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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Cold Spring Harb Protoc. Author manuscript; available in PMC 2010 October 1.
Published in final edited form as:
PMCID: PMC2916733

The Parasitoid Wasp Nasonia: An Emerging Model System With Haploid Male Genetics


Nasonia is a complex of four closely related species that is rapidly emerging as a model for evolutionary and developmental genetics. It has several features that make it an excellent genetic system, including a short generation time, ease of rearing, interfertile species, visible and molecular markers, and a sequenced genome. The form of sex determination, called haplodiploidy, makes Nasonia particularly suited as a genetic tool. Females are diploid and develop from fertilized eggs, whereas males are haploid and develop from unfertilized eggs. This allows geneticists to exploit many of the advantages of haploid genetics in an otherwise complex eukaryotic organism. Nasonia readily inbreeds, permitting production of isogenic lines, and the four species in the genus are inter-fertile (after removal of the endosymbiont Wolbachia), facilitating movement of genes between the species for efficient positional cloning of quantitative trait loci (QTL). Genome sequencing of the genetic model N. vitripennis and two interfertile species N. giraulti and N. longicornis is now completed. This genome project provides a wealth of interspecies polymorphisms (SNPs, indels, microsatellites) to facilitate positional cloning of genes involved in species differences in behavior, morphology and development. Advances in the genetics of this system also open a path for improvement of parasitoid insects as agents of pest control.


Nasonia vitripennis is the “lab rat” of parasitoid wasps, and has been the subject of genetic, ecological, developmental and behavioral research for over 60 years (Whiting 1950, Whiting 1967, Beukeboom and Desplan 2003). With the advent of a genome sequencing project for N. vitripennis and two sibling species (Werren et al 2004), and development of RNAi (Lynch and Desplan 2006) and transformation (C. Desplan, pers communication) techniques, the system is rapidly advancing as a new genetic model.

Nasonia (Hymenoptera: Pteromalidae) are small insects (approximately 2mm in length) commonly referred to as “jewel wasps” due to their iridescent coloration when viewed under the microscope (Fig. 1). There are currently four identified species in the genus Nasonia (Fig. 2): N. vitripennis, N. girualti, N. longcornis and N. oneida (Darling and Werren 1990, Raychoudhury et al 2009), but more are likely. Due to confusion in the naming of this genus, early work referred to the genus name Mormoniella; however, Nasonia is now recognized and widely used. Adult females sting and lay their eggs within the puparium of various fly species. Venoms injected into the host modify host physiology and eventually kill the host. Developing wasps typically emerge 2–3 weeks later (depending on temperature). Exposure of the mother to short photoperiod and cool temperatures results in a larval diapause that can overwinter or kept in long-term storage under refrigeration (Saunders 1965). Whereas N. vitripennis is a generalist that parasitized blowflies, fleshflies, houseflies and others, the other three species preferentially parasitize blood-sucking blowflies of the genus Protocalliphora, which are commonly known as “bird-nest” blowflies. N. vitripennis has a worldwide distribution, and has presumably spread around the world due to its use of human associated flies. The other species are endemic to temperate North America, with N. giraulti and N. oneida occurring in the east and N. longicornis typically in the west.

Figure 1
Nasonia vitripennis male and female engaged in courtship.
Figure 2
Phylogenetic relationships among Nasonia species based on 10 nuclear genes (NJ using Mega 3.1, 5000 reps). Time scale uses divergence Nv-Ng estimates based on earlier calculations (Campbell et al. 1993). The relationships are also highly supported by ...

Nasonia vitripennis was first established for genetic studies in the 1950s by P.W. Whiting and colleagues. They recognized the utility of haplodiploidy and Nasonia’s ease of laboratory maintenance. It was used for genetic studies in the 1960s and 1970s by a few groups, most notably Saul and colleagues, who developed a linkage map (Saul et al. 1967, Saul 1993). Also during that time a number of research groups used Nasonia in ecological studies (Cornell and Pimintel 1978), and van den Assem conducted detailed studies of behavior, especially courtship (e.g., van den Assem 1980). In the 1980 and 1990s Werren and colleagues used Nasonia species as a model for evolutionary genetics of sex ratio control, genetic conflict, speciation and host-parasite evolution. Pultz began genetic investigations of development, taking advantage of haploid males in mutagenesis screens for embryonic patterning genes (Pultz et al. 1999). Since 2000, there has been an increase in the number of laboratory groups using Nasonia, including Desplan (developmental genetics), Beukeboom and van de Zande (courtship behavior and sex determination), Shuker (ecology and sex ratio control) and Gadau (QTL studies, nuclear-mitochondrial interactions). The pace of new researchers entering the field is increasing rapidly with the advent of genomic and genetic tools in the system.

Recent studies in Nasonia reveal it to be a good system for comparative developmental genetics and the genetic basis of complex traits. Nasonia achieves a long germband embryonic development using different mechanisms than Drosophila, particularly in determination of the A–P axis (Pultz et al. 1999, Lynch et al 2006, Rosenberg et al. 2009). Quantitative trait loci (QTL) have been mapped that affect hybrid incompatibility (Gadau et al. 1999, Niehuis et al. 2008), wing size (Weston et al 1999, Gadau et al. 2002) and mate discrimination (Velthuis et al. 2005). QTL can be more precisely identified in Nasonia males because the confounding effects of dominance epistasis can be ignored, and traits requiring replicate phenotyping can be identified in the ‘clonal’ female offspring of haploid males (Velthuis et al. 2005; see below). In addition, QTL effects can be isolated and studied in detail by backcrossing from one species into the other (Weston et al. 1999). One a QTL has been introgressed, visible and molecular markers can then be used for positional cloning.

Normally, the Nasonia species are completely or partially reproductively isolated due to Wolbachia (Breeuwer et al 1990, Bordenstein et al 2001), widespread intracellular bacteria found in arthropods and nematodes (Werren et al 2008). However, antibiotic curing of the bacteria enables the production of hybrid progeny in the laboratory and movement of genes between the species by backcrossing (Breeuwer and Werren 1995, Weston et al. 1999). The presence of these interfertile closely related sibling species has made Nasonia an outstanding system to study the genetics of the speciation process (Gadau et al. 2002, Velthuis et al. 2005), including the role of intracellular bacteria in host speciation (Raychoudhury et al. 2009).


Strains of Nasonia can be obtained from a number of research groups, and a few can also be obtained from biological supply companies (Carolina Biological and Ward’s Scientific). The Werren laboratory currently has the largest repository of wild-type and mutant strains for the four Nasonia species, and a list is available through the website (

Field collections of Nasonia are readily made, either from birdnests or using baits (Beukeboom and Werren 2000, see protocol 7). Nasonia is extremely easy to culture in the laboratory. Wasps can be sexed during the immobile pupal stages (Fig 3). Adults can be handled without anaesthetization due to a low tendency for flight. Large numbers of strains can be maintained in relatively small space in test tubes or standard culture tubes. The generation time can be as short as fourteen days at 25C or extended to over 1 month at lower temperatures. Larval diapause is induced by maternal exposure to short photoperiod and cool temperatures (Protocol 1). These can then be stored safely under refrigeration for over 1.5 years.

