The Plasmodium transmission-blocking symbiont, Microsporidia MB, is vertically transmitted through Anopheles arabiensis germline stem cells

Thomas Ogao Onchuru; Edward Edmond Makhulu; Purity Cassandra Ronnie; Stancy Mandere; Fidel Gabriel Otieno; Joseph Gichuhi; Jeremy Keith Herren

doi:10.1371/journal.ppat.1012340

Abstract

Microsporidia MB is a promising candidate for developing a symbiont-based strategy for malaria control because it disrupts the capacity of An. arabiensis to transmit the Plasmodium parasite. The symbiont is predominantly localized in the reproductive organs and is transmitted vertically from mother to offspring and horizontally (sexually) during mating. Due to the contribution of both transmission routes, Microsporidia MB has the potential to spread through target vector populations and become established at high prevalence. Stable and efficient vertical transmission of Microsporidia MB is important for its sustainable use for malaria control, however, the vertical transmission efficiency of Microsporidia MB can vary. In this study, we investigate the mechanistic basis of Microsporidia MB vertical transmission in An. arabiensis. We show that vertical transmission occurs through the acquisition of Microsporidia MB by Anopheles cystocyte progenitors following the division of germline stem cells. We also show that Microsporidia MB replicates to increase infection intensity in the oocyte of developing eggs when mosquitoes take a blood meal suggesting that symbiont proliferation in the ovary is coordinated with egg development. The rate of Microsporidia MB transmission to developing eggs is on average higher than the recorded (mother to adult offspring) vertical transmission rate. This likely indicates that a significant proportion of An. arabiensis offspring lose their Microsporidia MB symbionts during development. The stability of germline stem cell infections, coordination of symbiont proliferation, and very high rate of transmission from germline stem cells to developing eggs indicate that Microsporidia MB has a highly specialized vertical transmission strategy in An. arabiensis, which may explain host specificity.

Author summary

Mosquito vectors of diseases are associated with a broad range of microbes. Some of the microbes significantly affect vector biology including pathogen transmission efficiency. Anopheles mosquitoes, which transmit the malaria parasite, Plasmodium falciparum, harbor a native microbe known as Microsporidia MB. This microbe interferes with the formation of transmissible stages of the parasite (sporozoites) that are transferred to humans by female mosquitoes when taking a blood meal. This phenotype can be exploited to develop a novel strategy for controlling malaria similar to the control of dengue fever using Aedes mosquitoes carrying Wolbachia bacteria. Mother to offspring transmission of protective microbes is important in sustainable application of microbe-based technologies to control vector-borne diseases because it ensures maintenance of the microbe in target vector populations across many generations. Here, we investigated stability of Microsporidia MB infections and efficiency of mother to offspring transmission during early stages of egg formation and development. We found that this microbe has a specialized transmission mechanism that involves infecting the germline cells that are important in egg production. We also demonstrated a very high transmission rate (97%) of the Microsporidia MB from infected germline cells into daughter cells during cell division. As the germline daughter cells developed into eggs, Microsporidia MB established itself in the egg yolk through active replication which only occured after the female mosquitoes had a blood meal. Our study gives insights into an efficient mother to offspring transmission route of Microsporidia MB that can be utilized sustainably in microbe-based intervention to control malaria.

Citation: Onchuru TO, Makhulu EE, Ronnie PC, Mandere S, Otieno FG, Gichuhi J, et al. (2024) The Plasmodium transmission-blocking symbiont, Microsporidia MB, is vertically transmitted through Anopheles arabiensis germline stem cells. PLoS Pathog 20(11): e1012340. https://doi.org/10.1371/journal.ppat.1012340

Editor: Marcelo U. Ferreira, University of Sao Paulo, BRAZIL

Received: June 13, 2024; Accepted: October 29, 2024; Published: November 11, 2024

Copyright: © 2024 Onchuru 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.

Data Availability: The relative area of the primary oocyte infected with Microsporidia MB across developmental stages of the ovariole, the primary follicle infection rate, and the vertical transmission rate of Microsporidia MB to sugar-fed and blood-fed adults data are available in the Figshare Digital Repository at https://doi.org/10.6084/m9.figshare.25846351.

Funding: The authors gratefully acknowledge the financial support for this research by the following organizations and agencies: Open Philanthropy (SYMBIOVECTOR Track A to JKH), the Bill and Melinda Gates Foundation (INV0225840 to JKH), and the Children’s Investment Fund Foundation (SMBV-FFT to JKH). This work was also supported by funds from the Swedish International Development Cooperation Agency (Sida); the Swiss Agency for Development and Cooperation (SDC); the Australian Centre for International Agricultural Research (ACIAR); the Government of Norway; the German Federal Ministry for Economic Cooperation and Development (BMZ); and the Government of the Republic of Kenya to icipe. 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.

Introduction

Vector-borne diseases, and in particular malaria, are of global importance because of the associated economic burden and high mortality rates, especially in developing countries [1–3]. Malaria is transmitted by Anopheles mosquitoes and is responsible for over half a million deaths annually [3]. The primary method for controlling this disease is vector management which includes the use of long-lasting insecticidal nets (LLINs), indoor residual spraying (IRS), and the management of mosquito breeding sites [4]. The application of these methods has significantly reduced transmission intensity and subsequently the malaria burden. However, the progress in malaria control has stalled suggesting that the current malaria control strategies have limitations in driving malaria cases to significantly low levels for successful elimination [3,5]. Therefore, the development of novel approaches is urgently needed to sustain the fight against malaria.

There are now several cases where it has been demonstrated that microbes associated with mosquito vectors can significantly alter their ability to efficiently transmit pathogens [6]. For instance, the vectorial capacity of the Aedes aegypti mosquito, which is responsible for the transmission of medically important viruses such as dengue, chikungunya, and zika, is significantly impaired in the presence of an intracellular endosymbiont, Wolbachia [6]. Based on this phenotype and the ability of Wolbachia to spread through insect populations, Wolbachia-based strategies have been developed and implemented to control dengue fever transmission successfully [7–9]. Similarly, it has been demonstrated that the association of malaria-transmitting mosquitoes and specific microbial symbionts reduces the ability of these vectors to transmit malaria-causing Plasmodium parasites [10–14]. The efficiency of malaria transmission-blocking and the stability of association with the vector are important parameters to consider when selecting a potential symbiont for malaria control. Notably, endosymbiotic microbes that have high-fidelity vertical transmission have an inherent ability to maintain their prevalence across mosquito generations and are therefore likely to be sustainable and cost-effective as malaria transmission blockers. Recently, Microsporidia MB, was discovered as an endosymbiont of Anopheles mosquitoes and shown to impair the transmission of the Plasmodium falciparum parasite by An. arabiensis mosquito [14]. The exact mechanisms involved in transmission blocking have not yet been characterized. The fastidious nature of endosymbionts presents challenges to establishing the precise molecular mechanisms behind pathogen protection. In other insect-symbionts systems where the mechanistic basis of protection has been elucidated, studies have shown that protection can involve the production of antagonistic compounds, competition for nutritional resources and space, and host immune priming [15]. Unlike many previously reported microsporidians which are pathogens of mosquitoes [16], Microsporidia MB does not have any apparent negative fitness effects on its host [14] and is located in a different group, clade IV, which is distinct from other microsporidia clades containing most of the known parasitic microsporidians of mosquitoes [17]. Microsporidia MB is vertically transmitted (mother to offspring) and horizontally transmitted (sexually) and therefore able to spread efficiently through mosquito populations [14,18]. Recently, we have shown that Microsporidia MB is predominantly localized in the reproductive organs which is relevant for vertical transmission from mother to offspring and horizontal transmission during mating [18–20]. However, the precise mechanisms involved in the vertical transmission of Microsporidia MB have not been elucidated. Understanding the strategies employed by this symbiont to vertically infect Anopheles mosquitoes will be important to determine the factors limiting infection rates and specificity to different malaria vector species and populations. Understanding these factors could facilitate strategies to sustainably increase Microsporidia MB infections in natural populations to significantly reduce malaria infections.

In this study, we investigated how the microsporidian symbiont is transferred from mother to offspring by studying its localization within the cells of the Anopheles mosquito ovary. We report that Microsporidia MB is present in the germarium and infects the germline stem cells from where it enters the cystocytes as a result of stem cell division. As the cystocytes develop into follicles, Microsporidia MB infection intensities increase first in the nurse cells and then in oocytes through active symbiont cell division. The average rate of transmission from stem cells to follicles (>96%) is higher than the average rate of observed adult to adult vertical transmission, suggesting that some Anopheles mosquitoes lose their symbiont infections over development.

