Sun Spiders of the Family Mummuciidae and the Importance of Natural History Collections

Ricardo Botero-Trujillo, Ph.D.

Theodore Roosevelt Postdoctoral Research Fellow, Department of Invertebrate Zoology and Richard Gilder Graduate School, American Museum of Natural History, N.Y., U.S.A.

Figure 1.  Dorsal aspect of an adult male of a new (still undescribed and unnamed) species of mummuciid solifuge. Note the three dark band pattern characteristic of the family, the very distinct morphology of the first pair of legs and pedipalps, and the big chelicerae. Photo: R. Botero Trujillo.

Figure 1. Dorsal aspect of an adult male of a new (still undescribed and unnamed) species of mummuciid solifuge. Note the three dark band pattern characteristic of the family, the very distinct morphology of the first pair of legs and pedipalps, and the big chelicerae. Photo: R. Botero Trujillo.

All of our readers surely know what a spider is. Loved by some, feared by others, they are found almost everywhere. Look carefully and you will find them living in your home or working place.

Spiders are the most famous order within the class, Arachnida, however, several other orders of arachnids exist. Some of them are reminiscent of spiders in certain aspects, whereas others have notably different appearances (for example, scorpions). In many languages, including English and Spanish, the word “spider” (or equivalent) is combined with other words that together refer to some of those other “spider-looking” arachnids. For example, the arachnid order Amblypygi is commonly referred to as whip spiders, while the lesser known arachnid order, Ricinulei, is called the hooded tick spiders, although neither of these orders are actually spiders.

For the Taxon of the Month, we focus on another arachnid order known as the sun spiders or camel spiders, but referred to in the scientific community as Solifugae. As you may have guessed, solifuges are not spiders (clearly not camels either). Most species in this >1100 species group are nocturnal (active at night). Therefore, the common name sun spider is only technically accurate for diurnal (active during the day) species (Cloudsley-Thompson 1977). One solifuge family that is active during the day is Mummuciidae (Figure 2). This is one out of 12 families of living solifuges, and one of only four present in the Americas (Maury 1984). Most Mummuciidae species are quite small, not surpassing 20 mm in total length although most individuals are much smaller than that! One species in this family, Vempironiella aguilari, described from Peru has the smallest males among all solifugaes with adult males not reaching 6 mm (Botero-Trujillo 2016). Despite their small size, mummuciids are ferocious and voracious animals, feeding on practically any prey that they can kill (Cloudsley-Thompson 1977).

Figure 2.  One of multiple arid to semi-arid landscapes where mummuciid Solifuges can be found active during daylight hours of the hottest days. Locality: Mendoza Province, Argentina. Photo: R. Botero Trujillo.

Figure 2. One of multiple arid to semi-arid landscapes where mummuciid Solifuges can be found active during daylight hours of the hottest days. Locality: Mendoza Province, Argentina. Photo: R. Botero Trujillo.

Mummucids are entirely diurnal animals. They are quite abundant in open areas, and can be found during the hottest months of the year (Figure 2). They can barely be spotted when they are running, for they are extremely fast, especially males. In contrast, females are most often found digging burrows in the substrate, which they use for refuge and egg deposition. When studied under a stereomicroscope (or even when seen in Figure 1) members of Mummuciidae can very easily be recognized by the presence of three longitudinal dark bands on the dorsal surface. This coloration pattern is very unique among South American solifuges, and only reminiscent of some other diurnal species in the Old World (Botero-Trujillo 2018).

Figure 3.  Detail on dorsal aspect of the anteriormost part (prosoma) of an adult female belonging to another new mummuciid species. The two massive structures are the chelicerae. With hard cuticle and powerful muscles attached inside of them, these serve the female to dig burrows in the substrate, and to kill and tear apart preys. Photo: R. Botero Trujillo.

Figure 3. Detail on dorsal aspect of the anteriormost part (prosoma) of an adult female belonging to another new mummuciid species. The two massive structures are the chelicerae. With hard cuticle and powerful muscles attached inside of them, these serve the female to dig burrows in the substrate, and to kill and tear apart preys. Photo: R. Botero Trujillo.

Like other Solifuges, mummuciids have very large chelicerae (jaws), which are among the largest and most powerful in Arachnida (Figure 3). Chelicerae are used to kill and tear apart prey, but also have other functions. Females use the chelicerae for digging burrows, whereas the chelicerae of males are morphologically and functionally modified to transfer sperm into females (Figure 4; Bird et al. 2015). For this reason, it is not surprising that males have developed taxon-specific morphologies for their chelicerae, and consequently, the chelicerae of males represent an extremely important structure for uncovering solifuge diversity (Bird et al. 2015; Botero-Trujillo et al. 2017). Despite their aggressiveness and high cheliceral strength relative to their body size, mummuciids, and solifuges in general, are harmless to humans. They lack venom, relying completely on speed and their strong chelicerae for hunting.

Figure 4.  Chelicerae of female (A) and male (B) of a mummuciid solifuge. Note the different morphologies between them, and that the male has a dorsal translucent structure, which is absent in females named the “flagellum,” that is involved in mating. Photos: R. Botero Trujillo.

Figure 4. Chelicerae of female (A) and male (B) of a mummuciid solifuge. Note the different morphologies between them, and that the male has a dorsal translucent structure, which is absent in females named the “flagellum,” that is involved in mating. Photos: R. Botero Trujillo.

Figure 5.  Scanning electron microscopy of the leg-like (but not walk-involved) pedipalps (A) and foremost leg (B), of a new mummuciid species. Photos: R. Botero Trujillo.

Figure 5. Scanning electron microscopy of the leg-like (but not walk-involved) pedipalps (A) and foremost leg (B), of a new mummuciid species. Photos: R. Botero Trujillo.

Another interesting fact about Mummuiciidae, and solifuges in general, is that despite being an arachnid and having eight legs, they actually only use six legs for walking (much like insects do; Punzo 1998). They use their first pair of legs as sensory appendages, raising them above the substrate, detecting cues, much like insect antennae (Figure 5). But that’s not all. The pedipalps (Figure 5), which have a common ancestry with scorpion “arms”, are also modified in solifuges. They are leg-like in appearance, serve a sensory function and have an adhesive structure, which is eversible, allowing them to literally climb on surfaces as smooth as glass, as well as capture jumping prey (Cushing et al. 2005; Klann et al. 2008; Willemart et al. 2011)!

Despite Mummuciidae being a widespread taxon in arid and semi-arid regions of most South American countries, little is known about this family. Very little information exists on the habitat requirements, diversity and reproduction of this family and no comprehensive examination of the group exists. For my Ph.D. research (Botero-Trujillo 2018), I studied the diversity of mummuciid solifuges, addressing their morphology, distribution, evolution, and classification at the Argentinean Museum of Natural Sciences, in Buenos Aires, while being funded by Argentinean government-sponsored doctoral fellowship.

Back into 2013, when I started my research, only 18 species were known in Mummuciidae. While conducting my research, I borrowed solifuge specimens from 27 museums and institutions from all around the world. This was only possible thanks to the institutional support that museums offer to scientific projects. Whilst the whole previous knowledge about Mummuciidae had been built on a handful of specimens (less than a hundred) (e.g., Roewer 1934), use of resources from all the collaborating institutions allowed me to gather over 6000 specimens belonging to this family, making this study the largest ever conducted on any group of solifuges. Likewise, field work conducted in Argentina, Chile, Brazil, Ecuador and Peru, contributed additional specimens and data.

Results were remarkable, although not necessarily unexpected considering the amount of material available and the scarce knowledge about this family. For instance, 40 new species were discovered, tripling the known diversity of the family. Of these, five new species have already been described and published in scientific journals (Botero-Trujillo 2016; Botero-Trujillo et al. 2017), whereas all others are currently in process of description and are expected to be formally named in the course of the next year.

My work with Mummuciidae would not have been possible without natural history museums. The current epoch we live in, with rapid industrial development and a growing human population, is accelerating the rate of species extinctions, by destroying habitats and causing global climate change. All of what is known about the diversity of Mummuciidae, including the recent discoveries here highlighted, have only been possible thanks to the biological collections and scientific museums around the globe. Museums serve as the pillar of a large inter-institutional and interdisciplinary network, connecting researchers from every country. They are also guardians and reservoirs of the planet’s past and present biodiversity. The existence of biological collections makes it possible that specimens and many other kinds of data sources will be available for study for many years to come.

Unfortunately, natural history collections are not indestructible, and at the beginning of September, a fire destroyed the collections of the Brazilian National Museum in Rio de Janeiro (see Greshko 2018). That terrible fire, destroyed millions of irreplaceable artifacts and animal specimens, including numerous type specimens. Type specimens are individuals to which the identity of the species is tied upon discovery, which makes them the reference specimen for the recognition of the species. Among the types destroyed in the fire at the Brazilian National Museum were specimens of two species of Mummuciidae: Mummucina exlineae and Cordobulgida bruchi. Sadly, Mummucina exlineae, was only known from the single type specimen housed at the Brazilian National Museum, so now, no other material of this species exists in the world and the only knowledge of these species exists in the form of publications and photographs. During my Ph.D., I was able to borrow the type material of these two mummuciid species and examine them only a few months before the fire occurred.

Museums and biological collections need to be taken care of. Their importance does not lie only in the exhibitions open to the public, but is limitless, on account of all the new information that is associated with the specimens stored in the collections. Several discoveries and contributions to science that collections can contribute will require many decades of work, and therefore, many decades are worth dedicating efforts to protect them.

About the author

Ricardo Botero-Trujillo completed his Biology B.S. at Javeriana University in Bogotá, Colombia. After a five-year period working in the pharmaceutical industry, Ricardo moved to Argentina where he earned a Ph.D. in Biological Sciences from the Universidad de Buenos Aires. His research mainly focusses on the systematics, taxonomy, and evolution of three arachnid groups: scorpions, solifuges and ricinuleids. Since 2006, he has described several new species of these three groups from different South and Central American countries. Ricardo joined the Division of Invertebrate Zoology at the American Museum of Natural History with a Theodore Roosevelt Postdoctoral Research Fellowship from the Richard Gilder Graduate School at the AMNH in August 2018. His current research covers aspects of Ricinulei evolution.

All photos taken by the author in the context of his doctoral thesis (Botero-Trujillo 2018).


Bird, T.L., R. Wharton & L. Prendini. 2015. Cheliceral morphology in Solifugae (Arachnida): primary homology, terminology and character survey. Bulletin of the American Museum of Natural History, 394, 355 pp.

Botero-Trujillo, R. 2016. The smallest known solifuge: Vempironiella aguilari, new genus and species of sun-spider (Solifugae: Mummuciidae) from the coastal desert of Peru. Journal of Arachnology, 44, 218–226.

