Daily Factoid: Some insects parasitise plants

In a brilliant bit of parasitism, galling insects drill into plants and affect their metabolism and development in such a way that galls form. The exact molecular mechanisms for this are still largely unknown, but what is known is that these galls are highly-advantageous for the insects. They can take various forms, from being mere modifications of the plant’s nutrient production (inducing greater protein and sugar production for the insect to feed on), to being local growths of nutrient-rich plant tissues on which the insect can munch, to full-blown houses for the insect to live in.

Other insects are leaf-miners, and some of these insects also have the ability to modify the plant’s development. Next autumn, observe the trees with yellow leaves about to drop off. Some of them might have green “islands” on them – areas that are still photosynthetically active even though the rest of the leaf is ageing and dying.

Biochemical analyses of both leaf mines and galls show higher levels of cytokinins, plant hormones that inhibit ageing, maintain chlorophyll, and enhance nutrient release. In other words, the green mined areas are much more nutritious than the rest of the leaf. Like galls, they’re induced by the leaf-mining insects to get an easy source of food – and at a critical time of the year, right before winter.

What’s as interesting as this modification of plant structure by parasitic insects (which is, in principle, similar to behavioural modifications by animal parasites), many studies suggest that the involved cytokinins are not of plant origin, but injected directly by the insects. Going even deeper, the cytokinins injected by the insects aren’t produced by the insects, but by bacterial symbionts. Draw that on a flowchart to see how cool this is: it’s a triple-layered interaction that’s been most likely moulded by millions of years of coevolution.

Further Reading:

General:

The evolution and adaptive significance of the leaf-mining habit

Geometrical Games between a Host and a Parasitoid

Cytokinins and insect galls

Galls:

Manipulation of the phenolic chemistry of willows by gall-inducing sawflies

Mines:

Are green islands red herrings? Significance of green islands in plant interactions with pathogens and pests

Cytokinins:

Extracellular Invertase Is an Essential Component of Cytokinin-Mediated Delay of Senescence

Pathological hormone imbalances

Toxoplasma gondii: Cause of schizophrenia and bad driving

Toxoplasma gondii is an apicomplexan intracellular parasite, most famous for causing toxoplasmosis, and is most probably the most successful protozoan parasite of vertebrates – some estimate that over one billion humans alone are infected. Its relatives include Plasmodium, the malaria-causing parasites, and Cryptosporidium, cause of many foetal deaths worldwide.

In the wild, Toxoplasma uses rodents and many mammals (including humans) as intermediate hosts, not producing reproductive stages in them. They live in these animals as bradyzoites in the tissues, waiting to be ingested by a felid. Once inside the cat’s alimentary tract, it reproduces and gets excreted. Millions of eggs, called sporozoites, can be found in an infected cat’s feces, and they’re infectuous for years. From the feces, it needs to find a route to infect an intermediary host, usually done via some feces-loving insect that picks it up inadvertently, or by getting directly ingested; they can also be waterborne. Once inside a mammal, it begins multiplying rapidly as the trachyzoite stage, forming clusters that then invade tissues and lie there as bradyzoites, and the cycle begins again.

This brings up the story of the success of Toxoplasma gondii. Obviously, the domestication of the cat was a big boon to Toxoplasma‘s evolutionary success, allowing it to spread worldwide, using humans as a readily-available intermediate host. But it also famously induces behavioural changes. It makes rats more active (Webster, 1994), and causes them to hang out next to cats more by getting rid of their built-in aversion to cat odour (Berdoy et al., 2000) and making them attracted to the pheromones in cat urine (Vyas et al., 2007). Such specificity is proof that this is the result of behavioural modification, not just a symptom of illness. The overall result favours the parasite because it only produces eggs in the cat, so it needs the rats to get eaten. By getting the rat to play with the cat urine, Toxoplasma turns it into an easy prey waiting to be eaten.

A much more devious and long-term behavioural modification is its ability to make female rats prefer mating with parasitised males (Dass et al., 2011), the complete opposite of the natural tendency of any animal to avoid mating with parasitised or otherwise diseased mates. This is achieved by the parasite nesting in the testes and getting ejaculated by the male into the female along with semen, thus infecting the female and allowing the parasite to work its neurochemical magic. All this results in more opportunities for transmission.

There are also some reports of human behavioural alteration (Holliman, 1997), namely irritation and aggression, with potentially dire consequences including more car crashes in infected people (Flegr et al., 2002). Infected women are also reportedly more prone to risky behaviour (Flegr et al., 2000). It’s also long been known that an unusual number of schizophrenics suffer from toxoplasmosis (Torrey et al., 2007), with current consensus being that latent toxoplasmosis is the most significant causative factor of schizophrenia (Torrey & Yolken, 1995). The mechanism for this is through dopamine, an important molecule in schizophrenia (Tandon et al., 2010) that’s also found in greater concentrations in toxoplasmosis sufferers (Flegr et al., 2003). Nevertheless, all these are of little evolutionary value for Toxoplasmosis since humans are a dead-end for the parasite’s life cycle, and are probably just a side-effect of its rodent control mechanism (Berdoy et al., 2000). As for an effect on human evolution, there’s a lot of hyperbole floating around about Toxoplasma. Disregard it, at least until we see a major rise in schizophrenia.

Such evolutionary success has made it the subject of an argument about keeping it preserved as a bioweapon to use against hostile extraterrestrials. You know, just in case. I hope you don’t think I’m joking.

Many humans are infected with Toxoplasma, with infection levels varying by geographical region. It’s mostly asymptomatic and kept in check by the immune system, with only acute cases of toxoplasmosis being treated. Infected cats are treatable with monensin and toltrazuril. Adult humans usually just get headaches, fever, and muscle pain, no different than a severe flu. Treatment has a very high success rate using a systemic antifolate antiobiotic combination of pyrimethamine and sulfadiazine.

Since the parasite is transferrable through the placenta (Holliman, 1995), babies are particularly susceptible and most end up stillborn or aborted. People with a compromised immune system (e.g. from AIDS) are also susceptible, so both these cases are treated immediately. Pregnant women can be treated with antiobiotics as above, or can be given a vaccine, Toxovax. Untreated babies that are born asymptomatic may later develop retinitis and learning difficulties as the Toxoplasma nests in the central nervous system (Guerina et al., 1994); if born already expressing symptoms, then they will be hydrocephalic, jaundiced, have low platelet counts, and have retinitis, among other disorders. However, even in these cases, treatment is still possible using the same antiobiotic cocktail as above, just for longer periods.

