332(6031):855–8. pmid:21566196
      • 69. Ramirez JL, Short SM, Bahia AC, Saraiva RG, Dong Y, Kang S, et al. Chromobacterium Csp_P reduces malaria and dengue infection in vector mosquitoes and has entomopathogenic and in vitro anti-pathogen activities. PLoS Pathog. 2014;10(10):e1004398. pmid:25340821
      • 70. Saraiva RG, Huitt-Roehl CR, Tripathi A, Cheng Y-Q, Bosch J, Townsend CA, et al. Chromobacterium spp. mediate their anti-Plasmodium activity through secretion of the histone deacetylase inhibitor romidepsin. Sci Rep. 2018;8(1):6176. pmid:29670144
      • 71. Bahia AC, Dong Y, Blumberg BJ, Mlambo G, Tripathi A, BenMarzouk-Hidalgo OJ, et al. Exploring Anopheles gut bacteria for Plasmodium blocking activity. Environ Microbiol. 2014;16(9):2980–94. pmid:24428613
      • 72. Gao H, Bai L, Jiang Y, Huang W, Wang L, Li S, et al. A natural symbiotic bacterium drives mosquito refractoriness to Plasmodium infection via secretion of an antimalarial lipase. Nat Microbiol. 2021;6(6):806–17. pmid:33958765
      • 73. Povelones M, Christophides G. The complement system of malaria vector mosquitoes. Adv Insect Physiol. 2016;51:223–42.
      • 74. Clayton AM, Dong Y, Dimopoulos G. The Anopheles innate immune system in the defense against malaria infection. J Innate Immun. 2014;6(2):169–81. pmid:23988482
      • 75. Garver LS, Dong Y, Dimopoulos G. Caspar controls resistance to Plasmodium falciparum in diverse anopheline species. PLoS Pathog. 2009;5(3):e1000335. pmid:19282971
      • 76. Garver LS, Bahia AC, Das S, Souza-Neto JA, Shiao J, Dong Y, et al. Anopheles Imd pathway factors and effectors in infection intensity-dependent anti-Plasmodium action. PLoS Pathog. 2012;8(6):e1002737. pmid:22685401
      • 77. Yassine H, Osta MA. Anopheles gambiae innate immunity. Cell Microbiol. 2010;12(1):1–9. pmid:19804484
      • 78. Luna C, Hoa NT, Lin H, Zhang L, Nguyen HLA, Kanzok SM, et al. Expression of immune responsive genes in cell lines from two different Anopheline species. Insect Mol Biol. 2006;15(6):721–9. pmid:17201765
      • 79. Dong Y, Das S, Cirimotich C, Souza-Neto JA, McLean KJ, Dimopoulos G. Engineered anopheles immunity to Plasmodium infection. PLoS Pathog. 2011;7(12):e1002458. pmid:22216006
      • 80. Claudio-Piedras F, Recio-Tótoro B, Condé R, Hernández-Tablas JM, Hurtado-Sil G, Lanz-Mendoza H. DNA methylation in Anopheles albimanus modulates the midgut immune response against Plasmodium berghei. Front Immunol. 2020;10:3025. pmid:31993053
      • 81. Dong S, Fu X, Dong Y, Simões ML, Zhu J, Dimopoulos G. Broad spectrum immunomodulatory effects of Anopheles gambiae microRNAs and their use for transgenic suppression of Plasmodium. PLoS Pathog. 2020;16(4):e1008453. pmid:32330198
      • 82. Kaur H, Pacheco M, Garber L, Escalante A, Vinetz J. Evolutionary insights into the microneme-secreted, chitinase-containing high-molecular-weight protein complexes involved in Plasmodium invasion of the mosquito midgut. Infect Immun. 2022;90(1):e0031421.
      • 83. Ghosh AK, Coppens I, Gardsvoll H, Ploug M, Jacobs-Lorena M. Plasmodium ookinetes coopt mammalian plasminogen to invade the mosquito midgut. Proc Natl Acad Sci U S A. 2011;108(41):17153–8.
      • 84. Tomas AM, Margos G, Dimopoulos G, van Lin LH, de Koning-Ward TF, Sinha R, et al. P25 and P28 proteins of the malaria ookinete surface have multiple and partially redundant functions. EMBO J. 2001;20(15):3975–83. pmid:11483501
      • 85. Molina-Cruz A, Canepa G, Barillas-Mury C. Plasmodium P47: a key gene for malaria transmission by mosquito vectors. Curr Opin Microbiol. 2017;40:168–74.