Botero-Trujillo, R. 2018. Revisión Sistemática y Filogenia de los Solífugos de la Familia Mummuciidae (Arachnida, Solifugae). PhD Thesis. Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, Argentina. Volumes I and II: 336 pp. + 209 pp. [unpublished].

Botero-Trujillo, R., R. Ott & L.S. Carvalho. 2017. Systematic revision and phylogeny of the South American sun-spider genus Gaucha Mello-Leitão (Solifugae: Mummuciidae), with description of four new species and two new generic synonymies. Arthropod Systematics & Phylogeny 75: 3–44.

Cloudsley-Thompson, J.L. 1977. Adaptational biology of Solifugae (Solpugida). Bulletin of the British Arachnological Society 4: 61–71.

Cushing, P.E., J.O. Brookhart, H.-J. Kleebe, G. Zito & P. Payne. 2005. The suctorial organ of the Solifugae (Arachnida, Solifugae). Arthropod Structure & Development 34: 397–406.

Greshko, M. (2018, September 6) Fire devastates Brazil's oldest science museum. "National Geographic Society," – Available at:

Klann, A.E., A.V. Gromov, P.E. Cushing, A.V. Peretti & G. Alberti. 2008. The anatomy and ultrastructure of the suctorial organ of Solifugae (Arachnida). Arthropod Structure & Development 37: 3–12.

Maury, E.A. 1984. Las familias de solífugos Americanos y su distribución geográfica (Arachnida, Solifugae). Physis (Buenos Aires), secc. C 42: 73–80.

Punzo, F. 1998. The Biology of Camel Spiders (Arachnida, Solifugae). Kluwer Academic Publishers, Norwell, Massachusetts, U.S.A., 301 pp.

Roewer, C.F. 1934. Solifuga, Palpigrada. In: Bronn, H.G. (Ed.), Klassen und Ordnungen des Tierreichs. V Band: Arthropoda. IV Abteilung: Arachnoidea und kleinere ihnen nahegestellte gruppen; Akademische Verlagsgesellschaft mbH. Leipzig, pp. 481–723.

Willemart, R.H., R.D. Santer, A.J. Spence, & E.A. Hebets. 2011. A sticky situation: Solifugids (Arachnida, Solifugae) use adhesive organs on their pedipalps for prey capture. Journal of Ethology 25: 177–180.

Darters (Family Percidae)

Kara Million

Ph.D. Candidate, Department of Biology (Evolution, Ecology, and Behavior), Indiana University, Bloomington, Indiana

Figure 1.  Greenside Darter ( Etheostoma blennioides ). Photo by Derek Wheaton.

Figure 1. Greenside Darter (Etheostoma blennioides). Photo by Derek Wheaton.

Figure 2.  Bluemask Darter ( Etheostoma akatulo ). Photo by Derek Wheaton.

Figure 2. Bluemask Darter (Etheostoma akatulo). Photo by Derek Wheaton.

Figure 3.  Rainbow Darter ( Etheostoma caeruleum ). Photo by Derek Wheaton.

Figure 3. Rainbow Darter (Etheostoma caeruleum). Photo by Derek Wheaton.

Figure 4.  Johnny Darter ( Etheostoma nigrum ). Photo by Derek Wheaton.

Figure 4. Johnny Darter (Etheostoma nigrum). Photo by Derek Wheaton.

When you look into an eastern North American stream in the springtime, you probably see little shapes flitting through the water here and there, and never give them a second thought. But if you look a little more closely, you might be astonished to see a parade of radiant colors and patterns displayed by feisty fish no larger than your finger. You might think that they were exotic tropical fish released into the water by irresponsible pet owners, but they are in fact native fish called Darters, who have inhabited those waters for millions of years.

Figure 5.  Fantail Darter ( Etheostoma flabellare ). Egg mimics are visible on the first dorsal fin. Photo by Derek Wheaton.

Figure 5. Fantail Darter (Etheostoma flabellare). Egg mimics are visible on the first dorsal fin. Photo by Derek Wheaton.

Darters (Family Percidae) are a large group of tiny, colorful freshwater fishes found only in North America. There are more than 200 described species, some of which were discovered just recently (Kuehne & Barbour 1983; Page 1983, Page & Burr 2010). Their common name refers to the fact that they are built for acceleration and can rapidly “dart” backwards or forwards in bursts of speed. Darters fall into four to five genera (Ammocrypta, Crystallaria, Percina, Etheostoma, and- according to some researchers- Nothonotus) (Near et al. 2011), the largest of which is by far Etheostoma, with 157 species. They are broadly distributed in streams across the eastern United States and parts of Canada, as well as Mexico. The geographic ranges of individual species vary widely (Page & Burr 2010); while species such as the Greenside Darter (Etheostoma blennioides) are found as far south as Alabama and as far north as Canada (Page 1983), species such as the Bluemask Darter (Etheostoma akatulo) (Laymen & Mayden 2009) are restricted to a single river system, others even to a single stream.

Figure 6.  Juvenile Blotchside Logperch ( Percina burtoni ) propagated at Conservation Fisheries. Photo by Derek Wheaton.

Figure 6. Juvenile Blotchside Logperch (Percina burtoni) propagated at Conservation Fisheries. Photo by Derek Wheaton.

Darters live primarily in stream habitats with clear, fast-flowing, oxygen rich water. However, microhabitat preferences can vary by species. For example, Rainbow Darters (Etheostoma caeruleum) prefer to inhabit riffles where water flows rapidly over cobble, while Johnny Darters (Etheostoma nigrum) prefer sandy substrate with some gravel, and can inhabit cloudy water. Darters primarily feed on stream invertebrates, and are prey items for larger fish, as well as some snakes and birds. Multiple species frequently co-occur in streams, although microhabitat preferences allow for effective niche partitioning between species (Kuehne & Barbour 1983). However, introgressive hybridization has been observed between some closely related species (Harrington et al. 2012).

Figure 7.  Candy Darters ( Etheostoma osburni ). The male (bottom right) displays striking nuptial coloration in contrast to the female (top left). Photo by Derek Wheaton.

Figure 7. Candy Darters (Etheostoma osburni). The male (bottom right) displays striking nuptial coloration in contrast to the female (top left). Photo by Derek Wheaton.

Darter species vary dramatically in their secondary sexual characters and reproductive behaviors. One particularly striking example are male nuptial coloration differences found between species. During the breeding season (which in most species occurs in the spring), males of many species undergo dramatic transformations in which they exhibit brilliant colors and patterns (Kuehne & Barbour 1983). Although this has long been thought to attract mates and appeal to female preferences, recent research shows that male breeding coloration might actually serve as a signal that mediates male-male competition: males recognize their own species colors and patterns and can direct aggressive behavior towards males of their own species, while avoiding aggressive interactions with heterospecific males who are unlikely to pose competition for mates (Moran & Fuller, in press). The extent to which females prefer bright male coloration is still under investigation.

Figure 8.  Candy Darter males in breeding condition evaluate each other as potential competition. Photo by Derek Wheaton.

Figure 8. Candy Darter males in breeding condition evaluate each other as potential competition. Photo by Derek Wheaton.

Darters also vary in their spawning behavior. In species where male breeding colors are more prominent, spawning typically occurs on the substrate or among aquatic plants, and the eggs are not tended by either parent. In species such as the Fantail Darter (Etheostoma flabellare), whose males are less colorful, however, eggs are attached to the undersides of rocks, and males provide all the parental care for the eggs. Males will fan the eggs with their tails and clean the eggs with their mouths until the fry hatch (Kuehne & Barbour 1983; Page 1983). Females are attracted to males who already possess egg clutches from other females, presumably because this suggests the males are capable caregivers (Knapp & Sargent, 1989). Males, meanwhile, will take over and “adopt” nests of genetically unrelated eggs to increase mating opportunities, since multiple females will contribute to a single existing nest. Males in breeding condition also exhibit small ornaments on their dorsal fins called “egg mimics” that closely resemble darter eggs. Several studies demonstrate that females prefer males who possess egg mimics (Strange, 2001; Knapp & Sargent, 1989).

Figure 9.  Brown Darter ( Etheostoma edwini ). Photo by Derek Wheaton.

Figure 9. Brown Darter (Etheostoma edwini). Photo by Derek Wheaton.

Although Darters are too small to be of interest to most game fishermen, they serve an important purpose as biological indicators of stream health. Darters are highly sensitive to habitat quality and water cleanliness. Many species are threatened or endangered, largely due to poor water management practices and habitat destruction (Grabarkiewicz & Davis 2008).

Figure 10.  Tangerine Darter ( Percina aurantiaca ) .  Photo by Derek Wheaton.

Figure 10. Tangerine Darter (Percina aurantiaca). Photo by Derek Wheaton.

Despite the many threats faced by these vulnerable fish, there is hope for their future. Organizations, such as Conservation Fisheries in Tennessee, are working to restore native fish habitats, while breeding threatened species in captivity to maintain “arks” against extinction. After successful habitat restoration, these fish will then be reintroduced to their natural habitats. Up to this point, Conservation Fisheries has successfully propagated more than 65 vulnerable native fish species, including many Darters ( 2018). While captive breeding programs are a beneficial step to preserve native biodiversity, long-term habitat management solutions are needed to ensure the continued survival of Darters in the wild.

Figure 11.  Redline Darter ( Etheostoma rufilineatum , or  Nothonotus rufilineatus ). Photo by Derek Wheaton.

Figure 11. Redline Darter (Etheostoma rufilineatum, or Nothonotus rufilineatus). Photo by Derek Wheaton.

Given their diverse characteristics and behaviors, darters are wonderful models for multiple areas of scientific research spanning ecology, evolution, animal behavior, and conservation biology. The extreme diversity of the darter group makes them excellent for studies in phylogenetics and diversification (Near et al. 2011). Their life histories have been documented extensively by ichthyologists for decades (Page 1983). The variation in reproductive behaviors and social structures have made them appealing to researchers interested in mate choice and behavioral isolation between species (Mendelson et al. 2018, Williams & Mendelson 2013). One research group has even recorded male vocalizations (“singing”) in several darter species (Noel & Johnston 2015).

Figure 12.  Tennessee Snubnose Darter ( Etheostoma simoterum ). Photo by Derek Wheaton.

Figure 12. Tennessee Snubnose Darter (Etheostoma simoterum). Photo by Derek Wheaton.

Additionally, darters are hosts to a variety of local parasites, including a genus of gill parasite (Aethycteron) that infects darters as their preferred host group. Many of the parasites in this genus have been found on only one darter species each, demonstrating a high degree of specialization on their hosts (Suriano & Beverly-Burton 1982). Thus, co-occurring darter species have presented with dramatically differing gill parasite burdens (Hanson & Stallsmith 2013, Million et al. unpublished data). However, we even find within-species parasite load variation: in Fantail Darters, peak gill parasite infection tends to occur during the hosts’ breeding season, and males are generally more heavily infected than females (Million et al. 2017). My current research focuses on the intersection of parasitism, mate choice, and immunogenetic diversity in Darters. For this, I use Darters of Indiana and several of their local parasites (including Aethycteron) as models to evaluate hypotheses designed to explain the high diversity of MHC genes (genes that code for a component of the vertebrate immune system). The variation in host reproductive behaviors and characteristics and the presence of multiple wild pathogens that interact closely with their hosts make Darters an excellent system to study the mechanisms that maintain immunogenetic diversity in vertebrates.