Although effective, this antibiotic mixture is a rather toxic combination, and doesn’t affect Toxoplasma cysts lying in tissues. I’m sure there is ongoing research for more efficient drugs, no doubt helped by the fact that we’ve been able to cultivate Toxoplasma in the lab for decades now, and we’ve also sequenced its genome, making research on it pretty easy.

One potential avenue for treating Toxoplasmosa is through disruption of its extrachromosomal DNAs. Besides the mitochondrial genome, there’s a 35kb large circular DNA in the apicoplast, a remnant of its photosynthetic ancestry (Funes et al., 2002) and with important functions in growth and development (Fichera & Roos, 1997) by being a biosynthetic center for many compounds (Ralph et al., 2004); the isoprenoid-producing function seems to be the most vulnerable and thus should make a great target for drugs (Nair et al., 2011).

One other factor that makes Toxoplasma relatively easy to target is that it’s highly-clonal, despite being a sexually-reproducing organism (Howe & Sibley, 1995). Such low genetic diversity is great for us, it means less adaptability to whatever we throw at it.

References:

Berdoy M, Webster JP & Macdonald JW. 2000. Fatal attraction in rats infected with Toxoplasma gondii. Proc. R. Soc. B 267, 1591-1594.

Dass SAH, Vasudevan A, Dutta D, Soh LJT, Sapolsky RM & Vyas A. 2011. Protozoan Parasite Toxoplasma gondii Manipulates Mate Choice in Rats by Enhancing Attractiveness of Males. PLoS ONE 6, e27229.

Fichera ME & Roos DS. 1997. A plastid organelle as a drug target in apicomplexan parasites. Nature 390, 407-409.

Flegr J, Kodym P & Tolarová V. 2000. Correlation of duration of latent Toxoplasma gondii infection with personality changes in women. Biological Psychology 53, 57-68.

Flegr J, Havlícek J, Kodym P, Malý M & Smahel Z. 2002. Increased risk of traffic accidents in subjects with latent toxoplasmosis: a retrospective case-control study. BMC Infectious Diseases 2, 11.

Flegr J, Preiss M, Klose J, Havlı́ček J, Vitáková M & Kodym P. 2003. Decreased level of psychobiological factor novelty seeking and lower intelligence in men latently infected with the protozoan parasite Toxoplasma gondii. Dopamine, a missing link between schizophrenia and toxoplasmosis? Biological Psychology 63, 253-268.

Funes S, Davidson E, Reyes-Prieto A, Magallón S, Herion P, King MP & González-Halphen D. 2002. A Green Algal Apicoplast Ancestor. Science 298, 2155.

Guerina NG, Hsu H-W, Meissner HC, Maguire JH, Lynfield R, Stechenberg B, Abroms I, Pasternack MS, Hoff R, Eateron RB, Grady GF & New England Regional Toxoplasma Working Group. 1994. Neonatal Serologic Screening and Early Treatment for Congenital Toxoplasma gondii Infection. NEJM 330, 1858-1863.

Holliman RE. 1995. Congenital toxoplasmosis: prevention, screening and treatment. Journal of Hospital Infection 30 Supplement, 179-190.

Holliman RE. 1997. Toxoplasmosis, behaviour and personality. Journal of Infection 35, 105-110.

Howe DK & Sibley LD. 1995. Toxoplasma gondii Comprises Three Clonal Lineages: Correlation of Parasite Genotype with Human Disease. The Journal of Infectious Diseases 172, 1561-1566.

Meléndez RD. 1996. Toxoplasma gondii: The best terrestrial biological weapon against extraterrestrial invaders? Parasitology Today 12, 166.

Nair SC, Brooks CF, Goodman CD, Strurm A, McFadden GI, Sundriyal S, Anglin JL, Song Y, Moreno SNJ & Striepen B. 2011. Apicoplast isoprenoid precursor synthesis and the molecular basis of fosmidomycin resistance in Toxoplasma gondii. The Journal of Experimental Medicine 208, 1547-1559.

Ralph SA, van Dooren GG, Waller RF, Crawford MJ, Fraunholz MJ, Foth BJ, Tonkin CJ, Roos DS & McFadden GI. 2004. Tropical infectious diseases: Metabolic maps and functions of the Plasmodium falciparum apicoplast. Nature Reviews Microbiology 2, 203-216.

Tandon R, Nasrallah HA & Keshavan MS. 2010. Schizophrenia, “Just the Facts” 5. Treatment and prevention. Past, present, and future. Schizophrenia Research 122, 1-23.

Torrey EF & Yolken RH. 1995. Could Schizophrenia Be a Viral Zoonosis Transmitted From House Cats? Schizophrenia Bulletin 21, 167-171.

Torrey EF, Bartko JJ, Lun Z-R & Yolken RH. 2007. Antibodies to Toxoplasma gondii in Patients With Schizophrenia: A Meta-Analysis. Schizophrenia Bulletin 33, 729-736.

Vyas A, Kim S-K, Giacomini N, Boothroyd JC & Sapolsky RM. 2007. Behavioral changes induced by Toxoplasma infection of rodents are highly specific to aversion of cat odors. PNAS 104, 6442-6447.

Webster JP. 1994. The effect of Toxoplasma gondii and other parasites on activity levels in wild and hybrid Rattus norvegicus. Parasitology 109, 583-589.

Wolbachia: The Ubiquitous Male-Killing, Feminising Parasite

I mentioned Wolbachia in my parthenogenesis post; here I will talk about it in more detail, because it’s a really cool parasite, as well as in the center of much research nowadays.

Index:

• Introduction
• Parasitism
• Transfer
• Biocontrol
• Positive Effects of Wolbachia
• Evolutionary Theory
• Practical Problems
• References

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Introduction

Wolbachia are Gram-negative ⍺-Proteobacteria (order Rickettsiales, family Anaplasmataceae; Dumler et al.,2001), systematically split into 6 supergroups, lettered from A to F (Lo et al., 2002). Those in supergroups C and D have coevolved with their nematodan hosts and are mutualistic (Bandi et al., 1998). Those in supergroups A and B are parasites in arthropods. Morphologically, they come in two groups: ~1µm large rods, and ~0.5µm diameter coccoids. They can be found in a membrane-bound vacuole in the host cell’s cytoplasm (Yen & Barr, 1971); in arthropods, this will be in egg and ovarian cells (see diagram above, from Weiss et al. (2009)), although it has also been found in other tissues, including the Malpighian tubules, and muscle and nervous tissue (Stouthamer et al., 1999). When found in sperm cell cytoplasm, it disappears during spermatogenesis (Clark et al., 2002).

