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. Author manuscript; available in PMC: 2024 Jul 1.
Published in final edited form as: Methods Mol Biol. 2024;2739:55–68. doi: 10.1007/978-1-0716-3553-7_4

Examining Wolbachia-Induced Parthenogenesis in Hymenoptera

Laura C Fricke 1, Amelia R I Lindsey 1
PMCID: PMC11216367  NIHMSID: NIHMS2001303  PMID: 38006545

Abstract

The maternally transmitted reproductive manipulator Wolbachia can impact sex ratios of its arthropod host by different mechanisms, ultimately promoting the spread of infection across a population. One of these reproductive phenotypes, parthenogenesis induction (PI), is characterized by the asexual production of female offspring, which in many cases results in an entirely female population. Cases of Wolbachia-mediated PI are most common in the order Hymenoptera, specifically in parasitoid wasps. The complex sex determination pathways of hymenopterans, their diverse life histories, the multiple cytogenetic mechanisms of PI, and the lack of males make functional studies of parthenogenesis induction challenging. Here, we describe the mechanisms of PI, outline methods to recognize and cure PI-Wolbachia infection, and note possible complications when working with PI-Wolbachia strains and their parthenogenetic hosts.

Keywords: Asexual, Thelytoky, Arrhenotoky, Parthenogenesis, Parasitoids, Haplodiploidy

1. Introduction

1.1. Parthenogenesis Induction by Wolbachia

Parthenogenesis induction (PI), characterized by the asexual reproduction of female offspring, is one of the four reproductive phenotypes caused by Wolbachia infections [1, 2]. By varying cellular mechanisms, Wolbachia induces diploidy, thus eliminating the need for a second set of chromosomes typically provided by a male through fertilization [3] (Fig. 1).

Fig. 1.

Fig. 1

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Haplodiploid sex determination and parthenogenesis induction mechanisms. (a) Arrhenotokous parthenogenesis where haploid males develop from unfertilized eggs and diploid females develop from fertilized eggs. (b) Thelytokous parthenogenesis results in diploid females from unfertilized eggs. (c) Wolbachia-mediated thelytokous parthenogenesis via gamete duplication. A failed anaphase during the first mitotic division of the embryo results in diploidization (e.g., Trichogramma sp., Leptopilina clavipes [9, 11]). (d) An alternative mechanism of Wolbachia-mediated gamete duplication seen in Muscidifurax uniraptor in which nuclei of the second mitotic prophase fuse [10]. (Schematics created with BioRender.com)

A population fixed for infection with PI Wolbachia may become entirely asexual and eventually become dependent on Wolbachia to produce female offspring [4–6]. While instances of PI are found across arthropod taxa, it is most pervasive within Hymenoptera, especially in parasitoid wasps of the supergroup Chalcidoidea [3]. Mechanisms for Wolbachia-mediated PI are diverse and include both manipulation of meiotic processes during oogenesis and various forms of altered mitosis during the first stages of embryogenesis [7–11]. In hymenopterans, all recorded occurrences of Wolbachia-mediated PI are achieved through some form of gamete duplication in the embryo. For example, this can occur through a failed first anaphase or via fusion of nuclei during the second embryonic prophase (Fig. 1c, d) [10, 11].

1.2. Haplodiploid Sex Determination and Occurrences of PI

Among arthropods, the best-understood cases of Wolbachia-mediated PI are associated with a haplodiploid sex determination system [3]. Haplodiploidy is an asymmetrical genetic reproductive system in which males are haploid and females are diploid [12]. However, how diploid offspring are generated varies across lineages. The ancestral mode within Hymenoptera is termed arrhenotoky (Fig. 1a), where unfertilized eggs (i.e., haploid and produced asexually) are laid, giving rise to male offspring. Under arrhenotoky, diploid zygotes destined to develop female are produced sexually. In contrast, thelytoky is the asexual production of females (Fig. 1b): Diploidy is achieved through a sex-independent mechanism. The sex of offspring produced by unmated females offers distinction between these two modes of haplodiploid parthenogenesis: Unmated arrhenotokous females produce male offspring, and unmated thelytokous females will produce female offspring.