About the author

Kara Million is a Ph.D. candidate in Ecology, Evolution, and Behavior at Indiana University. She is advised by Dr. Curtis Lively. Her research focuses on parasitism, mate choice, and immunogenetic diversity in Darters.


Grabarkiewicz, J.D. & W.D. Davis. 2008. An introduction to freshwater fishes as biological indicators. Washington, DC: Environmental Protection Agency. 96 p.

Hanson, R.G. & B.W. Stallsmith. 2013. Patterns of infection by monogenoideans in an assemblage of darters. J. Freshw. Ecol. Available at:

Harrington, R.C., E. Benavides & T. J. Near. 2012. Phylogenetic inference of nuptial trait evolution in the context of asymmetrical introgression in North American darters (Teleostei). Evolution 67(2): 388-402.

Knapp, R.A & R.C. Sargent. 1989. Egg-mimicry as a mating strategy in the fantail darter, Etheostoma flabellare: females prefer males with eggs. Behav. Ecol. Sociobiol. 1989(25) :321-326.

Kuehne, R.A. & R.W. Barbour. 1983. The American darters. Lexington, KY: University Press of Kentucky. 208 p.

Layman, S.R. & R.L. Mayden. 2009. A new species of the darter subgenus Doration (Percidae: Etheostoma) from the Caney Fork River System, Tennessee. Archived October 2, 2011, at the Wayback Machine. Copeia 2009(1): 157-170.

Mendelson, T.C., J.M. Gumm, M.D. Martin & P.J. Ciccotto. 2018. Preference for conspecifics evolves earlier in males than females in a sexually dimorphic radiation of fishes. Evolution 72(2): 337-347.

Million, K.M., C.L. Tarver & B.W. Stallsmith. 2017. Does Infection by the Monogenoidean Gill Parasite Aethycteron moorei Affect Reproductive Ecology of the Darter Etheostoma flabellare in Mill Creek, Tennessee? Copeia 105(1):75-81.

Moran, R.L. & R.C. Fuller. In press. Agonistic character displacement of genetically based male color patterns across darters. Proc. R. Soc. London, B. doi:10.1098/rspb.2018.1248 (bioRxiv preprint)

Near, T.J., C.M. Bossu, G.S. Bradburd, R.L. Carlson, R.C. Harrington, P.R. Hollingsworth, B.P. Keck & D.A. Etnier. 2011. Phylogeny and temporal diversification of darters (Percidae: Etheostomatinae). Syst. Biol. 60 (5): 565-595.

Noel, P.S. & C. Johnston. 2015. Geographic variation in acoustic signaling in the guardian darter Etheostoma oophylax: effects of contemporary versus historical isolation. Environ. Biol. Fishes 98:1355–1363.

Page, L.M. 1983. Handbook of Darters. Neptune, NJ: TFH Publications. 272 p.

Page, L.M. & B.M. Burr. 2010. A field guide to freshwater fishes of North America north of Mexico. Boston, MA: Houghton Mifflin Company. 432 pp.

Strange, R.M. 2001. Female preference and the maintenance of male fin ornamentation in three egg-mimic darters (Pisces: Percidae). J Freshw Ecol 16(2):267-271.

Suriano, D.M. & M. Beverly-Burton. 1982. Aethycteron n.g. (Monogenea: Ancyrocephalinae) from darters (Percidae: Etheostomatini) in Ontario, Canada with descriptions of A. caerulei n.sp., A. micropercae n.sp., and A. nigrei n.sp. from Etheostoma spp. Can. J. Zool. 60:1397-1407.

Williams, T.H. & T.C. Mendelson. 2013. Male and female responses to species-specific coloration in darters (Percidae: Etheostoma). Anim Behav. 85:1251-1259.

Poxvirus Evolution

Sara Reynolds, Ph.D.

Assistant Professor, Fairleigh Dickinson University, Madison, New Jersey

Figure 1.  Images of poxvirus infections in humans. (Left) A young Bangladeshi villager in 1974 is one of the last known infections of a human with variola virus ( McFadden et al. 2010 ). (Middle) Severe, generalized cowpox virus infection of a 4 year-old girl in Finland ( Pelkonen et al. 2003 ). (Right) A 7 year-old girl displaced from the Democratic Republic of the Congo in 2010 shows a rash confirmed as Monkeypox. Her uncle died of the infection ( Reynolds et al. 2013 ). (Obtained from: National Library of Medicine, History of Medicine Division, Image Database. Open Source.

Figure 1. Images of poxvirus infections in humans. (Left) A young Bangladeshi villager in 1974 is one of the last known infections of a human with variola virus (McFadden et al. 2010). (Middle) Severe, generalized cowpox virus infection of a 4 year-old girl in Finland (Pelkonen et al. 2003). (Right) A 7 year-old girl displaced from the Democratic Republic of the Congo in 2010 shows a rash confirmed as Monkeypox. Her uncle died of the infection (Reynolds et al. 2013). (Obtained from: National Library of Medicine, History of Medicine Division, Image Database. Open Source.

It is widely accepted that the basic unit of all life is a cell, a single entity composed of a double-stranded DNA genome containing regions called genes. These genes encode the information necessary to make ribosomes, enzymes that synthesize proteins and act as the core of the cell’s metabolism, as well as other proteins and enzymes that respond to the cell’s environment. Surrounding the genome is a gelatinous cytoplasm where much of the cell’s metabolism takes place. All of this is bounded within an outer cell membrane.

Some organisms have evolved complexity above this basic cellular structure (check out some of the previous Taxon of the Month posts). For example, eukaryotes have extra membranes to organize and separate cellular components.

Figure 2.   Vaccinia  virus   particles. Poxvirus particle morphology demonstrates the structural adaptations that allow for the poxvirus lifecycle. These include the presence of a dumbbell shaped core (C), two protein-filled lateral bodies (LB) and multiple host-derived membranes that wrap around the outside of the particle (M). (Obtained from: Gray et al., 2016 in National Library of Medicine, History of Medicine Division, Image Database. Open Source.  Creative Commons, Attribution 4.0 International (CC By 4.0 License . Modified with figure cropped. Original  here .)

Figure 2. Vaccinia virus particles. Poxvirus particle morphology demonstrates the structural adaptations that allow for the poxvirus lifecycle. These include the presence of a dumbbell shaped core (C), two protein-filled lateral bodies (LB) and multiple host-derived membranes that wrap around the outside of the particle (M). (Obtained from: Gray et al., 2016 in National Library of Medicine, History of Medicine Division, Image Database. Open Source. Creative Commons, Attribution 4.0 International (CC By 4.0 License. Modified with figure cropped. Original here.)

In contrast to cells, the basic unit of a virus is a genome composed of either single- or double-stranded DNA or RNA surrounded by a protein coat called a capsid. That’s it - no ribosomes or gelatinous cytoplasm and no metabolism of their own. The number of genes found on a virus is surprisingly small. For example, while the bacteria E. coli has about 500 (Lukjancenko, 2010) and you (a human) have about 19,000 -25,000 (Ezkurdia 2014), the ever burdensome HIV virus has only 9 genes (Li 2015) in its genome.

While viruses might lack morphological complexity, they certainly have evolved a wide range of behaviors. For example, some viruses wrap themselves in membranes derived from host cells (Figure 2; Buchmann 2015). Most viruses have a narrow host range, but some viruses have accumulated enough adaptations that they actually develop the ability to infect new host species or types of host cells, broadening their host range (Haller 2014).

As our understanding of the genomic material of humans and viruses has increased one very interesting aspect of evolution has emerged; viruses and their hosts sometimes exchange genetic material. Examples of viral genetic material has been found in the genomes of animals (Blanc 2015), plants (Maumus 2014) and more (Sharma 2014). As a matter of fact, the encounters that eukaryotes, including humans, have with viruses appears to provide an “evolutionary push” (Enard 2016; Moeling 2013; Yi 2004). Similarly, viruses can capture host genes and adapt them for use in the viral life cycle (Filée 2014). In poxviruses, many of these genes are altered to inhibit the host defense against the viral infection (Ouyang 2014; Martin 2012).

The discovery of larger, complex viruses has added new fuel to the debate regarding whether viruses are alive (like cells) and the implications that has regarding the origin of viruses. Recently discovered Klosneuviruses have genomes that are more cell-like than any previously identified virus with the largest known panel of enzymes critical for ribosome activity (Schulz 2017), but they still require host ribosomes. These viruses belong to the Nucleo-Cytoplasmic Large DNA Viruses (NCLDV), a monophyletic clade of large and complex viruses that contain up to 2500 genes (Koonin 2010). Members of the NCLDV family infect a wide range of eukaryotic cells and the adaptations that allow infection also demonstrate virus evolution, just like how living hosts have evolved to defeat them.

Another family found within NCLDVs are Poxviruses, some of which famously use humans as their host. They have a double-stranded DNA genome containing between 130 and 360 genes (Hughes, 2010). The family is divided into two subfamilies; one that infects insects (entomopoxvirinae), and another that infects a variety of vertebrates (chordopoxvirinae, Figure 3). Within the chordopox subfamily is the orthopoxvirus genus, which includes numerous species important to humans (Figure 1). Variola virus causes smallpox, one of the most devastating diseases in human history; and vaccinia virus, the weaker vaccine virus that eradicated smallpox from the natural world (Henderson 2009; Smithsom 2014). Closely related is monkeypox, an emerging zoonotic virus in Africa. In 2003, it made its way to the U.S. and infected at least 37 people (Reed 2004). While no one died, monkeypox has emerged as a potential bioterrorism threat with up to 10% mortality in some regions of Africa (McCollum 2014). Monkeypox, and related cowpox, both infect a wide range of hosts but cause a milder form of the disease than variola (Haller 2014; McCollum 2014). Cases of CPXV continue to appear in humans and animals in Europe and Asia (Vorou 2008), and even the smallpox vaccine virus has emerged as a pathogen in South American cattle (Franco-Luiz 2014). Molluscum contagiosum virus is a human-specific non-orthopoxvirus that, although it shows no mortality today (Mguyen 2014), could potentially evolve and adapt to better parasitize its human host.

Figure 3.  One possible poxvirus phylogenetic tree ( Hendrickson et al. 2010 ). (Source: National Library of Medicine, History of Medicine Division, Image Database.  Creative Commons Attribution 3.0 Unported (CC BY 3.0) Open Source .