Figure 3
Diagram of Nasonia vitripennis pupae. Characteristics used to distinguish males and females (wing size, ovipositor) are highlighted. Pupal wing size is not as reliable for sexing the other species. A, female. B, male.

N. vitripennis can be reared on various hosts, including flesh flies, blow flies and house flies. The other species prefer “birdnest” blowflies of the genus Protocalliphora, but can be reared on flesh flies or blow flies. Host pupae are either reared in the laboratory (protocol 2) or purchased from several supply companies (e.g. Carolina Biological, Wards Scientific, Grubco) and bait shops. Sarchophaga bullata are a popular laboratory host, due to their large size and because pupae can be placed under refrigeration (4 C) for several months and remain suitable for parasitization. Host quality is checked by cracking open the puparium at the head region of a few hosts (to be discarded). Hosts are suitable up to the brownish eye stage, although are preferable for use in the white-eye to yellow-eye stage. Once bristles begin to form on the body or the body begins to darken, the hosts are unsuitable.

The easiest way to collect Nasonia eggs is to allow females to lay eggs for a prescribed period of time on a host to which their access is restricted to one end (Protocol 4). This can be accomplished by placing the host into a foam plug with a hole in one end to restrict the oviposition site. After an oviposition period (the narrower the time, the more synchronized the eggs), hosts are removed, the puparial end is “popped off” with a probe, and eggs are collected with a fine brush.


The genus Nasonia belongs to the superfamily Chalcidoidea, or chalcid wasps. It is a member of the family Pteromalidae, a widespread group of parasitoid wasps, many of which are important in controlling insect populations in natural and agricultural communities. Many but not all of the close relatives of Nasonia are parasitoids of fly pupae. Noteworthy among these are members of the genera Trichomalopsis, Urolepis, and Muscidifurax. Muscidifurax includes a parthenogenetic species M. uniraptor, which has female parthenogenesis induced by Wolbachia bacteria (Stouthamer et al 1993). A strain of Wolbachia closely related to M. uniraptor is found in N. vitripennis, but does not induce parthenogenetic development of females (Raychoudhury et al. 2009). Trichomalopsis contains a number of diverse species that have not been well characterized. The closest known species to members of the genus Nasonia is Trichomalopsis sarcophagae. Although there are currently only 4 members of the genus Nasonia, three have been discovered in the past 20 years. There are likely additional Nasonia species, particularly in Eurasia and Africa, where widespread sampling has not been conducted.


Nasonia has served as a model for genetics, evolution, and comparative development. Genome sequences for three species (currently in annotation and analysis phase) and new genomic and genetic tools promise to quickly advance the system as a model for these research areas.


Nasonia is an exceptional organism for genetic research. The important features that make it so are (a) short generation time (b) large family sizes, (c) ease of handling (including virgin collection), (d) ability to inbreed and produce healthy inbred isogenic lines, (e) availability of visible and molecular markers (f) ease of complete genome screening for mutations in the haploid sex, (g) four closely related and interfertile species, which provide a wealth of phenotypic and molecular marker differences, (h) ability to produce hundreds of genetically identical recombinant inter-species hybrids, (i) availability of genomic resources for efficient genotyping and (j) genetic resources for manipulation, including systemic RNAi and (recently) transformation. These features make Nasonia an excellent organism for basic studies in genetics, including developmental genetics, evolutionary genetics, molecular evolution and comparative genomic research. Nasonia is particularly suited for the study of complex genetic traits, due to advantages provided by haploid males and the ability to easily produce inbred lines and genetically identical recombinant hybrids. Positional cloning is practical in Nasonia, due to the high recombination rate, abundance of molecular marker differences between the interfertile species, and simplicity of phenotyping and genotyping haploid males. Several labs are currently cloning QTL for interspecies differences in morphology, behavior and hybrid breakdown.

All four species of Nasonia have 5 chromosomes, corresponding to 5 linkage groups. A visible mutant map of Nasonia has been produced containing about 20 actively cultured mutant strains (Table 1), most of which are eye color, body color, morphological and embryonic lethal mutations (Saul 1993). Screening for new mutations in Nasonia is straightforward, given that the complete genome can be screened for recessive mutations in the haploid sex. With genome sequence and linkage maps now available, there are also opportunities for cloning existing mutations in Nasonia. For example dant (distantennapedia) is a recessive homeotic mutation that converts antennae to legs; it has not been cloned, but morphological and mapping evidence suggest that it may be homologous to spineless-aristapedia in Drosophila (Werren and Perrot-Minnot 1999, Loehlin et al. 2009).

Table 1
Visible mutant map of Nasonia. Note that chromosome 2 currently has no visible mutants. Earlier maps that placed bk424 on Chromosome 2 were found to be in error. Except for 2, at least one mutant marker for each linkage group has been mapped onto the ...

Production and mapping of molecular markers in Nasonia is easy. This is because there is a high incidence of sequence differences between the species, and polymorphisms can be quickly mapped in haploid F2 males from an interspecies cross without the need to optimize for heterozygosity. In addition to a visible mutant map, microsatellite, SNP and AFLP marker maps have been generated and used to map features onto the Nasonia linkage groups (Niehuis et al 2008). For positional cloning genes involved in species differences, new markers between map locations are easily generated by comparing two species’ genome sequences at a particular region and looking for indel or SNP differences. Most recently, an interspecies mapping microarray has been developed using a Nimblegen platform that simultaneously genotypes individuals or strains for ~19,000 loci (Desjardins, in prep), as well as an Illumina SNP array for genotyping individuals at ~1,000 loci (Niehuis et al in prep). These are being used to map the genome assembly scaffolds onto the linkage map of Nasonia, and will be publicly available soon.


Nasonia has become a powerful alternative model to understand early mechanisms in development, in part because male haploidy achieves the same effect as the balancer chromosomes typically used to identify early embryonic patterning mutant genes in Drosophila (Pultz et al. 1999). Further experiments have established that although Nasonia and Drosophila both have a long-germ-band program of early development, in which gradients of localized maternal RNAs establish the anterior-posterior axis, some of the critical genetic mechanisms are different (Rosenberg et al. 2009). For example, the earliest patterning mechanisms in Drosophila, such as the maternal bicoid gradient, are not well conserved. In Nasonia, the conserved gene orthodenticle takes the place of bicoid in establishing an anterior (and posterior) gradient (Lynch et al. 2006).

The mechanism by which haplodiploid sex determination is achieved is another area of active research. In the honey bee sex is determined by heterozygosity at a single complementary sex determiner (CSD) locus (Beye et al 2003). Individuals that are heterozygous at the CSD locus develop into females, while hemizygotes and homozygotes develop into haploid and diploid males. Thus inbreeding results in diploid males. Nasonia lacks CSD, as it readily inbreeds without production of diploid males. Current data suggest that sex is determined by an interaction between maternal effect and zygotic gene products, and maternal genetic imprinting has been proposed (Kamping et al. 2007). The genome sequence has uncovered a number of loci known to be involved in sex determination in diverse organisms including doublesex (Oliveira et al 2009,Oliveira et al in press?) and transformer, and experiments are underway to unravel the mechanism of sex determination in this insect.