The haplodiploid sex determination system is found in several arthropod groups, including Acari (e.g., mites), Hemiptera (e.g., whiteflies), rare cases within Coleoptera (e.g., some bark beetles), and it is ubiquitous in the orders Thysanoptera (thrips) and Hymenoptera (ants, bees, and wasps) [13, 14]. Across these groups, unambiguous involvement of Wolbachia in thelytokous parthenogenesis (the asexual production of females) has been confirmed in Hymenoptera, Thysanoptera, and Acari [3]. It is suggested that haplodiploid sex determination provides conditions most conducive for symbiont-mediated thelytokous parthenogenesis, as unfertilized eggs are viable [15–17] due to the existing cellular mechanisms for activating eggs and forming centrosomes independent of fertilization [18].

Though the regulatory cascades of sexual differentiation are not fully understood across Hymenoptera, it certainly is of great consequence in the context of compatible PI mechanisms. While all hymenopterans use haplodiploid sex determination, there is lineage-specific variation on the mechanism by which ploidy determines sex. In Hymenoptera, complementary sex determination (CSD) is thought to be the ancestral mode, where zygosity, in addition to ploidy, is important for determining sex. CSD involves sex determining loci, which when hemizygous or homozygous results in fertile haploid or sterile diploid males, respectively. Females develop from diploid eggs heterozygous at the sex determining locus (or loci). Ultimately, the CSD mechanism punishes inbreeding. However, entire lineages lost this CSD mechanism including the supergroup Chalcidoidea [12]. Alternatively, instances of paternal genomic imprinting have been identified in other hymenopterans; a mechanism that ensures viable diploid female development is only possible with a paternally derived set of chromosomes [19]. Because many of the mechanisms for Wolbachia-mediated PI result in homozygous diploid embryos (which would develop into sterile males in many CSD linages), CSD and paternal imprinting are thought to pose significant barriers to PI induction. This is likely the reason why Chalcidoidea, where CSD is lost, is enriched for PI-microbes.

1.3. Difficulties of Working with PI-Wolbachia and Hymenopterans

Hymenoptera are well known for having populations with significant sex ratio biases, ranging between 23% and 100% female [20]. Indeed, the presence of Wolbachia is correlated with thelytokous reproduction in many groups of Hymenoptera, as well as a range of other arthropods. However, neither Wolbachia infection nor a sex ratio bias are necessarily indicative of symbiont-mediated reproductive effects [12, 20, 21]. There are many examples of thelytokous lineages that have no reproductive symbionts, thelytokous lineages infected with reproductive symbionts where symbionts are not the cause of thelytoky, and there are arrhenotokous lineages infected with reproductive symbionts [21–24].

There are special considerations for curing thelytokous hymenopterans of their parthenogenesis-inducing symbionts and establishing a sexual, arrhenotokous colony (that produces haploid males and diploid females). Not only do you need to remove the bacterial infection, but you also need to restore sexual reproduction. If a thelytokous population is cured too quickly, the result is a population of only male offspring. To establish sexual reproduction, curing requires at least one generation in which you have reduced, but not eliminated thelytokous reproduction to provide the co-occurrence of males and females (Fig. 2), which allows for sexual production of female offspring in the following generations. Furthermore, re-establishing sex is not always possible. Sexual behaviors, structures, and processes can be significantly impaired in lineages that have long been reproducing exclusively via thelytokous parthenogenesis [4–6, 25, 26].

Fig. 2.

Fig. 2

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Antibiotic curing of PI-Wolbachia and establishing an arrhenotokous line. (a) Present honey and/or host food combined with antibiotics (“abx”) to newly emerged adults. After 24 h, provide hosts for parasitization. Each consecutive day replace the antibiotic honey and hosts with fresh aliquots. (b) Inspect F1 sex ratios and select vials with males and females for sibling mating, antibiotic treatment, and F2 generation. (c) Present antibiotics and hosts to F2 progeny and repeat the curing treatment in B-C to generate F3 wasps. (d) Collect individual mated F3 females, and (e) establish cured isofemale lines. (For * see Note 4.6 when curing fails). Honey without antibiotics can be given to cured adults, as is standard for most parasitoid rearing. (Schematics created with BioRender.com)