Figure 3. One possible poxvirus phylogenetic tree (Hendrickson et al. 2010). (Source: National Library of Medicine, History of Medicine Division, Image Database. Creative Commons Attribution 3.0 Unported (CC BY 3.0) Open Source.

Without traditional fossil records, viral evolution is investigated primarily through genomics. Genomic data indicate that the parasitic lifecycle of viruses also makes their evolution highly dependent upon host evolution and availability (Hughes 2010). Indeed, the very thing that made variola virus so capable of devastating human populations, its strict human host specificity (6), also made smallpox the first disease ever eradicated. Variola virus only infected humans and lacked any animal reservoirs, which meant that the implementation of a vaccine in humans was able to break the chain of transmission from person-to-person and thus ending all new infections (Henderson 2009; Smithson 2014).

The culmination of competitive and host-derived pressures on viruses have shaped how viruses evolve, yet describing the exact nature of that evolution can be challenging. Understanding how the deadly smallpox virus (VARV) evolved to be the human-specific killer that plagued civilization for centuries is an example of that challenge. Yet, understanding the nature of how VARV evolved from ancestral poxviruses could help researchers to understand how other deadly pathogens could arise in the future. Here we will explore just a few hypotheses.

First, other poxviruses such as ectromelia virus (ECTV), an orthopoxvirus which infects mice, and myxoma virus (MYXV), a leporipoxvirus which infects rabbits, can have high mortality (death) rates in non-natural host species. Some have suggested that a host switch under specific circumstances of an ancestral, highly lethal poxvirus could explain the evolutionary origin of smallpox (Smithson 2014).

A similar, second hypothesis is derived from the belief that cowpox virus (CPXV) is the ancestral orthopoxvirus, containing all the genes found in any other orthopoxvirus. It is possible that the gradual restriction of host range on CPXV could lead to gene loss and evolution that created highly-specific and highly-lethal pathogens such as VARV (Smithson 2014; Haller 2014).

In contrast to these first two hypotheses, the phylogenetic analysis of poxviral genomes indicate that VARV is significantly more similar to camelpox (CMLV) and taterapox virus (TATV), than to other members of the orthopoxvirus family. However the presence of genetic recombination, or the swapping of gene material, likely played a significant role in the subsequent evolution of VARV which complicates the ability to examine phylogenetic relationships based on genomes sequence (Hughes 2010; Smithson 2014).

Researchers may never know who came first, the virus or the cell, but a better understanding of how viruses infect hosts and the influences that drive viral evolution will better prepare us to predict and understand the epidemics that emerge in the future. The parasitic nature of virus-host interactions makes it particularly challenging to study evolution. Viruses must adapt to defeat the host, but also to the adaptations that the host is making to defeat the virus. Strategies such as host gene capture attest to these adaptations, yet complicate our interpretations of viral relationships or the timelines of speciation. Regardless of the evolutionary path that has led to the diversity of poxviruses today, one fact is abundantly clear; poxviruses are not done evolving!

Useful Resources

Interested in learning more about viruses and their impact on humans? I highly recommend the following books:

D. A. Henderson, M.D. 2014. Smallpox: the death of a disease.

C. J. Peters, M.D. 1997. Virus Hunter: thirty years of battling hot viruses around the world.

 About the Author

Sara Reynolds received her Ph.D. in 2016 from the University of Maryland and in cooperation with the National Institutes of Health (NIH). Her dissertation research focused on a single large gene in cowpox, monkeypox and ectromelia poxviruses. Interestingly, this gene reduced the severity of disease during infections. Dr. Reynolds is currently an Assistant Professor at Fairleigh Dickinson University in Madison, New Jersey. Her present work focuses on using PCR technology to detect the presence of viral and bacterial species in the waters of Northern New Jersey. Correspondence can be directed to


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The Eimeriid Coccidia

Alex Knight

Ph.D. Student, School of Biological Sciences, The University of Auckland, New Zealand

If you were asked to think of a prolific pathogenic parasite of humans and wildlife, malaria or maybe a helminth worm might come to mind, but probably few people would consider coccidia. In fact, in my experience many people have never even heard of this group of organisms, which is somewhat surprising as coccidia are, perhaps, the greatest hitchhikers the natural world has ever known. Coccidia are recognized as a class of protozoa that are known to parasitize a vast range of animals including birds, amphibians, reptiles, arthropods and mammals, including humans. This broad range of potential hosts has enabled this taxon to become globally distributed. The radiation of these freeloaders was probably an ancient affair, and their long co-evolutionary histories with their varied hosts have resulted in a complex and extremely speciose phylogenetic (evolutionary) tree. This phylogenetic complexity has led to taxonomical controversy that has been deftly reviewed by Tenter et al. (2002).        

Figure 1. Coccidia extracted from hihi, a threatened passerine bird endemic to New Zealand. The coccidia on the left and right have undergone the process of sporulation and formed two sporocysts each containing the sporozoites, which are required to infect the next host. The coccidium in the centre has yet to sporulate.

Figure 1. Coccidia extracted from hihi, a threatened passerine bird endemic to New Zealand. The coccidia on the left and right have undergone the process of sporulation and formed two sporocysts each containing the sporozoites, which are required to infect the next host. The coccidium in the centre has yet to sporulate.

The Eimeriidae are a family within the coccidia, whose membership - although often revised - includes some of the taxon’s most notorious members. Different taxonomic classifications based on morphological characters of this family can include upward of 10 genera and thousands of species. I continue the focus of this article on two genera, Eimeria and Isospora. The two genera were classically distinguished using morphological features such as the number of sporocysts and sporozoites, structures that develop during the exogenous stage of the parasite (Fig. 1). Isospora spp. have two sporocysts containing four sporozoites, the invasive stage of the parasite, while Eimeria spp. have four sporocysts with two sporozoites. 

Coccidia of the genera Isospora and Eimeria live by invading the cells of their host and reproducing asexually and then sexually within the cell before being excreted back into the environment. It appears that some coccidia are restricted to invading epithelial cells of the gastro-intestinal tract of their host. Others, however, are capable of invading immune system cells and finding their way to the viscera (Box 1977). This latter type of infection may be much more pathogenic, but what causes this difference in infection location is not well understood. The development of infection at different sites in the host could be related to different strains of coccidia that are capable of infecting differing cell types or be influenced by the physical state of the host. One of the major symptoms of coccidiosis (the collective term for the pathologies these parasites cause) is an inability to absorb nutrients. This affects the body condition of the host by reducing their mass and inhibiting proper development or maintenance of the musculature. Severe infections with coccidia can be lethal, and pathogenicity varies among coccidial species.

Eimeria are often associated with domestic fowl and can cause substantial economic losses to the poultry industry. Indeed, if anyone is familiar with coccidia and coccidiosis, it is probably poultry farmers. The cost that these parasites impose on the poultry industry is estimated to be in the hundreds of millions of dollars annually (Allen & Fetterer 2002). Isospora are closely related to Eimeria; species of this genus are frequently found in wild birds and have been linked to reduced fitness and high mortality rates leading to concerns about their impact on species of conservation concern (Knight et al. 2018). For example, a captive population of the Critically Endangered blue-crowned laughingthrush had failed to rear chicks past 1 year-of-age for 17 consecutive years (Mohr et al. 2017). Investigations identified coccidia to be present in both adults and chicks and was potentially the cause of such high mortality among the chicks of the population. 

Notwithstanding the dangers associated with this taxon and their more cherished hosts, these organisms are a remarkable example of adaptation: they are passed from host to host via a faecal-oral pathway, i.e. one organism has to excrete it before another consumes it. While in the high density environments of chicken farms, frequent host to host transmission is easily imaginable, it is an incredible feat of transference that an organism about 20 to 30 microns in diameter with no locomotive ability in its exogenous stage can find its way among hosts in the great outdoors. Coccidia that infect passerine birds have evolved a clever adaptation to achieve such a feat. They appear to control the timing of their excretion from their hosts, so that the majority of coccidia are expelled in the late afternoon or evening (reviewed in Knight et al. 2018). This has a two-fold advantage, it increases the concentration of coccidia in the environment prior to peaks in avian feeding activity, the afternoon and the following morning, while avoiding the harmful UV rays of the midday sun. 

Knowledge regarding this taxon has remained somewhat esoteric to date. If the global biodiversity crisis continues, understanding the epidemiology of this parasite will become increasingly pertinent for managing threatened hosts. Parasites, despite having the potential to regulate host populations, are also a component of biodiversity that will be lost along with their hosts in the event of their extinction (Spencer & Zuk 2016). For more information please see our recent review of coccidia in passerines: “The Evolutionary Biology, Ecology and Epidemiology of Coccidia of Passerine Birds” published this year in Advances in Parasitology (Knight et al. 2018). 

About the Author

Alex Knight is currently working on the epidemiology of coccidian infection in the hihi (Notiomystis cintca), a threatened passerine bird endemic to New Zealand. Alex is a Ph.D. candidate in conservation biology at the University of Auckland whose professional interests include wildlife epidemiology, population genetics and population biology. He is supervised by Dr Anna Santure at the University of Auckland, and co-supervised by Dr John Ewen and Patricia Brekke at the Zoological Society of London.


Allen, P.C. & Fetterer, R.H. 2002. Recent advances in biology and immunobiology of Eimeria species and in diagnosis and control of infection with these coccidian parasites of poultry. Clinical Microbiology Reviews 15: 58–65.

Box, E.D. 1977. Life cycles of two Isospora species in the canary, Serinus canarius Linnaeus. Journal of Protozoology 24: 57–67.

Knight, A., J.G. Ewen, P. Brekke & A.W. Santure. 2018. The Evolutionary Biology, Ecology and Epidemiology of Coccidia of Passerine Birds. Advances in Parasitology. 99: 35–60.

Mohr, F., M. Betson & B. Quintard. 2017. Investigation of the presence of Atoxoplasma spp. in blue-crowned laughingthrush (Dryonastes courtoisi) adults and neonates. Journal of Zoo and Wildlife Medicine 48: 1–6.

Spencer, H.G. & M. Zuk. 2016. For Host's Sake: The Pluses of Parasite Preservation. Trends in Ecology and Evolution 31: 341–343.

Tenter, A.M., J.R. Barta, I. Beveridge, D.W. Duszynski, H. Mehlhorn, D.A. Morrison, R.C.A. Thompson & P.A. Conrad. 2002. The conceptual basis for a new classification of the coccidia. International Journal for Parasitology 32: 595–616.


Pūkeko (Porphyrio melanotus)

Aileen Sweeny

Ph.D. Student, School of Biological Sciences, The University of Auckland, New Zealand

Fig. 1.  Banded adult Pūkeko at the study site in Auckland, New Zealand. Photo credit Aileen Sweeney.

Fig. 1. Banded adult Pūkeko at the study site in Auckland, New Zealand. Photo credit Aileen Sweeney.