Note that in this post and in the literature, what’s referred to as Wolbachia is Wolbachia pipientis. Before DNA sequencing, when bacteria were classified morphologically, W. persica and W. melaphagi were also thought to be related, but these have nothing to do with the famous Wolbachia (W. persica is a γ-proteobacterium, W. melaphagi is a rhizobacterium).

Being present endosymbiotically in 20+% of all known insect species (Werren & Windsor, 2000), and given that some estimate that over half of all insect species are infected by it (Hilgenboecker et al., 2008), it’s obviously extremely successful, although it should be noted that the virulence of each Wolbachia strain is different (Min & Benzer, 1997), and each species is has a different susceptibility. In Australian and Panamanian fig wasps, Wolbachia has been found to infect over 70% of all species (Haine & Cook, 2005). A host database can be found here.

Parasitism

The biggest reason for its success is its parasitic effect: it’s always transferred maternally in the cytoplasm of the eggs, and in order to ensure maximal transfer rates, it alters its host’s reproductive abilities to favour their own reproductive success (Charlat et al., 2003). It has four ways to do this, in contrast to the one or two ways present in other such parasites:

• Inducing thelytokous parthenogenesis in haplodiploid organisms, e.g. hymenopterans, thus making sure that only female offspring are produced (e.g. Stouthamer et al., 1993). It occurs by forcing the fusion of the two nuclei of the first mitotic division (Huigens et al., 2000).
• Feminizing males, i.e. males reproduce as females (Wilkinson, 1998). The mechanism for this varies; for example, in the pillbug, Wolbachia blocks the formation of the androgenic gland which produces the masculinising androgenic hormone (Martin et al., 1990).
• Killing males, either in the embryonic stages or later (Hurst, 1991). The mechanisms of action are still largely unknown, although recent pioneering research by Riparbelli et al. (2012) shows that it happens due to Wolbachia (purposely?) messing around with male chromosomes at various stages in development, leading to defective embryos and death.
• Inducing cytoplasmic incompatibility (CI) between infected males and uninfected females (Hurst & Werren, 2001), causing sterility (Bordenstein & Werren, 2007) and eventually reproductive isolation within a species.

The latter was the first of Wolbachia‘s effects to be discovered, by Yen & Barr (1971) in mosquitoes; Wolbachia itself was first spotted by Hertig & Wolbach (1924). What happens is that the sperm enters the cell as normal, but its chromosomes don’t decondense and fuse with maternal chromosomes due to a delay in the breakdown of the nuclear envelope in the male’s pronucleus (Tram & Sullivan (2002); see Landmann et al. (2009) for more molecular details), and so can’t enter the first mitosis, meaning they get discarded (Lassy & Karr, 1996). The embryo either becomes a haploid female (in haplodiploid organisms), or it dies. In evolutionary theory terms, in a population susceptible to Wolbachia-induced CI, uninfected females become more unfit, therefore giving a fitness advantage to infected females (Bourtzis et al., 2003), hence explaining how the phenotype persists.

Which manipulation happens cannot be predicted even within the same species, as demonstrated by Hornett et al. (2008), who found that in their North American Hypolimnas bolina butterflies, Wolbachia was a male killer while in Southeast Asian ones, it was a CI-inducer. The difference came from a dominant allele in the SE Asian butterfly genome suppressing the male-killing. Charlat et al. (2007) showed that such a mutation can become fixed in a population in under 10 generations due to the extreme selection pressure to maintain a decent sex ratio, a testament to the ecological power of Wolbachia.

However, in some cases, Wolbachia can be so prominent that the entire affected population gets a very female-biased sex ratio, which in the case of the butterfly Acarea encedon has led to females reversing their sexual roles and behaving like males (Jiggins et al., 2000). In other cases, Wolbachia is counteracted by other elements, for example the B chromosome in the parasitoid wasp Trichogramma kaykai which turns Wolbachia-feminised males back into regular males (Stouthamer et al., 2001).

Transfer

It must be mentioned that part of their success is their ability to be transmitted horizontally across different species (Raychoudhury et al., 2009) – they aren’t host-specific. This has been shown as happening through parasitoids (Heath et al., 1999) or through the environment (e.g. by sharing a common food source (Huigens et al., 2000)). This also means that a single individual may have multiple Wolbachia species (or, better said, strains) co-existing and mingling inside it; the largest number I know of is eight, in the fire ant Solenopsis daguerrei (Dedeine et al., 2005). Note that this ability only comes in the arthropod-associated Wolbachia, whose genomes are more plastic, with recombination and phage-derived elements (Wu et al., 2004), none of which are characteristics present in their nematodan counterparts (Foster et al., 2005).

One study also reported the possibility of Wolbachia having transferred part or all of its genome to its hosts, albeit with only 2% of the genes able to be transcribed and none of them having any described effect (Hotopp et al., 2007).

Biocontrol

The negative effects of Wolbachia are obviously of great interest for biocontrol of pests and disease vectors. For example, Alam et al. (2011) discuss the possibility of using Wolbachia to control the tsetse fly, a vector of trypanosomiasis; Atyame et al. (2011) do the same for mosquitoes. Such biocontrol would work by allowing a chosen genotype to dominate the population by infecting the undesired genotype (e.g. Xi et al. (2005)), or by shortening lifespans to prevent sexual maturity (e.g. Moreira et al. (2009)). Wolbachia can also lead to population bottlenecks with very few individuals becoming able to reproduce (Nice et al., 2009), which is another way to control a pest population.

In non-pest studies, Wolbachia leads to increased susceptibility to parasitoids in Drosophila (Fytrou et al., 2006). It also leads to a less effective immune system in the pillbug Armadillidum vulgare, as seen by a lower density of haemocytes and higher density of bacteria (Bracquart-Vanier et al., 2008). These would be other avenues for pest control if a similar effect is seen in pest groups.

Positive Effects of Wolbachia

Interestingly, the effects of Wolbachia aren’t all negative. In the Cimicidae (bed bugs), Wolbachia is a mutualist; getting rid of it with antibiotics reduces the amount of food the host gets (Hosokawa et al., 2010). In mosquitoes, Wolbachia was found to boost their immune system and cause resistance to dengue virus (Bian et al., 2010). Pinto et al. (2012) describe how this happens at the genetic level. This is another potential use of Wolbachia as a biocontrol agent for disease vectors. In Drosophila, Wolbachia has been shown to confer resistance to several RNA viruses (Teixeira et al., 2008). In a Drosophila lab culture, Weeks et al. (2007) showed that the Wolbachia went from being a parasite to being a mutualist within two decades.