Results

Microsporidia MB is vertically transmitted in the An. arabiensis germarium

Microsporidia MB was localized in An. arabiensis female ovaries using fluorescence in situ hybridization (FISH) microscopy with a Microspordia MB-specific probe [14]. An. arabiensis ovaries comprise tens to hundreds of polytrophic ovarioles with each ovariole composed of a germarium, the secondary, and the primary follicles. The germarium is located at the distal tip of the ovariole and contains 2–4 germline stem cells (GSCs). As each GSC divides, it produces an identical GSC and differentiating germline cell called a cystocyte. As a cystocyte progresses through the germarium, it undergoes mitotic divisions in combination with incomplete cytokinesis resulting in a follicle that has seven nurse cells connected through cytoplasmic bridges and one oocyte (Fig 1A and 1B). Through the detection of newly synthesized DNA, we found evidence of newly formed cystocytes following stem cell divisions in the germarium of newly emerged two-day-old mosquitoes (Fig 1C). We further demonstrated the localization of Microsporidia MB in the germarium (Figs 1D–1F and S1), inside the germline stem cells from where it is transferred to cystocytes during cell division. We also found Microsporidia MB infections in the secondary and primary follicles (Figs 1D–1F and S1). In the primary follicles, the symbiont was present in the developing oocytes and the nurse cells in high abundance but not in the follicular epithelial cells (Figs 1D–1F and S1).

Download:

Fig 1. Microsporidia MB localization in the polytrophic An. arabiensis ovariole.

(A) Schematic representation of the ovariole structure showing the germarium, the secondary follicle, and the primary follicle (with the oocyte and nurse cells in one chamber surrounded by the epithelial follicular cells). The germline stem cells (GSCs) and cystocytes are located in the germarium. (B & C) Images of a Microsporidia MB negative control ovariole obtained from a two-day-old An. arabiensis mosquito showing the localization of stem cell daughter cells, cystocytes (in blue), following cell divisions within the germarium (the blue signal represents newly synthesized DNA of cystocytes as detected by Click-iT Edu staining). (D-F) Images of a Microsporidia MB-infected An. arabiensis ovariole showing the distribution of the symbiont within the ovariole. Microsporidia MB is present in the germarium (germ) inside the GSCs (white arrowheads pointing to the GSC nucleus) and cystocytes (white arrow pointing to the cystocyte nucleus), the secondary follicle (SF), and the primary follicle (PF) i.e., the nurse cells (nc) and the oocyte (Oc). (Green represents Sytox Green DNA staining, image D, red represents Microsporidia MB-specific FISH probe staining, Image E, and a merged image of the green and red channel, Image F). Scale = 10μm. (Images are representative pictures of >100 independent observations).

https://doi.org/10.1371/journal.ppat.1012340.g001

Microsporidia MB intensities increase in the oocyte during egg development and maturation

Differentiation of the primary follicle into a mature egg within the ovariole in Anopheline mosquitoes is characterized by seven stages; stages I-III (previtellogenesis) and stages IV-VII (vitellogenesis). The vitellogenic stages IV-VII only occur after the female mosquito takes a blood meal. To determine Microsporidia MB growth dynamics in the ovariole during development, we performed FISH microscopy for ovarioles at different stages of development. We analyzed Microsporidia MB localization patterns in the germarium, the secondary follicle, and the primary follicle and found them to be infected across all ovariole developmental stages (Figs 2A, a-g and S2A). In the early previtellogenic stages (stages I and II), the symbiont was evenly distributed in the nurse cells and oocyte of the primary follicle (Fig 2A, a and b). In the late previtellogenic stage and vitellogenic (stages III-VI), however, Microsporidia MB accumulated in the developing oocyte but decreased in the nurse cells (Fig 2A, c-f). Once the egg developed to maturity and established a typical cigar shape following the degeneration of the nurse cells, an even distribution of Microsporidia MB cells throughout the egg was observed (Fig 2A, g).

Download:

Fig 2. Localization of Microsporidia MB within the germarium and follicles across the different developmental stages of An. arabiensis ovariole.

(A) Images showing Microsporidia MB localization in the germarium, secondary, and primary follicles in all ovariole developmental stages. Microsporidia MB is spread throughout the primary follicle during the previtellogenic phase of development (stages I-III). As development progresses to the vitellogenic phase (stages III-VI), microsporidia symbiont cells accumulate in the anterior end of the primary follicle oocyte in high densities. The upper panels (a-g) represent the Sytox Green DNA channel signal, the second row of panels (a’-g’) represents Microsporidia MB-specific CY-5 channel signal and the panels in the third row (a”-g”) represent merged images of both signals. Scale = 25μm. (B) The relative area of the primary follicle oocyte occupied by Microsporidia MB symbiont as measured using the Sytox Green DNA channel signal. Microsporidia MB occupies a small area of the primary follicle oocyte in the previtellogenic stages of development and increases significantly during the vitellogenic stages of development (χ² [5] = 57.13, p < 0.01).

https://doi.org/10.1371/journal.ppat.1012340.g002

We also quantified relative Microsporidia MB infection intensities in the oocytes of ovarioles at different developmental stages by measuring the area covered by Microsporidia MB signal (based on SytoxGreen general DNA staining and Microsporidia MB-specific probe staining) in relation to the entire area covered by the primary follicle oocyte. Both measurements showed low Microsporidia MB infection intensities in previtellogenic stage oocytes (Figs 2B and S2B). However, the infection intensities exponentially increased during vitellogenic stages of ovariole development. (χ² [5] = 57.13, p < 0.01). Previously, we observed a similar increase in Microsporidia MB infection intensities based on qPCR quantifications in An. arabiensis vitellogenic ovaries following blood feeding [20]. This increase of Microsporidia MB intensities inside the primary follicle oocyte suggests active proliferation of the symbiont or its deposition from the nurse cells.

Microsporidia MB lifecycle and proliferation in the oocyte

We observed an increase of Microsporidia MB intensities in oocytes of the primary follicles across the vitellogenic developmental stages (Fig 2). To establish how Microsporidia MB intensities increase in the developing primary follicles, we investigated the physiological states of Microsporidia MB cells in the oocyte and compared them to typical microsporidian developmental stages which are mainly divided into three stages i.e., infective, proliferative, and sporogonic stages (Fig 3A) [21]. The infective, also known as the environmental phase, is the only extracellular part of the cycle where microsporidia exist only as inactive spores until they inject their sporoplasm into a host cell to establish an infection. The proliferative stage involves the intracellular division of microsporidia cells through the process of merogony. The cells then enter the sporogonic developmental stage to form spores that are eventually released into the environment. Using the Microsporidia MB-specific FISH probe, we identified different sizes of Microsporidia MB cells inside the primary follicles ranging from smaller cells with a size of about 2μm to larger cells with a size of about 5μm (Fig 3B–3D and S3 Fig) suggesting that the symbiont exists in multiple forms in the follicle. We also observed proliferative Microsporidia MB cells in the process of fission (merogony) in the oocyte (Fig 3E–3G), which indicated that the symbiont was actively proliferating within the developing oocyte. The pattern of Microsporidia MB development in the Anopheles oocyte is consistent with merogony since intracellular proliferation was observed and spores were not detected. To confirm Microsporidia MB proliferation in the ovariole, we stained for actively dividing Microsporidia MB cells in infected ovarioles using Click-iT EdU cell proliferation assay that detects a nucleus with newly synthesized DNA following cell division. Microsporidia MB nuclei containing newly synthesized DNA can be differentiated from the mosquito’s newly synthesized nuclei (as observed in Fig 1C) because of its small size which is consistent with the Sytox Green DNA staining. Using the Click-iT EdU cell proliferation assay we were able to detect new Microsporidia MB cells mostly in the oocyte of the primary follicle but also the secondary follicle and the germarium (Fig 3H–3J) of infected ovarioles.

Download:

Fig 3. Microsporidia MB development in the ovariole.