The Australasian swamp hen is known by its native Maori name “Pūkeko” in New Zealand. This brightly coloured rail (Fig. 1) is found throughout Oceania, having first arrived in Australia around 600,000 years ago (Garcia-R & Trewick, 2015). New Zealand, however, is one of its more recent conquests, with estimates of self-colonisation (from Australia) between 500-1000 years ago (Trewick & Worthy, 2001; Worthy & Holdaway, 1996). Pūkeko are found throughout New Zealand in a range of habitats, however they favour open grass areas near vegetated water systems (Dey & Jamieson, 2013). The endemic and endangered Takahē (Porphyrio hochstetteri) is closely related to the Pūkeko and, interestingly, this flightless bird is believed to have evolved from a previous colonisation of New Zealand by a Porphyrio species ~2.5 million years ago (Garcia-R & Trewick, 2015).

Pūkeko are one of the few native New Zealand species which haven’t experienced significant population declines since the introduction of two major contributors to ecosystem change: agriculture and invasive species. Indeed not only have Pūkeko been unaffected by these changes, they have proliferated in response to them, to the point that they are often regarded as a pest (Dey & Jamieson, 2013). Unlike endemic New Zealand species, Pūkeko evolved in the presence of a variety of terrestrial marsupial predators in Australia and therefore exhibit appropriate anti-predator behaviours (Jamieson, 1994). For this reason, they generally thrive in habitats where introduced predators such as brushtail possum (Trichosurus vulpecula), stoat (Mustela ermine) and rats (Rattus spp.) are present (Bunin & Jamieson, 1995).  

Fig. 2.  Adult Pūkeko displaying its frontal shield. Photo taken during the banding process. Photo credit Aileen Sweeney.

Fig. 2. Adult Pūkeko displaying its frontal shield. Photo taken during the banding process. Photo credit Aileen Sweeney.

Pūkeko have complex social structures. Generally, they live in polygynandrous groups (both, males and females have multiple mating partners), which defend a shared territory (Craig, 1980). They do, however, show considerable inter-population variability in their mating systems and may also be monogamous, polyandrous or polygynous (Jamieson, 1997, 1999). Both sexes can be highly philopatric, which is unusual in avian species (Craig & Jamieson, 1988). Within these social groups, adults form mixed-sex dominance hierarchies which influence access to reproductive opportunities (Craig, 1980; Jamieson & Craig, 1987) as well as non-sexual resources (Craig, 1977). Males and females are morphologically very similar (low sexual dimorphism) and both sexes have a bright red fleshy frontal shield (figure 1 & figure 2). This ornament acts as a “badge of status” and its color is correlated with dominance (Dey, Dale, & Quinn, 2014; Dey, Quinn, King, Hiscox, & Dale, 2017). It is likely that the color is carotenoid-based, given that carotenoids are common components in red, orange and yellow ornaments in other animals (Goodwin, 1984). Similarly, as it is found for the frontal shields of the closely related moorhen (Gallinula chloropus), it is likely to be testosterone-mediated (Dey et al., 2017; Eens, Van Duyse, Berghman, & Pinxten, 2000).

One of the most intriguing facts about Pūkeko is that they are cooperative breeders, meaning that the entire group contributes to raising chicks regardless of parentage (Craig, 1980). Cooperative breeding is rare in avian species; only 8.9% of non-marine bird species are known to engage in this breeding system (Jetz & Rubenstein, 2011). When there is more than one reproductive male and reproductive female in a Pūkeko group, chicks are always of mixed parentage (Lambert, Millar, Jack, Anderson, & Craig, 1994). Furthermore, if there are multiple breeding females present in a group they all lay in a single nest, a phenomenon known as joint-laying (Craig, 1980; Vehrencamp & Quinn, 2004). Females typically lay 4-6 eggs each, meaning that clutch sizes of up to 18 eggs in a single nest can be observed when multiple females lay together (Dey & Jamieson, 2013) (figure 3). Interestingly, conspecifics in Australia have not been reported to exhibit joint-laying, which suggests that this behaviour has evolved relatively recently in New Zealand populations (Dey & O’Connor, 2010).

Fig. 3.  Pūkeko nest containing 17 eggs. Note the varying patterns on the eggs. Photo credit Aileen Sweeney.

Fig. 3. Pūkeko nest containing 17 eggs. Note the varying patterns on the eggs. Photo credit Aileen Sweeney.

Joint-laying is a puzzling behaviour as it can lead to intragroup conflict due to the fact that hatching success decreases with increasing total clutch size (Dey, O’Connor, & Quinn, 2014; Quinn, Haselmayer, Dey, & Jamieson, 2012). However, no such conflict appears present in Pūkeko despite the fact that dominant females experience a reproductive cost when a subordinate breeding female is present (Dey, O’Connor, Balshine, & Quinn, 2014; Quinn et al., 2012). There are two possible reasons for this: first, there is an apparent lack of egg recognition in individuals, despite the fact that eggs have distinctive colouring/patterning that is unique to each female (making it relatively easy for researchers to identify separate clutches within a single nest) (Craig, 1974; Quinn et al., 2012). If females are unable to distinguish between eggs then the risk of accidentally ejecting or destroying their own egg is high. Secondly, males have been shown to decrease their incubation investment, occasionally to the point of complete nest abandonment, in response to female-female competition, and so it can be assumed that the threat of this may influence females to tolerate co-breeders (Dey, O’Connor, Balshine, et al., 2014). It is interesting to note that territory size, and therefore quality, is correlated with the number of males present in a group (Craig & Jamieson, 1990). A breeding female benefits from having access to a higher quality territory, which therefore means having more than one breeding males present. However, breeding males benefit from having more than one breeding female present as it increases their potential reproductive output. As such, this may also explain the presence of joint-laying in Pūkeko (Vehrencamp & Quinn, 2004).

Having previously worked with primates, I enjoy referring to Pūkeko as feathered baboons; they are noisy, aggressive, territorial, and have social ranks within their groups. Due to their complex and variable social structure, Pūkeko are excellent models for examining cognition and social learning in the wild. Cognition can be defined as “the acquisition, processing, storage and use of information”, and cognitive abilities allow animals to adjust their behaviour in response to new situations (Shettleworth, 2010). Traditionally, it has been studied almost exclusively in the lab, however studying cognition in the field has become more common in recent years, providing numerous benefits including reduced stress, ecologically relevant settings and a wider array of species (Pritchard, Hurly, Tello-Ramos, & Healy, 2016). There have been fascinating results emerging from recent cognition studies on wild bird populations (for examples please refer to: Cauchard, Angers, et al., 2017; Cauchard, Doucet, Boogert, Angers, & Doligez, 2017; Preiszner et al., 2017), and therefore there is huge potential for further research in this field. My research interests encompass establishing any potential relationships between cognitive ability, dominance, testosterone and ornamentation. The project is still in its early stages (designing and trialling experiments, as well as banding/PIT tagging individual birds), but I have no doubt that the coming years will uncover new insights into these rails, and indeed - hopefully - into animal cognition in general.

About the Author

Aileen Sweeney is a Ph.D. student in the School of Biological Sciences at The University of Auckland in New Zealand. She is co-supervised by Dr Kristal Cain and Dr Gregory Holwell. Hailing from Ireland, she spent time researching Vervet monkey (Chlorocebus pygerythrus) and Kinda baboon (Papio kindae) behaviour in South Africa and Zambia before emigrating to New Zealand. Her Ph.D. is focused on Pukeko behaviour, specifically their cognition and social learning in relation to dominance, testosterone and ornamentation.


Bunin, J.S. & I.G. Jamieson. 1995. New Approaches Toward a Better Understanding of the Decline of Takahe (Porphyrio mantelli) in New Zealand. Conservation Biology 9: 100–106.

Cauchard, L., B. Angers, N.J. Boogert, M. Lenarth, P. Bize & B. Doligez. 2017. An Experimental Test of a Causal Link between Problem-Solving Performance and Reproductive Success in Wild Great Tits. Frontiers in Ecology and Evolution 5.

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Photo credit Adam Brody.

Photo credit Adam Brody.

The House Cricket - Acheta domesticus

Adam Brody

In German, this little insect is known as Heimchen, in Dutch Huiskrekel, in French Le Grillon Domestique, and in English The House Cricket. To Carl Linnaeus and his scientific kin, it is known as Acheta domesticus, and to the MSNH it is known as the Taxon of the Month.

As I write, I am listening to roughly 500 house crickets chirping in my apartment. 

It is a chilly late-April evening, 51°F outside. It is a bit warmer in my apartment, but the crickets are still not chirping very fast. They would prefer if it was in the high 70s, or even 80s. I am able to single out one cricket’s chirps amidst the chorus and count how many chirps he makes in fourteen seconds: twenty. I add the number forty to this, which tells me that it is roughly 60°F in my apartment. This equation for figuring out the temperature based on crickets was published in The Farmer’s Almanac, starting in the late 1800s. 

I started raising house crickets in my Brooklyn apartment in December 2016. Since then, I’ve raised six generations of crickets, with an average colony size of 1000 crickets. I raise crickets to eat, to educate and inspire people, and to hold cricket-listening concerts. I also write essays and poetry about them. One of my goals is to open up a storefront living-museum where people can step off of the city street and into an oasis of cricket songs. I hope this happens soon, because my apartment is getting crowded. My appreciation for these amazing, often unnoticed creatures, has inspired me to learn more about their origins, diversity, behavior, ecology, thereby creating this Taxon of the Month post.

Photo credit Adam Brody.

Photo credit Adam Brody.

Origins and Distribution The origin of the house cricket is debated among scientists. They may have originated in Southeast Asia, Southwest Asia, and North Africa. It is thought that they arrived to Europe in the Middle Ages, through the spice trade. Once in Europe, they became a common housemate to humans, taking advantage of the year-round warmth of homes, hearths, and garbage dumps. House crickets arrived to North America in the 18th century and have since spread across the continent, establishing populations in southern Canada, eastern United States (except the Florida peninsula), southern California, and northern Mexico. Much of the recent spread of house crickets is thought to be a result of feral populations that have escaped from cricket farms where they are bred for reptile-feed, a high volume industry, which ships millions of crickets to pet stores and zoos each week.

Appearance House crickets are yellowish-brown, with a distinctive dark band between their eyes, and an additional dark spot between their long antennae (emerging beneath their eyes), which I think looks like a dog nose. Their bodies are countershaded, which means that their backs are darker than their undersides. A series of symmetrical, geometric patterns adorn the top of their abdomens. Along the midline, these patterns resemble a pair of abutting keystones, which decrease in size until reaching the cerci. They have two compound eyes and grinding mouthparts. Before reaching maturity, they molt between eight and ten times. Each phase of molting is called an instar. On emerging from a molt, their skin is often whitish and translucent. Once they reach maturity, both males and females develop four wings, two tough forewings that protect two softer hindwings on the thorax. They do not use these wings to fly. Rather, the males use their wings to chirp. Like all insects they have three pairs of legs. The hindmost legs are the most noticeable, since they are adapted for jumping. Crickets rarely jump as a means of decisive locomotion, but rather do so as a response to fear or agitation. Crickets also have two sensory appendages called cerci at the tip of the abdomen. The cerci looks like a flying V. Mature females have an ovipositor, a needle-like egg-laying organ, between the cerci, and are very easy to identify for this reason.