At the extreme end, nematode-infecting Wolbachia are needed for nematode development and fertility (Foster et al., 2005), so Wolbachia antibiotics could be used to control their populations (Taylor & Hoerauf, 1999). This is useful knowledge, given that some of the affected nematodes are vectors for very serious diseases like elephantiasis and onchocerciasis. It’s a similar story with the wasp Asobara tabida, wherein no ovocyte can even be produced when Wolbachia isn’t present (Dedeine et al., 2001) because its absence promotes excessive apoptosis in the ovarioles (Pannebakker et al., 2007).

Other effects of Wolbachia on sexual physiology have been documented, for example an increase in sperm competition in Tribolium beetles (Wade & Chang, 1995), or changes to the spermathecal duct in female Allonemobius crickets (Marshall, 2007).

Evolutionary Theory

An interesting point can be made about the process of molecular evolution in Wolbachia. As I said in the introductory paragraph, Wolbachia is a nematode mutualist and arthropod parasite (generally speaking). One of the Wolbachia genes involved in interaction with the host is wsp, which codes for cell membrane proteins. It was found to be undergoing divergent selection when it is in a parasitic relationship, but not when it’s in a mutualistic relationship (Jiggins et al., 2002). This is in line with what we expect: wsp being involved in host recognition means it theoretically should experience heightened evolutionary rates, and this is confirmed by the empirical data.

Many, if not all, negative and positive effects of Wolbachia have evolved by natural selection in order to maximise the transmission of the strain, either by allowing the bacterium to survive in the host (depressed immune system), or reducing competition by blocking the transmission of other pathogens (as Teixeira et al. (2008) suggest for the viral resistance effect). By extension, this means that parthenogenic arthropods aren’t expected to be Wolbachia hosts, since the manipulations are useless there. In terms of evolutionary theory, they can be treated as nothing more than selfish genetic elements.

When I first heard of Wolbachia, my intuition was that it played a sizeable role in speciation, since it promotes reproductive isolation, or by selecting for subdivided populations (Hatcher et al., 2000). Some analyses showed it not to be true (Rousset & Raymond, 1991), but more and more recent studies are supportive of the idea (Bordenstein, 2003), so it’s accepted as a cause of speciation. It definitely has been demonstrated (Thompson, 1987), and in some cases has also induced rapid speciation (Bordenstein et al., 2001).

On a general evolutionary synthesis level, Wolbachia is pretty interesting as a very recognisable case of inheritable symbiosis, one of the few proper examples that lend credence to the view that symbioses are a driving force behind evolution.

Milder effects of large-scale Wolbachia infection and sex ratio-skewing include altering dispersal ability – many insects have dispersing females and non-dispersing males, or vice versa. On a more influential level, there is also evidence that they can play a role in sexual selection (Jiggins et al., 2000), since sexual conflict gets reduced when levels of polyandry fall (Arnqvist & Rowe, 2005).

Practical Problems

Wolbachia can sometimes present a methodological stumbling block for molecular phylogenies based on mitochondrial DNA, since mtDNA will also hitchike maternally, favouring the maternal mtDNA haplotype, eventually leading to the entire dataset being worthless; see Ballard & Rand (2005) for more information. However, Arthofer et al. (2010) tested this idea using infected bark beetles and found no significant effect from Wolbachia, so it is still unsure just how significant this slight inaccuracy is.

Where it is definitely a problem is in barcoding initiatives using mtDNA. For successful barcoding of a species, a stable molecular marker needs to be used that is guaranteed not to vary across individuals, populations, or ecomorphs. However, there are some studies that show that Wolbachia causes divergences in mtDNA sequences even among individuals of the same species, e.g. in the butterfly genus Hypolimnas (Galtier et al., 2009). The reason for this is that mitochondrial genes are transferred only maternally, so only the maternal set plays a role in evolution. Given the ubiquity of Wolbachia, this is definitely a large problem that should be studied carefully before proceeding with mtDNA barcoding.

Wolbachia alone can’t be cultivated, but it is possible to keep a Wolbachia line using host cell lines (Noda et al., 2002), so experimental evolution studies are possible with them.

For the entomologists among you, make sure to check any colonies for Wolbachia infections, as they could invalidate your results, especially if you’re doing population biology. They can be gotten rid of using any antibiotic. I hear that tetracycline is recommended; if that’s not possible, high heat is enough, since Wolbachia is sensitive to temperature. If you’re sequencing your insects as well, using DNA from the legs is probably the safest way to avoid getting contaminating Wolbachia DNA amplified (this is standard procedure anyway).

Other symbionts that alter the reproduction of their arthropod hosts include Buchnera and Cardinium – but I’ll leave them for other posts.

References:

Alam U, Medlock J, Breisfoard C, Pais R, Lohs C, Balmand S, Carnogursky J, Heddi A, Takac P, Galvani A & Aksoy S. 2011. Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly Glossina morsitans. PLoS Pathogens 7, e1002415.

Arnqvist G & Rowe L. 2005. Sexual Conflict.

Arthofer W, Avtzis DN, Riegler M & Stauffer C. 2010. Mitochondrial phylogenies in the light of pseudogenes and Wolbachia: re-assessment of a bark beetle dataset. ZooKeys 56, 269-280.

Atyame CM, Pasteur N, Dumas E, Tortosa P, Tantely ML, Pocquet N, Licciardi S, Bheecarry A, Zumbo B, Weill M & Duron O. 2011. Cytoplasmic Incompatibility as a Means of Controlling Culex pipiens quinquefasciatus Mosquito in the Islands of the South-Western Indian Ocean. PLoS Neglected Tropical Diseases 5, e1440.

Ballard JWO & Rand DM. 2005. The population biology of mitochondrial DNA and its phylogenetic implications. Annual Review of Ecology, Evolution, and Systematics 36, 621-642.

Bandi C, Anderson TJC, Genchi C & Blaxter ML. 1998. Phylogeny of Wolbachia in Filarial Nematodes. Proc. R. Soc. B 265, 2407-2413.

Bian G, Xu Y, Lu P, Xie Y & Xi Z. 2010. The Endosymbiotic Bacterium Wolbachia Induces Resistance to Dengue Virus in Aedes aegypti. PLoS Pathogens 6, e1000833.

Bordenstein SR. 2003. Symbiosis and the origin of species. In: Bourtzis K & Miller TA (eds.). Insect Symbiosis, Vol. 1.

Bordenstein SR & Werren JH. 2007. Bidirectional incompatibility among divergent Wolbachia and incompatibility level differences among closely related Wolbachia in Nasonia. Heredity 99, 278-287.