(A) A typical microsporidia lifecycle showing the three microsporidian developmental stages i.e., the infective stage, proliferative, and sporogonic stages (figure adapted from [21]). (B-H) Confocal microscopy images showing the developmental stages of Microsporidia MB observed in infected An. arabiensis ovarioles. (B-D) Images of an ovariole with different sizes of Microsporidia MB cells ranging from 2μm in the germarium and secondary follicle (white arrowhead) to 5μm in the nurse cells and oocyte of the primary follicle, green and blue arrowheads, respectively. (E-G) Images showing Microsporidia MB cells in the merogony stage of proliferation in the oocyte (white arrowheads) and the presence of newly synthesized DNA in actively proliferating cells (H-J). The first column (panels B, E, and H) in green represents the Sytox Green DNA staining signal, the second column (panels C and F) in red represents the Microsporidia MB-specific CY-5 staining signal while (panel I) in blue represents Click-iT Edu staining signal and the third column (panels D, G, J) represent merged images of two channels. Scale = 10μm.

https://doi.org/10.1371/journal.ppat.1012340.g003

Vertical transmission rate of Microsporidia MB into primary follicles

We quantified Microsporidia MB infection rates of primary follicles in the ovaries to establish the vertical transmission rate of Microsporidia MB into developing eggs in infected ovaries. We found that a small number of primary follicles (3%) were not infected with Microsporidia MB while a majority (97%) of the primary follicles in infected ovaries had Microsporidia MB (Fig 4A). Interestingly, in ovarioles whose primary follicles were not infected with Microsporidia MB (3%), the symbiont was consistently present in the germarium and the secondary follicles (Fig 4B) suggesting that early stem cell divisions could occasionally not transfer Microsporidia MB to the follicles. To establish whether this phenomenon could explain the imperfect vertical transmission rate (45–100%) of Microsporidia MB [10], we compared the primary follicle infection rate and Microsporidia MB prevalence in sugar-fed adult F1 offspring (males and females) and blood-fed adult F1 offspring (females) obtained from Microsporidia MB-infected isofemale mothers. We found that whereas 97% of the primary follicles were infected with Microsporidia MB, only 61% of sugar-fed adult F1 offspring and 63% of blood-fed adult F1 females were infected with Microsporidia MB (Fig 4A), suggesting that Microsporidia MB prevalence is not influenced by nutritional status in adults and is likely lost during pre-imaginal stages of development. In all Microsporidia MB-infected An. arabiensis female gonads we investigated, we found the symbiont to be localized in both ovaries indicating even distribution of the symbiont in infected gonads (Fig 4C).

Download:

Fig 4. Microsporidia MB infection rates in primary follicles and vertical transmission (VT) rates.

Microsporidia MB distribution in infected female ovaries was investigated and compared to vertical transmission rates. (A-B) A majority (97%) of the primary follicles within infected ovarioles had Microsporidia MB while the symbiont was completely absent in 3% of the primary follicles (B, white arrowhead). However, the germarium (germ) and secondary follicles (SF) of the 3% primary follicles that lacked Microsporidia MB were consistently infected with Microsporidia MB. Microsporidia MB was detected in 61% of sugar-fed (male & female) and 63% of blood-fed (female) adult offspring of Microsporidia MB-infected mothers. (C) Microsporidia MB was found to be present in both ovaries of infected female gonads investigated. Green represents Sytox Green general DNA staining while red represents Microsporidia MB-specific CY-5 probe staining.

https://doi.org/10.1371/journal.ppat.1012340.g004

Discussion

Successful deployment of a novel Microsporidia MB-based approach for controlling malaria will depend on the ability of the microsporidian symbiont to invade, spread, and be maintained in target An. arabiensis vector populations. While Microsporidia MB has high rates of maternal transmission in An. arabiensis, these rates are variable. The rate of maternal transmission is critical for determining the prevalence of Microsporidia MB in a population of Anopheles mosquitoes and the impact on Plasmodium [22]. In this study, we established the mechanistic basis of Microsporidia MB vertical transmission, which occurs through the acquisition of Microsporidia MB by Anopheles cystocyte progenitors from the germline stem cells following cell division. As the daughter stem cell cystocytes differentiate into follicles, Microsporidia MB establishes itself in the nurse cells and ultimately the oocyte and mature egg. Our findings suggest that Microsporidia MB becomes established in the Anopheles germline early during development and this seems to be highly efficient, as only 3% of primary follicles do not acquire a Microsporidia MB infections. We didn’t find any evidence of the germline becoming re-infected from somatic cell infections, but we do not rule out that this could be occurring. In particular, we have previously observed that Microsporidia MB infections acquired via sexual transmission can subsequently be vertically transmitted [18]. Vertical transmission of microsporidian symbionts through egg infections has been reported in different species of mosquitoes and in the amphipod Gammarus duebeni [23–27]. However, symbiont infections in these hosts were only observed in the oocyte and epithelial follicular cells but not in the germline stem cells suggesting that the mechanism of vertical transmission is through somatic cell infections and not germline stem cell infections as we found for Microsporidia MB. Somatic cell infections and subsequent transovarial transmission has been demonstrated for Edhazardia aedis in Aedes aegypti and Amblyospora sp. in Aedes cantator, Aedes triseriatus and Culex salinarius when horizontally (orally) acquired spores infect oenocytes and spread to the ovaries where they proliferate and form transovarial spores [16,23,25,28]. Similarly, vertical transmission of a microsporidian symbiont of Gammarus duebeni occurs when the symbiont invades and proliferates in follicular cells before spreading into the developing oocytes for transmission to offspring [24].

The mechanistic basis of Microsporidia MB transmission in An. arabiensis is strikingly similar to Wolbachia infections in other insects including D. melanogaster and Aedes mosquito species where the endosymbiont is in the germline stem cells and is vertically transferred to the progeny following stem cell divisions [12,29–31]. For Wolbachia, there is apparently an additional vertical transmission route, which involves infection of the follicle from the surrounding somatic cells. This additional infection route may be important for maintaining near-perfect rates of vertical transmission, which are required for an endosymbiont that is entirely dependent on vertical transmission [31,32]. In this study, Microsporidia MB was not observed in the somatically derived epithelial follicular cells that surround the egg chamber. Wolbachia infections of the germline stem cells of Drosophila are capable of influencing mitotic processes to promote more egg production [33,34]. Although Microsporidia MB does not apparently affect egg production in An. arabiensis [14], its high infection intensity in the germline stem cells and developing oocytes may have consequences for cellular processes and embryonic development. It is notable that the mechanistic basis of Microsporidia MB vertical transmission is very different from another insect endosymbiont, Spiroplasma. Spiroplasma is absent from the germline stem cells but gains access to the vitellogenic oocyte during vitellogenesis by co-opting the yolk transport system [35]. The existence of Microsporidia MB in the stem cells and oocyte of developing eggs in An. arabiensis provides strong evidence for a highly specialized vertical transmission strategy for Microsporidia MB in An. arabiensis. This high level of specialization may explain why Microsporidia MB has so far only been found in Anopheles mosquitoes [18].

Similar to other microsporidians, Microsporidia MB is a unicellular obligate intracellular organism that can only replicate and survive inside the host cells and only exist as spores outside the host cells [36]. Our findings demonstrate that cystocytes, nurse cells, and oocytes are among the suitable host cell types for this symbiont within the An. Arabiensis host. We identified different sizes of Microsporidia MB cells in the ovariole suggesting that Microsporidia MB is transitioning through different life cycle stages in this tissue. Further, we found evidence that An. arabiensis ovarioles provide an optimal environment for the proliferative phase of Microsporidia MB development. We observed that this microsporidian symbiont actively proliferates inside the primary follicle oocyte through the detection of Microsporidia MB cells undergoing merogony and the presence of newly synthesized Microsporidia MB DNA. This proliferation was observed after mosquitoes had a successful blood meal suggesting that Microsporidia MB proliferation is coordinated with host cellular processes. It is notable that in a typical microsporidian lifecycle, sporogeny follows merogony. In the case of Microsporidia MB, it is not clear if sporogeny plays a role in vertical transmission since spores enable survival in the environment and invasion of cells and we observe that vertical transmission occurs without Microsporidia MB leaving an intracellular environment. Sporulation of Amblyospora species of microsporidia in Culex salinarius and Culiseta incidens only occurs after the mosquitoes take a blood meal and it coincides with the maturation of the oocytes [23,37]. The development of the microsporidian symbiont of Gammarus duebeni is also highly coordinated with the host reproductive cycle [24]. The controlled proliferation of symbionts within insect hosts is important in mitigating any fitness costs to the host [24,38,39]. Unlike other pathogenic microsporidian symbionts that kill their hosts [36], Microsporidia MB has no apparent fitness costs to the host despite attaining high infection intensities in the oocyte of the developing egg [14]. For vertically transmitted symbionts, there’s a strong selection for avirulence to the host because the symbiont’s success is associated with the host’s reproductive success [40,41]. Therefore, the specialized vertical transmission of Microsporidia MB through infection of oocytes of developing eggs of An. arabiensis suggests that avirulence has been selected for during evolution leading to its low pathogenicity to the host.