      • 86. Ukegbu CV, Giorgalli M, Tapanelli S, Rona LDP, Jaye A, Wyer C, et al. PIMMS43 is required for malaria parasite immune evasion and sporogonic development in the mosquito vector. Proc Natl Acad Sci U S A. 2020;117(13):7363–73. pmid:32165544
      • 87. Ukegbu CV, Akinosoglou KA, Christophides GK, Vlachou D. Plasmodium berghei PIMMS2 promotes ookinete invasion of the Anopheles gambiae mosquito midgut. Infect Immun. 2017;85(8):e00139-17. pmid:28559405
      • 88. Shahabuddin M, Toyoshima T, Aikawa M, Kaslow DC. Transmission-blocking activity of a chitinase inhibitor and activation of malarial parasite chitinase by mosquito protease. Proc Natl Acad Sci U S A. 1993;90(9):4266–70. pmid:8483942
      • 89. Tsai YL, Hayward RE, Langer RC, Fidock DA, Vinetz JM. Disruption of Plasmodium falciparum chitinase markedly impairs parasite invasion of mosquito midgut. Infect Immun. 2001;69(6):4048–54. pmid:11349075
      • 90. Dessens JT, Sidén-Kiamos I, Mendoza J, Mahairaki V, Khater E, Vlachou D, et al. SOAP, a novel malaria ookinete protein involved in mosquito midgut invasion and oocyst development. Mol Microbiol. 2003;49(2):319–29. pmid:12828632
      • 91. Templeton TJ, Kaslow DC, Fidock DA. Developmental arrest of the human malaria parasite Plasmodium falciparum within the mosquito midgut via CTRP gene disruption. Mol Microbiol. 2000;36(1):1–9. pmid:10760158
      • 92. Kariu T, Ishino T, Yano K, Chinzei Y, Yuda M. CelTOS, a novel malarial protein that mediates transmission to mosquito and vertebrate hosts. Mol Microbiol. 2006;59(5):1369–79. pmid:16468982
      • 93. Ecker A, Pinto SB, Baker KW, Kafatos FC, Sinden RE. Plasmodium berghei: plasmodium perforin-like protein 5 is required for mosquito midgut invasion in Anopheles stephensi. Exp Parasitol. 2007;116(4):504–8. pmid:17367780
      • 94. Kadota K, Ishino T, Matsuyama T, Chinzei Y, Yuda M. Essential role of membrane-attack protein in malarial transmission to mosquito host. Proc Natl Acad Sci U S A. 2004;101(46):16310–5. pmid:15520375
      • 95. Jennison C, Armstrong J, Dankwa D, Hertoghs N, Kumar S, Abatiyow B, et al. Plasmodium GPI-anchored micronemal antigen is essential for parasite transmission through the mosquito host. Mol Microbiol. 2023.
      • 96. Ramakrishnan C, Dessens JT, Armson R, Pinto SB, Talman AM, Blagborough AM, et al. Vital functions of the malarial ookinete protein, CTRP, reside in the A domains. Int J Parasitol. 2011;41(10):1029–39. pmid:21729699
      • 97. Deligianni E, Silmon de Monerri NC, McMillan PJ, Bertuccini L, Superti F, Manola M, et al. Correction: essential role of Plasmodium perforin-like protein 4 in ookinete midgut passage. PLoS One. 2018;13(9):e0204083. pmid:30208115
      • 98. Zhang G, Niu G, Franca C, Dong Y, Wang X, Butler N. Anopheles midgut FREP1 mediates Plasmodium invasion. J Biol Chem. 2015;290(27):16490–501.
      • 99. Niu G, Franc A, Zhang G, Roobsoong W, Nguitragool W, Wang X. The fibrinogen-like domain of FREP1 protein is a broad-spectrum malaria transmission-blocking vaccine antigen. J Biol Chem. 2017;292(28):11960–9.
      • 100. Wang X, Zhao Q, Christensen BM. Identification and characterization of the fibrinogen-like domain of fibrinogen-related proteins in the mosquito, Anopheles gambiae, and the fruitfly, Drosophila melanogaster, genomes. BMC Genomics. 2005;6:114.
      • 101. Zhang G, Niu G, Hooker-Romera D, Shabani S, Ramelow J, Wang X, et al. Targeting plasmodium alpha-tubulin-1 to block malaria transmission to mosquitoes. Front Cell Infect Microbiol. 2023;13:1132647.
      • 102. Dinglasan RR, Kalume DE, Kanzok SM, Ghosh AK, Muratova O, Pandey A, et al. Disruption of Plasmodium falciparum development by antibodies against a conserved mosquito midgut antigen. Proc Natl Acad Sci U S A. 2007;104(33):13461–6. pmid:17673553
      • 103. Atkinson SC, Armistead JS, Mathias DK, Sandeu MM, Tao D, Borhani-Dizaji N, et al. The Anopheles-midgut APN1 structure reveals a new malaria transmission-blocking vaccine epitope. Nat Struct Mol Biol. 2015;22(7):532–9. pmid:26075520
      • 104. Bender NG, Khare P, Martinez J, Tweedell RE, Nyasembe VO, López-Gutiérrez B, et al. Immunofocusing humoral immunity potentiates the functional efficacy of the AnAPN1 malaria transmission-blocking vaccine antigen. NPJ Vaccines. 2021;6(1):49. pmid:33824336
      • 105. Volohonsky G, Paul-Gilloteaux P, Štáfková J, Soichot J, Salamero J, Levashina EA. Kinetics of Plasmodium midgut invasion in Anopheles mosquitoes. PLoS Pathog. 2020;16(9):e1008739. pmid:32946522
      • 106. Gao H, Yang Z, Wang X, Qian P, Hong R, Chen X, et al. ISP1-Anchored polarization of GCβ/CDC50A complex initiates malaria ookinete gliding motility. Curr Biol. 2018;28(17):2763-2776.e6. pmid:30146157
      • 107. Sinden RE, Alavi Y, Raine JD. Mosquito—malaria interactions: a reappraisal of the concepts of susceptibility and refractoriness. Insect Biochem Mol Biol. 2004;34(7):625–9. pmid:15242703
      • 108. MacDonald NJ, Singh K, Reiter K, Nguyen V, Shimp R Jr, Gittis AG, et al. Structural and immunological differences in Plasmodium falciparum sexual stage transmission-blocking vaccines comprised of Pfs25-EPA nanoparticles. NPJ Vaccines. 2023;8(1):56. pmid:37061547
