Malaria transmission to mosquitoes depends on the presence of mature gametocytes in human peripheral blood that are ingested by a mosquito during blood feeding. Ingested parasites undergo several developmental transformations in a process called sporogony. After ingestion, gametocytes transform into male and female gametes that fuse to form a zygote. The zygote differentiates into a motile ookinete that penetrates the midgut epithelium to form an oocyst. Multiple rounds of mitotic replication result in the formation of sporozoites inside an oocyst. Upon oocyst rupture, sporozoites are released into the haemocoel and invade the salivary glands (Mueller et al., 2010). These sporozoites penetrate the distal portion of the two lateral and medial lobes of the glands and accumulate extracellularly inside secretory cavities before entering the salivary ducts (Wells and Andrew, 2019). Despite the large number of sporozoites in the cavities, only a small proportion pass through the proximal part of the lobes where the salivary ducts become narrow (Frischknecht et al., 2004; Bradley et al., 2018), and only tens or low hundreds of sporozoites are assumed to be inoculated per mosquito bite (Graumans et al., 2020). Developmental bottlenecks during sporogony, as well as the size of the sporozoite inoculum, remain incompletely understood (Graumans et al., 2020).

The density of gametocytes in human peripheral blood is an important determinant of human to mosquito transmission. Though infections with parasite densities below the microscopic threshold for detection can infect mosquitoes (Schneider et al., 2007; Bousema et al., 2006), the likelihood and infection intensity increase with the number of ingested gametocytes (Bradley et al., 2018). Because of the abundance of low-density gametocyte carriers among infected populations, these are considered important drivers of malaria transmission (Andolina et al., 2021; Slater et al., 2019; Ouédraogo et al., 2016). Importantly, this conclusion is based on the assumption that all infected mosquitoes are equally infectious regardless of oocyst densities.

In apparent support of this assumption, single oocyst infections can result in thousands of salivary gland sporozoites (Rosenberg and Rungsiwongse, 1991; Pringle, 1965). A positive correlation between oocyst densities and salivary gland sporozoites was previously observed in Plasmodium falciparum (Vaughan et al., 1992; Miura et al., 2019) as well as in Plasmodium vivax (Solarte et al., 2011), suggesting that on average of 1000–2000 sporozoites reach the salivary glands per single ruptured oocyst. The few studies that quantified sporozoite inoculum by allowing mosquitoes to salivating into capillary tubes containing mineral oil, sucrose solution, or blood (Beier et al., 1991a; Beier et al., 1991b; Rosenberg et al., 1990) estimated median inocula ranging between 8 and 39 sporozoites with a minority of mosquitoes expelling >100 sporozoites (reviewed in Graumans et al., 2020).

While these studies provide some insights into sporozoite expelling and transmission dynamics, they do not reflect natural feeding conditions. Also, microscopy techniques used to quantify sporozoites may have underestimated the number of sporozoites (Medica and Sinnis, 2005). Subsequent studies with rodent Plasmodium models allowed natural mosquito probing through skin and showed a positive association between sporozoite density in salivary glands and expelled sporozoite numbers (Medica and Sinnis, 2005; Aleshnick et al., 2020). A recent study using a Plasmodium yoelii malaria model demonstrated that mosquitoes with over 10,000 salivary gland sporozoites were 7.5 times more likely to initiate a malaria infection in mice (Aleshnick et al., 2020). This finding has not been replicated for human malarias but is broadly consistent with post hoc analysis of infection likelihood in controlled human malaria infections (CHMI), where only mosquitoes that had >1000 P. falciparum salivary gland sporozoites remaining after probing (the residual sporozoite load) were capable of establishing an infection in malaria-naïve volunteers (Churcher et al., 2017).

If low oocyst/low sporozoite densities in mosquitoes are unlikely to initiate infections in humans, this may have profound consequences for our understanding of transmission (WHO, 2017). If mosquitoes with low-infection burdens have limited transmission potential, then the rationale for targeting low-density infections in humans that give rise to low-infection burdens in mosquitoes (Bradley et al., 2018) may be diminished.

Here, we examined the progression of sporozoite development and the number of sporozoites expelled into artificial skin by individual Anopheles mosquitoes infected either with P. falciparum gametocyte cultures or in experimental infections with naturally circulating parasite strains. We directly assessed the association between oocyst burden, salivary gland infection intensity, and the number of sporozoites expelled.

We examined P. falciparum sporogony in different mosquito species by employing sensitive molecular and staining techniques in conjunction with mosquito probing experiments on artificial skin. Through the visualization of ruptured oocysts and the simultaneous quantification of sporozoites that are expelled by individual mosquitoes, we observed that 95% of oocysts rupture to release sporozoites and that higher salivary gland sporozoite load is associated with shorter time to colonization of salivary glands and larger inoculum size.