The presence of the microsporidian symbiont in most (97%) of the developing eggs implies a potential for high vertical transmission rates. However, in our experiments, we observed an average of 61% and 63% vertical transmission rates from infected mothers to sugar-fed and blood-fed adult F1s, respectively, which was lower than the infection rate in eggs that were developing in the ovary. Previously, we reported marginally higher transmission rates of Microsporidia MB from infected mothers to their offspring [14]. The absence of the symbiont in some of the primary follicles only partially explains the imperfect vertical transmission rate. Additionally, other factors such as the ability to gain access to target host cells and tissues (possibly the germline stem cells), host immune responses, and environmental factors during embryogenesis and larval stages may be responsible for the loss of infections during mosquito development. As demonstrated in other insect-symbiont associations [42–44], the presence of Microsporidia MB may elicit An. arabiensis’ immune response which then directly interferes with the establishment of the symbiont before it colonizes the primary sites of infection where it may be protected from the host’s immune responses. Also, environmental factors such as temperature and humidity that are known to influence the occurrence and transmission of different endosymbionts including microsporidian symbionts in other insects are likely to affect the establishment of Microsporidia MB [45–47].

Our study has shown a specialized vertical transmission route of Microsporidia MB in An. arabiensis that is relevant in developing a symbiont-based strategy to control malaria. These findings will serve as a basis for investigations on Microsporidia MB transmission phenotypes in Microsporidia MB found in diverse geographical regions and Anopheline vector species aimed at maximizing Microsporidia MB vertical transmission rates for the development of a viable strategy for symbiont-based malaria transmission blocking.

Methods

Mosquito collection and rearing

Gravid An. gambiae s.l mosquitoes were collected from the Ahero irrigation scheme in Kenya in 2022 and 2023 and transported to icipe’s Duduville campus in Nairobi Kenya. The mosquitoes were then maintained in the insectary at 27 ± 2.5°C, humidity 60–80%, 12-h day and 12-h night cycles and induced to oviposit through the provision of oviposition cups with a wet cotton wool lined with a filter paper. After ovipositing, DNA was extracted from the females and used for species identification and screening for Microsporidia MB infections using PCR [14]. Offspring from Microsporidia MB-infected and uninfected female An. arabiensis mosquitoes were then reared in the insectary as described above for experimental investigations. Blood feeding of mosquitoes to initiate the processes of vitellogenesis and oogenesis was conducted as previously described [20].

Fluorescence in situ hybridization (FISH) staining protocol

FISH was conducted to localize Microsporidia MB in the ovarioles of newly emerged 2-day-old and 2–5 days post-blood-fed An. arabiensis mosquitoes. Ovaries of Microsporidia MB-infected and uninfected (controls) were dissected in 1x PBS and fixed overnight in 4% Paraformaldehyde (PFA) at 4°C then rehydrated in 50% ethanol for 30 minutes, followed by a washing step in 1x PBS for 30 minutes. FISH was conducted as previously described to localize Microsporidia MB within the ovarioles [14,20]. Briefly, hybridization was done by incubating the tissues at 50˚C overnight in 100μl of hybridization mix (hybridization buffer i.e., dH2O, 5M NaCl, 1M Tris/HCl [pH = 8], and 10% SDS), 0.5μM final concentration of the Microsporidia MB specific CY5 probe (5′-CY5-CCCTGTCCACTATACCTAATGAACAT-3′) [10], and 0.025μM Sytox Green general DNA staining. After staining, the samples were washed twice with 100μl of wash buffer prewarmed at 50˚C (dH2O, 5M NaCl, 1M Tris/HCl [pH = 8], 0.5M EDTA, and 10% SDS). The tissues were subsequently mounted on glass slides using fluoromount mounting medium for confocal microscopy.

Click-iT EdU staining of actively dividing cells in An. arabiensis germarium

Ovaries of An. arabiensis females were dissected in 10μM EdU in 1x PBS solution and labeled in this solution for one hour before they were fixed in a 3.7% PFA solution for 30 minutes. Subsequently, they were washed twice with 500μL of 3% BSA in 1x PBS and permeabilized using 0.5% Triton X-100 in 1x PBS for 20 minutes at room temperature before washing twice with 500μL of 3% BSA in 1x PBS solution. The ovaries were then stained and protected from light for 2 hours in a freshly prepared 500μL Click-iT Plus reaction cocktail prepared as per the manufacturer’s specifications. Once the Click-iT staining process was complete, the tissues were washed in with 500μL BSA in 1x PBS solution and counterstained with Sytox Green fluorescent general DNA stain (5 μg/ml) before mounting on a slide using fluoromount mounting medium for imaging.

Confocal microscopy and image processing

The FISH and Click-iT EdU stained tissues were visualized immediately after mounting using a Leica SP5 confocal microscope (Leica Microsystems, USA), and images were taken using LAS AF software (Leica Microsystems, USA). Each stain was detected and images were captured independently using unique laser wavelength settings that were standardized across all samples. The images were analyzed using the ImageJ 1.50i software package [31] to identify Microsporidia MB and the host. Microsporidia MB was detected and quantified using a Microsporidia MB specific CY-5 FISH probe that targeted Microsporidia MB 18s rDNA and rRNA (red signal) while Microsporidia MB and An. arabiensis DNA in the nuclei was identified using Sytox Green general DNA stain (green signal). Based on multiple examinations of general DNA staining of Microsporidia MB infected and uninfected tissues, we identified the host cell nucleus from Microsporidia MB nucleus based on their size, location, and shape. This method has been used to identify Wolbachia in infected insect tissues [48]. Additionally, in all Microsporidia MB infected samples, there was a Sytox Green general DNA channel signal within the Microsporidia MB specific CY-5 FISH probe channel signal in the cell cytoplasm. This was used to establish the size and location of the Microsporidia MB nucleus and distinguish it from An. arabiensis nucleus. Subsequently, the distinct and consistent small size of Microsporidia MB nucleus in the cytoplasm as determined by the Sytox Green channel signal and Microsporidia MB specific probe staining was then used to distinguish between actively dividing Microsporidia MB and An. arabiensis cells after Click-iT Edu staining that targets nuclei with newly synthesized DNA.

Quantification of Microsporidia MB in An. arabiensis ovaries

ImageJ software was used to measure the area occupied by Microsporidia MB in the primary follicle oocyte by measuring the fluorescence intensity signal captured by the Microsporidia MB-specific probe channel. Measurements based on the Sytox Green general DNA channel were also performed to avoid overestimating infection intensities using the Microsporidia MB-specific probe because it targets both 18s rDNA and 18s rRNA and therefore likely to be influenced by 18s rRNA expression patterns depending on the stage of Microsporidia MB development. The images acquired using the Microsporidia MB-specific probe channel and Sytox Green DNA channel were independently processed in ImageJ and then converted into an 8-bit grayscale binary scale. The regions of interest (ROI) around each of these structures were defined using the polygon selection tool in ImageJ based on the position of the follicular epithelial cells so that only the pixels within the area of interest were selected and included in the final measurements. Subsequently, we measured the proportion of the area that was occupied by Microsporidia MB cells within each of the ROIs as previously described [49]. The area occupied by the host nucleus was also calculated and removed from the total area obtained when the analysis was performed using the Sytox Green DNA signal channel that detects both Microsporidia MB and An. arabiensis DNA. Images where the symbiont and host nuclei staining were not clearly distinguishable were not included in the analysis. The relative oocyte area covered by Microsporidia MB across the different stages of egg development was then compared using Kruskal-Wallis H test. Rstudio software (2023.12.1.402) was used for statistical analysis [50].

Microsporidia MB infection rates in the ovaries

To determine Microsporidia MB infection rates in the ovaries, a complete set of eight ovaries was obtained from select offspring of wild-caught Microsporidia MB-infected mothers that were collected from different locations within Ahero. The ovaries were stained by FISH as described above and the presence or absence of Microsporidia MB in both ovaries was recorded. Microsporidia MB infection rates in the primary follicles were determined by randomly selecting and counting the number of infected and uninfected follicles in the eight pairs of ovaries.

Vertical transmission rate of Microsporidia MB

Field-collected gravid females were induced to oviposit as described above and then screened for Microsporidia MB infections using PCR [14]. Eggs from Microsporidia MB-infected mothers were hatched in the insectary and maintained at a temperature of 27 degrees and the larvae were fed on tetramin fish food until they pupated. The isofemale line pupae were subsequently transferred into cages where they emerged. We then performed two experiments to determine the vertical transmission rates. In one experiment, emerged F1s from 32 isofemale lines were maintained on a 10% sugar diet for two days and then immediately screened for Microsporidia MB. In another experiment, emerged F1s from 5 isofemale lines were maintained on a 10% sugar diet for two days and then given a blood meal for two consecutive days as previously described [20] before they were screened for Microsporidia MB 3 days post blood feeding. DNA was extracted from these two sets of experiments and used to screen for Microsporidia MB with PCR and qPCR as previously described [14,20]. We used controls, a plasmid construct with low copy number concentrations (10–100 copies) of the target gene to determine the detection limits of our qPCR protocol to ensure that low Microsporidia MB infections were measured. The number of infected adult offspring per isofemale line was expressed as a percentage.