1.4. Outline of Procedures and Strategies for Removing PI Symbionts

The following protocols are aimed at interrogating the role of Wolbachia in hymenopteran reproduction with considerations for their unique developmental and reproductive biology. Such methods should be easily adaptable for other putative reproductive symbionts that also induce PI (e.g., Cardinium, Rickettsia), as well as other arthropods in which microbe-mediated thelytoky is suspected. Here we outline (1) methods for distinguishing between arrhenotokous and thelytokous parthenogenesis, (2) generating appropriate samples for Wolbachia screening, (3) testing the role of symbionts in thelytoky, and (4) procedures for curing thelytokous lines of their PI-Wolbachia and establishing sexual lines.

2. Materials

1. Antibiotic honey:

Weigh out an appropriate amount of antibiotic and combine with 1 mL of honey or other substrate based on insect life history (Table 1). Mix and protect from light and store at 4 °C for up to 2 weeks. If there are no protocols for your species of interest, a 5% rifampicin in honey solution (50 mg/mL) is broadly effective for a range of PI-Wolbachia (Table 1; see Notes 1 and 2).

Table 1:

Taxon-specific PI-Wolbachia curing protocols

Host Host Biology1 Curing Protocols and Notes
Trichogramma sp. (Trichogrammatidae) Idiobiont endoparasitoids of lepidopteran eggs 10% tetracycline hydrochloride, 10% sulfamethoxazole, or 10% rifampicin in honey, each worked for curing across several generations of various Trichogramma species. 1% tetracycline used for the majority of experiments as 10% resulted in some toxic effects. Gentamycin, penicillin G, and erythromycin were not effective [45].
Encarsia sp. (Aphelinidae) Koinobiont parasitoid of third or fourth instar whitefly nymphs 5% tetracycline for 24 hours prior to host parasitization, followed by access to a second batch of hosts three days later [46].
Aphytis sp. (Aphelinidae) Synovigenic ectoparasitoids of scale insects. 5% rifampicin in honey for six hours prior to parasitization of hosts [47].
Megastigmus sp. (Megastigmidae) Phytophagous: specialist seed feeders. Ectoparasitoids of larval and pupal gall wasps. Wasps provided cotton ball soaked in a “sugared solution” with 0.2% tetracycline hydrochloride for five days prior to oviposition into host plants. Very long life cycle, so offspring sex determined via flow cytometry of juveniles [48].
Eretmocerus mundus (Aphelinidae) Koinobiont parasitoid of 3rd or 4th instar whitefly Bemisia tabaci 3% rifampicin (50 mg rifampicin, 0.18 ml glycerin, 0.5 ml water, 1 ml honey) for 24 hours prior to parasitization of hosts. Repeated for three generations [49].
Diaphorencyrtus aligarhensis (Encyrtidae) Koinobiont endoparasitoid of 2nd - 4th instar nymphs of Asian citrus psyllid 10% tetracycline hydrochloride in honey for three consecutive generations [50].
Apoanagyrus diversicornis (Encyrtidae) Late-nymphal cassava mealybug endoparasitoid 1% or 5% tetracycline or rifampicin for 2 days, then access to two hosts per day each day. Sulphadiazine was not effective [51].
Muscidifurax uniraptor (Pteromalidae) Ectoparasitic on pupal muscoid dipterans 10% rifampicin in honey [43].
Asobara japonica (Braconidae) Endoparasitoid of 1st or 2nd instar Drosophila larvae 150 μl of a 2% rifampicin solution added to 1.5 g of Drosophila diet on which 70 flies were reared from egg to second instar. Wasps parasitized antibiotic-fed larvae, generating a population of antibiotic-treated adult females [52].
Telenomus nawai (Platygastridae) Egg endoparasitoid of Spodoptera sp. Cotton ball soaked with 5% tetracycline hydrochloride in honey provided to newly emerged females for 24-hours, followed by a 24-hour parasitism period [53].
Leptopilina clavipes (Figitidae) Proovigenic koinobiont endoparasitoid of 2nd instar Drosophila virilis larvae 0.5% tetracycline or rifampicin in honey for adults, plus host larvae were fed yeast with 0.2% tetracycline or rifampicin [54].
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1

Idiobiont parasitoids arrest host development upon parasitization, whereas koinobiont parasitoids allow the host to continue developing, and parasitoid often stays dormant until the host is larger and more mature.