The proportion of infectious mosquitoes is a central component of malariometric indices both in terms of quantifying the force of infection and the human infectious reservoir. The entomological inoculation rate (EIR) is defined as the number of infectious bites per person per time unit and is the product of human biting rate and the proportion of sporozoite-positive mosquitoes (Tusting et al., 2014; Shaukat et al., 2010). While EIR is a common measure of human malaria exposure, mosquito infection prevalence is used in this calculation and thus assumes all sporozoite-positive mosquitoes are equally infectious (Smith et al., 2012). Similarly, assessments of the human infectious reservoir for malaria typically take the number of oocyst-positive mosquitoes as a measure of transmission (Ouédraogo et al., 2016; Gonçalves et al., 2017) and thereby not only assume that all oocysts will lead to salivary gland sporozoites but also that all oocyst-positive mosquitoes have equal transmission potential. Recent work with a rodent malaria model challenged this central assumption (Aleshnick et al., 2020); we provide the first direct evidence for P. falciparum that sporozoite burden may indeed be a relevant determinant in efficient sporozoite expelling.

In this study, we assessed sporozoite expelling by mosquitoes carrying low- and high-infection burdens. Our findings confirm that the vast majority (~95%) of oocysts rupture to release sporozoites. This estimate is higher than a previous study with cultured gametocytes (~72%) (Stone et al., 2013) that did not provide a second bloodmeal that may accelerate oocyst maturation (Shaw et al., 2020); a second bloodmeal also better mimics natural feeding habits where multiple bloodmeals are taken within the period required for sporogony. Moreover, we observed a strong positive association between sporozoite salivary gland load and ruptured oocyst density in mosquitoes infected with both cultured and naturally circulating gametocytes, with similar median numbers of 4951 (IQR, 3016–8318) and 6350 (IQR, 4225–8475) salivary gland sporozoites per ruptured oocyst in An. stephensi and An. coluzzii, respectively. Previous studies have shown that despite the substantial release of sporozoites per oocyst, only a proportion of sporozoites successfully reach the salivary glands (Rosenberg and Rungsiwongse, 1991; Hillyer et al., 2007). In line with this, we observed considerably higher sporozoite estimates in intact oocysts (10,485–15,390 sporozoites per oocyst). Whilst these sporozoite estimates are higher than commonly reported (Graumans et al., 2020), an early study from the 1960s reported microscopy-detected sporozoite densities above 9000 sporozoites per oocyst (Pringle, 1965) and a recent study using qPCR similarly observed up to 12,583 sporozoites from a single oocyst (Wang et al., 2018). These high sporozoite numbers per oocyst also mean that, different from P. yoelii (Aleshnick et al., 2020) and P. berghei (Churcher et al., 2017), only a minority of infected mosquitoes had sporozoite densities below the threshold values previously reported in relation to a very low likelihood of achieving secondary infections. Even among infected An. stephensi mosquitoes with only one ruptured oocyst, 92.9% (26/28) had >1000 and 35.7% (10/28) had >10,000 sporozoites in their salivary glands. In our experiments in natural gametocyte carriers in Burkina Faso, only two mosquitoes were observed with single oocysts and both had >10,000 salivary gland sporozoites. Although P. falciparum sporozoite densities below these lower thresholds were thus uncommon, we observed that 39% of sporozoite-positive mosquitoes across a wide range of infection densities failed to expel sporozoites upon probing. This finding broadly aligns with an earlier study of Medica and Sinnis that reported that 22% of P. yoelii-infected mosquitoes failed to expel sporozoites (Medica and Sinnis, 2005). For highly infected mosquitoes, this inefficient expelling has been related to a decrease in apyrase in the mosquito saliva (Rossignol et al., 1984; Thiévent et al., 2019).

Importantly, we observed a positive association between salivary gland sporozoite density and the number of expelled sporozoites. For unknown reasons, this association was markedly stronger in experiments where An. coluzzii mosquitoes were infected using blood from gametocyte carriers who were naturally infected with P. falciparum in Burkina Faso compared to An. stephensi mosquitoes infected with cultured gametocytes. In our experiments with natural gametocyte carriers, sporozoite density appeared associated with both the likelihood of expelling any sporozoites and the inoculum size. The most heavily infected mosquitoes expelled ~14-fold higher sporozoite numbers compared to the lowest quartile. In addition, we found evidence that the EIP – the period required for a mosquito to become salivary gland sporozoite positive – is shorter for heavily infected mosquitoes. Previous studies have not found consistent effects of parasite burden on EIP (Guissou et al., 2023; Stopard et al., 2021) but, unlike our experiments, were also not specifically designed to examine this association across a broad range of infection intensities. Our associations of a shorter EIP in highly infected mosquitoes and, in separate experiments, a larger sporozoite inoculum size for highly infected mosquitoes make mosquito infection intensity a plausible factor in determining onward transmission potential to humans. Heavily infected mosquitoes may be infectious sooner and be more infectious. On the other side of the infection spectrum, it is conceivable that submicroscopic gametocyte carriers that typically result in low oocyst burdens in mosquitoes give rise to infected mosquitoes with a reduced transmission potential. This would greatly reduce their importance for sustaining malaria transmission and make it less important to identify individuals with low parasite densities for malaria elimination purposes (Slater et al., 2015). At the same time, we occasionally observed high sporozoite inocula from mosquitoes with low infection intensity. This argues against a clear threshold sporozoite or oocyst density below which mosquitoes are truly irrelevant for transmission. Moreover, the high number of sporozoites per oocyst makes it plausible that even the low oocyst burdens that are typically observed from asymptomatic parasite carriers (in the range of 1–5 oocysts/infected gut; Andolina et al., 2021; Gonçalves et al., 2017) would be sufficient to result in salivary gland sporozoite loads that are sufficient to result in secondary infections.