Supporting information

S1 Fig. Representative microscopic images of Microsporidia MB localized in An. arabiensis ovariole.

(A) Microsporidia MB negative control ovariole of a two-day-old An. arabiensis mosquito showing the localization of stem cell daughter cells, cystocytes, (in blue as detected by Click-iT Edu staining) within the germarium. (B) Image of a Microsporidia MB infected ovarioles of An. arabiensis mosquito showing localization of Microsporidia MB in the germline stem cells (white arrowheads) and daughter cystocyte cells (white arrows) in the germarium (germ), secondary follicles (SF) and the nurse cells (nc) and oocyte (oc) of the primary follicle (PF). (Green represents Sytox Green DNA staining, red represents Microsporidia MB-specific FISH probe staining). Scale = 10μm.

https://doi.org/10.1371/journal.ppat.1012340.s001

(TIF)

S2 Fig. Representative microscopic images of Microsporidia MB localization during egg development.

(A-G) Images of microscopic observations of Microsporidia MB in ovariolesduring egg maturation. (B) The relative area of the primary follicle oocyte occupied by Microsporidia MB symbiont as measured using the Microsporidia MB specific CY5 probe signal. Microsporidia MB occupies a small area of the primary follicle oocyte in the previtellogenic stages of development and increases significantly during the vitellogenic stages of development as observed with the Sytox Green DNA signal (χ² [5] = 75.26, p < 0.01). Scale = 10μm.

https://doi.org/10.1371/journal.ppat.1012340.s002

(TIF)

S3 Fig. Representative images of microscopic observations showing the different sizes of Microsporidia MB cells existing in the ovarioles ranging from 2μm in the germarium, secondary follicle, and oocyte (blue arrowheads) to 5μm in the nurse cells and oocyte of the primary follicle (white arrows).

Green represents Sytox Green DNA staining signal, Red represents the Microsporidia MB-specific CY-5 staining signal, and blue Click-iT Edu staining signal. Scale = 10μm.

https://doi.org/10.1371/journal.ppat.1012340.s003

(TIF)

Acknowledgments

We acknowledge the support provided by the icipe insectary team (Jeniffer Thiong’o, Peris Wambui), fieldwork facilitation by Gerald Ronoh, and project administration by Faith Kyengo.