      • 109. Jagannath DK, Valiyaparambil A, Viswanath VK, Hurakadli MA, Kamariah N, Jafer AC, et al. Refolding and characterization of a diabody against Pfs25, a vaccine candidate of Plasmodium falciparum. Anal Biochem. 2022;655:114830. pmid:35944694
      • 110. Kefi M, Cardoso-Jaime V, Saab SA, Dimopoulos G. Curing mosquitoes with genetic approaches for malaria control. Trends Parasitol. 2024;40(6):487–99. pmid:38760256
      • 111. Isaacs AT, Jasinskiene N, Tretiakov M, Thiery I, Zettor A, Bourgouin C, et al. Transgenic Anopheles stephensi coexpressing single-chain antibodies resist Plasmodium falciparum development. Proc Natl Acad Sci U S A. 2012;109(28):E1922-30. pmid:22689959
      • 112. Dong Y, Simões ML, Marois E, Dimopoulos G. CRISPR/Cas9-mediated gene knockout of Anopheles gambiae FREP1 suppresses malaria parasite infection. PLoS Pathog. 2018;14(3):e1006898. pmid:29518156
      • 113. Kumar S, Molina-Cruz A, Gupta L, Rodrigues J, Barillas-Mury C. A peroxidase/dual oxidase system modulates midgut epithelial immunity in Anopheles gambiae. Science. 2010;327(5973):1644–8. pmid:20223948
      • 114. Oliveira G de A, Lieberman J, Barillas-Mury C. Epithelial nitration by a peroxidase/NOX5 system mediates mosquito antiplasmodial immunity. Science. 2012;335(6070):856–9. pmid:22282475
      • 115. Kumar S, Gupta L, Han YS, Barillas-Mury C. Inducible peroxidases mediate nitration of anopheles midgut cells undergoing apoptosis in response to Plasmodium invasion. J Biol Chem. 2004;279(51):53475–82. pmid:15456781
      • 116. Gupta L, Molina-Cruz A, Kumar S, Rodrigues J, Dixit R, Zamora RE, et al. The STAT pathway mediates late-phase immunity against Plasmodium in the mosquito Anopheles gambiae. Cell Host Microbe. 2009;5(5):498–507. pmid:19454353
      • 117. Garver LS, de Almeida Oliveira G, Barillas-Mury C. The JNK pathway is a key mediator of Anopheles gambiae antiplasmodial immunity. PLoS Pathog. 2013;9(9):e1003622. pmid:24039583
      • 118. Molina-Cruz A, Garver LS, Alabaster A, Bangiolo L, Haile A, Winikor J, et al. The human malaria parasite Pfs47 gene mediates evasion of the mosquito immune system. Science. 2013;340(6135):984–7. pmid:23661646
      • 119. Canepa GE, Molina-Cruz A, Barillas-Mury C. Molecular analysis of Pfs47-mediated plasmodium evasion of mosquito immunity. PLoS One. 2016;11(12):e0168279. pmid:27992481
      • 120. Molina-Cruz A, Canepa GE, Alves E Silva TL, Williams AE, Nagyal S, Yenkoidiok-Douti L, et al. Plasmodium falciparum evades immunity of anopheline mosquitoes by interacting with a Pfs47 midgut receptor. Proc Natl Acad Sci U S A. 2020;117(5):2597–605. pmid:31969456
      • 121. Alves E, Canepa G, Sweeney B, Hessab Alvarenga P, Zhao M, Vega-Rodriguez J. The heat shock protein Hsc70-3 mediates an anti-apoptotic response critical for Plasmodium evasion of Anopheles gambiae immunity. Microbiol Spectr. 2023;11(6):e0094023.
      • 122. Dong Y, Aguilar R, Xi Z, Warr E, Mongin E, Dimopoulos G. Anopheles gambiae immune responses to human and rodent Plasmodium parasite species. PLoS Pathog. 2006;2(6):e52. pmid:16789837
      • 123. Herrera-Ortiz A, Martínez-Barnetche J, Smit N, Rodriguez MH, Lanz-Mendoza H. The effect of nitric oxide and hydrogen peroxide in the activation of the systemic immune response of Anopheles albimanus infected with Plasmodium berghei. Dev Comp Immunol. 2011;35(1):44–50. pmid:20708028
      • 124. Castillo JC, Ferreira ABB, Trisnadi N, Barillas-Mury C. Activation of mosquito complement antiplasmodial response requires cellular immunity. Sci Immunol. 2017;2(7):eaal1505. pmid:28736767
      • 125. Blandin S, Shiao S-H, Moita LF, Janse CJ, Waters AP, Kafatos FC, et al. Complement-like protein TEP1 is a determinant of vectorial capacity in the malaria vector Anopheles gambiae. Cell. 2004;116(5):661–70. pmid:15006349
      • 126. Yassine H, Kamareddine L, Osta MA. The mosquito melanization response is implicated in defense against the entomopathogenic fungus Beauveria bassiana. PLoS Pathog. 2012;8(11):e1003029. pmid:23166497
      • 127. Kwon H, Smith RC. Chemical depletion of phagocytic immune cells in Anopheles gambiae reveals dual roles of mosquito hemocytes in anti-Plasmodium immunity. Proc Natl Acad Sci U S A. 2019;116(28):14119–28. pmid:31235594
      • 128. Volohonsky G, Hopp A-K, Saenger M, Soichot J, Scholze H, Boch J, et al. Transgenic expression of the anti-parasitic factor TEP1 in the malaria mosquito Anopheles gambiae. PLoS Pathog. 2017;13(1):e1006113. pmid:28095489
      • 129. Levashina EA, Moita LF, Blandin S, Vriend G, Lagueux M, Kafatos FC. Conserved role of a complement-like protein in phagocytosis revealed by dsRNA knockout in cultured cells of the mosquito, Anopheles gambiae. Cell. 2001;104(5):709–18. pmid:11257225
      • 130. Baxter R, Steinert S, Chelliah Y, Volohonsky G, Levashina E, Deisenhofer J. A heterodimeric complex of the LRR proteins LRIM1 and APL1C regulates complement-like immunity in Anopheles gambiae. Proc Natl Acad Sci U S A. 2010;107(39):16817–22.