Bordenstein SR, O’Hera FP & Werren JH. 2001. Wolbachia- induced incompatibility precedes other hybrid incompatibility in Nasonia. Nature 409, 707-710.

Bourtzis K, Braig HR & Karr TL. 2003. Cytoplasmic incompatibility. In: Bourtzis K & Miller TM (eds). Insect Symbiosis.

Bracquart-Vanier C, Lachat M, Herbinière J, Johnson M, Caubet Y, Bouchon D & Sicard M. 2008. Wolbachia media variation in host immunocompetence. PLoS ONE 3, e3286.

Charlat S, Hurst GDD, Merçot H. 2003. Evolutionary consequences of Wolbachia infections. Trends in Genetics 19, 217-223.

Charlat S, Hornett EA, Fullard JH, Davies N, Roderick GK, Wedell N & Hurst GDD. 2007. Extraordinary Flux in Sex Ratio. Science 317, 214.

Clark ME, Veneti Z, Bourtzis K & Karr TL. 2002. The distribution and proliferation of the intracellular bacteria Wolbachia during spermatogenesis in Drosophila. Mechanisms of Development 111, 3-15.

Dedeine F, Vavre F, Fleury F, Loppin B, Hochberg ME & Boulétreau M. 2001. Removing symbiotic Wolbachia bacteria specifically inhibits oogenesis in a parasitic wasp. PNAS 98, 6247-6252.

Dedeine F, Ahrens M, Calcaterra L, Shoemaker DD. 2005. Social parasitism in fire ants (Solenopsisspp.): a potential mechanism for interspecies transfer of Wolbachia. Molecular Ecology 14, 1543-1548.

Dumler JS, Barbet AF, Bekker CP, Dasch GA, Palmer GH, Ray SC, Rikihisa Y & Rurangirwa FR. 2001. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and ‘HGE agent’ as subjective synonyms of Ehrlichia phagocytophila. International Journal of Systematic and Evolutionary Microbiology 51, 2145-2165.

Foster J, Ganatra M, Kamal I, Ware J, Makarova K, Ivanova N, Bhattachryya A, Kapatral V, Kumar S, Posfai J, Vincze T, Ingram J, Moran L, Lapidus A, Omelchenko M, Kyprides N, Ghedin E, Wang S, Goltsman E, Joukov V, Ostrovskaya O, Tsukerman K, Mazur M, Comb D, Koonin E & Slatko B. 2005. The Wolbachia Genome of Brugia malayi: Endosymbiont Evolution within a Human Pathogenic Nematode. PLoS Biology 3, e121.

Fytrou A, Schofield PG, Kraaljeveld AR & Hubbard SF. 2006. Wolbachia infection suppresses both host defence and parasitoid counter-defence. Proc. R. Soc. B 273, 791-796.

Galtier N, Nabholz B, Glémin S & Hurst GDD. 2009. Mitochondrial DNA as a marker of molecular diversity: a reappraisal. Molecular Ecology 18, 4541-4550.

Haine ER & Cook JM. 2005. Convergent incidences of Wolbachia infection in fig wasp communities from two continents. Proc. R. Soc. B 272, 421-429.

Hatcher MJ, Dunn AM & Tofts C. 2000. Co-existence of hosts and sex ratio distorters in structured populations. Evolutionary Ecology Research 2, 185-205.

Heath BD, Butcher RDJ, Whitfield WGF & Hubbard SF. 1999. Horizontal transfer of Wolbachia between phylogenetically distant insect species by a naturally occurring mechanism. Current Biology 9, 313-316.

Hertig M & Wolbach SB. 1924. Studies on Rickettsia-Like Micro-Organisms in Insects. Journal of Medical Research 44, 329-374.

Hilgenboecker K, Hammerstein P, Schlatmann P, Telschow A & Werren JH. 2008. How many species are infected with Wolbachia? – a statistical analysis of current data. FEMS Microbiology Letters 281, 215-220.

Hornett EA, Duplouy AM, Davies N, Roderick GK, Wedell N, Hurst GDD & Charlat S. 2008. YOU CAN’T KEEP A GOOD PARASITE DOWN: EVOLUTION OF A MALE-KILLER SUPPRESSOR UNCOVERS CYTOPLASMIC INCOMPATIBILITY. Evolution 62, 1258-1263.

Hosokawa T, Koga R, Kikuchi Y, Meng X-Y & Fukatsu T. 2010. Wolbachia as a bacteriocyte-associated nutritional mutualist. PNAS 107, 769-774.

Hotopp JCD, Clark ME, Oliveira DCSG, Foster JM, Fischer P, Torres MCM, Giebel JD, Kumar N, Ishmael N, Wang S, Ingram J, Nene RV, Shepard J, Tomkins J, Richards S, Spiro DJ, Ghedin E, Slatko BE, Tettelin HE & Werren JH. 2007. Widespread Lateral Gene Transfer from Intracellular Bacteria to Multicellular Eukaryotes. Science 317, 1753-1756.

Huigens ME, Luck RF, Klaassen RHG, Maas MFPM, Timmermans MJTM & Stouthamer R. 2000. Infectious parthenogenesis. Nature 405, 178-179.

Hurst LD. 1991. The Incidences and Evolution of Cytoplasmic Male Killers. Proc. R. Soc. B 244, 91-99.

Hurst GDD & Werren JH. 2001. The role of selfish genetic elements in eukaryotic evolution. Nature Reviews Genetics 2, 597-606.

Jiggins FM, Hurst GDD & Majerus MEN. 2000. Sex-ratio-distorting Wolbachia causes sex-role reversal in its butterfly host. Proc. R. Soc. B 267, 69-73.

Jiggins FM, Hurst GDD & Yang Z. 2002. Host-Symbiont Conflicts: Positive Selection on an Outer Membrane Protein of Parasitic but not Mutualistic Rickettsiaceae. Molecular Biology and Evolution 19, 1341-1349.

Landmann F, Orsi GA, Loppin B & Sullivan W. 2009. Wolbachia-Mediated Cytoplasmic Incompatibility Is Associated with Impaired Histone Deposition in the Male Pronucleus. PLoS Pathogens 5, e1000343.

Lassy CW & Karr TL. 1996. Cytological analysis of fertilization and early embryonic development in incompatible crosses of Drosophila simulans. Mechanisms of Development 57, 47-58.

Lo N, Casiraghi M, Salati E, Bazzocchi C & Bandi C. 2002. How Many Wolbachia Supergroups Exist? Molecular Biology and Evolution 19, 341-346.