References

1. Bhatt S, Gething PW, Brady OJ, Messina JP, Farlow AW, Moyes CL, et al. The global distribution and burden of dengue. Nature. 2013;496(7446). pmid:23563266
- View Article
- PubMed/NCBI
- Google Scholar
2. Lessler J, Chaisson LH, Kucirka LM, Bi Q, Grantz K, Salje H, et al. Assessing the global threat from Zika virus. Vol. 353, Science. 2016. pmid:27417495
- View Article
- PubMed/NCBI
- Google Scholar
3. WHO. World Malaria Report 2022. World Health Organization. 2022.
4. Nalinya S, Musoke D, Deane K. Malaria prevention interventions beyond long-lasting insecticidal nets and indoor residual spraying in low- and middle-income countries: a scoping review. Malar J. 2022;21(1). pmid:35109848
- View Article
- PubMed/NCBI
- Google Scholar
5. Epstein A, Maiteki-Sebuguzi C, Namuganga JF, Nankabirwa JI, Gonahasa S, Opigo J, et al. Resurgence of malaria in Uganda despite sustained indoor residual spraying and repeated long lasting insecticidal net distributions. PLOS Global Public Health. 2022;2(9). pmid:36962736
- View Article
- PubMed/NCBI
- Google Scholar
6. Moreira LA, Iturbe-Ormaetxe I, Jeffery JA, Lu G, Pyke AT, Hedges LM, et al. A Wolbachia symbiont in Aedes aegypti limits infection with Dengue, Chikungunya, and Plasmodium. Cell. 2009;139(7). pmid:20064373
- View Article
- PubMed/NCBI
- Google Scholar
7. Nazni WA, Hoffmann AA, NoorAfizah A, Cheong YL, Mancini M V., Golding N, et al. Establishment of Wolbachia strain wAlbB in Malaysian populations of Aedes aegypti for Dengue control. Current Biology. 2019;29(24). pmid:31761702
- View Article
- PubMed/NCBI
- Google Scholar
8. Ribeiro dos Santos G, Durovni B, Saraceni V, Souza Riback TI, Pinto SB, Anders KL, et al. Estimating the effect of the wMel release programme on the incidence of dengue and chikungunya in Rio de Janeiro, Brazil: a spatiotemporal modelling study. Lancet Infect Dis. 2022;22(11). pmid:36182679
- View Article
- PubMed/NCBI
- Google Scholar
9. Indriani C, Tanamas SK, Khasanah U, Ansari MR, Rubangi Tantowijoyo W, et al. Impact of randomised wmel Wolbachia deployments on notified dengue cases and insecticide fogging for dengue control in Yogyakarta City. Glob Health Action. 2023;16(1). pmid:36700745
- View Article
- PubMed/NCBI
- Google Scholar
10. 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 (1979). 2011;332(6031):855–8. pmid:21566196
- View Article
- PubMed/NCBI
- Google Scholar
11. 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
- View Article
- PubMed/NCBI
- Google Scholar
12. Bian G, Joshi D, Dong Y, Lu P, Zhou G, Pan X, et al. Wolbachia invades Anopheles stephensi populations and induces refractoriness to Plasmodium infection. Science (1979). 2013;340(6133):748–51. pmid:23661760
- View Article
- PubMed/NCBI
- Google Scholar
13. 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). pmid:33958765
- View Article
- PubMed/NCBI
- Google Scholar
14. Herren JK, Mbaisi L, Mararo E, Makhulu EE, Mobegi VA, Butungi H, et al. A microsporidian impairs Plasmodium falciparum transmission in Anopheles arabiensis mosquitoes. Nat Commun. 2020 May 4;11(1):2187. pmid:32366903
- View Article
- PubMed/NCBI
- Google Scholar
15. Gerardo NM, Parker BJ. Mechanisms of symbiont-conferred protection against natural enemies: An ecological and evolutionary framework. Curr Opin Insect Sci. 2014;4(1):8–14. Available from: pmid:28043411
- View Article
- PubMed/NCBI
- Google Scholar
16. Becnel JJ, White SE, Shapiro AM. Review of microsporidia-mosquito relationships: From the simple to the complex. In: Folia Parasitologica. 2005. https://doi.org/10.14411/fp.2005.006
17. Vossbrinck CR, Debrunner-Vossbrinck BA. Molecular phylogeny of the Microsporidia: Ecological, ultrastructural and taxonomic considerations. In: Folia Parasitologica. 2005. https://doi.org/10.14411/fp.2005.017 pmid:16004372
18. Nattoh G, Maina T, Makhulu EE, Mbaisi L, Mararo E, Otieno FG, et al. Horizontal transmission of the symbiont Microsporidia MB in Anopheles arabiensis. Front Microbiol. 2021;12:647183. pmid:34394019
- View Article
- PubMed/NCBI
- Google Scholar
19. Nattoh G, Onyango B, Makhulu EE, Omoke D, Ang’ang’o LM, Kamau L, et al. Microsporidia MB in the primary malaria vector Anopheles gambiae sensu stricto is avirulent and undergoes maternal and horizontal transmission. Parasit Vectors. 2023 Sep 25;16(1):335. pmid:37749577
- View Article
- PubMed/NCBI
- Google Scholar
20. Makhulu EE, Onchuru TO, Gichuhi J, Otieno FG, Wairimu AW, Muthoni JN, et al. Localization and tissue tropism of the symbiont Microsporidia MB in the germ line and somatic tissues of Anopheles arabiensis. mBio. 2024;15(1). pmid:38063396
- View Article
- PubMed/NCBI
- Google Scholar
21. Cali A, Becnel JJ, Takvorian PM. Microsporidia. In: Handbook of the Protists. Cham: Springer International Publishing; 2017. p. 1559–618. https://doi.org/10.1007/978-3-319-28149-0_27
22. Mfangnia CNT, Tonnang HEZ, Tsanou B, Herren J. Mathematical modelling of the interactive dynamics of wild and Microsporidia MB-infected mosquitoes. Mathematical Biosciences and Engineering. 2023;20(8). pmid:37679176
- View Article
- PubMed/NCBI
- Google Scholar
23. Andreadis TG, Hall DW. Development, ultrastructure, and mode of transmission of Amblyospora sp. (Microspora) in the mosquito. J Protozool. 1979;26(3).
- View Article
- Google Scholar
24. Terry RS, Dunn AM, Smith JE. Cellular distribution of a feminizing microsporidian parasite: A strategy for transovarial transmission. Parasitology. 1997;115(2). pmid:10190171
- View Article
- PubMed/NCBI
- Google Scholar
25. Becnel JJ, Hazard EI, Fukuda T. Fine structure and development of Pilosporella chapmani (Microspora: Thelohaniidae) in the mosquito, Aedes triseriatus (Say). J Protozool. 1986;33(1). pmid:3959008
- View Article
- PubMed/NCBI
- Google Scholar
26. Avery SW, Undeen AH. Horizontal transmission of Parathelohania anophelis to the copepod, Microcyclops varicans, and the mosquito, Anopheles quadrimaculatus. J Invertebr Pathol. 1990;56(1). pmid:2115902
- View Article
- PubMed/NCBI
- Google Scholar
27. Dunn AM, Adams J, Smith JE. Transovarial transmission and sex ratio distortion by a microsporidian parasite in a shrimp. J Invertebr Pathol. 1993;61(3).
- View Article
- Google Scholar
28. Andreadis TG. Life cycle and epizootiology of Amblyospora sp. (Microspora: Amblyosporidae) in the mosquito, Aedes cantator. J Protozool. 1983;30(3).
- View Article
- Google Scholar
29. Werren JH, Baldo L, Clark ME. Wolbachia: Master manipulators of invertebrate biology. Vol. 6, Nature Reviews Microbiology. 2008. pmid:18794912
- View Article
- PubMed/NCBI
- Google Scholar
30. Zouache K, Voronin D, Tran-Van V, Mousson L, Failloux AB, Mavingui P. Persistent Wolbachia and cultivable bacteria infection in the reproductive and somatic tissues of the mosquito vector Aedes albopictus. PLoS One. 2009;4(7). pmid:19633721
- View Article
- PubMed/NCBI
- Google Scholar
31. Hughes GL, Dodson BL, Johnson RM, Murdock CC, Tsujimoto H, Suzuki Y, et al. Native microbiome impedes vertical transmission of Wolbachia in Anopheles mosquitoes. Proc Natl Acad Sci U S A. 2014;111(34):12498–503. pmid:25114252
- View Article
- PubMed/NCBI
- Google Scholar
32. Xi Z, Khoo CCH, Dobson SL. Wolbachia establishment and invasion in an Aedes Aegypti laboratory population. Science (1979). 2005 Oct 14;310(5746):326–8. pmid:16224027
- View Article
- PubMed/NCBI
- Google Scholar
33. Fast EM, Toomey ME, Panaram K, Desjardins D, Kolaczyk ED, Frydman HM. Wolbachia enhance Drosophila stem cell proliferation and target the germline stem cell niche. Science (1979). 2011;334(6058). pmid:22021671
- View Article
- PubMed/NCBI
- Google Scholar
34. Toomey ME, Panaram K, Fast EM, Beatty C, Frydman HM. Evolutionarily conserved Wolbachia-encoded factors control pattern of stem-cell niche tropism in Drosophila ovaries and favor infection. Proc Natl Acad Sci U S A. 2013;110(26). pmid:23744038
- View Article
- PubMed/NCBI
- Google Scholar
35. Herren JK, Paredes JC, Schüpfer F, Lemaitre B. Vertical transmission of a Drosophila endosymbiont via cooption of the yolk transport and internalization machinery. mBio. 2013;4(2). pmid:23462112
- View Article
- PubMed/NCBI
- Google Scholar
36. Han B, Weiss LM. Microsporidia: Obligate intracellular pathogens within the fungal kingdom. Microbiol Spectr. 2017;5(2). pmid:28944750
- View Article
- PubMed/NCBI
- Google Scholar
37. Dickson DL, Barr AR. Development of Amblyospora campbelli (Microsporida: Amblyosporidae) in the mosquito Culiseta incidens (Thomson). J Protozool. 1990;37(2). pmid:1969479
- View Article
- PubMed/NCBI
- Google Scholar
38. Becnel JJ, Andreadis TG. Microsporidia in insects. Microsporidia: Pathogens of Opportunity: First Edition. 2014;521–70.
- View Article
- Google Scholar
39. Herren JK, Paredes JC, Schüpfer F, Arafah K, Bulet P, Lemaitre B. Insect endosymbiont proliferation is limited by lipid availability. Elife. 2014;3:e02964. pmid:25027439
- View Article
- PubMed/NCBI
- Google Scholar
40. Bull JJ, Molineux IJ, Rice WR. Selection of benevolence in a host-parasite system. Evolution (N Y). 1991;45(4). pmid:28564051
- View Article
- PubMed/NCBI
- Google Scholar
41. Ewald PW. Transmission modes and evolution of the parasitism‐mutualism continuum. Ann N Y Acad Sci. 1987;503(1). pmid:3304078
- View Article
- PubMed/NCBI
- Google Scholar
42. Maire J, Vincent-Monégat C, Masson F, Zaidman-Rémy A, Heddi A. An IMD-like pathway mediates both endosymbiont control and host immunity in the cereal weevil Sitophilus spp. Microbiome. 2018;6(1):6. pmid:29310713
- View Article
- PubMed/NCBI
- Google Scholar
43. Maire J, Vincent-Monégat C, Balmand S, Vallier A, Hervé M, Masson F, et al. Weevil pgrp-lb prevents endosymbiont TCT dissemination and chronic host systemic immune activation. Proc Natl Acad Sci U S A. 2019;116(12):5623–32. pmid:30819893
- View Article
- PubMed/NCBI
- Google Scholar
44. Kim JK, Han SH, Kim CH, Jo YH, Futahashi R, Kikuchi Y, et al. Molting-associated suppression of symbiont population and up-regulation of antimicrobial activity in the midgut symbiotic organ of the Riptortus-Burkholderia symbiosis. Dev Comp Immunol. 2014;43(1):10–4. pmid:24201132
- View Article
- PubMed/NCBI
- Google Scholar
45. Trzebny A, Nahimova O, Dabert M. High temperatures and low humidity promote the occurrence of microsporidians (Microsporidia) in mosquitoes (Culicidae). Parasit Vectors. 2024 Apr 11;17(1):187. pmid:38605410
- View Article
- PubMed/NCBI
- Google Scholar
46. Martins M, César CS, Cogni R. The effects of temperature on prevalence of facultative insect heritable symbionts across spatial and seasonal scales. Vol. 14, Frontiers in Microbiology. 2023. pmid:38143870
- View Article
- PubMed/NCBI
- Google Scholar
47. Cai W-J, Moriyama M, Fukastu T. High temperature treatment optimized for symbiont suppression in an obligatory gut bacterial symbiosis in the stinkbug Plautia stali. BioRxiv. 2024;
- View Article
- Google Scholar
48. Serbus LR, Sullivan W. A cellular basis for Wolbachia recruitment to the host germline. PLoS Pathog. 2007;3(12). pmid:18085821
- View Article
- PubMed/NCBI
- Google Scholar
49. Shihan MH, Novo SG, Le Marchand SJ, Wang Y, Duncan MK. A simple method for quantitating confocal fluorescent images. Biochem Biophys Rep. 2021;25. pmid:33553685
- View Article
- PubMed/NCBI
- Google Scholar
50. Posit team. RStudio: Integrated Development for R. 2024. [accessed 15 Oct 2024] Available from: http://www.posit.co/.

[ref1] 1. Bhatt S, Gething PW, Brady OJ, Messina JP, Farlow AW, Moyes CL, et al. The global distribution and burden of dengue. Nature. 2013;496(7446). pmid:23563266
View Article
PubMed/NCBI
Google Scholar

[2] View Article

[3] PubMed/NCBI

[4] Google Scholar

[ref2] 2. Lessler J, Chaisson LH, Kucirka LM, Bi Q, Grantz K, Salje H, et al. Assessing the global threat from Zika virus. Vol. 353, Science. 2016. pmid:27417495
View Article
PubMed/NCBI
Google Scholar

[6] View Article

[7] PubMed/NCBI

[8] Google Scholar

[ref3] 3. WHO. World Malaria Report 2022. World Health Organization. 2022.

