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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.
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 (http://www.rochester.edu/College/BIO/labs/WerrenLab/nasonia/).
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.
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).
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).
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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.
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
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.
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.
JHW and DL acknowledge support from the NIH 1 R24 GM084917-01 and assistance from Rachel Edwards, Jon Giebel, Michael Clark and Rhitoban Raychoudhury.