2. Appropriate hosts:

Plan out the time series of your experiments with consideration for the dates on which you will need to feed or host (provide suitable eggs for oviposition) your insects. Egg parasitoids typically require fresh (never frozen) host eggs, which can be UV-irradiated to prevent hatching. Host eggs can often be kept at 4 °C for a week or more. For larval parasitoids, pupal parasitoids, or phytophagous (seed feeding or galling) wasps, rear appropriate hosts ahead of time such that you have the correct host stage when needed.

3. Methods

3.1. Determining Reproductive Mode of Haplodiploid Insects

  1. Isolate individual insects into separate rearing vials during the late pupal stage to collect unmated females (see Note 3). Be gentle as you transfer the developing insects. If needed, use a wet paintbrush to detach hosts containing pupae from substrates.

  2. Return isolated pupae to standard rearing conditions until adult eclosion. If you are working with an iso-female line, isolate sufficient pupae to have a minimum of ten individual unmated females for offspring sex ratio analysis. If you are working with a genetically diverse population in which there may be multiple cytoplasmic backgrounds present, increase your sample size accordingly.

  3. Upon adult eclosion, determine the sex of each individual according to species-specific criteria (e.g., antennal morphology, ovipositor presence). Males can be discarded at this step.

  4. Provide the isolated, unmated, females (G0) access to hosts and maintain them at standard rearing temperatures.

  5. Remove G0 females before offspring hatch to ensure G0 mothers are not miscounted as F1 female offspring.

  6. Collect F1 offspring and determine sex of each individual. If any female offspring are present, that indicates some level of thelytokous reproduction for that mother. If all offspring of the unmated females are male, the insects are arrhenotokous. Typically, broods will be 100% male or 100% female. However, in some symbiont-mediated thelytokous lines, occasional males appear (oftentimes the “younger siblings” [27–29]). See Table 1 for notes on long-lived species (e.g., Megastigmus sp.) and the use of flow cytometry to determine sex.

  7. If you have a line with mixed thelytokous and arrhenotokous individuals, initiate a separate isofemale line using the female offspring derived from one thelytokous mother. After a few generations, test for the absence of males.

3.2. Determine Wolbachia Infection Status

  1. Isolate unmated females as per methods above, but do not give them access to hosts (see Note 4 regarding the importance of testing unhosted females).

  2. Extract DNA from unmated, unhosted females using standard methods.

  3. PCR screen for symbionts. Use the Wolbachia specific primers W-Specf 5′-CATACCTATTCGAAGGGATA −3′ and W-Specr 5′-AGCTTCGAGTGAAACCAATTC −3′ to amplify a 438 bp region of 16S rRNA [30, 31].

  4. Conduct gel electrophoresis to determine if a sample is positive for Wolbachia (see Note 5 on other PI-symbionts).

3.3. Determine Wolbachia’s Role in PI

  1. Isolate symbiont-infected, unmated female pupae as per Subheading 3.1. Streak the vials with antibiotic or control honey so newly eclosed insects can begin feeding (see Note 3 on unmated females).

  2. Check for eclosion daily and provide fresh antibiotic or control honey to newly emerged females.

  3. After 24-h of feeding on antibiotic or control honey, transfer unmated females to a vial containing an excess of hosts, and a fresh streak of the appropriate honey.

  4. Every 24-h transfer the wasps to a fresh vial of hosts with fresh antibiotic honey, until you generate three or four 24-h oviposition periods. Maintain all vials under standard rearing conditions as they develop.

  5. When F1 offspring start to eclose, inspect sex ratios from each 24-h period of oviposition. The thelytokous mothers that received control honey should produce all-female broods. If Wolbachia is mediating thelytoky, males should significantly increase in number in the antibiotic-treated vials (see Note 2 on proovigeny).