[ref4] 4. Nalinya S, Musoke D, Deane K. Malaria prevention interventions beyond long-lasting insecticidal nets and indoor residual spraying in low- and middle-income countries: a scoping review. Malar J. 2022;21(1). pmid:35109848
View Article
PubMed/NCBI
Google Scholar

[11] View Article

[12] PubMed/NCBI

[13] Google Scholar

[ref5] 5. Epstein A, Maiteki-Sebuguzi C, Namuganga JF, Nankabirwa JI, Gonahasa S, Opigo J, et al. Resurgence of malaria in Uganda despite sustained indoor residual spraying and repeated long lasting insecticidal net distributions. PLOS Global Public Health. 2022;2(9). pmid:36962736
View Article
PubMed/NCBI
Google Scholar

[15] View Article

[16] PubMed/NCBI

[17] Google Scholar

[ref6] 6. Moreira LA, Iturbe-Ormaetxe I, Jeffery JA, Lu G, Pyke AT, Hedges LM, et al. A Wolbachia symbiont in Aedes aegypti limits infection with Dengue, Chikungunya, and Plasmodium. Cell. 2009;139(7). pmid:20064373
View Article
PubMed/NCBI
Google Scholar

[19] View Article

[20] PubMed/NCBI

[21] Google Scholar

[ref7] 7. Nazni WA, Hoffmann AA, NoorAfizah A, Cheong YL, Mancini M V., Golding N, et al. Establishment of Wolbachia strain wAlbB in Malaysian populations of Aedes aegypti for Dengue control. Current Biology. 2019;29(24). pmid:31761702
View Article
PubMed/NCBI
Google Scholar

[23] View Article

[24] PubMed/NCBI

[25] Google Scholar

[ref8] 8. Ribeiro dos Santos G, Durovni B, Saraceni V, Souza Riback TI, Pinto SB, Anders KL, et al. Estimating the effect of the wMel release programme on the incidence of dengue and chikungunya in Rio de Janeiro, Brazil: a spatiotemporal modelling study. Lancet Infect Dis. 2022;22(11). pmid:36182679
View Article
PubMed/NCBI
Google Scholar

[27] View Article

[28] PubMed/NCBI

[29] Google Scholar

[ref9] 9. Indriani C, Tanamas SK, Khasanah U, Ansari MR, Rubangi Tantowijoyo W, et al. Impact of randomised wmel Wolbachia deployments on notified dengue cases and insecticide fogging for dengue control in Yogyakarta City. Glob Health Action. 2023;16(1). pmid:36700745
View Article
PubMed/NCBI
Google Scholar

[31] View Article

[32] PubMed/NCBI

[33] Google Scholar

[ref10] 10. 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 (1979). 2011;332(6031):855–8. pmid:21566196
View Article
PubMed/NCBI
Google Scholar

[35] View Article

[36] PubMed/NCBI

[37] Google Scholar

[ref11] 11. 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
View Article
PubMed/NCBI
Google Scholar

[39] View Article

[40] PubMed/NCBI

[41] Google Scholar

[ref12] 12. Bian G, Joshi D, Dong Y, Lu P, Zhou G, Pan X, et al. Wolbachia invades Anopheles stephensi populations and induces refractoriness to Plasmodium infection. Science (1979). 2013;340(6133):748–51. pmid:23661760
View Article
PubMed/NCBI
Google Scholar

[43] View Article

[44] PubMed/NCBI

[45] Google Scholar

[ref13] 13. 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). pmid:33958765
View Article
PubMed/NCBI
Google Scholar

[47] View Article

[48] PubMed/NCBI

[49] Google Scholar

[ref14] 14. Herren JK, Mbaisi L, Mararo E, Makhulu EE, Mobegi VA, Butungi H, et al. A microsporidian impairs Plasmodium falciparum transmission in Anopheles arabiensis mosquitoes. Nat Commun. 2020 May 4;11(1):2187. pmid:32366903
View Article
PubMed/NCBI
Google Scholar

[51] View Article

[52] PubMed/NCBI

[53] Google Scholar

[ref15] 15. Gerardo NM, Parker BJ. Mechanisms of symbiont-conferred protection against natural enemies: An ecological and evolutionary framework. Curr Opin Insect Sci. 2014;4(1):8–14. Available from: pmid:28043411
View Article
PubMed/NCBI
Google Scholar

[55] View Article

[56] PubMed/NCBI

[57] Google Scholar

[ref16] 16. Becnel JJ, White SE, Shapiro AM. Review of microsporidia-mosquito relationships: From the simple to the complex. In: Folia Parasitologica. 2005. https://doi.org/10.14411/fp.2005.006

[ref17] 17. Vossbrinck CR, Debrunner-Vossbrinck BA. Molecular phylogeny of the Microsporidia: Ecological, ultrastructural and taxonomic considerations. In: Folia Parasitologica. 2005. https://doi.org/10.14411/fp.2005.017 pmid:16004372

[ref18] 18. Nattoh G, Maina T, Makhulu EE, Mbaisi L, Mararo E, Otieno FG, et al. Horizontal transmission of the symbiont Microsporidia MB in Anopheles arabiensis. Front Microbiol. 2021;12:647183. pmid:34394019
View Article
PubMed/NCBI
Google Scholar

[61] View Article

[62] PubMed/NCBI

[63] Google Scholar

[ref19] 19. Nattoh G, Onyango B, Makhulu EE, Omoke D, Ang’ang’o LM, Kamau L, et al. Microsporidia MB in the primary malaria vector Anopheles gambiae sensu stricto is avirulent and undergoes maternal and horizontal transmission. Parasit Vectors. 2023 Sep 25;16(1):335. pmid:37749577
View Article
PubMed/NCBI
Google Scholar

[65] View Article

[66] PubMed/NCBI

[67] Google Scholar

[ref20] 20. Makhulu EE, Onchuru TO, Gichuhi J, Otieno FG, Wairimu AW, Muthoni JN, et al. Localization and tissue tropism of the symbiont Microsporidia MB in the germ line and somatic tissues of Anopheles arabiensis. mBio. 2024;15(1). pmid:38063396
View Article
PubMed/NCBI
Google Scholar

[69] View Article

[70] PubMed/NCBI

[71] Google Scholar

[ref21] 21. Cali A, Becnel JJ, Takvorian PM. Microsporidia. In: Handbook of the Protists. Cham: Springer International Publishing; 2017. p. 1559–618. https://doi.org/10.1007/978-3-319-28149-0_27

[ref22] 22. Mfangnia CNT, Tonnang HEZ, Tsanou B, Herren J. Mathematical modelling of the interactive dynamics of wild and Microsporidia MB-infected mosquitoes. Mathematical Biosciences and Engineering. 2023;20(8). pmid:37679176
View Article
PubMed/NCBI
Google Scholar

[74] View Article

[75] PubMed/NCBI

[76] Google Scholar

[ref23] 23. Andreadis TG, Hall DW. Development, ultrastructure, and mode of transmission of Amblyospora sp. (Microspora) in the mosquito. J Protozool. 1979;26(3).
View Article
Google Scholar

[78] View Article

[79] Google Scholar

[ref24] 24. Terry RS, Dunn AM, Smith JE. Cellular distribution of a feminizing microsporidian parasite: A strategy for transovarial transmission. Parasitology. 1997;115(2). pmid:10190171
View Article
PubMed/NCBI
Google Scholar

[81] View Article

[82] PubMed/NCBI

[83] Google Scholar

[ref25] 25. Becnel JJ, Hazard EI, Fukuda T. Fine structure and development of Pilosporella chapmani (Microspora: Thelohaniidae) in the mosquito, Aedes triseriatus (Say). J Protozool. 1986;33(1). pmid:3959008
View Article
PubMed/NCBI
Google Scholar

[85] View Article

[86] PubMed/NCBI

[87] Google Scholar

[ref26] 26. Avery SW, Undeen AH. Horizontal transmission of Parathelohania anophelis to the copepod, Microcyclops varicans, and the mosquito, Anopheles quadrimaculatus. J Invertebr Pathol. 1990;56(1). pmid:2115902
View Article
PubMed/NCBI
Google Scholar

[89] View Article

[90] PubMed/NCBI

[91] Google Scholar

[ref27] 27. Dunn AM, Adams J, Smith JE. Transovarial transmission and sex ratio distortion by a microsporidian parasite in a shrimp. J Invertebr Pathol. 1993;61(3).
View Article
Google Scholar

[93] View Article

[94] Google Scholar

[ref28] 28. Andreadis TG. Life cycle and epizootiology of Amblyospora sp. (Microspora: Amblyosporidae) in the mosquito, Aedes cantator. J Protozool. 1983;30(3).
View Article
Google Scholar