Sex determination is of particular interest because Nasonia’s sex determining and sex allocation systems are frequently hijacked by reproductive parasites. Nasonia is one of the most well developed systems for studies of Wolbachia-host interactions. The mechanism of Wolbachia cytoplasmic incompatibility appears to depend on host, not bacterial, genotype (Bordenstein et al. 2003) The bacterial genus Arsenophonus was originally described in Nasonia (Gherna et al 1991), and is now known to be widespread in insects. Arsenophonua nasoniae kills sons of infected females by disrupting the formation of maternally derived centrosomes in unfertilized haploid embryos (Ferree et al 2008). Nasonia is also known to harbor the most extreme example of selfish DNA found in any organism, Paternal Sex Ratio (PSR). PSR is a supernumerary chromosome found in some populations that is transmitted through sperm, but disrupts proper condensation of the other sperm derived chromosomes, thus converting the embryo into a haploid and therefore male carrier of PSR (Werren and Stouthamer 2003). PSR has been used to decouple the signals of fertilization and ploidy as a way to dissect the primary signal of sex determination (e.g., Ferree et al. 2008).

Ecology & Behavior

Nasonia has been used quite extensively for behavioral and ecological research. Its parasitoid lifestyle allows investigations of questions relating to parasitoid-host dynamics, host preference, specialist versus generalist biology, et cetera. In terms of behavior, there are many interesting questions about courtship behavior, male aggression and territoriality, female dispersal, and sex ratio control. Perhaps the most important quality of Nasonia as a behavioral and ecological model is it has the tools to dissect the genetic basis of these species differences (e.g., Velthuis et al 2005).

Courtship involves stereotypic displays that differ between the species (van den Assem and Werren 1994) as well as the release of pheromones from the male’s mandibular region that play an important role in female receptivity (van den Assem et al 1980). Courtship occurs quickly (typically it is completed within 1–2 minutes) making it a popular subject of study in undergraduate teaching laboratories and for undergraduate research. Other experimentally tractable behavioral differences are ripe for further investigation. Females of N. giraulti often mate within the host, whereas this is uncommon or absent in N vitripennis and N. longicornis. After mating, females disperse from the natal patch in search of new hosts. Dispersal behavior of females differs between strains and species. Males of N. vitripennis have vestigial wings and are incapable of flying. Males of N. longicornis have intermediate sized wings and N. giraulti males have large wings similar in size to those of females. The latter two species are capable of flying, although they do not do so as readily as females. Males will defend hosts that contain adult females that have not yet emerged, and have a number of aggressive displays associated with this territorial behavior (Leonard and Boake 2006).

Most mating occurs locally within the natal patch, and sibling mating is common. Therefore, Nasonia is subject to local mate competition, and has been shown to alter sex ratio among progeny in response to the number of females in a group of hosts or as a consequence of superparasitism in patterns consistent with local mate competition theory (Werren 1980, 1983, but see Parker and Orzack 1985, Orzack and Parker 1986). Single ovipositing females typically produce strongly female-biased sex ratios (80–95% daughters), whereas when in groups they produce more equal ratios. The haplodiploid sex determination provides a mechanism for control of the sex ratio among offspring, and female reproductive anatomy suggests that they can control individual fertilization of eggs (Whiting 1967).

The four species differ in their host preferences. N. vitripennis is a generalist and will parasitize a wide range of fly hosts, including blowflies, fleshflies and houseflies. The other three species appear to be specialists, and are found parasitizing Protocalliphora, blowflies that specialize as ectoparasites in birds’ nests (Darling and Werren 1990). N. giraulti, N. longicornis and N. oneida prefer these hosts, although they will parasitize S. bullata in the lab. Studies of hybrids have identified a genetic region that strongly influences host preference, and recently developed genetic tools, including the dense SNP and indel map of Nasonia, are being used in efforts to clone the gene(s) involved in host preference.

Nasonia is a tractable system for field research. Wasps can be collected from bird nests and from the vicinity of carcasses (N. vitripennis). Baits using meat that has been fed upon by blowfly larvae placed in mesh bags can be efficiently used to sample natural populations. Field studies have uncovered a variety of th e important features of this system, including sex ratio distorters, new species, and intraspecific differences in behavior and morphology. A set of strains collected from North America and Europe are available to interested researchers.

Evolutionary Genetics & Speciation

Given the existence of closely related and interfertile Nasonia species, there are excellent opportunities for evolutionary genetic studies, particularly those focused on the genetic basis of speciation and adaptation. A core set of strains from different populations in North America and Eurasia exists for all four species (the NasCore) and can be obtained from the Werren lab. Analysis of mitochondrial CO1 sequences suggests some population subdivision in N. vitripennis and N. longicornis (Raychoudhury et al. 2009). Studies are underway to clone the genetic basis of some phenotypic differences (e.g. wing size and female mate preference) between the species. The tools for detailed evolutionary genetic studies are now in place, and this promises to be a growth area in the near future.

Genes involved in hybrid incompatibility have been investigated in Nasonia. Hybrid incompatibility genes tend to be recessive, and are immediately uncovered in the haploid F2 hybrid males (Breeuwer and Werren 1995, Gadau et al 1999, Niehuis et al 2008). Strong nuclear-mitochondrial incompatibilities occur in Nasonia hybrids (Breeuwer and Werren 1995), likely due to the exceptionally high mitochondrial mutation rate (Oliveira et al 2008). Nuclear genes involved in the OXPHOS pathway (the electron transport chain) have been implicated in these incompatibilities (Niehuis et al 2008), and efforts are now underway to clone these incompatible loci.

The ability to produce isogenic inbred lines in Nasonia is an advantage for quantitative genetic studies. Unlike the honey bee, Nasonia readily inbreeds both in nature and in the lab without the deleterious effects found in many diploid organisms. This form of sex determination permits the generation of very healthy highly inbred strains because harmful recessives have been purged in the haploid males. Isogenic females can be placed in different environments to investigate genotype × environment interactions and norms of reaction. Toward this end, sets of recombinant inbred lines have been produced within and between Nasonia species (Velthuis et al 2005, Werren, unpublished), which can be screened for phenotypes involved in species differences.

As mentioned, hybrid females can be set as virgins, and these virgins produce haploid recombinant males. In addition, a unique aspect of the combination of haplodiploidy and highly isogenic strains is that large sets of genetically identical females can be produced in the F3 generation. The approach works as follows (Figure 4). A cross is made between two strains (or species) that differ in a phenotype of interest (e.g. female mate preference). F1 females produce recombinant haploid male progeny. Because of haplodiploidy, all the sperm from each individual recombinant male are genetically identical. Therefore, when he is backcrossed to an isogenic line of females, all the F3 daughters will be genetically identical with 1 recombinant genome (from the male) – these are referred to as F3 clonal recombinant females (see Figure 4). Hundreds of clonal females can be produced by this message. This is advantageous for subtle or variable phenotypes because genotype – to – phenotype matching can be more confidently accomplished through replication of clonal females. Furthermore, sets of clonal females can be placed into different environmental conditions to investigate genotype × environment interactions (e.g. norms of reaction). Finally, individual F2 males can mate with many dozens of females, allowing crossing of the same haplotype into many different genetic backgrounds, each then producing hundreds of females for phenotypic characterization. The F2 recombinant males are readily be genotyped (e.g. using molecular markers) without marker codominance problems, and the genotype of the F3 females is known by also genotyping the maternal inbred line. This approach was used effectively to map genes involved in female mate preference in N. longicornis (Velthuis et al. 2005). These features make Nasonia almost uniquely adapted among higher eukaryotes for the study of complex genetic traits.