3.4. Establishing Cured Sexually Reproducing Lines

  1. Create a replicate stock of the line you wish to cure of infection.

  2. Upon adult eclosion, provide the insects with honey containing antibiotics, streaked onto cardstock (Fig. 2a).

  3. After 24-h of feeding on antibiotic honey, give the insects access to hosts, and replace the cardstock with a fresh aliquot of antibiotic honey (Fig. 2a).

  4. Every 24-h transfer the wasps to a fresh vial of hosts with fresh antibiotic honey (Fig. 2a).

  5. Maintain all vials under standard rearing conditions as F1 offspring develop.

  6. As F1 offspring start to eclose, inspect sex ratios from each 24-h period of oviposition. Typically, older siblings will be female, and the younger siblings derived from mothers who had been feeding on antibiotics for several days are more likely to be male (see Note 2 on proovigeny) (Fig. 2b).

  7. Select F1 offspring to reestablish sexual reproduction and continue curing. Select an F1 vial that contains both male and female offspring. Alternatively, take male offspring from any vial and combine them with female siblings. Ideally, choose the youngest possible females, as they are more likely to have lower symbiont titers, making curing success higher (Fig. 2b, c).

  8. Set up this F1 cross and provide fresh antibiotic honey. It is helpful at this step to inspect these F1 insects and confirm copulation (see Note 6) (Fig. 2b, c).

  9. Every 24-h transfer mated F1s to a fresh batch of hosts and antibiotic honey (Fig. 2c).

  10. Maintain all developing F2s at standard temperature.

  11. Inspect F2s as they begin to hatch. You should find a mix of male and female offspring in each vial (Fig. 2d).

  12. Select F2s to continue curing (Fig. 2d). While the insects may be sexually reproducing at this point, it is not uncommon for them to still contain Wolbachia. Select the F2s that emerged from the last vial as they are less likely to contain Wolbachia due to a longer treatment period of their mother.

  13. Repeat steps 8–12 to generate an F3 generation (Fig. 2b–d).

  14. Upon eclosion of F3 adults, initiate 10–20 isofemale lines by isolating a single mated female and providing her hosts and standard honey (Fig. 2e).

  15. Let the isolated F3 females parasitize hosts for 48-h.

  16. Remove F3 females from hosts and use PCR to screen them for Wolbachia infection (consider using a strain-specific primer: see Note 4 on host-feeding).

  17. If any of those F3 mothers screen positive for Wolbachia, do not maintain that line.

  18. Conduct routine screenings with the antibiotic treated line to ensure Wolbachia infection is completely eliminated (see Note 7).

Footnotes

1.

Antibiotic treatments.

A range of antibiotic concentrations and treatment regimens have been used for curing of PI-Wolbachia (Table 1). The specific antibiotic, the concentration, time of antibiotic treatment prior to hosting, and numbers of generations of curing will likely vary according to insect biology. Such factors may need to be optimized for new species to account for differences including how long adults live, when their eggs develop, their life-time fecundity, and potential sensitivities to antibiotics. Finally, the goal of antibiotic treatment will likely inform your selection. If you wish to test for the role of Wolbachia in PI and are simply looking for the appearance of males (see Subheading 3.3), higher concentrations of antibiotic are likely to suffice. This is in contrast to attempts at curing where you need some female offspring in the first generation (see Subheading 3.4).

2.

Proovigenic versus synovigenic species.

Conditions for antibiotic treatment may also differ depending on life history of the parasitoid. Proovigenic species emerge as adults with a full complement of eggs [32]. This is in contrast to synovigenic species in which egg development continues through adulthood [32]. Because the eggs develop exclusively during the pupal period in proovigenic species, antibiotic treatment of adults can reduce bacterial titers, but it might be too late to reverse the effects of the reproductive manipulation established in the developed oocytes. For example, the pupal period is critical for penetrance of symbiont reproductive phenotypes in some parasitoids [33]. For that reason, additional generations of antibiotic treatment may be required to produce males. If you suspect proovigeny, repeat the antibiotic treatment protocol in Subheading 3.3, steps 1–4, isolate the unmated F1 females during their pupal period (instead of letting them eclose as per step 5), and then initiate a second round of antibiotic treatment following steps 1–5. You may likewise need to modify the antibiotic treatment during curing (see Subheading 3.4). Another approach is to include antibiotics in the media of the host food (see Table 1, Asobara and Leptopilina) such that Wolbachia titers are reduced or eliminated prior to pupation.