Marshall JL. 2007. Rapid evolution of spermathecal duct length in the Allonemobius socius complex of crickets: species, population and Wolbachia effects. PLoS ONE 2, e720.

Martin G, Juchault P, Sorokine O & van Dorsselaer. 1990. Purification and characterization of androgenic hormone from the terrestrial isopod Armadillidium vulgare Latr. (Crustacea, Oniscidea). General and Comparative Endocrinology 80, 349-354.

Min K-T & Benzer S. 1997. Wolbachia, normally a symbiont of Drosophila, can be virulent, causing degeneration and early death. PNAS 94, 10792-10796.

Moreira LA, Iturbe-Ormaetxe I, Jeffery JA, Lu G, Pyke AT, Hedges LM, Rocha BC, Hall-Mendelin S, Day A, Riegler M, Hugo LE, Johnson KN, Kay BH, McGraw EA, van den Hurk AF, Ryan PA & O’Neill SL. 2009. A Wolbachia Symbiont in Aedes aegypti Limits Infection with Dengue, Chikungunya, and Plasmodium. Cell 139, 1268-1278.

Nice CC, Gompert Z, Forister ML & Fordyce JA. 2009. An unseen foe in arthropod conservation efforts: the case of Wolbachia infections in the Karner blue butterfly. Biological Conservation 142, 3137-3146.

Noda H, Miyoshi T & Koizumi Y. 2002. In vitro cultivation of Wolbachia in insect and mammalian cell lines. In Vitro Cellular & Developmental Biology – Animal 38, 423-427.

Pannebakker BA, Loppin B, Elemans CPH, Humblot L & Vavre F. 2007. Parasitic inhibition of cell death facilitates symbiosis. PNAS 104, 213-215.

Pinto SB, Mariconti M, Bazzocchi C, Bandi C & Sinkins SP. 2012. Wolbachia surface protein induces innate immune responses in mosquito cells. BMC Microbiology 12 (Suppl. 1), S11.

Raychoudhury R, Baldo L, Oliveira DCSG & Werren JH. 2009. Modes of acquisition of Wolbachia: horizontal transfer, hybrid introgression, and codivergence in the Nasonia species complex. Evolution 63, 165-183.

Riparbelli MG, Giordano R, Ueyama M & Callaini G. 2012. Wolbachia-Mediated Male Killing Is Associated with Defective Chromatin Remodeling. PLoS ONE 7, e30045.

Rousset F & Raymond M. 1991. Cytoplasmic incompatibility in insects: Why sterilize females? TrEE 6, 54-57.

Stouthamer R, Breeuwer JAJ, Luck RF & Werren JH. 1993. Molecular identification of microorganisms associated with parthenogenesis. Nature 361, 66-68.

Stouthamer R, Breeuwer JAJ, Hurst GDD. 1999. WOLBACHIA PIPIENTIS: Microbial Manipulator of Arthropod Reproduction. Annual Review of Microbiology 53, 71-102.

Stouthamer R, van Tilborg M, de Jong JH, Nunney L & Luck RF. 2001. Selfish element maintains sex in natural populations of a parasitoid wasp. Proc. R. Soc. B 268, 617-622.

Taylor MJ & Hoerauf A. 1999. Wolbachia Bacteria of Filarial Nematodes. Parasitology Today 15, 437-442.

Teixeira L, Ferreira Á & Ashburner M. 2008. The Bacterial Symbiont Wolbachia Induces Resistance to RNA Viral Infections in Drosophila melanogaster. PLoS Biology 6, e1000002.

Thompson JN. 1987. Symbiont induced speciation. Biological Journal of the Linnean Society 32, 385-393.

Tram U & Sullivan W. 2002. Role of delayed nuclear envelope breakdown and mitosis in Wolbachia-induced cytoplasmic incompatibility. Science 296, 1124-1126.

Wade MJ & Chang NW. 1995. Increased male fertility in Tribolium confusum beetles after infection with the intracellular parasite Wolbachia. Nature 373, 72-74.

Weeks AR, Turelli M, Harcombe WR, Reynolds KT & Hoffmann AA. 2007. From Parasite to Mutualist: Rapid Evolution of Wolbachia in Natural Populations of Drosophila. PLoS Biology 5, e114.

Weiss BL, Attardo GM & Aksoy S. 2009. Insect–Protozoa–Bacteria Associations: a Model System for Investigating Host–Parasite Interactions. In: Stock SP, Vandenberg J, Glazer I & Boemare N. Insect Pathogens: Molecular Approaches and Techniques, pp. 223-240.

Werren JH & Windsor DM. 2000. Wolbachia infection frequencies in insects: evidence of a global equilibrium? Proc. R. Soc. B 267, 1277-1285.

Wilkinson T. 1998. Wolbachia come of age. Trends in Ecology & Evolution 13, 213-214.

Wu M, Sun LV, Vamathevan J, Riegler M, Deboy R, Brownile JC, McGraw EA, Martin W, Esser C, Ahmadinejad N, Wiegand C, Madupu R, Beanan MJ, Brinkac LM, Daugherty SC, Durkin AC, Kolonay JF, Nelson WC, Mohamoud Y, Lee P, Berry K, Young MB, Utterback T, Weidman J, Nierman WC, Paulsen IT, Nelson KE, Tettelin H, O’Neill SL & Eisen JA. 2004. Phylogenomics of the Reproductive Parasite Wolbachia pipientis wMel: A Streamlined Genome Overrun by Mobile Genetic Elements. PLoS Biology 2, e69.

Xi Z, Dean JL, Khoo C & Dobson SL. 2005. Generation of a novel Wolbachia infection in Aedes albopictus (Asian tiger mosquito) via embryonic microinjection. Insect Biochemistry and Molecular Biology 35, 903-910.

Yen JH & Barr AR. 1971. New Hypothesis of the Cause of Cytoplasmic Incompatibility in Culex pipiens L. Nature 232, 657-658.

Research Blogging necessities :)

R. Stouthamer, J. A. J. Breeuwer, & G. D. D. Hurst (1999). WOLBACHIA PIPIENTIS: Microbial Manipulator of Arthropod Reproduction Annual Review of Microbiology DOI: 10.1146/annurev.micro.53.1.71

Tongue Biters and Deep Sea Giants: The Cymothoida (Crustacea: Isopoda)

The above picture shows a member of the Cymothoida suborder of isopod (Wägele, 1989), containing over 2700 species according to the Smithsonian’s world list of isopods. Cymothoidae (the family, not the suborder; notice the endings!) are well-known across the internet for their wacky parasitic lifestyle (some call it gruesome).