House cricket molting. Photo credit Adam Brody.

House cricket molting. Photo credit Adam Brody.

Songs Crickets use sounds and vibrations to communicate with each other. They have tympanic membrane on their front legs. This "ear" is like a human eardrum, and is very sensitive to sound vibrations. Only male crickets can stridulate, or chirp. They have a file and scraper body part on their wings, with which they produce their songs. They do not use their legs to chirp. Caelifera, the other suborder of Orthoptera which includes grasshoppers, do use their legs for chirping. Crickets have different songs for different purposes. The chirping that is most common is that of a male trying to attract females. It is rhythmic and quite loud. Temperature affects the rate at which they chirp. Often, male crickets will lock into rhythms with one another as they collectively attract females. Once a female approaches a male cricket, he woos her with a courting song that is very quiet and sweet sounding, almost like a purring cat. Once the male and female crickets have finished mating, he sings another song to keep her nearby and guard her from being mated with another male. When one male invades another male's territory and they encounter one another, they produce an aggressive song, which is a loud trill. This song is often made before or after an instance of combat.

Photo credit Adam Brody.

Photo credit Adam Brody.

Mating It is worth noting that a female cricket mounts the male cricket and remains on top throughout copulation. Mating is concluded when the male emits a spermatophore, a whitish gelatinous capsule that contains sperm, and deposits the capsule under the female’s genital opening. Once the pair have separated, the female rips the spermatophore open, by herself, at which point the sperm travel toward her reproductive organs. Within a few days, she will begin laying her fertilized eggs in any available, damp substrate. Estimates vary widely on how many eggs a female cricket will lay in her lifetime, with numbers ranging between 100 and 700. 

Cricket paralysis virus Throughout the eighties and into the early aughts, a "cricket paralysis virus" was reported to be involved in catastrophic collapses in American and European cricket rearing facilities. This virus killed an estimated 90 percent of all A. domesticus crickets in breeding facilities. There is some debate as to whether the disease in question was, in fact, a DNA containing virus called Acheta domesticus densovirus (AdDNV), as opposed to the RNA virus referred to a CrPV. Regardless, over 60 million crickets died as a result.

Crickets and Humans I'm not the only Brooklynite who loves crickets. The connection between human and crickets extends across the globe. For example, crickets were once ubiquitous residents of the Paris subway system, but in recent years their presence diminished. “The Protection League for the Crickets of Paris Metro” was established in 1992 and blamed transit strikes for decimating the population, saying that service freezes led to actual freezing temperatures, which the crickets could not endure. They also blamed increased sanitation for depriving the crickets of food. Additionally, they claimed that a smoking ban in the metro deprived crickets of a prime food source, cigarette butts, though this claim has been contested. "Ideally, we'd like the two Metro lines where there are the most crickets to be declared a Natural Park for them," said the president of the Protection League.

Crickets and Entomophagy I began farming crickets with the intention of spreading the gospel of entomophagy. Crickets are a viable food source for humans, and require significantly less food and water than vertebrate livestock. Crickets even require less water than most kinds of plant-based protein. Growing one pound of almonds require 1,929 gallons of water. Growing one pound of lentils require 704 gallons. One pound of cricket requires  <drumroll please>  1/2 gallon of water. And don’t forget greenhouse gases. Crickets aren’t running tiny furnaces inside of their bodies to stay warm, which means they don’t fart as much as warm-blooded livestock. And let's not forget about land use. Crickets can fit into small spaces; cows, not so much.

Cricket farm. Photo credit Adam Brody.

Cricket farm. Photo credit Adam Brody.

What do I suggest then? Become a cricket farmer. Small is beautiful. Do it yourself! Supplement your diet with crickets, while sharing your home with a fascinating creature that will fill your heart with songs and your mind with cool thoughts about animal behavior, evolution, cultural taboos and insect sex.

All you need is : 

A container (I bend sheet cardboard into a tube, see picture)

Some housing material (I use egg flats that I pick up from outside of bakeries)

Food (I feed my crickets oat bran, flax meal, and veggie scraps, though I plan on shifting to an entirely waste-stream diet soon)

Water (crickets drown easily, so I often use a shallow dish with rice in it when they’re small)

& some kind of heating element if your house runs cold.

I’ll be authoring something more thorough about home cricket farming in the future. Until then, stay in touch with me via my poorly-updated website : or my more frequently updated instagram : or email me at :

About the Author

Adam Brody has been farming crickets in his Brooklyn apartment since 2016. His cricket project explores inter-species relationships, crickets as an accessible and environmentally low-impact food source, and the therapeutic benefit of cricket songs. He performs music with his crickets along with fellow cricket farmer Jude Tallichet. Brody has presented his crickets at The Free Library of Philadelphia, Spring Sessions Residency, The Jordan National Gallery of Fine Art, Dixon Place and Open Source Gallery. You can more about his work at:


Behavior of the House Cricket, Acheta domesticus. 2012. Cricket Lecture, Annimal Behavior Course (BIO 342). Reed College.

Boehrer, K. Oct. 13, 2014. This is How Much Water It Takes to Make Your Favorite Foods. HuffPost. 

Cousteaux, G. & J. Cousteaux. 2003. Les grillons. Insectes 129: 27–30.

The Crickets of the Paris Metro. Ligue de Protection des Grillons du Métro Parisien.

Marshall, J.A. 1983. The orthopteroid insects described by Linnaeus, with notes on the Linnaean collection. Zoological journal of the Linnean Society 78: 375–396.

McGavin, G.C. 2001. Essential Entomology: An Order-by-Order Introduction. Oxford: Oxford University Press. pp. 20.

Otte, D. 1992. Evolution of Cricket Songs. Journal of Orthoptera Research 1: 25–49.

Pener, M.P. 2016. Allergy to Crickets: A Review. Journal of Orthoptera Research 25: 91–95.

Peters, A. Aug. 21. 2017. This Giant Automated Cricket Farm Is Designed To Make Bugs A Mainstream Source Of Protein. Fast Company.

Resh, V.H. & R.T. Cardé. 2009. Encyclopedia of Insects. Academic Press. p. 232. ISBN 978-0-08-092090-0

Walker, T.J. 1999. House Cricket, Acheta domesticus (Linnaeus) (Insecta: Orthoptera: Gryllidae). University of Florida: IFAS Extension p. 1–2. 

Walker, T.J. Singing Insects of North America: House Cricket, Acheta domesticus Linnaeus 1758. University of Florida: IFAS, Entomology Department.

Ascetosporea: Ornate spore-bearers of doom

Yana Eglit

Ph.D. Candidate, Department of Biology, Dalhousie University, Halifax, Nova Scotia, Canada

Figure 1 . Tree of Eukaryotes (modified from  Simpson &amp;&nbsp;Eglit 2016 ) showing Plants, Animals, and Fungi in blue and Ascetosporea in red. Everything not in blue text is typically considered a ‘protist’.

Figure 1. Tree of Eukaryotes (modified from Simpson & Eglit 2016) showing Plants, Animals, and Fungi in blue and Ascetosporea in red. Everything not in blue text is typically considered a ‘protist’.

If you studied any biology, you have probably heard of the three domains of life: the eukaryotes, i.e. a group of organisms, or taxon, with a true nucleus (plants, animals, fungi, and “protists”), as well as those without a nucleus, the bacteria and the lesser known archaea (latest data suggest eukaryotes are essentially a very derived ‘type of archaea’). It might surprise many, that plants, animals and fungi are but twigs on the eukaryote tree of life (Fig. 1; in blue). In fact, the eukaryotes are nothing but a small twig in the largely bacteria-dominated diversity of life (Simpson & Eglit 2016). Most posts in this Taxon of the Month series, most diversity of life you’re intuitively familiar with, most nature you see around you by the naked eye, exists entirely on these three ‘tiny’ twigs. All other eukaryotes are “protists”, and the vast majority of these “protists” are small, and thus invisible to the naked eye. However, in the ocean, protists can get big, in the form of seaweeds. Kelps, red algae, and green algae are all considered “protists” (see Phaeophyta, Rhodophytes, and Chlorophyceae in Fig. 1, respectively), and some of these can also be quite tasty!

Now, the point isn’t that plant and animal diversity is somehow dull, but rather that if this is just a very small part of the overall diversity within eukaryotes. Just try to imagine the breathtaking diversity of the rest of it! And with new major branches of this tree found or established each year (Brown et al. 2013; Seenivasan et al. 2013Janouškovec et al. 2017), one can only dream of what alien-like lifeforms remain unknown to humankind! Developments in sequencing technology have accelerated this rate of discovery to unprecedented levels – and at least a handful more entirely new types of eukaryotes will be published in the next few years.

Today, we will look at one such neglected and poorly-known group that is so esoteric, its formal scientific name means “elaborately-crafted spore bearers” – Ascetosporea (from Greek asketos: ‘well-made, elaborate, ornate’). Their ornateness probably brings little comfort to the hundreds if not thousands of livelihoods that can be wiped out by these parasites – they are highly efficient parasites of economically-important shellfish, sometimes causing absolute mortality and total collapse of the fishery.

Ascetosporea (Fig. 1; in red) find their phylogenetic home within a little-known supergroupnamed Rhizaria. Previously a superficially eccentric assemblage of vastly different-looking organisms only supported by molecular data, this group is also becoming increasingly riddled with parasites. Ascetosporeans comprise two groups: Haplosporidia, known for spores like lidded jars, and Paramyxids, with an incomprehensible inward division resulting in a cellular matryoshka.

Figure 2 . Haplosporidian spores:  A ) transmission electron microscopy (TEM) image showing a cross section of a  Haplosporidium  sp. spore ( Feist et al. 2009 );  B ) diagram of a mature  Haplosporidium montforti  spore ( Azevedo et al. 2006 );  C ) scanning electron microscopy image of a  Minchinia mercenariae  spore, arrow pointing to the opened “lid” and the opening below ( Ford et al. 2009 ).

Figure 2. Haplosporidian spores: A) transmission electron microscopy (TEM) image showing a cross section of a Haplosporidium sp. spore (Feist et al. 2009); B) diagram of a mature Haplosporidium montforti spore (Azevedo et al. 2006); C) scanning electron microscopy image of a Minchinia mercenariae spore, arrow pointing to the opened “lid” and the opening below (Ford et al. 2009).