      • 131. Reyes Ruiz VM, Sousa GL, Sneed SD, Farrant KV, Christophides GK, Povelones M. Stimulation of a protease targeting the LRIM1/APL1C complex reveals specificity in complement-like pathway activation in Anopheles gambiae. PLoS One. 2019;14(4):e0214753. pmid:30958840
      • 132. Fraiture M, Baxter RHG, Steinert S, Chelliah Y, Frolet C, Quispe-Tintaya W, et al. Two mosquito LRR proteins function as complement control factors in the TEP1-mediated killing of Plasmodium. Cell Host Microbe. 2009;5(3):273–84. pmid:19286136
      • 133. Povelones M, Waterhouse RM, Kafatos FC, Christophides GK. Leucine-rich repeat protein complex activates mosquito complement in defense against Plasmodium parasites. Science. 2009;324(5924):258–61. pmid:19264986
      • 134. Riehle MM, Xu J, Lazzaro BP, Rottschaefer SM, Coulibaly B, Sacko M, et al. Anopheles gambiae APL1 is a family of variable LRR proteins required for Rel1-mediated protection from the malaria parasite, Plasmodium berghei. PLoS One. 2008;3(11):e3672. pmid:18989366
      • 135. Mitri C, Jacques J-C, Thiery I, Riehle MM, Xu J, Bischoff E, et al. Fine pathogen discrimination within the APL1 gene family protects Anopheles gambiae against human and rodent malaria species. PLoS Pathog. 2009;5(9):e1000576. pmid:19750215
      • 136. Zmarlak NM, Lavazec C, Brito-Fravallo E, Genève C, Aliprandini E, Aguirre-Botero MC, et al. The Anopheles leucine-rich repeat protein APL1C is a pathogen binding factor recognizing Plasmodium ookinetes and sporozoites. PLoS Pathog. 2024;20(2):e1012008. pmid:38354186
      • 137. Povelones M, Bhagavatula L, Yassine H, Tan LA, Upton LM, Osta MA, et al. The CLIP-domain serine protease homolog SPCLIP1 regulates complement recruitment to microbial surfaces in the malaria mosquito Anopheles gambiae. PLoS Pathog. 2013;9(9):e1003623. pmid:24039584
      • 138. Yassine H, Kamareddine L, Chamat S, Christophides GK, Osta MA. A serine protease homolog negatively regulates TEP1 consumption in systemic infections of the malaria vector Anopheles gambiae. J Innate Immun. 2014;6(6):806–18. pmid:25012124
      • 139. Collins FH, Sakai RK, Vernick KD, Paskewitz S, Seeley DC, Miller LH, et al. Genetic selection of a Plasmodium-refractory strain of the malaria vector Anopheles gambiae. Science. 1986;234(4776):607–10. pmid:3532325
      • 140. Osta MA, Christophides GK, Kafatos FC. Effects of mosquito genes on Plasmodium development. Science. 2004;303(5666):2030–2. pmid:15044804
      • 141. Simões ML, Dong Y, Mlambo G, Dimopoulos G. C-type lectin 4 regulates broad-spectrum melanization-based refractoriness to malaria parasites. PLoS Biol. 2022;20(1):e3001515. pmid:35025886
      • 142. Nakhleh J, El Moussawi L, Osta MA. The melanization response in insect immunity. Adv Insect Physiol. 2017;52:83–109.
      • 143. Cao X, Gulati M, Jiang H. Serine protease-related proteins in the malaria mosquito, Anopheles gambiae. Insect Biochem Mol Biol. 2017;88:48–62. pmid:28780069
      • 144. Volz J, Müller H-M, Zdanowicz A, Kafatos FC, Osta MA. A genetic module regulates the melanization response of Anopheles to Plasmodium. Cell Microbiol. 2006;8(9):1392–405. pmid:16922859
      • 145. El Moussawi L, Nakhleh J, Kamareddine L, Osta MA. The mosquito melanization response requires hierarchical activation of non-catalytic clip domain serine protease homologs. PLoS Pathog. 2019;15(11):e1008194. pmid:31765430
      • 146. Nakhleh J, Christophides GK, Osta MA. The serine protease homolog CLIPA14 modulates the intensity of the immune response in the mosquito Anopheles gambiae. J Biol Chem. 2017;292(44):18217–26. pmid:28928218
      • 147. Volz J, Osta MA, Kafatos FC, Müller H-M. The roles of two clip domain serine proteases in innate immune responses of the malaria vector Anopheles gambiae. J Biol Chem. 2005;280(48):40161–8. pmid:16188883
      • 148. Zhang X, Li M, El Moussawi L, Saab S, Zhang S, Osta MA, et al. CLIPB10 is a terminal protease in the regulatory network that controls melanization in the African malaria mosquito Anopheles gambiae. Front Cell Infect Microbiol. 2021;10585986. pmid:33520733
      • 149. Sousa GL, Bishnoi R, Baxter RHG, Povelones M. The CLIP-domain serine protease CLIPC9 regulates melanization downstream of SPCLIP1, CLIPA8, and CLIPA28 in the malaria vector Anopheles gambiae. PLoS Pathog. 2020;16(10):e1008985. pmid:33045027
      • 150. Saab SA, Zhang X, Zeineddine S, Morejon B, Michel K, Osta MA. Insight into the structural hierarchy of the protease cascade that regulates the mosquito melanization response. Microbes Infect. 2024;26(1–2):105245. pmid:37918462
      • 151. Zhang X, Zhang S, Kuang J, Sellens KA, Morejon B, Saab SA, et al. CLIPB4 is a central node in the protease network that regulates humoral immunity in Anopheles gambiae mosquitoes. J Innate Immun. 2023;15(1):680–96. pmid:37703846
      • 152. An C, Budd A, Kanost MR, Michel K. Characterization of a regulatory unit that controls melanization and affects longevity of mosquitoes. Cell Mol Life Sci. 2011;68(11):1929–39. pmid:20953892
      • 153. Povelones M, Upton LM, Sala KA, Christophides GK. Structure-function analysis of the Anopheles gambiae LRIM1/APL1C complex and its interaction with complement C3-like protein TEP1. PLoS Pathog. 2011;7(4):e1002023. pmid:21533217
      • 154. Simoes M, Mlambo G, Tripathi A, Dong Y, Dimopoulos G. Immune regulation of Plasmodium is Anopheles species specific and infection intensity dependent. mBio. 2017;8(5):e00710-17.