Figure 4
Crossing Scheme for Haploid Recombinant Males and Clonal Females. Haploid recombinant males can be produced between interfertile species (vitripennis and giraulti) or inbred strains within a species. F3 clonal females are produced when individual haploid ...

The widespread endosymbiotic bacterium Wolbachia has been implicated in reproductive isolation between Nasonia species (Breeuwer et al 1990, Bordenstein et al 2001, Bordenstein et al 2007). The general pattern is that each species harbors a different set of Wolbachia, and these induce complete to nearly complete incompatibility between sperm and eggs in most interspecies crosses. Two significant exceptions are that N. giraulti and N. oneida have apparently identical Wolbachia (and mitochondria), while N. longicornis is polymorphic for multiple B-group Wolbachia (Raychoudhury et al. 2009). Antibiotic curing results in a dramatic increase in the number of F1 hybrids. Nasonia is one of the best illustrations of the possible role of Wolbachia in speciation.


Genetic resources in Nasonia include mapped and unmapped visible mutants (Table 1.), wild-type strains from all four species collected from different geographic regions, and interspecies hybrid inbred lines. Visible markers present on each chromosome come from a long history of classical genetics and molecular marker maps produced using a combination of SNP, RAPD, AFLP markers from interspecies and intraspecies crosses are available through the Baylor Nasonia website. Information on the Nimblegen ~20K locus and Illumina SNP mapping arrays will soon be available. Approaches to genetic crosses are described in Protocol 4. Larval RNAi methods are described in protocol 5 and pupal RNAi methods are described in Lynch and Desplan (2006). Gene transformation has been demonstrated by the Desplan lab (unpublished).

Genomic resources include the assembled and partially annotated genome of N. vitripennis as well as genome sequences of two sibling species N. giraulti and N. longicornis (assembled to the reference N. vitripennis genome), EST and full length cDNA sequences, SNP, indel and Microsat marker locations in the genome, and a scaffold linkage map. All these resources are available or soon will be through the Human Genome Sequencing Center (Baylor College of Medicine) Nasonia website (see below) and the NCBI Wasp Genome Resource site (see below). BAC libraries for N. vitripennis and N. giraulti are available from the Clemson University Genomics Institute. Genome resources that will soon become available through the Indiana Center for Genomics and Bioinformatics are data on sex and life-stage specific gene expression developed from a Nasonia tiling microarray and design information for Nasonia expression arrays and comparative genomic hybridization mapping arrays.


Here we provide protocols on (1) Nasonia strain maintenance, (2) Rearing of Sarcophaga bullata hosts, (3) Egg collection, (4) Virgin collection and haplo-diploid crossing methods, (5) Larval RNAi injection, (6) Curing Wolbachia infections in Nasonia, and (7) collecting Nasonia with baits. Several DNA extraction methods work. The “squish” DNA extraction protocol (Gloor and Engels 1992) may be easiest, though a full strength extraction from females occasionally inhibits PCR. Substituting 0.2 uL female genomic DNA for the usual 1.0 uL in PCR works reliably for us.

Protocol 1: N. vitripennis strain maintenance

This protocol describes standard rearing of N. vitripennis strains on Sarcophaga bullata hosts. By using incubators at different temperatures, Nasonia’s development rate can be adjusted to conform to the investigator’s schedule. Nasonia will produce diapause larvae when reared at 18C with a short-day light cycle (Saunders 1965); these can be archived at 4C for over a year. Diapausing Nasonia may lose their Wolbachia infections (Perrot-Minnot et al. 1996). N. longicornis, N. giraulti and N. oneida can be reared under similar conditions, with the caveat that N. giraulti and N. oneida lay all-diapause broods more frequently and are therefore best reared at 25C.


  • Sarcophaga bullata pupae (see Protocol 2 – fly rearing) or other calliphorid fly pupae (stored at 4C)
  • 25mm diameter Drosophila vials and plugs (cotton, foam, rayon etc.)
  • 12mm diameter plastic vials and cotton plugs
  • honey water (40% honey: 60 % H2O aliquots in 1.5mL tubes)

  • Incubators: 25C and 21C (constant light), 18C (8h light: 16h dark)
  • Refrigerator: 4C (preferably two for redundancy).
  • Small paintbrushes (number 2 or similar)
  • Rubber or foam pad


  1. 25C and 21C strain maintenance
    1. Transfer 20 mated females and 4 or fewer males to new 25mm Drosophila vial. If transferring from same sized vial, place the new vial upside-down over the old vial. Mated females will climb up into the new vial. Otherwise, use a brush.
    2. Cover vial opening with two fingers and knock vial bottom against rubber pad. This will make most Nasonia fall to the bottom.
    3. Add 20 Sarcophaga hosts (or fill the vial 1/4 full if hosts are small).
    4. Plug with foam or cotton plugs and incubate new vial in 25C or 21C incubator. Used foam plugs can be autoclaved and reused.
    5. Backup emerged cultures in 4C refrigerator. Mated females will survive and remain fertile for approximately 1–3 weeks.
    6. Adults will emerge approximately 14 days later (25C) or 21 days later (21C). Nasonia females will mate immediately upon emergence.
  2. 18C strain maintenance
    1. Follow the same procedure as for 25C maintenance, except incubate at 18C using a 8h light: 16h dark cycle. Nasonia reared at 18C will emerge in approximately one month and live for several weeks.
    2. If no Nasonia have emerged after six weeks, or after all emerged Nasonia have died, check the vial for diapause larvae. Crack open all hosts at the midline using thumbnail or a probe. Diapause larvae are as large or larger than the average Nasonia and are distinguished by prominent white fat cells visible through the cuticle.
    3. Transfer hosts containing diapause larvae to a new 12mm plastic vial, plug with a cotton ball, label, and store in a 4C refrigerator for 6–24 months. Store in box with air holes, such as empty pipet tip box. Record strain and deposit time in a diapause log. If possible, split tubes between two 4C refrigerators to reduce the impact of equipment failure.
  3. Recovery of diapause larvae
    1. Remove tubes of diapausing larvae from 4C refrigerator. Strains are most likely to recover after refrigeration for 6–18 months.
    2. Remove cotton plug. Take new cotton plug and gently moisten with one or two drops of distilled water (do not saturate or mold will grow). Cover all tubes with a sheet of aluminum foil to reduce loss of humidity.
    3. Incubate recovering diapause larvae at 25C until wasps pupate, eclose and mate (about 14 days). If greater than 50% males emerge, separate approximately 20 females into a new vial using a paintbrush, add 2–4 males, paint a dot of honey water on the wall of the tube, and allow mating to occur for 4h or overnight.
    4. For long-term strain maintenance, pull 2–3 full 12mm vials from diapause at 18–20 months. After rebuilding the population for a generation or two at 25C, return the strain to 18C culture to produce more diapause larvae.
    5. Remove from 18C culture only when several batches of diapause larvae have been collected and stored (generally >6 vials worth).