Haplosporidia are parasites to a variety of marine invertebrates like molluscs (including the oysters), crustaceans, echinoderms, and even other parasites (Azevedo & Hine 2017). The infection starts out with solitary uninucleate cells invading and gradually forming a multinucleate ooze (plasmodium) growing among the host tissues. The plasmodium splits up into smaller pieces (sporonts), then forming walls of membrane centred on each nucleus (up to hundreds!), and (in most species) proceeds to build spores (Azevedo & Hine 2017). Haplosporidian spores feature jars or sometimes handle-less amphorae with hinged ‘lids’ and various surface decorations like long tails or tassels and surface reliefs (Fig. 2, Ford et al. 2009). Inside each vessel is a nucleus, haplosporosomes (some harpoon-like organelles of uncertain function) and a peculiar “spherulosome”, a complex clump of membranes. Once inside an appropriate next victim, the lid then opens (Fig. 2c, arrow), releasing newborn parasites to graze happily on their new host. Haplosporidians are mysterious and fascinating in their own right, just remember that somewhere out there are cellular oozes currently making little microscopic pottery in which to spit out its progeny.

Paramyxids are parasites of various marine invertebrates ranging from molluscs and crustaceans, including many commercial ‘shellfish’, to polychaete worms and even tunicates. Once inside their host, paramyxids proliferate throughout tissues, devouring their host from within, and then sporulate primarily in digestive and reproductive organs (Fig. 3a, Lester & Hine 2017). This sporulation is distinctive for paramyxids, namely their remarkable formation of spores inside cells. The process is quite complicated and not at all how we learned mitosis in school, as the daughter cell is formed inside the mother, like inverse budding (Fig. 3b,c). As far as I know, the only other cells that like that are pollen forming cells in plants. Species differ in how many inverse buddings they undergo (Feist et al. 2009), and to keep things simple I’ll just go over the most complex case: Paramyxa paradoxa (Audemard et al. 2002; Stentiford et al. 2017; Fig. 4).

Figure 3 . Paramyxid appearance:  A ) an infected oyster ( Desportes 1984 );  B ) general view (TEM) of  Paramyxa nephtys  cells in various stages of spore formation  within  polychaete gut lining cells – this species develops inside host cells rather than between them! Pink outlines roughly correspond to host cells, yellow/orange and blue concentric outlines show layers of paramyxid cells (modified from  Audemard et al. 2002 );  C ) detail of  P. nephtys  tertiary cell (C3) with C4-C5 internal cells nested inside ( Audemard et al. 2002 ). For images of mature spores see  Audemard et al. 2002 &nbsp;(free access).  D ) Primary cell (C1) of  Paramarteilia canceri  with sporonts and spores (colours corresponding to those in Fig. 4; modified from  Larsson &amp; Køie 2005 ). Note the dark blob and rod-like structures in C3: those are thought to be related to the dark-staining structures in haplosporidia called haplosporosomes labelled ‘Hs’ in Fig. 2a.

Figure 3. Paramyxid appearance: A) an infected oyster (Desportes 1984); B) general view (TEM) of Paramyxa nephtys cells in various stages of spore formation within polychaete gut lining cells – this species develops inside host cells rather than between them! Pink outlines roughly correspond to host cells, yellow/orange and blue concentric outlines show layers of paramyxid cells (modified from Audemard et al. 2002); C) detail of P. nephtys tertiary cell (C3) with C4-C5 internal cells nested inside (Audemard et al. 2002). For images of mature spores see Audemard et al. 2002 (free access). D) Primary cell (C1) of Paramarteilia canceri with sporonts and spores (colours corresponding to those in Fig. 4; modified from Larsson & Køie 2005). Note the dark blob and rod-like structures in C3: those are thought to be related to the dark-staining structures in haplosporidia called haplosporosomes labelled ‘Hs’ in Fig. 2a.

An amoeboid primary cell (C1) crawls between the host cells and multiplies until it is ready to host the spore construction action. It then divides endogenously (inward) to form a secondary cell (C2), which itself divides ‘normally’ twice to form four (Fig. 4a). The precise mechanism of this idiosyncratic endogenous division remains mysterious!  Now, each of the secondary cells repeats the process above to form four internal tertiary cells (C3; Fig. 4a). From this point on, we will no longer see conventional mitosis, only inward budding: each of the tertiary cells divides endogenously four times, forming a four-cell-layered spore (C4-C6; Fig. 4a). To tally up the math: C1 + 4C2*4C3*4(C4-C6) = 4x(4x4+1)+1 = 69 cells and nuclei total! (Fig. 4b) And to top it all off, sometimes the primary cell can have a parasite of its own (Stentiford et al. 2017)!

The primary and secondary cells then decay away to release the C3s (spores) (Fig. 4c), which in some cases undergo further maturation by forming a cell wall and a striated “tail” (Fig. 4d; Larsson & Køie 2005). After such convoluted origins, where do these newborn spores go next? As the paramyxid life cycle progresses, our understanding of it drops approximately exponentially, and it was only in the oyster parasite Marteilia refringens when the next stage for only one species was determined to be a copepod – by cleverly taking advantage of a minimalist (low species-richness) lagoon ecosystem and molecularly testing each species of invertebrate for signs of the parasite (Audemard et al. 2002). For the rest of paramyxids – the few that are discovered – we have no idea. It is unclear how the spores from copepod return to the oyster, as experimental infection attempts have failed (Carrasco et al. 2008), and there does appear to be a seasonal component (Boyer et al. 2013).

Figure 4 . Spore formation ( A , B ) and maturation ( C , D ) in among the most complicated known paramyxids,  Paramyxa (= Paramyxoides )  nephtys  (based on  Audemard et al. 2002 ;  Stentiford et al. 2017 ). Can you count all 69 cells in the final sporont?

Figure 4. Spore formation (A,B) and maturation (C,D) in among the most complicated known paramyxids, Paramyxa(=Paramyxoides) nephtys (based on Audemard et al. 2002; Stentiford et al. 2017). Can you count all 69 cells in the final sporont?

There are about a couple dozen or so people in the world who work on Paramyxids, largely government scientists who monitor the health of fisheries. Perhaps another dozen work on Haplosporidia. Despite obvious economic importance, obscure protist parasites do not receive much attention or funding. Even oomycetes, a group containing a plant pathogen from which the population of Ireland has yet to recover (Potato Famine), are an acquired taste.

These off-the-beaten path protist parasites are likely relevant to a variety of lesser-studied crops outside Europe, North America and East Asia, and thus potentially important for food security. Without well-understood evolutionary contexts and basic biology, it would be difficult to manage these organisms when they turn to being our nightmares. I hope to have piqued a little bit of interest in these odd and diverse parasites, and maybe even in their just-as-fascinating free-living relatives too. Just remember that every bit of awe-some biodiversity you learn about comes with its own assortment of parasites, some perhaps equipped with their very own surrealist spores.

About the Author

Yana Eglit is a protistology Ph.D. Candidate in the lab of Alastair Simpson (Department of Biology, Dalhousie University, Canada) studying evolution and cell biology of understudied and novel deep lineages of eukaryotes (i.e., “very weird protists”).


I would like to thank Noèlia Carrasco (IRTA), IFREMER, and the paramyxid community for the 2015 workshop invite immersing me in the Paramyxean world, and Aaron L. Beek (University of Memphis) for clarifying the nuanced meanings of asketos in Ancient Greek.


Audemard, C. 2002. Needle in a haystack: involvement of the copepod Paracartia grani in the life-cycle of the oyster pathogen Marteilia refringens. Parasitol. 124: 315–323.

Azevedo, C. & P. M. Hine. 2017. Haplosporidia. In Handbook of the Protists (J. Archibald, A.G.B. Simpson, C. Slamovits, Eds.).

Azevedo, C., P. Balseiro, G. Casal, C. Gestal, R. Aranguren, N.A. Stokes, R.B. Carnegie, B. Novoa, E.M. Burreson & A. Figueras. 2006. Ultrastructural and molecular characterization of Haplosporidium montforti n. sp., parasite of the European abalone Haliotis tuberculate. J. Invert Pathol. 92: 23–32.

Boyer, S., B. Chollet, D. Bonnet & I. Arzul. 2013. New evidence for the involvement of Paracartia grani (Copepoda, Calanoida) in the life cycle of Marteilia refringens (Paramyxea). Int. J. Parasitol. 43: 1089–1099.

Brown, M.W., S.C. Sharpe, J.D. Silberman, A.A. Heiss, B.F. Lang, A.G. Simpson & A.J. Roger. 2013. Phylogenomics demonstrates that breviate flagellates are related to opisthokonts and apusomonads. Proc. Biol. Sci. 289: 20131755.

Carrasco, N., I. Arzul, I. B. Chollet, M. Robert, J.P. Joly, M.D. Furones & F.C.J. Berthe. 2008. Comparative experimental infection of the copepod Paracartia grani with Merteilia refringens and Marteilia maurini. J. Fish Dis. 31: 497–504.

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Feist, S.W., P.M. Hine, K.S. Bateman, G.D. Stentiford & M. Longshaw. 2009. Paramerteilia canceri sp. n. (Cercozoa) in the European edible crab (Cancer pagurus) with a proposal for the revision of order Paramyxids Chatton, 1911. Folia Parasitol. 56: 73–85.

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Grauer’s Broadbill (Pseudocalyptomena graueri)

Elise Morton, Ph.D.

Ph.D. Student, University of Florida, Gainesville, Florida

 I spent this past summer conducting field research in Bwindi Impenetrable National Park, one of the only remaining areas in the world where one can safely observe the Grauer’s Broadbill (Pseudocalyptomena graueri), also known as the African Green Broadbill, endemic to the montane forests of the Albertine Rift. The Albertine Rift is a mountainous region in East Africa formed by the splitting of the Somali and African plates. Owing in part to its long evolutionary history, diversity of habitats, and steep elevational gradient, it is the most biodiverse region in Africa. The region supports more vertebrate species than anywhere else on the continent (Plumptre et al. 2007) including nearly 50% of Africa’s bird species, many of which are endemic and globally threatened (Carr et al. 2013). Because to this diversity, the region has been identified as a top priority area for biodiversity conservation (WWF 2016) and is thought to be one of the most vulnerable regions on the planet due to human-caused environmental changes (Carr et al. 2013).

Grauer's Broadbill. Photo Credit: Michael Todd.

Grauer's Broadbill. Photo Credit: Michael Todd.

Bwindi Impenetrable National Park (BINP) in southwest Uganda is a 331 square km area of protected montane forest supporting an incredible diversity of species and habitats including over 380 species of birds, 120 mammal species, and half of the world’s critically endangered Mountain gorilla population (Plumptre et al. 2007). In 2002, avian diversity was characterized by the Wildlife Conservation Society across the Albertine Rift, revealing higher species richness and the presence of more endemic and threatened species in BINP compared to other forests in the region (Plumptre et al. 2012). I was there to generate new rigorous estimates of avian diversity and to examine the relationship between bird communities and habitat along elevational and disturbance gradients.