      • 155. Bishnoi R, Sousa GL, Contet A, Day CJ, Hou C-FD, Profitt LA, et al. Solution structure, glycan specificity and of phenol oxidase inhibitory activity of Anopheles C-type lectins CTL4 and CTLMA2. Sci Rep. 2019;9(1):15191. pmid:31645596
      • 156. Raddi G, Barletta ABF, Efremova M, Ramirez JL, Cantera R, Teichmann SA, et al. Mosquito cellular immunity at single-cell resolution. Science. 2020;369(6507):1128–32. pmid:32855340
      • 157. Pinto SB, Lombardo F, Koutsos AC, Waterhouse RM, McKay K, An C, et al. Discovery of Plasmodium modulators by genome-wide analysis of circulating hemocytes in Anopheles gambiae. Proc Natl Acad Sci U S A. 2009;106(50):21270–5. pmid:19940242
      • 158. Severo M, Landry J, Lindquist R, Goosmann C, Brinkmann V, Collier P, et al. Unbiased classification of mosquito blood cells by single-cell genomics and high-content imaging. Proc Natl Acad Sci U S A. 2018;115(32):E7568–77.
      • 159. Kwon H, Mohammed M, Franzén O, Ankarklev J, Smith RC. Single-cell analysis of mosquito hemocytes identifies signatures of immune cell subtypes and cell differentiation. Elife. 2021;10:e66192. pmid:34318744
      • 160. Lombardo F, Ghani Y, Kafatos FC, Christophides GK. Comprehensive genetic dissection of the hemocyte immune response in the malaria mosquito Anopheles gambiae. PLoS Pathog. 2013;9(1):e1003145. pmid:23382679
      • 161. Lombardo F, Christophides GK. Novel factors of Anopheles gambiae haemocyte immune response to Plasmodium berghei infection. Parasit Vectors. 2016;9:78. pmid:26858200
      • 162. Smith RC, King JG, Tao D, Zeleznik OA, Brando C, Thallinger GG, et al. Molecular profiling of phagocytic immune cells in anopheles gambiae reveals integral roles for hemocytes in mosquito innate immunity. Mol Cell Proteomics. 2016;15(11):3373–87. pmid:27624304
      • 163. Dong Y, Dimopoulos G. Anopheles fibrinogen-related proteins provide expanded pattern recognition capacity against bacteria and malaria parasites. J Biol Chem. 2009;284(15):9835–44. pmid:19193639
      • 164. Simões ML, Dong Y, Hammond A, Hall A, Crisanti A, Nolan T, et al. The Anopheles FBN9 immune factor mediates Plasmodium species-specific defense through transgenic fat body expression. Dev Comp Immunol. 2017;67:257–65. pmid:27667688
      • 165. Li J, Wang X, Zhang G, Githure JI, Yan G, James AA. Genome-block expression-assisted association studies discover malaria resistance genes in Anopheles gambiae. Proc Natl Acad Sci U S A. 2013;110(51):20675–80.
      • 166. Aly ASI, Vaughan AM, Kappe SHI. Malaria parasite development in the mosquito and infection of the mammalian host. Annu Rev Microbiol. 2009;63:195–221. pmid:19575563
      • 167. Ukegbu CV, Christophides GK, Vlachou D. Identification of three novel plasmodium factors involved in ookinete to oocyst developmental transition. Front Cell Infect Microbiol. 2021;11:634273. pmid:33791240
      • 168. Smith RC, Barillas-Mury C. Plasmodium oocysts: overlooked targets of mosquito immunity. Trends Parasitol. 2016;32(12):979–90. pmid:27639778
      • 169. Srinivasan P, Fujioka H, Jacobs-Lorena M. PbCap380, a novel oocyst capsule protein, is essential for malaria parasite survival in the mosquito. Cell Microbiol. 2008;10(6):1304–12. pmid:18248630
      • 170. Sasaki H, Sekiguchi H, Sugiyama M, Ikadai H. Plasmodium berghei Cap93, a novel oocyst capsule-associated protein, plays a role in sporozoite development. Parasit Vectors. 2017;10(1):399. pmid:28841886
      • 171. Nakayama K, Kimura Y, Kitahara Y, Soga A, Haraguchi A, Hakozaki J, et al. Role of Plasmodium berghei ookinete surface and oocyst capsule protein, a novel oocyst capsule-associated protein, in ookinete motility. Parasit Vectors. 2021;14(1):373. pmid:34289894
      • 172. Preira CMF, Pizzi E, Fratini F, Grasso F, Boccolini D, Mochi S, et al. A time point proteomic analysis reveals protein dynamics of Plasmodium oocysts. Mol Cell Proteomics. 2024;23(3):100736. pmid:38342407
      • 173. Zhu F, Zheng H, Chen S, Zhang K, Qin X, Zhang J, et al. Malaria oocysts require circumsporozoite protein to evade mosquito immunity. Nat Commun. 2022;13(1):3208. pmid:35680915
      • 174. Nyasembe VO, Hamerly T, Lopez-Gutierrez B, Leyte-Vidal AM, Coatsworth H, Dinglasan RR. Adipokinetic hormone signaling in the malaria vector Anopheles gambiae facilitates Plasmodium falciparum sporogony. Commun Biol. 2023;6(1):171.
      • 175. Saeed S, Tremp A, Dessens J. Plasmodium sporozoite excystation involves local breakdown of the oocyst capsule. Sci Rep. 2023;13(1):22222.