Problem: low sex ratio (mostly males)

Solution: Crowding by multiple males may prevent some females from mating, who then produce all male broods and repeat the cycle. Remove as many females as possible using paintbrush, or, if necessary, knock out with CO2 and sort. Separate up to 20 females into a new vial, add 2–4 males, paint a dot of honey water on the wall of the tube, and allow mating to occur for 4h or overnight. All-male broods means no female mated. Recover strains from a culture backed up at 4C.

  • Perrot-Minnot MJ, Guo LR, Werren JH. Single and Double Infections with Wolbachia in the Parasitic Wasp Nasonia vitripennis: Effects on Compatibility. Genetics. 1996;43:961–972. [PubMed]
  • Saunders DD. Larval Diapause of Maternal Origin: Induction of Diapause in Nasonia vitripennis (Walk.) (Hymenoptera: Pteromalidae) Journal of Experimental Biology. 1965;42:495.

PROTOCOL 2: Rearing Sarcophaga bullata fly hosts


Nasonia vitripennis is a parasitoid of a number of calliphorid flies, such as S. bullata. S. bullata are relatively large, increasing the offspring yield that a single N. vitripennis female can produce. They are also easily reared in the lab if proper ventilation is available.


  • Beef liver, cut in 2″ cubes
  • Dry fly food (half dehydrated milk, half granulated sugar)
  • 12mm glass test tubes and cotton plugs
  • Paper towels
  • Bleach

  • Wire mesh fly cage
  • Ventilation hood
  • 30C incubator
  • 10×14″ plastic tubs with lids (cut large hole in lid)
  • 16″×16″ cotton cloth squares (such as bed sheet)
  • 8×8×3″ plastic containers
  • 8×8×3″ plastic containers with 4mm drainage holes drilled in bottom
  • Steel colander
  • Plastic wash basin
  • Paper drinking cups with 1mm air holes poked in them
  • Rubber bands


    1. Start and maintain a population of >50 flies (5 female: 1 male). Cover the bottom of the cage with a layer of paper towels. Change water and liver baits every day (see below).
    2. Each day, change water: fill two 250mL Erlenmyer flasks with water and insert 5 rolled-up paper towels in each as a wick.
    3. Each day, remove old liver baits and add 2 new pieces of liver (on Petri plates or similar).
    4. Each day, add flies to the cage in a ratio of 5 females to 1 male (aim for 10:2).
    5. Change fly food three times a week: two Petri plates of 50% sugar, 50% dehydrated milk.
    6. Set up 32 fly pupae individually in 12mm glass vials with cotton plugs three times a week. Use the oldest hosts in the refrigerator, but crack a couple open to ensure they are good and will develop into flies. Healthy flies will have firm shells and clear eyes.
    7. Clean the cage once a week. Remove paper toweling, sweep up debris, replace paper towels.
    1. LIVER
      1. Remove liver from refrigerator. Place pieces onto paper towels to remove excess liquid.
      2. Take necessary amount of liver (usually one 10lb bag) from freezer to defrost for later. Poke holes in bottom of bag and place bag in colander. Place this in the plastic wash basin. This allows drainage of blood from liver.
      3. Always ensure that there is enough thawed liver for the next two days.
    2. DAY 0 LARVAE
      1. Add liver to cover the bottom of a 8×8×3 inch plastic container.
      2. Add larvae from the fly cage liver baits.
      3. Place into 10 × 14 inch larger holding tub with paper toweling in the bottom (about 5 sheets).
      4. Place and seal lid and cloth cover over the large holding container. Make absolutely sure that the lid is completely closed.
      5. Label with date. Place into ventilation hood.
    3. DAY 1 LARVAE
      1. Open container. Remove excess liquid with paper towels. Turn over liver if dry and sprinkle with water.
      2. Close container, making sure to seal properly, and return to hood.
    4. DAY 2 LARVAE
      1. Open container.
      2. Transfer liver and larvae into 8×8×3 inch plastic container with drainage holes. Stack in same size container without holes.
      3. If larvae are crowded, split between two bins, adding additional fresh liver to each.
      4. Close container and return to hood.
    5. DAY 3 LARVAE
      1. Open. Remove bottom container that catches the drippings.
      2. Place drainage container on the paper towels in the larger tub. Prop up one edge of the container so it will drain.
      3. Close container, making sure to seal properly, and return to hood.
    6. DAY 4 LARVAE
      1. Open. Living larvae should have congregated at the high end of the drainage container.
      2. Scoop larvae over the side of the drainage container onto paper towels. Only move those larvae that are grouped away from the liver. Remaining larvae will follow.
      3. Close container, making sure to seal properly, and return to hood. Larvae at this stage are most likely to crawl out of the bin and escape.
      1. Open. Remove smaller container with liver. Dispose of liver and any remaining larvae (put in plastic bag for freezing).
      2. Remove old paper towels from tub, making sure to pick out larvae and pupae.
      3. Distribute larvae evenly into two cleaned tubs, in between layers of paper towels. Cover with cloth and lid.
      4. Place in 30C incubator for 3 days.
    8. DAY 8 PUPAE
      1. Check day 6 and day 7 pupae. If they are wet, change the paper towels.
      2. Collect day 8 pupae into paper cups. Check pupa quality by cracking puparium with thumbnail in head region and examining quality of host. Firm puparia and clear eyes are good.
      3. Write date on paper towel and cover cup using paper towel and rubber band.
      4. Place cups into the refrigerator. Use a different refrigerator than that used to rear Nasonia, which can crawl through the air holes.


Problem: larvae appear dry, sticky, or covered with foam

Solution: wet hands and sift through larvae. Usually a small amount of water is sufficient for them to begin to clean themselves off. Use paper towel to mop up foam or standing liquid. Change paper towels under bin if they are wet.

PROTOCOL 3: Nasonia egg collection


This protocol describes methods to count and/or collect Nasonia eggs. Fly hosts are placed in a foam plug such that only one oviposition site is available to the female wasp.


  • 12mm Culture vials
  • Sarcophaga hosts
  • 15mm foam plugs
  • #12 syringe needles


  1. Heat dissecting needle using a laboratory burner. Use this to burn holes in the end of foam plugs. Do not insert the needle more than 75% of the way through the plug. Plugs can be reused.
  2. Collect females of strain of interest. To maximize egg production, give females hosts for 2–3 days to host-feed. Virgin females will produce all-male offspring, while mated females will produce about 90% female offspring.
  3. Transfer emerged females to 12mm culture vials by dumping onto lab bench and placing inverted tubes over females. Allow females to climb up into the vials.
  4. Check host quality. Roll hosts on flat surface to check for dessication. Discard any which do not roll evenly. Also discard any very dark or misshapen hosts.
  5. Place host pupae anal-end first into host plug and insert into the culture vial, allowing only the head-end of the host to be accessible to the wasp. If the plug is loose around the pupa, do not use that plug.
  6. Allow females to oviposit for 2 hours, and then remove females.
  7. Crack open host at the base of the head segment using thumbnail or dissecting needle. Practice so that you can do this without breaking the host.
  8. Count eggs or collect eggs using a fine brush or sterilized syringe needle. Eggs will be arranged in a circular pattern around the sting site. The sting site can be identified as a brown/black dot, possibly with a protruding feeding tube.