[96] View Article

[97] Google Scholar

[ref29] 29. Werren JH, Baldo L, Clark ME. Wolbachia: Master manipulators of invertebrate biology. Vol. 6, Nature Reviews Microbiology. 2008. pmid:18794912
View Article
PubMed/NCBI
Google Scholar

[99] View Article

[100] PubMed/NCBI

[101] Google Scholar

[ref30] 30. Zouache K, Voronin D, Tran-Van V, Mousson L, Failloux AB, Mavingui P. Persistent Wolbachia and cultivable bacteria infection in the reproductive and somatic tissues of the mosquito vector Aedes albopictus. PLoS One. 2009;4(7). pmid:19633721
View Article
PubMed/NCBI
Google Scholar

[103] View Article

[104] PubMed/NCBI

[105] Google Scholar

[ref31] 31. Hughes GL, Dodson BL, Johnson RM, Murdock CC, Tsujimoto H, Suzuki Y, et al. Native microbiome impedes vertical transmission of Wolbachia in Anopheles mosquitoes. Proc Natl Acad Sci U S A. 2014;111(34):12498–503. pmid:25114252
View Article
PubMed/NCBI
Google Scholar

[107] View Article

[108] PubMed/NCBI

[109] Google Scholar

[ref32] 32. Xi Z, Khoo CCH, Dobson SL. Wolbachia establishment and invasion in an Aedes Aegypti laboratory population. Science (1979). 2005 Oct 14;310(5746):326–8. pmid:16224027
View Article
PubMed/NCBI
Google Scholar

[111] View Article

[112] PubMed/NCBI

[113] Google Scholar

[ref33] 33. Fast EM, Toomey ME, Panaram K, Desjardins D, Kolaczyk ED, Frydman HM. Wolbachia enhance Drosophila stem cell proliferation and target the germline stem cell niche. Science (1979). 2011;334(6058). pmid:22021671
View Article
PubMed/NCBI
Google Scholar

[115] View Article

[116] PubMed/NCBI

[117] Google Scholar

[ref34] 34. Toomey ME, Panaram K, Fast EM, Beatty C, Frydman HM. Evolutionarily conserved Wolbachia-encoded factors control pattern of stem-cell niche tropism in Drosophila ovaries and favor infection. Proc Natl Acad Sci U S A. 2013;110(26). pmid:23744038
View Article
PubMed/NCBI
Google Scholar

[119] View Article

[120] PubMed/NCBI

[121] Google Scholar

[ref35] 35. Herren JK, Paredes JC, Schüpfer F, Lemaitre B. Vertical transmission of a Drosophila endosymbiont via cooption of the yolk transport and internalization machinery. mBio. 2013;4(2). pmid:23462112
View Article
PubMed/NCBI
Google Scholar

[123] View Article

[124] PubMed/NCBI

[125] Google Scholar

[ref36] 36. Han B, Weiss LM. Microsporidia: Obligate intracellular pathogens within the fungal kingdom. Microbiol Spectr. 2017;5(2). pmid:28944750
View Article
PubMed/NCBI
Google Scholar

[127] View Article

[128] PubMed/NCBI

[129] Google Scholar

[ref37] 37. Dickson DL, Barr AR. Development of Amblyospora campbelli (Microsporida: Amblyosporidae) in the mosquito Culiseta incidens (Thomson). J Protozool. 1990;37(2). pmid:1969479
View Article
PubMed/NCBI
Google Scholar

[131] View Article

[132] PubMed/NCBI

[133] Google Scholar

[ref38] 38. Becnel JJ, Andreadis TG. Microsporidia in insects. Microsporidia: Pathogens of Opportunity: First Edition. 2014;521–70.
View Article
Google Scholar

[135] View Article

[136] Google Scholar

[ref39] 39. Herren JK, Paredes JC, Schüpfer F, Arafah K, Bulet P, Lemaitre B. Insect endosymbiont proliferation is limited by lipid availability. Elife. 2014;3:e02964. pmid:25027439
View Article
PubMed/NCBI
Google Scholar

[138] View Article

[139] PubMed/NCBI

[140] Google Scholar

[ref40] 40. Bull JJ, Molineux IJ, Rice WR. Selection of benevolence in a host-parasite system. Evolution (N Y). 1991;45(4). pmid:28564051
View Article
PubMed/NCBI
Google Scholar

[142] View Article

[143] PubMed/NCBI

[144] Google Scholar

[ref41] 41. Ewald PW. Transmission modes and evolution of the parasitism‐mutualism continuum. Ann N Y Acad Sci. 1987;503(1). pmid:3304078
View Article
PubMed/NCBI
Google Scholar

[146] View Article

[147] PubMed/NCBI

[148] Google Scholar

[ref42] 42. Maire J, Vincent-Monégat C, Masson F, Zaidman-Rémy A, Heddi A. An IMD-like pathway mediates both endosymbiont control and host immunity in the cereal weevil Sitophilus spp. Microbiome. 2018;6(1):6. pmid:29310713
View Article
PubMed/NCBI
Google Scholar

[150] View Article

[151] PubMed/NCBI

[152] Google Scholar

[ref43] 43. Maire J, Vincent-Monégat C, Balmand S, Vallier A, Hervé M, Masson F, et al. Weevil pgrp-lb prevents endosymbiont TCT dissemination and chronic host systemic immune activation. Proc Natl Acad Sci U S A. 2019;116(12):5623–32. pmid:30819893
View Article
PubMed/NCBI
Google Scholar

[154] View Article

[155] PubMed/NCBI

[156] Google Scholar

[ref44] 44. Kim JK, Han SH, Kim CH, Jo YH, Futahashi R, Kikuchi Y, et al. Molting-associated suppression of symbiont population and up-regulation of antimicrobial activity in the midgut symbiotic organ of the Riptortus-Burkholderia symbiosis. Dev Comp Immunol. 2014;43(1):10–4. pmid:24201132
View Article
PubMed/NCBI
Google Scholar

[158] View Article

[159] PubMed/NCBI

[160] Google Scholar

[ref45] 45. Trzebny A, Nahimova O, Dabert M. High temperatures and low humidity promote the occurrence of microsporidians (Microsporidia) in mosquitoes (Culicidae). Parasit Vectors. 2024 Apr 11;17(1):187. pmid:38605410
View Article
PubMed/NCBI
Google Scholar

[162] View Article

[163] PubMed/NCBI

[164] Google Scholar

[ref46] 46. Martins M, César CS, Cogni R. The effects of temperature on prevalence of facultative insect heritable symbionts across spatial and seasonal scales. Vol. 14, Frontiers in Microbiology. 2023. pmid:38143870
View Article
PubMed/NCBI
Google Scholar

[166] View Article

[167] PubMed/NCBI

[168] Google Scholar

[ref47] 47. Cai W-J, Moriyama M, Fukastu T. High temperature treatment optimized for symbiont suppression in an obligatory gut bacterial symbiosis in the stinkbug Plautia stali. BioRxiv. 2024;
View Article
Google Scholar

[170] View Article

[171] Google Scholar

[ref48] 48. Serbus LR, Sullivan W. A cellular basis for Wolbachia recruitment to the host germline. PLoS Pathog. 2007;3(12). pmid:18085821
View Article
PubMed/NCBI
Google Scholar

[173] View Article

[174] PubMed/NCBI

[175] Google Scholar

[ref49] 49. Shihan MH, Novo SG, Le Marchand SJ, Wang Y, Duncan MK. A simple method for quantitating confocal fluorescent images. Biochem Biophys Rep. 2021;25. pmid:33553685
View Article
PubMed/NCBI
Google Scholar

[177] View Article

[178] PubMed/NCBI

[179] Google Scholar

[ref50] 50. Posit team. RStudio: Integrated Development for R. 2024. [accessed 15 Oct 2024] Available from: http://www.posit.co/.

Figures

Abstract

Author summary

Introduction

Results

Microsporidia MB is vertically transmitted in the An. arabiensis germarium

Microsporidia MB intensities increase in the oocyte during egg development and maturation

Microsporidia MB lifecycle and proliferation in the oocyte

Vertical transmission rate of Microsporidia MB into primary follicles

Discussion

Methods

Mosquito collection and rearing

Fluorescence in situ hybridization (FISH) staining protocol

Click-iT EdU staining of actively dividing cells in An. arabiensis germarium

Confocal microscopy and image processing

Quantification of Microsporidia MB in An. arabiensis ovaries

Microsporidia MB infection rates in the ovaries

Vertical transmission rate of Microsporidia MB

Supporting information

S1 Fig. Representative microscopic images of Microsporidia MB localized in An. arabiensis ovariole.

S2 Fig. Representative microscopic images of Microsporidia MB localization during egg development.

Acknowledgments

References