The one pictured at the top of this post is quite unique. It’s either a Ceratothoa or a Cymothoa species, the two most common genera with species exhibiting this tongue-biter phenotype, in which the female attaches to the tongue of the fish. It’s fascinating not only for its strangeness, but also because it’s the only known case of a parasite effectively replacing any of its host’s organs. What happens is the cymothoid attaches to the tongue to feed on blood, but ends up drinking so much that the tongue atrophies and falls off. The cymothoid stays, and the host uses it as its tongue (Brusca & Gilligan, 1983).

As all parasites, parasitic Cymothoida either converge at similar morphologies or get so simplified or convoluted that species recognition is very hard, and the possibility/number of cryptic species is very high. The tongue-biter phenotype described above is quite unique, but the majority of them are so similar even in lifestyle that they can only be recognised by detailed morphological analysis and host identification. If you need ID help with parasitics or non-parasitics, contact me or comment.

I will avoid a discussion of the higher-level relationships of the cymothoids within the isopods, as this is an area of the tree with a lot of confused history and disparate taxon names that have been extensively redefined or have faded into nonexistence. See the introduction in Brandt & Poore (2003) if you’re interested. Current consensus is that Cymothoida is a valid suborder characterised by carnivorous- and parasitic lifestyle-adapted mouthparts (particularly a thin, blade-like slicing mandibular molar process). It contains four superfamilies according to Ahyong et al. (2011) (which itself is most probably based on Brandt & Poore (2003)): Anthuroidea (6 fam., 600+ spp.), Bopyroidea (5 fam., 700+ spp.), Cryptoniscoidea (7 fam., 100+ spp.), and Cymothooidea (9 fam., 1300+ spp.).

I will highlight some of the more well-studied families now. Species numbers come from Ahyong et al. (2011). Here’s what you should look out for: the diversity of lifestyles evident in the Cymothoida. They’re not all parasitic, not even in the same superfamily. There is conservation of a certain lifestyle within each family, and this makes them potentially very interesting as a study system for the evolution of lifestyles at the higher level (and not just of parasitism); I’d be particularly interested in how they became associated with their particular hosts. Of course, this needs a solid phylogenetic framework that doesn’t yet exist. I’m not sure if there’s any ongoing project investigating this (anyone wanna hire me for it?), but it would make a neat research theme for an entire lab.

Cymothooidea

Gnathiidae is a 200+ species-rich family known from all depths, even down to the abyss (Cohen & Poore, 1994), although most species are shallow-marine. Their larvae are ectoparasitic on fish, feeding on blood and tissue fluids. They’re most notable for being the most common coral reef fish parasite (Grutter et al., 2000), and they play a large role in the cleaner fish-host fish mutualism (Grutter, 1999). Infection is always temporary, lasting a couple of minutes and maximum a couple of hours (Paperna & Por, 1977). Adults are non-feeding, often found in association with sponges, polychaete tubes, coral rubble, barnacle nests, or otherwise on the benthos (Wägele, 1988), where they reproduce. The resultant juveniles, the zuphea 1 stage, start the parasitic cycle, finding and feeding on the fish to become the bloated praniza 1 larvae. They detach, becoming the zuphea 2 stage, which finds another host, and the cycle continues until the praniza 3 stage, at which point they migrate to molt to the adult stage (Smit & Davis, 2004). You can tell adult males and females apart by the mandibles, which are greatly enlarged in the male.

The Cymothoidae (380+ spp.) contains the species most commonly pictured on the internet, including the one from the beginning of the post. They’re ectoparasites attaching themselves to the skin, fins, gills or mouth of fish, both marine and freshwater (Brusca, 1981); sometimes they also drill into the musculature, to feed on blood (Trilles, 1991). What fish is infected and where depends on the cymothoid species – they are highly host- and site-specific. The juveniles (mancae; these shouldn’t be confused with larvae, which don’t exist in Cymothoidae) produce anticoagulants to help their blood-feeding, and temporarily attach themselves to any fish they find to feed, since they can only feed on blood (they can’t hunt or scavenge). In effect, they use any fish as intermediate hosts (although some are not so loose, see Tsai et al. (1999)) until they find the correct host fish species, on which they will stay and develop into the male. Cymothoids are protandrous hermaphrodites, so if a female isn’t also present, the male will subsequently turn into a female. Once female, a cymothoid can’t deattach from the host. An environmental sexual determination system is also found, whereby females can secrete pheromones that prolong the masculinity of nearby males by stimulating androgen production (Raibaut & Trilles, 1993). The damage done by cymothoids on their host can be considerable: from decreased growth, weight and size (Lester & Roubal, 1995), through to anemia and tissue damage (Bunkley-Williams & Williams, 1998a), to death (Williams & Bunkley-Williams, 1994). As with the Gnathiidae, a cleaning mutualism exists to get rid of them, except here it’s with cleaner shrimp (Bunkley-Williams & Williams, 1998b). They’ve also radiated in some freshwater rivers, most notably the Amazon.

Aegidae (150+ spp.) are large, opportunistic parasites, attaching themselves to fish temporarily to feed on blood then deattaching and digesting on the benthos. Otherwise, they act as carnivorous scavengers.

Cirolanidae (480+ spp.) includes another famous-on-the-internet creature, the enormous Bathynomus giganteus, pictured above. Bathynomus lives in the deep sea, where it feeds on fish, cephalopods, crabs, and polychaetes. Other cirolanids are also scavengers or predators, most notably attacking trapped fish in nets (I doubt this is of any great economic importance).

Corallanidae (80+ spp.) are predominantly marine, coral reef inhabitants (cf. name), but some are also known from brackish and fresh waters. They attach themselves to turtles, fish, shrimp, and rays, but are predators, not parasites.

Bopyroidea

The Bopyridae (600+ spp.) are ectoparasites, similar to the various Cymothoidea, except they infect decapod crustaceans instead of fish. They attach to the branchial chamber, below the carapace. They can be very effectively used to demonstrate the drastic effects of parasitism on morphological evolution, with the miniaturisation effect extending all the way to the heart, which is reduced to nothing more than a tiny globular organ in the first pleonal segment (Dohrn, 1870). They also lack all lateral cardiac arteries (other isopods have 3-6 of them). It should be noted that similar reductions can also be seen in other parasitics (Cymothoidea also have a smaller heart size), but bopyrids are the most extreme.

The Dajidae (50+ spp.) also infect crustaceans, but have a wider scope than the bopyrids by infecting euphausiids and mysids in addition to shrimp. They also differ by attaching to the carapace, although some will also attach on gills. Examining the Dajidae and Bopyridae alone in the context I mentioned previously would make for a cool study, since they’re closely related and have similar hosts.