      • 176. Graumans W, Jacobs E, Bousema T, Sinnis P. When is a Plasmodium-infected mosquito an infectious mosquito? Trends Parasitol. 2020;36(8):705–16. pmid:32620501
      • 177. Zeineddine S, Jaber S, Saab SA, Nakhleh J, Dimopoulos G, Osta MA. Late sporogonic stages of Plasmodium parasites are susceptible to the melanization response in Anopheles gambiae mosquitoes. Front Cell Infect Microbiol. 2024;14:1438019. pmid:39149419
      • 178. Smith RC, Barillas-Mury C, Jacobs-Lorena M. Hemocyte differentiation mediates the mosquito late-phase immune response against Plasmodium in Anopheles gambiae. Proc Natl Acad Sci U S A. 2015;112(26):E3412-20. pmid:26080400
      • 179. Barletta A, Smith J, Burkart E, Bondarenko S, Sharakhov I, Criscione F. Mosquito midgut stem cell cellular defense response limits Plasmodium parasite infection. Nat Commun. 2024;15(1):1422.
      • 180. Habtewold T, Sharma AA, Wyer CAS, Masters EKG, Windbichler N, Christophides GK. Plasmodium oocysts respond with dormancy to crowding and nutritional stress. Sci Rep. 2021;11(1):3090. pmid:33542254
      • 181. Kwon H, Simoes M, Reynolds R, Dimopoulos G, Smith R. Additional feeding reveals differences in immune recognition and growth of Plasmodium parasites in the mosquito host. mSphere. 2021;6(2).
      • 182. Shaw WR, Holmdahl IE, Itoe MA, Werling K, Marquette M, Paton DG, et al. Multiple blood feeding in mosquitoes shortens the Plasmodium falciparum incubation period and increases malaria transmission potential. PLoS Pathog. 2020;16(12):e1009131. pmid:33382824
      • 183. Arrighi RBG, Lycett G, Mahairaki V, Siden-Kiamos I, Louis C. Laminin and the malaria parasite’s journey through the mosquito midgut. J Exp Biol. 2005;208(Pt 13):2497–502. pmid:15961736
      • 184. Warburg A, Shtern A, Cohen N, Dahan N. Laminin and a Plasmodium ookinete surface protein inhibit melanotic encapsulation of Sephadex beads in the hemocoel of mosquitoes. Microbes Infect. 2007;9(2):192–9. pmid:17224290
      • 185. Goulielmaki E, Sidén-Kiamos I, Loukeris TG. Functional characterization of Anopheles matrix metalloprotease 1 reveals its agonistic role during sporogonic development of malaria parasites. Infect Immun. 2014;82(11):4865–77. pmid:25183733
      • 186. Arrighi RBG, Hurd H. The role of Plasmodium berghei ookinete proteins in binding to basal lamina components and transformation into oocysts. Int J Parasitol. 2002;32(1):91–8. pmid:11796126
      • 187. Atella GC, Bittencourt-Cunha PR, Nunes RD, Shahabuddin M, Silva-Neto MAC. The major insect lipoprotein is a lipid source to mosquito stages of malaria parasite. Acta Trop. 2009;109(2):159–62. pmid:19013123
      • 188. Rono MK, Whitten MMA, Oulad-Abdelghani M, Levashina EA, Marois E. The major yolk protein vitellogenin interferes with the anti-plasmodium response in the malaria mosquito Anopheles gambiae. PLoS Biol. 2010;8(7):e1000434. pmid:20652016
      • 189. Jaramillo-Gutierrez G, Rodrigues J, Ndikuyeze G, Povelones M, Molina-Cruz A, Barillas-Mury C. Mosquito immune responses and compatibility between Plasmodium parasites and anopheline mosquitoes. BMC Microbiol. 2009;9:154. pmid:19643026
      • 190. Shaw WR, Marcenac P, Catteruccia F. Plasmodium development in Anopheles: a tale of shared resources. Trends Parasitol. 2022;38(2):124–35. pmid:34548252
      • 191. Matuschewski K. Getting infectious: formation and maturation of Plasmodium sporozoites in the Anopheles vector. Cell Microbiol. 2006;8(10):1547–56. pmid:16984410
      • 192. James AA. Blocking malaria parasite invasion of mosquito salivary glands. J Exp Biol. 2003;206(Pt 21):3817–21. pmid:14506217
      • 193. Wells M, Andrew D. Anopheles salivary gland architecture shapes Plasmodium sporozoite availability for transmission. mBio. 2019;10(4):.
      • 194. Dundas K, Shears MJ, Sinnis P, Wright GJ. Important extracellular interactions between Plasmodium sporozoites and host cells required for infection. Trends Parasitol. 2019;35(2):129–39. pmid:30583849
      • 195. Loubens M, Marinach C, Paquereau C-E, Hamada S, Hoareau-Coudert B, Akbar D, et al. The claudin-like apicomplexan microneme protein is required for gliding motility and infectivity of Plasmodium sporozoites. PLoS Pathog. 2023;19(3):e1011261. pmid:36928686
      • 196. Rodrigues J, Oliveira GA, Kotsyfakis M, Dixit R, Molina-Cruz A, Jochim R, et al. An epithelial serine protease, AgESP, is required for Plasmodium invasion in the mosquito Anopheles gambiae. PLoS One. 2012;7(4):e35210. pmid:22509400
      • 197. Pinheiro-Silva R, Borges L, Coelho LP, Cabezas-Cruz A, Valdés JJ, do Rosário V, et al. Gene expression changes in the salivary glands of Anopheles coluzzii elicited by Plasmodium berghei infection. Parasit Vectors. 2015;8:485. pmid:26395987
      • 198. Jacobs-Lorena M. Interrupting malaria transmission by genetic manipulation of anopheline mosquitoes. J Vector Borne Dis. 2003;40(3–4):73–7. pmid:15119075
      • 199. Hillyer JF, Barreau C, Vernick KD. Efficiency of salivary gland invasion by malaria sporozoites is controlled by rapid sporozoite destruction in the mosquito haemocoel. Int J Parasitol. 2007;37(6):673–81. pmid:17275826
      • 200. Rosenberg R, Rungsiwongse J. The number of sporozoites produced by individual malaria oocysts. Am J Trop Med Hyg. 1991;45(5):574–7. pmid:1951866
      • 201. King JG, Hillyer JF. Infection-induced interaction between the mosquito circulatory and immune systems. PLoS Pathog. 2012;8(11):e1003058. pmid:23209421
      • 202. Hernandez-Martinez S, Lanz H, Rodriguez M, Gonzalez-Ceron L, Tsutsumi V. Cellular-mediated reactions to foreign organisms inoculated into the hemocoel of Anopheles albimanus (Diptera: Culicidae). J Med Entomol. 2002;39(1):61–9.