PROTOCOL 4: Nasonia virgin collection and haplo-diploid crossing methods


This protocol covers most procedures used by the Nasonia geneticist, including virgin collection. Nasonia are easily sexed as pupae, so collecting virgins is easily done with a window of several days. The female-specific ovipositor is visible at even the earliest pupal stages as a longitudinal ridge on the ventral posterior abdomen. To prolong generations, yellow pupae can be stored at 4–8C for up to two months and adults for two to three weeks. Haplo-diploid crossing methods are introduced in the example of making a new Nasonia vitripennis mutant homozygous, using an aunt-nephew mating approach. The protocol can be extended to inbreeding strains. Inbreeding depression is generally not a problem in Nasonia unless the mutation itself is female sterile. Mother-son mating and sibmating are also reasonable approaches to inbreeding.


  • 12mm culture vials and cotton plugs
  • Fly hosts
  • Honey water (40% honey, 60% sterile H2O)

  • Dissecting microscope
  • CO2 diffuser
  • Dissecting needle bent at 45 degrees 2cm from tip (“hockey stick”)
  • Index card (fold long edges to 90 degrees, 1cm from edge, to form a tray)
  • 25C Incubator
  • 21C Incubator
  • 4C refrigerator


    1. Start with the offspring of a mated female. Nasonia pupae can be collected between days 10 and 13 if cultured at 25C, days 14–20 if cultured at 21C, and usually after day 27 if cultured at 18C, varying with the strain’s development time.
    2. Open fly hosts at midline using thumbnail or bent dissecting needle. Pour wasp pupae onto index card under dissecting scope.
    3. Sex pupae and separate using bent-angle dissecting needle. Females will have a longitudinal ovipositor ridge at the ventral posterior of the abdomen (Fig XX). The ovipositor ridge interrupts the transverse abdominal segments. Males will have uninterrupted abdominal segments. Male Nasonia vitripennis also have noticeably smaller wing pads (less obvious in other species).
    4. Put virgin pupae in 12mm culture vials and plug with cotton. Aim for 10 females per vial to reduce risks from sorting mistakes. Females should emerge in 14 days from original hosting at 25C or 21 days at 21C.
    1. Document phenotype of male mutant. (If the mutant appears in a female, host female as virgin for 2 days, then mate to a male of a wildtype strain and host).
    2. Identify wildtype stock to use for mating. If new mutant is Wolbachia-free, use an uninfected strain such as the genome strain AsymCX. Otherwise, or if in doubt, use an infected strain such as LabII.
    3. If adult virgin females are not available, place a drop of honey water on the side of the tube containing the mutant using a fine brush, allow to feed for 1 hour, and refrigerate until adult virgin females are available (See Virgin Collection above).
    4. Place male mutant in culture vial with 2–5 virgin wildtype females and drop of honey water. Observe copulation and wait 3 hours for sperm incorporation before hosting, or else allow to mate overnight.
    5. Host mated females with 1 fly host per female at 25C.
    6. 10 days later, collect three sets of ten F1 virgin females. The first set will be virgin-hosted to produce F2 males. The second set will be aunt-nephew mated to the F2 males. The third set will be virgin hosted to produce males to mate to the F2 females. Back up the rest of the yellow pupae at 4C. All F1 females will be heterozygous.
    7. Allow F1 virgin females to eclose. Check to see if mutation is dominant.
    8. Host first set of virgin females individually, with two hosts each, and incubate at 25C. Give honey water as above to the second and third set of virgins and allow to feed for 1 hour. Then store at 4C.
    9. Allow F2 males to emerge (Day 15). Place males on CO2 diffuser under dissecting microscope and check that phenotype was inherited. Some mutant phenotypes may be unable to escape the host; crack open the hosts to check..
    10. Sort 2–4 F2 mutant males and then mate them to one of the tubes of emerged F1 heterozygous females which was stored at 4C. Add new drop of honey water and allow mating to proceed overnight.
    11. Host mated females individually at 25C. Also host remaining tube of virgin F1 females.
    12. 10 days after hosting, collect 20 F2 virgins. These will be a mix of heterozygotes and homozygotes. Back up the rest at 4C.
    13. Host F2 virgins individually with two hosts at 25C. Label each vial to keep track of the females.
    14. 48 hours later, transfer virgin females to new tubes containing a drop of honey water. Add in a mutant male from the virgin setting and allow the mating to proceed overnight in the 21C incubator.
    15. After mating, add two fly hosts and incubate the cross at 21C.
    16. 15 days later, F3 males will emerge. Sort through male populations using CO2 diffuser, making sure to crack open the hosts. Identify homozygous mothers.
    17. Collect 20 virgins from F2 homozygous mothers. Also collect 10 virgins from heterozygous mothers. Back up the rest at 4C.
    18. Repeat steps 12–15 to confirm that the strain is homozygous. Continue maintaining strain as in Protocol 1.


Problem: No female pupae seen.

Solution: Female parent may not have mated, or asymmetric Wolbachia infections caused cytoplasmic incompatibility. Unmated (virgin) Nasonia females will produce all-male broods. Recently emerged females (< 3 hr) from stocks may not have mated. Collect virgin females from 4C backup and resume cross. Check Wolbachia infection status of stocks. In general, infected males can only successfully fertilize eggs that have the same Wolbachia types. Virgins from an infected lab strain such as LabII can be used if male infection status is unknown.

Problem: homozygous females do not lay eggs or do not mate.

Solution: The mutation may be female-sterile. Maintain in heterozygous fashion (Steps 12–16 above) or discard.

Protocol 5: Larval RNAi in Nasonia


This protocol describes a method to use RNA interference (RNAi) to knock down genes in Nasonia larvae. Unlike Drosophila, RNAi in Nasonia is systemic. Lynch and Desplan (2006) have injected double stranded RNA (dsRNA) in female pupae and successfully knocked down genes in offspring embryos. By injecting dsRNA against the eye color gene cinnabar in last-instar Nasonia larvae, we have successfully produced adult red-eye-color phenotypes.


  • Taq Polymerase
  • 5′ and 3′ Gene-specific primers with attached T7 polymerase binding sites at the 5″ ends.
  • 5′ and 3′ cinnabar primers with T7 sites:
  • Megascript T7 RNAi Kit (Ambion cat. no. 1626)
  • 2% Agar in 1×PBS Plates
  • Capillary Tubes: OD 1.0mm, ID 0.75mm, Length 100mm (World Precision Instruments cat. no. TW100F-4)
  • Food Coloring, preferably red or blue (McCormick brand)
  • Microloader Pipet Tips (Eppendorf cat. no. 5242 956.003)
  • Gelatin Capsules such as Coni-Snap #4 Clear/Clear (Medisca INC. cat. no. NDC 38779-1125-9)
  • Nasonia to inject (such as wildtype genome strain AsymCX)
  • Nasonia with known mutant phenotype (such as bl13, or123)
  • Fly hosts (Sarcophaga bullata, etc.)