If you enjoy this post and the rest of the blog, please help support my research (not cymothoid-related) by sharing and/or donating here! Thanks.

References:

Ahyong ST, Lowry JK, Alonso M, Bamber RN, Boxshall GA, Castro P, Gerken S, Karaman GS, Goy JW, Jones DS, Meland K, Rogers DC & Svavarsson J. 2011. Subphylum Crustacea Brünnich, 1772. In: Zhang Z-Q (ed.) Animal biodiversity: An outline of higher-level classification and survey of taxonomic richness.

Brandt A & Poore GCB. 2003. Higher classification of the flabelliferan and related Isopoda based on a reappraisal of relationships. Invertebrate Systematics 17, 893-923.

Brusca RC. 1981. A monograph on the Isopoda Cymothoidae (Crustacea) of the eastern Pacific. Zoological Journal of the Linnean Society 73, 117-199.

Brusca RC & Gilligan MR. 1983. Tongue replacement in a marine fish (Lutjanus guttatus) by a parasitic isopod (Crustacea: Isopoda). Copeia 3, 813-816.

Bunkley-Williams L & Williams Jr EH. 1998a. Isopods associated with fishes: a synopsis and corrections. Journal of Parasitology 84, 893-896.

Bunkley-Williams L & Williams Jr EH. 1998b. Ability of Pederson Cleaner Shrimp to Remove Juveniles of the Parasitic Cymothoid Isopod, Anilocra haemuli, from the Host. Crustaceana 71, 862-869.

Cohen BF & Poore GCB. 1994. Phylogeny and biogeography of the Gnathiidae (Crustacea: Isopoda) with descriptions of new genera and species, most from south-eastern Australia. Memoirs of the Museum of Victoria 54, 271-397.

Dohrn A. 1870. Untersuchungen über Bau und Entwicklung der Arthropoden, 5. Zur Kenntniss des Baues von Paranthura costana. Zeitschrift für wissenschaftliche Zoologie 20, 81-93.

Grutter AS. 1999. Cleaner fish really do clean. Nature 398, 672-673.

Grutter AS, Lester RJG & Greenwood J. 2000. Emergence rates from the benthos of the parasitic juveniles of gnathiid isopods. Marine Ecology Progress Series 207, 123-127.

Lester RJG & Roubal FR. 1995. Phylum Arthropoda. In: Woo PTK (ed.). Fish Diseases and Disorders, Vol 1. Protozoan and metazoan infections.

Paperna I & Por FD. 1977. Preliminary data on the Gnathiidae (Isopoda) of the Northern Red Sea, the Bitter Lakes, and the Mediterranean and the biology of Gnathia piscivora n. sp. Rapports et Proces-Verbaux des Reunions – Commission Internationale pour l’Exploration Scientifique de la Mer Mediterranee 24, 195–197.

Raibaut A & Trilles JP. 1993. The Sexuality of Parasitic Crustaceans. Advances in Parasitology 32, 367-444.

Romestand MB. 1979. Étude écophysiologique des parasitoses à Cymothoadiens. Annales de Parasitologie 54, 423-448.

Smit NJ & Davies AJ. 2004. The Curious Life-Style of the Parasitic Stages of Gnathiid Isopods. Advances in Parasitology 58, 289-391.

Trilles JP. 1991. Present researches and perspective on Isopoda (Cymothoidae and Gnathiidae) parasites of fishes (systematics, faunistics, ecology, biology and physiology). Wiadomosci Parazytologiczne 37, 141-143.

Tsai M-L, Li J-J & Dai C-F. 1999. Why selection favors protandrous sex change for the parasitic isopod, Ichthyoxenus fushanensis (Isopoda: Cymothoidae). Evolutionary Ecology 13, 327-338.

Wägele JW. 1988. Aspects of the life-cycle of the Antarctic fish parasite Gnathia calva Vanhöffen (Crustacea: Isopoda). Polar Biology 8, 287-291.

Wägele JW. 1989. Evolution und phylogenetisches System der Isopoda: Stand der Forschung und neue Erkenntnisse. Zoologica 140, 1-262.

Williams Jr EH & Bunkley-Williams L. 1994. Four cases of unusual crustacean-fish associations and comments on parasitic processes. Journal of Aquatic Animal Health 6, 202-208.

Research Blogging necessities :)
N. J. Smit, & A. J. Davies (2004). The Curious Life-Style of the Parasitic Stages of Gnathiid Isopods Advances in Parasitology DOI: 10.1016/S0065-308X(04)58005-3
Lucy Bunkley-Williams, & Ernest H. Williams, Jr. (1998). Isopods Associated with Fishes: A Synopsis and Corrections The Journal of Parasitology DOI: 10.2307/3284615

Dryinidae (Hymenoptera)

Dryinids are parasitoid wasps with larvae that infect auchenorrhynchs (Guglielmino & Olmi, 1997). The adult wasps can sometimes be predaceous: the last segments of many dryinid females have long spines on them used to impale prey. These aren’t just the typical parasitoid wasp, they can be active predators.

Myxozoa

The Myxozoa count as some of the most enigmatic organisms known. There’s around 1350 species of them, all tissue and cellular parasites. The majority infect aquatic and marine fish, while some use platyhelminths, annelids, reptiles, amphibians (e.g. Hartigan et al., 2011) or moles as primary hosts. One freshwater bryozoan parasite whose affinity has been debated since 1850, Buddenbrockia plumatella, has now also been placed as a myxozoan (Monteiro et al., 2002). The problem with it is that it looks completely different from other myozoans: it’s a wormish animal with muscles, while the rest of the myxozoans are sporeish.

Freaky Parasites: Chelonus

I’ve already written a post about parasites that affect their hosts’ behaviour, but it’s such a cool subject that I don’t think anyone will mind another example of it :)

Parasites Affecting Insect Behaviour

1) Infected Cephalotes ant.

Parasites need to have adaptations for colonisation, defeating immune systems and transmission. Often, parasitism involves very complex interactions between not-so-closely-related organisms. For example, the jungle-dwelling turtle ant above is infected by nematodes that gather and mate in the gaster (hind part of the ant). They deteriorate the cuticle of the gaster (turning it red), as well as the petiole joint (which joins the gaster to the rest of the ant). Not only that, they somehow mess around with the ant’s brain, making it stay on top of trees and wave its gaster around. The only reason for this is to make the ant’s gaster attractive to birds: they look like fruit and are swinging around, asking to be eaten; the nematodes need the birds to complete their life cycle.