      • 203. Hillyer J, Schmidt S, Christensen B. Rapid phagocytosis and melanization of bacteria and Plasmodium sporozoites by hemocytes of the mosquito Aedes aegypti. J Parasitol. 2003;89(1):62–9.
      • 204. Glenn JD, King JG, Hillyer JF. Structural mechanics of the mosquito heart and its function in bidirectional hemolymph transport. J Exp Biol. 2010;213(4):541–50. pmid:20118304
      • 205. Sigle LT, Hillyer JF. Mosquito hemocytes preferentially aggregate and phagocytose pathogens in the periostial regions of the heart that experience the most hemolymph flow. Dev Comp Immunol. 2016;55:90–101. pmid:26526332
      • 206. Hernández-Martínez S, Lanz-Mendoza H, Martínez-Barnetche J, Rodríguez MH. Antimicrobial properties of Anopheles albimanus pericardial cells. Cell Tissue Res. 2013;351(1):127–37. pmid:23229355
      • 207. Cardoso-Jaime V, Maya-Maldonado K, Celestino-Montes A, Tsutsumi V, Hernández-Martínez S. Lysozyme c-1 gene is overexpressed in Anopheles albimanus pericardial cells after an immune challenge. Dev Comp Immunol. 2021;114:103830. pmid:32805306
      • 208. Cardoso-Jaime V, Maya-Maldonado K, Tsutsumi V, Hernández-Martínez S. Mosquito pericardial cells upregulate Cecropin expression after an immune challenge. Dev Comp Immunol. 2023;147:104745. pmid:37268262
      • 209. Estevez-Lao T, Sigle L, Gomez S, Hillyer J. Nitric oxide produced by periostial hemocytes modulates the bacterial infection-induced reduction of the mosquito heart rate. J Exp Biol. 2020;223(Pt 15):.
      • 210. Bergmann S, Graf E, Hoffmann P, Becker SC, Stern M. Localization of nitric oxide-producing hemocytes in Aedes and Culex mosquitoes infected with bacteria. Cell Tissue Res. 2024;395(3):313–26. pmid:38240845
      • 211. Dimopoulos G, Seeley D, Wolf A, Kafatos FC. Malaria infection of the mosquito Anopheles gambiae activates immune-responsive genes during critical transition stages of the parasite life cycle. EMBO J. 1998;17(21):6115–23. pmid:9799221
      • 212. Rosinski-Chupin I, Briolay J, Brouilly P, Perrot S, Gomez SM, Chertemps T, et al. SAGE analysis of mosquito salivary gland transcriptomes during Plasmodium invasion. Cell Microbiol. 2007;9(3):708–24. pmid:17054438
      • 213. Roth A, Adapa S, Zhang M, Liao X, Saxena V, Goffe R. Unraveling the Plasmodium vivax sporozoite transcriptional journey from mosquito vector to human host. Sci Rep. 2018;8(1):12183.
      • 214. Arca B, Lombardo F, Valenzuela J, Francischetti I, Marinotti O, Coluzzi M. An updated catalogue of salivary gland transcripts in the adult female mosquito, Anopheles gambiae. J Exp Biol. 2005;208(20):3971–86.
      • 215. Bevivino G, Arcà B, Lombardo F. Effects of local and systemic immune challenges on the expression of selected salivary genes in the malaria mosquito Anopheles coluzzii. Pathogens. 2021;10(10):1300. pmid:34684249
      • 216. Barillas-Mury C, Ribeiro JMC, Valenzuela JG. Understanding pathogen survival and transmission by arthropod vectors to prevent human disease. Science. 2022;377(6614):eabc2757. pmid:36173836
      • 217. Arora G, Chuang Y-M, Sinnis P, Dimopoulos G, Fikrig E. Malaria: influence of Anopheles mosquito saliva on Plasmodium infection. Trends Immunol. 2023;44(4):256–65. pmid:36964020
      • 218. Ribeiro JM, Francischetti IM. Role of arthropod saliva in blood feeding: sialome and post-sialome perspectives. Ann Rev Entomol. 2003;48:73–88.