  • Micro-injection Pump (e.g. PicoSpritzer III from Parker Instrumentation)
  • Needle Puller (e.g. Flaming Brown Micropipette Puller Model no. P.80/PC, Sutter Instrument co.)
  • Small paintbrushes


    1. Prepare control and experimental dsRNA according to the protocol in the MEGAscript RNAi Kit. The PCR approach to making dsRNA templates is sufficient with some PCR condition optimization. Run 3 to 5 100μl reactions, pool them and ethanol precipitate the products to get a high enough concentration of the dsRNA template. Do the final elution using the elution buffer supplied with the kit. This buffer alone has no adverse effects on the larva. It is best to elute with the smallest recommended volume as it takes only a very small amount to do many injections and having a higher concentration of dsRNA can be beneficial. dsRNA against cinnabar can be used as a positive control.
    2. Prepare pulled capillary needles using a needle pulling machine. The best needles will come to a very fine point (only visible under magnification) without having “wispy” ends. The same program in different machines (even of the same model) will produce variable needles, so the protocol will have to be optimized. We used the above listed capillary tubes with a ramp value of 395 (see machine manual).
    1. Host Nasonia females to produce larvae for injection. To standardize larval body size, females can be hosted individually on two hosts for 24h–48h and then rehosted. Virgin females will produce all-male sex ratios, while mated females will produce approximately 95% female and 5% male offspring.
    2. Collect 6–7 day old larvae raised at 25°C. At this stage larvae should be finished feeding, detached from the host, and their gut material should appear to be gray and condensed. These are referred to as fourth instar pre-defecation or gray-gut larvae. Injections can be done as much as three days earlier or 1–2 days later than this, into the pupal stage. In either case the mortality may go up. When injecting fourth instar pre-defecation larvae the survival rate can be as high as 70%.
    3. Carefully open the host and spread larvae onto a 2% agar in 1× PBS plate using a small paint brush.
    4. Transfer larvae of the proper stage to fresh plates, dividing into experimental and control groups. On each plate organize the larvae into a line or other pattern.

  1. Mix 4.5uL dsRNA (experimental, cn positive control, elution buffer negative control) with 0.5uL food coloring.
  2. Backload injection mix into a needle using a microloader pipette tip, creating as few air bubbles as possible.
  3. Mount the filled needle in a micro-injection pump.
  4. Hold the needle at a 30°–45° angle with respect to the posterior end of the larva. Gently pin the larva down along its length with the tip of the needle facing the posterior end. The posterior end of Nasonia larvae is more pointed than the anterior end (Fig. xx).
  5. With the top half of the larvae depressed, slide the needle forward just beneath the cuticle, and inject. Be careful not to puncture the gut. (Fig. xx).
  6. If using gray-gut larvae (day 7) or later, these will have stopped feeding and can be left to develop on the PBS plate. Collect pupae at day 11 and transfer to glass vials if desired.

Procedure for injecting earlier larval stages

Injected larvae of earlier stages must be returned to parasitized hosts to feed. Foster larvae will continue to eat the host and reduce the risk of injected larvae drowning. To avoid ambiguous results, set up foster hosts with females marked by known mutants that differ from the expected results of RNAi knockdown. We used bl13, or123 which have blue bodies and orange eyes (readily distinguished from the scarlet eyes of the cn knockdown). Desiccated or rotted foster hosts are the main causes of larval loss after injection. To prevent this, use sterile techniques when handling foster hosts and be very careful not to puncture the fly carcass.

  1. Set up foster hosts with mutant-marked females at the same time that the strain to be injected is set up (PREPARE LARVAE, #1, above).
  2. Remove the anterior end of the fly puparium using a heavy gauge needle or sharp forceps to crack around the edges, being careful not to puncture the fly within.
  3. Before placing the injected larvae into the foster host (5–10 per foster host), remove approximately the same number of the foster larvae.
  4. Re-seal the puparium with an empty gelatin capsule, creating a proper seal. Size four capsules are appropriate for large Sarcophaga. Create a proper seal over the open foster host.
  5. Place the capped pupa in a vial and allow it to develop at 25°C. Mature Nasonia (day 14–15) should make their way out of the capsule but when scoring the progeny be sure to open the capsule and check that all of the eclosed Nasonia have escaped.


Problem: Injected wasps do not pupate.

Solution: dsRNA treatment or injection stress may have killed the larva or induced diapause. Compare survivorship of treatment and buffer-injected controls. Diapause larvae will continue to wiggle. Diapause can be broken by refrigeration for >8 weeks.

  • Lynch JA, Desplan C. A method for parental RNA interference in the wasp Nasonia vitripennis. Nature Protocols. 1:486–494. [PubMed]

PROTOCOL 6: Curing Wolbachia infections from Nasonia

This protocol describes curing a Nasonia strain of its Wolbachia symbionts (Breeuwer and Werren 1990, 1993). Wolbachia cause cytoplasmic incompatibility (CI) in Nasonia. Crosses between infected and uninfected wasps, or between infected wasps of different species, will fail because of Wolbachia. The sperm modification which induces CI appears to be established before the adult stage, so it is generally not possible to cure an infection and break the CI barrier in the same generation. A Wolbachia infection can be removed from a strain over a number of generations, however, by feeding antibiotics and selecting on females who show partial CI. The current genome sequenced strains of Nasonia (AsymCX, IV7(u), RV2X(u)) are all Wolbachia-free.


  • Sucrose
  • Tetracycline
  • Culture vials
  • Filter paper
  • Wasp Cultures


  1. Create 1% tetracycline in 10% sugar water solution.
    • 1ml ddH2O
    • 0.1g sucrose
    • 0.01g tetracycline
  2. Vortex solution for 20–30 sec or until all sugar crystals have dissolved.
  3. Cut filter paper into small squares (1cm × 1cm)
  4. Dip filter paper into antibiotic solution and place 1–2 squares into a small glass/plastic vial with 5 mated females of chosen strain. Repeat for about 5–10 replicates per strain.
  5. Allow females to feed on filter paper overnight.
  6. Remove filter paper and host females individually in 12mm culture vials with 2 hosts each. Re-host females 3–4 times in new vials in two day increments.
  7. Once offspring emerge, select mated female offspring from a family which shows a progressively reduced sex ratio (increasing proportion males over time). This is evidence of CI due to reduced Wolbachia levels in eggs produced after antibiotic treatment. Females from the later hostings with high proportion males are therefore more likely to have reduced Wolbachia loads. Select females from one of these later hostings.
  8. Repeat curing procedure for at least 3 generations in order to completely eliminate the infection. Low levels of Wolbachia may not show CI but can recover over subsequent generations.
  9. Test treated strain for Wolbachia. There are two options:
    1. Phenotypically test by setting up crosses to test for cytoplasmic incompatibility. Set up replicate matings between uninfected virgin females and males from the supposedly cured strain.
    2. Test for Wolbachia with PCR (e.g. Werren and Windsor 2000)