3.

Ensuring unmated status of females.

It is standard practice with parasitic hymenopterans to isolate insects during the pupal stage to ensure unmated status. This is in contrast to insects such as Drosophila melanogaster where unmated females are typically collected post-eclosion during a maturation time when they are not receptive to mating. In parasitic hymenopterans, male siblings typically eclose earlier than females and will often mate with females as they emerge. While all females in a thelytokous colony are likely to be unmated, stray males can occur when PI is lower penetrance, so it is best practice to isolate females during the pupal stage. Additionally, in rare occasions under solitary developmental conditions (one wasp per host), two offspring will emerge from a single host. It is best practice to ensure that only a single female is present in each vial you are using to test for reproductive mode.

4.

Considerations for PCR screening.

Standard PCR protocols are a good first step for determining the presence of Wolbachia or other reproductive symbionts. In addition to typical caveats of PCR-based screens for presence (e.g., divergent primer binding regions leading to false negatives, horizontal transfer of Wolbachia DNA into host chromosomes causing false positives [34]), there are additional considerations for working with parasitic hymenopterans. Some parasitic hymenopteran females will host feed as adults [35]; feeding on a symbiont-infected host can result in false positives for Wolbachia infection. To avoid this problem, you can either use validated uninfected hosts, PCR screen females that have not had access to fresh hosts, or design a strain-specific primer that will not detect host-derived Wolbachia. Aculeate Hymenoptera void all larval gut contents as meconium at pupation (prior to adult emergence), so freshly eclosed females should not screen positive for host-derived symbionts. In some species, you can screen for Wolbachia presence in the stray males (which do not host feed) that can occur in asexual lines (see Note 3), but this is not recommended as there is a strong correlation between Wolbachia titer and male presence [28, 36]. Additionally, some primer sets for Wolbachia detection are not compatible with certain PI-Wolbachia lineages due to divergence in the priming region (e.g., wsp primers and Trichogramma-infecting Wolbachia [37]).

5.

Other PI symbionts.

This protocol focuses on Wolbachia, but the bacteria Cardinium and Rickettsia can also induce parthenogenesis in Hymenopterans [3]. If antibiotic treatment of thelytokous females results in the production of males, but PCR screens for Wolbachia were negative, consider testing for the presence of other microbial reproductive symbionts [38, 39].

6.

Failed attempts at curing and establishing arrhenotoky.

Full curing of Wolbachia and generating a sexual line is not always possible for several reasons, so difficulties in establishing arrhenotoky might appear at various points in the curing process (Fig. 2). There are documented cases of some Wolbachia infections that are required for reproduction such that curing induces sterility [40, 41], in which case G0 females would not produce offspring. Lineages that have long been thelytokous are subject to various forms of sexual decay [25, 26, 42]. Male-specific traits may be impaired to the point that males do not develop, or they are not capable of carrying out mating behaviors, copulation, or fertilization [43]. Female-specific sexual traits often degrade much faster than the male traits in these thelytokous systems and take various forms including reduced propensity to mate, low rates of egg fertilization, and vestigialization of sperm storage structures (e.g., spermatheca) [43, 44]. Inspecting F1 wasps to determine if copulation is happening can aid in determining if sexual behaviors are impaired. Both male sterility and female non-fertilization traits (e.g., “virginity mutations” [5, 6]) would appear in the F2 generation wherein only male offspring would be produced.

7.

Recovery of Wolbachia titers and thelytoky after curing.

There are recorded instances of incomplete removal of PI-symbionts in which arrhenotokous reproduction is restored, but over time, Wolbachia titers recover, and the colony will again become thelytokous [45]. Antibiotic treatments can reduce Wolbachia titers to levels that PCR will not detect an infection, but once the antibiotics are removed, the infection is detectable again after a few host generations. We recommend screening cured colonies 1, 2, 5, and 10 generations after initiating cured isofemale lines, as well as prior to any major experiments, to ensure the infection status of the colony.

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