      • 219. Kanatani S, Stiffler D, Bousema T, Yenokyan G, Sinnis P. Revisiting the Plasmodium sporozoite inoculum and elucidating the efficiency with which malaria parasites progress through the mosquito. Nat Commun. 2024;15(1):748. pmid:38272943
      • 220. Schleicher TR, Yang J, Freudzon M, Rembisz A, Craft S, Hamilton M, et al. A mosquito salivary gland protein partially inhibits Plasmodium sporozoite cell traversal and transmission. Nat Commun. 2018;9(1):2908. pmid:30046053
      • 221. Hopp CS, Sinnis P. The innate and adaptive response to mosquito saliva and Plasmodium sporozoites in the skin. Ann N Y Acad Sci. 2015;1342(1):37–43. pmid:25694058
      • 222. Chuang Y-M, Agunbiade TA, Tang X-D, Freudzon M, Almeras L, Fikrig E. The effects of a mosquito salivary protein on sporozoite traversal of host cells. J Infect Dis. 2021;224(3):544–53. pmid:33306099
      • 223. Dragovic SM, Agunbiade TA, Freudzon M, Yang J, Hastings AK, Schleicher TR, et al. Immunization with AgTRIO, a protein in anopheles saliva, contributes to protection against Plasmodium infection in mice. Cell Host Microbe. 2018;23(4):523-535.e5. pmid:29649443
      • 224. Kebaier C, Voza T, Vanderberg J. Neither mosquito saliva nor immunity to saliva has a detectable effect on the infectivity of Plasmodium sporozoites injected into mice. Infect Immun. 2010;78(1):545–51. pmid:19884338
      • 225. Arora G, Sajid A, Chuang Y-M, Dong Y, Gupta A, Gambardella K, et al. Immunomodulation by mosquito salivary protein AgSAP contributes to early host infection by Plasmodium. mBio. 2021;12(6):e03091-21.
      • 226. McDowell MA. Vector-transmitted disease vaccines: targeting salivary proteins in transmission (SPIT). Trends Parasitol. 2015;31(8):363–72. pmid:26003330
      • 227. Arcà B, Colantoni A, Fiorillo C, Severini F, Benes V, Di Luca M. MicroRNAs from saliva of anopheline mosquitoes mimic human endogenous miRNAs and may contribute to vector-host-pathogen interactions. Sci Rep. 2019;9(1):2955.
      • 228. Fiorillo C, Yen P-S, Colantoni A, Mariconti M, Azevedo N, Lombardo F, et al. MicroRNAs and other small RNAs in Aedes aegypti saliva and salivary glands following chikungunya virus infection. Sci Rep. 2022;12(1):9536. pmid:35681077
      • 229. Accoti A, Damiani C, Nunzi E, Cappelli A, Iacomelli G, Monacchia G, et al. Anopheline mosquito saliva contains bacteria that are transferred to a mammalian host through blood feeding. Front Microbiol. 2023;14:1157613. pmid:37533823
      • 230. Klug D, Gautier A, Calvo E, Marois E, Blandin SA. The salivary protein Saglin facilitates efficient midgut colonization of Anopheles mosquitoes by malaria parasites. PLoS Pathog. 2023;19(3):e1010538. pmid:36862755
      • 231. Alves E, Radtke A, Balaban A, Pascini T, Pala Z, Roth A. The fibrinolytic system enables the onset of Plasmodium infection in the mosquito vector and the mammalian host. Sci Adv. 2021;7(6):eabe1234.
      • 232. Pala Z, Alves E, Minai M, Crews B, Patino-Martinez E, Carmona-Rivera C. Anopheles salivary apyrase regulates blood meal hemostasis and drives malaria parasite transmission. bioRxiv. 2023.
      • 233. Pascini TV, Jeong YJ, Huang W, Pala ZR, Sá JM, Wells MB, et al. Transgenic Anopheles mosquitoes expressing human PAI-1 impair malaria transmission. Nat Commun. 2022;13(1):2949. pmid:35618711
      • 234. Li M, Zhou Y, Cheng J, Wang Y, Lan C, Shen Y. Response of the mosquito immune system and symbiotic bacteria to pathogen infection. Parasit Vectors. 2024;17(1):69. pmid:38368353
      • 235. Contreras-Garduño J, Rodríguez MC, Hernández-Martínez S, Martínez-Barnetche J, Alvarado-Delgado A, Izquierdo J, et al. Plasmodium berghei induced priming in Anopheles albimanus independently of bacterial co-infection. Dev Comp Immunol. 2015;52(2):172–81. pmid:26004500
      • 236. Maya-Maldonado K, Cardoso-Jaime V, Hernández-Martínez S, Recio-Tótoro B, Bello-Garcia D, Hernández-Hernández F de la C, et al. Plasmodium exposure alters midgut epithelial cell dynamics during the immune memory in Anopheles albimanus. Dev Comp Immunol. 2022;133:104424. pmid:35447160
      • 237. Ramirez JL, de Almeida Oliveira G, Calvo E, Dalli J, Colas RA, Serhan CN, et al. A mosquito lipoxin/lipocalin complex mediates innate immune priming in Anopheles gambiae. Nat Commun. 2015;6:7403. pmid:26100162
      • 238. Rodrigues J, Brayner FA, Alves LC, Dixit R, Barillas-Mury C. Hemocyte differentiation mediates innate immune memory in Anopheles gambiae mosquitoes. Science. 2010;329(5997):1353–5. pmid:20829487
      • 239. Schmucker D, Clemens JC, Shu H, Worby CA, Xiao J, Muda M, et al. Drosophila Dscam is an axon guidance receptor exhibiting extraordinary molecular diversity. Cell. 2000;101(6):671–84. pmid:10892653
      • 240. Dong Y, Taylor HE, Dimopoulos G. AgDscam, a hypervariable immunoglobulin domain-containing receptor of the Anopheles gambiae innate immune system. PLoS Biol. 2006;4(7):e229. pmid:16774454
      • 241. Dong Y, Cirimotich CM, Pike A, Chandra R, Dimopoulos G. Anopheles NF-κB-regulated splicing factors direct pathogen-specific repertoires of the hypervariable pattern recognition receptor AgDscam. Cell Host Microbe. 2012;12(4):521–30. pmid:23084919
      • 242. Carballar-Lejarazú R, Dong Y, Pham TB, Tushar T, Corder RM, Mondal A, et al. Dual effector population modification gene-drive strains of the African malaria mosquitoes, Anopheles gambiae and Anopheles coluzzii. Proc Natl Acad Sci U S A. 2023;120(29):e2221118120. pmid:37428915