Protocol 6 References
  • Breeuwer JAJ, Werren JH. Microorganisms Associated With Chromosome Destruction and Reproductive Isolation Between Two Insect Species. Nature. 1990;346:558–560. [PubMed]
  • Breeuwer AJ, Werren JH. Cytoplasmic incompatibility and bacterial density in Nasonia vitripennis. Genetics. 1993;135:565–574. [PubMed]
  • Werren JH, Windsor DM. Wolbachia infection frequencies in insects: evidence of a global equilibrium? Proc R Soc Lond B Biol Sci. 2000;267:1277–1285. [PMC free article] [PubMed]

Protocol 7: Field collecting Nasonia from baits

Here we describe our standard method for collecting Nasonia using baits. The “bait” in this case is liver that has been fed on by Sarcophaga bullata larvae, collected after the mature larvae have dispersed and then kept frozen until use (called “flyliver”). This is attractive to the wasps. It is placed in a large (~ 15cm × 15cm) mesh bag that is hung in appropriate collection spots (such as near birds’ nests or under culverts). Within the large mesh bag is a smaller mesh bag containing 4–6 Sarcophaga pupae. These retain the wasps, which will go into the bag and begin stinging the hosts. This smaller bag is place abutting the flyliver. The mesh bags can be made with standard nylon window screening, or any other material with mesh width large enough to permit entry of the wasps.



Flyliver - liver remains fed upon by fly maggots (such as Sarcophaga), collected after the maggots have left the liver (From Protocol 2, Rearing Sarcophaga bullata fly hosts). They contain a volatile substance that is very attractive to the wasps. The flyliver is kept in a freezer (−20 or −80) for storage until approaching the time of use. For transport, we place the flyliver in Falcon tubes or other impermeable container, which prevents odor leakage. When you arrive at your site, keep this under refrigeration or in the freezer or cooler. Do not use fresh liver or rotten liver that has not been fed upon by fly larvae. It does not attract wasps.

Sarcophaga bullata (or other blowfly or fleshfly) pupae. Store under refrigeration (4–10 C) and in a cooler away from the sun when in the field. Do not let the hosts get wet! Make sure they are in a container that can “breathe”. Otherwise they will die and be of little use. Before using in the baits, a few hosts should be cracked open to check whether they are alive (not deflated) and not too old (eyes white to slightly pink is preferred – no body coloration).

Resealable plastic bags, small “sandwich bags”: These are to place the host mesh bag in, once it is removed from the bait. Wasps will be within this.

Resealable plastic bags, quart size – These are to place the bait mesh bag in when it is collected.

Collecting tubes. 12 × 75mm polypropylene tubes for collection of wasps.

Cotton plugs

Sealable shipping tubes, such as 2mL screw-cap vials, if you plan on preserving insects in ethanol (as opposed to live collecting) for shipment.

95% ethanol (for preserving collected insects). See note below about air travel with ethanol.


Large mesh bags – around 20 × 20 cm. Made from nylon/plastic window screening. This bag contains the flyliver attractant and the smaller mesh bag that contains Sarcophaga bullata fly pupae. The bag can be simply made by folding the window screeing and stapling it on two sides, leaving the top open.

Small mesh bags – around 2 by 3 inches. Host pupae (Sarcophaga or other fleshfly or blowfly) will be placed in this bag. It is made in the same way as the large one.

Tape and wire or string (for hanging the bags). The wire used to close plastic bags is well suited for this task


A. Setting up the Baits

Put a piece of flyliver in the large mesh bag.

Place 3 to 6 hosts in the small mesh bag.

Place the small mesh bag inside the large mesh bag so that the small bag contacts the flyliver.

Set the bait hanging in an appropriate place. The bait should be in a location protected from rain if there is any chance of rain (e.g. under bridges, eaves of buildings, culvert, underside of branches). If you wish to collect N. longicornis, N. giraulti or N. oneida, place the baits near bird nests (e.g. barnswallow nests under culverts or in buildings, within or near bird boxes, against trees with nest holes). Baits can also be placed near or under dead carcasses that have fly pupae under them, or larvae about to disperse. The bait can be tied or taped to an object. If rain is a concern, you can also put a cup or some protecting structure above the bait.

Collect the baits

Wasps will arrive at the bait sometimes quickly (e.g. within an hour or so) or later (by the next day). There are two basic ways to collect the baits. First, you can remove the smaller host mesh bag, quickly put it into a resealable bag, and replace it with a new one (leaving the bait hanging). This can be repeated over several days. Alternatively (or at the end of your collection), you can remove the bait bag with the host bag within it, placing both together in a resealable bag. Keep the bags with hosts/wasps out of direct sunlight. Transport the material back to where you will be processing it.

Screen the bait or host mesh bag for wasps. Place them individually in a polypropylene tube with 1–2 fresh hosts and plug the tube with cotton. Let the wasps sting the hosts for 1+ days. You can then ship the wasps/hosts back to your lab, or transport them with you. The offspring will not emerge for 14+ days. Note, you can also keep the hosts from the host bag and use what emerges. Female wasps collected directly from the field are preferred because they represent a “random” sampling of genotypes from the location. Note, if you do not want live wasps, just place the collected females into sealable tubes with 95% ethanol for transport back to the lab. Also note that restrictions in shipping ethanol on airplanes may require you to decant most of this ethanol before shipping, and restoring it back at the lab.

Protocol 5 Figure 1
RNAi injection in Nasonia larvae. A. fourth instar pre-defecation larvae have a slightly contracted gut with gray contents. The posterior end is more tapered than the anterior. B. double-stranded RNA (dsRNA) marked with food coloring is injected posterior ...
Protocol 5 Figure 2
Phenotype of adult wasps from RNAi of cinnabar. Adult wasps on the left were treated as 3rd instar larvae by injection with doublestranded RNA from the eye color gene cinnabar, whereas those on the right received control injections. As can be seen, a ...


JHW and DL acknowledge support from the NIH 1 R24 GM084917-01 and assistance from Rachel Edwards, Jon Giebel, Michael Clark and Rhitoban Raychoudhury.

WWW Resources

  1. Baylor Nasonia Genome Project: ( The site provides information on the Nasonia genome projects, assembly, gene prediction and annotation information, and a geneboree gene and sequence browser.
  2. NCBI Wasp Genome Resources ( This page offers a gateway to Nasonia genome resources at and beyond NCBI. The site will include the Nasonia vitripennis genome mapped onto the linkage groups, expression and other data, gene annotations, et cetera.
  3. Werren Lab Nasonia Site: ( This site provides general background information on Nasonia, including literature information and lists of mutant, wild-type strains.
  4. NasoniaBase ( NasoniaBase is part of the Hymenoptera Genome Database project (, which will provide comparative genomic resources for Hymenoptera species. NasoniaBase will include a genome browser and gene pages with information about transcripts, expression, functional annotation and orthologs.
  5. Indiana Center for Genomics & Bioinformatics ( This site will provides information on and data from the Nasonia expression, genotyping and tiling arrays. The site will be populated with information as it comes available.


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