Our study leaves a number of questions and has several limitations. While the use of two mosquito and gametocyte sources was a relevant strength of our study, an uncertainty relates to the choice of artificial skin that has a realistic 1.33 mm thickness but is arguably less natural than microvascularized skin with all the natural cues for mosquito probing. Whilst genuine skin might have improved natural feeding behavior, probing and blood feeding were highly efficient in our model and we see no reasons to assume bias in the comparison between high- and low-infected mosquitoes. Our assessments of EIP and sporozoite expelling did not demonstrate the viability of sporozoites. Whilst the infectivity of sporozoites at different time points PI has been examined previously (Yang et al., 2017), these experiments have never been conducted with individual mosquitoes. In vitro experiments that aim to determine the infectivity of single mosquito bites would ideally retain the skin barrier that may be a relevant determinant for invasion capacity and use primary hepatocytes. These experiments were beyond the scope of the current work and would also not provide conclusive evidence on the likelihood of achieving secondary infections. Given striking differences in sporozoite burden between different Plasmodium species – low sporozoite densities appear considerably more common in mosquitoes infected with P. yoelii and P. berghei (Graumans et al., 2020; Churcher et al., 2017; Ahmed et al., 2021), and the inherent limitations of in vitro studies – the association between sporozoite inoculum size and the likelihood of achieving secondary infections may be best examined in controlled human infection studies. CHMI experiments can be specifically designed to estimate the likelihood that probing by (individual) high- and low-infected mosquitoes results in blood-stage infection in malaria-naïve volunteers. Whilst laborious, CHMI studies are unique in allowing definitive evidence on the possible differences in infectiousness between high- and low-infected mosquitoes.

In conclusion, we observed that the majority of oocysts rupture and contribute to salivary gland infection load. We further observe that this sporozoite load is highly variable and an important determinant of the number of sporozoites that is expelled into the skin upon probing.

All data generated or analysed during this study are deposited on Dryad: https://doi.org/10.5061/dryad.dbrv15f89.

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Author details

1. Chiara Andolina

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing – original draft

   Contributed equally with

   Wouter Graumans

   Competing interests

   No competing interests declared

2. Wouter Graumans

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing – original draft

   Contributed equally with

   Chiara Andolina

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3952-6491

3. Moussa Guelbeogo

   Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina Faso

   Contribution

   Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

4. Geert-Jan van Gemert

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2740-5337

5. Jordache Ramijth

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Formal analysis, Visualization, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

6. Soré Harouna

   Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina Faso

   Contribution

   Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

7. Zongo Soumanaba

   Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina Faso

   Contribution

   Writing – review and editing

   Competing interests

   No competing interests declared

8. Rianne Stoter

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Investigation

   Competing interests

   No competing interests declared

9. Marga Vegte-Bolmer

   Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

   Contribution

   Investigation

   Competing interests

   No competing interests declared

10. Martina Pangos

    Department of Plastic and Reconstructive Surgery, Azienda Ospedaliero Universitaria GiulianoIsontina Trieste, Trieste, Italy

    Contribution

    Resources, Methodology

    Competing interests

    No competing interests declared

11. Photini Sinnis

    Department of Molecular Microbiology and Immunology, Johns HopkinsBloomberg School of Public Health, Baltimore, United States

    Contribution

    Conceptualization, Writing – review and editing

    Competing interests

    No competing interests declared

12. Katharine Collins

    Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

    Contribution

    Supervision, Writing – review and editing

    Competing interests

    No competing interests declared

13. Sarah G Staedke

    Liverpool School of Tropical Medicine, Liverpool, United Kingdom

    Contribution

    Supervision, Writing – review and editing

    Competing interests

    No competing interests declared

14. Alfred B Tiono

    Centre National de Recherche et de Formation sur le Paludisme, Ouagadougou, Burkina Faso

    Contribution

    Supervision, Investigation, Writing – review and editing

    Competing interests

    No competing interests declared

15. Chris Drakeley

    Department of Immunology and Infection, London School of Hygiene and Tropical Medicine, London, United Kingdom

    Contribution

    Conceptualization

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-4863-075X

16. Kjerstin Lanke

    Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands

    Contribution

    Investigation, Methodology, Writing – original draft

    Competing interests

    No competing interests declared

17. Teun Bousema

    1. Department of Medical Microbiology, Radboud University Nijmegen Medical Centre, Nijmegen, Netherlands
    2. Department of Immunology and Infection, London School of Hygiene and Tropical Medicine, London, United Kingdom

    Contribution

    Conceptualization, Formal analysis, Supervision, Funding acquisition, Methodology, Writing – original draft, Writing – review and editing

    For correspondence

    teun.bousema@radboudumc.nl

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-2666-094X

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