One major predictor of species extinction risk, particularly in the tropics is having a limited elevation range (Harris & Pimm 2008; Lee & Jetz 2011). This is concerning given that approximately 30% of the world’s threatened bird species are restricted to narrow elevational ranges in montane regions (Jankowski et al. 2013), yet, determinants of species elevation ranges, however, are still poorly understood. One mechanism proposed for elevational specialization for tropical montane species includes physiological constraints due to narrow thermal optima (Janzen 1967; Colwell et al. 2008; McCain 2009). However, biotic factors are also important and vegetation heterogeneity and productivity, which varies along elevational gradients and with disturbance, has been shown to be positively correlated with avian species diversity (MacArthur & MacArthur 1961; Terborgh 1977; Rahbek 1995).

Downscaled climate models predict continued warming and declines in precipitation for lower elevations coupled with increased precipitation and cooler temperatures at higher elevations (Phillipps & Seimon 2010), potentially reducing the elevational ranges of many species. Alternatively, or in addition, in the absence of biotic constraints (e.g., competition or specialized habitat requirements), species should be able to maintain their climate niches by moving relatively short distances up or downslope. However, if habitat requirements, specific to an elevational range, constrain species beyond their physiological limitations, such species should be more vulnerable to changing climate. In either scenario, this shrinking of species ranges or reshuffling of avian communities would potentially lead to a disruption of ecological networks and new species interactions, accompanied by changes in abundance for many species. For this reason, understanding what factors determine species’ elevation ranges is necessary to predict how those species will respond to anthropogenic change, including climate and land disturbance.

One of Albertine Rift’s most range-restricted and vulnerable species is the Grauer’s Broadbill. This rare little bird has been a coveted species for birders and ornithologists since it was first collected in 1908 in the Democratic Republic of Congo by Rudolf Grauer, a naturalist from Australia. The first account of its discovery read, “the tropics are full of birds that are green, but this was different” (Rockefeller & Murphy 1933). Prior to its second discovery in 1929, Sterling Rockefeller and Charles Murphy wrote “It was an object of envy among naturalists, yet none of the collectors who had visited Kivu were able to find a single individual.” Still today, avid birders flock to this region in hopes of glimpsing one of these special birds and only few are lucky enough to do so.

Occurring only in the Itombwe Mountains and Kahuzi-Biéga National Park of the Democratic Republic of Congo and BINP, the Grauer’s Broadbill is listed as vulnerable by IUCN due to its limited distribution coupled with fragmentation and deforestation throughout its range. Recent surveys have been unsuccessful in locating the species in Kahuzi-Biéga National Park (A. Plumptre in litt. 2007; BirdLife International 2016) and although the species has been reported to be more common in the Itombwe Mountains, the region is unprotected. The current population trend is thought to be decreasing, however, rigorous quantitative assessments of population size have yet to be generated and extinction risk is unknown.

The Grauer’s Broadbill is small in size (~13.6–15.6 cm and 29–32.5 g), predominantly bright grass green in color with a varied pattern of pale blue on the chest and throat, ear coverts, and under the tail. Adults have a black-streaked buff crown with a narrow black eye stripe. In foliage, its coloring provides exceptional camouflage, but in direct light it’s a striking little bird. Little is known about its breeding behavior but birds are in breeding condition in July and August, and one active nest was observed in April. The species forms dome nests made of lichen and mosses, generally suspended in the outer branches at about mid-canopy height. Observational accounts from Kahuzi-Biéga National Park and the Itombwe Mountains of DRC and BINP suggest potential ecological differentiation between the populations (Friedmann 1970). In BINP, it prefers forest edges and isolated trees in open areas and has been found between 1,760 and 2,480 m in DRC and between 1,060 and 2,285 m in Uganda. In Uganda, the species is often found in the upper portions of the understory in stands of Chrysophyllum gorungosanum. In DRC, however, it is found predominantly in the upper canopy below the bamboo zone, favoring areas of dense foliage along forest edges and near cultivation.

Despite its status as a rare, endemic and vulnerable species, specific habitat requirements, behavior and demographic parameters are still largely unknown. Originally thought to be a flycatcher (Lowe 1929), it has been observed gleaning insects from foliage or catching them in the air (Rockefeller & Murphy, 1933). Unlike other eurylaimids, individuals have also been seen moving like woodpeckers along the undersides of horizontal branches (Bruce 2018). The stomach contents of five specimens revealed small beetles, snails, insects, insect larvae, seeds, flowers, buds and fruits (Friedmann 1970). It has been observed foraging singly or forming multi-species flocks of up to ten birds (del Hoyo et al. 2003), a behavior common to many tropical insectivore species (Bruce 2018). The most common explanations invoked for this behavior are protection from predators and increased foraging efficiency (e.g., flushing insects) coupled with reduced competition with conspecifics relative to heterospecifics (Terborgh 1990; Beauchamp 2004; Darrah & Smith 2014). These mixed flocks can be very specific in their composition, but this has yet to be studied for the Grauer’s Broadbill.

While doing point counts in the highland Mubwindi swamp of Bwindi, I was very fortunate to see one active Grauer’s Broadbill nest, with young chicks. Given the biological diversity that exists in this incredible forest and its ecological importance, it is shocking how little we know about the birds that inhabit it. My educational background is in Microbiology but I changed career paths because I wanted to do applied conservation research. As I was warned would be the case by every conservation biologist and wildlife ecologist I spoke with for advice, the potential and actual impact of my work feels at times, perhaps most times, discouragingly small. E. B. White wrote, “I arise in the morning torn between a desire to improve (or save) the world and a desire to enjoy (or savor) the world. This makes it hard to plan the day.” Switching careers to wildlife ecology and conservation, was one an attempt to alleviate the struggle between these desires, to merge them into one. I cannot say that I have accomplished this goal. However, waking up in a wonderful place like Bwindi, so privileged to hear and observe this special little bird that we know so little about, it is impossible not to be moved by the beauty of this world and feel satisfied with the savoring alone.

About the Author

Elise Morton is a Ph.D. student in the School of Natural Resources and Environment and Wildlife Ecology and Conservation at the University of Florida ( She is co-advised by Professors Madan K. Oli and Scott K. Robinson. Her research is focused on spatial and temporal patterns of avian diversity along elevation and disturbance gradients in tropical montane forests.


Beauchamp, G. 2004. Reduced flocking by birds on islands with relaxed predation. Proceedings of the Royal Society of London, Series B: Biological Sciences 271: 1039–1042.

Bruce, M.D. 2018. Grauer's Broadbill (Pseudocalyptomena graueri). In: del Hoyo, J., Elliott, A., Sargatal, J., Christie, D.A. & de Juana, E. (eds.). Handbook of the Birds of the World Alive. Lynx Edicions, Barcelona. (retrieved from on 22 January 2018).

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Carr, J., Outhwaite, W., Goodman, G., Oldfield, T. & Foden, W. 2013. Vital but vulnerable: Climate change vulnerability and human use of wildlife in Africa’s Albertine Rift. IUCN.

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Colwell, R.K., Brehm, G., Cardelús, C.L., Gilman, A.C. & Longino, J.T. (2008). Global warming, elevational range shifts, and lowland biotic attrition in the wet tropics. Science 322: 258–261.

Darrah, A.J. & Smith, K.G. 2014. Ecological and behavioral correlates of individual flocking propensity of a tropical songbird. Behavioral Ecology 25: 1064–1072.

del Hoyo, J., Elliott, A. and Sargatal, J. 2003. Handbook of the Birds of the World. Vol.8: Broadbills to Tapaculos. Lynx Edicions, Barcelona.

Friedmann, H. 1970. The status and habits of Grauer's Broadbill in Uganda (Aves: Eurylaimidae). Contrib. Sci. Los Angeles Co. Mus. 176: 1–4.

Jankowski, J.E., Londoño, G.A., Robinson, S.K. & Chappell, M.A. 2013. Exploring the role of physiology and biotic interactions in determining elevational ranges of tropical animals. Ecography 36: 1–12.

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Potamopyrgus antipodarum

Deanna M. Soper, Ph.D.

Assistant Professor of Biology, University of Dallas, Irving, Texas

Fig.&nbsp;1.  Male  P  otamopyrgus antipodarum.

Fig. 1. Male Potamopyrgus antipodarum.

In the Northern Hemisphere, the month of January means colder temperatures and shorter days, while in the Southern Hemisphere, where the Taxon of the Month is native, it is summer. Potamopyrgus antipodarum is a small (2–5mm), freshwater snail species native to New Zealand and can be quite abundant in lakes and streams on both the North and South Islands of New Zealand. This species is being used across the world to answer a wide variety of different biological questions including: the evolution of sexual reproduction, evolutionary genetics, ecotoxicology, invasive biology, and reproductive behavioral evolution. Its current range includes not only New Zealand, but also invasive populations across Europe and North America. Potamopyrgus antipodarum, sometimes referred to as the New Zealand Mud Snail, was first described by John Edward Gray in 1843 and later characterized by Michael Winterbourn (1970, 1972). In 1974, Michael Winterbourn documented infection of the snails by several sterilizing trematode parasites (worms), but one genus, Microphallus, is particularly common in lake populations (Lively, 1987).  This sterilizing trematode first infects snail hosts, where the parasite develops into cercariae. Infected snails are then eaten by ducks where the parasite develops into the adult worm stage. The worms undergo sexual reproduction whereby eggs are produced and then released with the duck’s feces. This gives the snails an opportunity to eat the eggs and become infected with the parasite starting the life cycle over again.

Reading about this snail, the famed evolutionary biologist John Maynard Smith (1978) proposed that this species could serve as a model system to solve a long-standing riddle: the evolution of sex. Why sexual reproduction evolves and persists in populations has been a question since Charles Darwin’s time. In 1859, Darwin published the first edition of On the Origin of Species and in it expressed doubt that long-standing asexual lineages existed when he said that “Finally then, we may conclude that in many organic beings, a cross between two individuals is an obvious necessity for each birth; in many others it occurs perhaps only at long intervals; but in none, as I suspect, can self-fertilisation go on for perpetuity.” (Darwin, 1859, page 101). Sexual females are required to produce males, which means that they have a reduced growth rate compared to asexual females because males cannot produce offspring (see figure 1). This means that all else being equal, asexual females should take over sexual females in the same population quickly. And yet, sexuality is ubiquitous among all higher order plants and animals. 

Fig. 2.  Sexual reproduction versus asexual reproduction. Color representative of genetic background.

Fig. 2. Sexual reproduction versus asexual reproduction. Color representative of genetic background.

This snail species provides an ideal opportunity to provide answers to the question of sex because many endemic populations contain ecologically non-distinct sexual and asexual lineages. Consequently, Curtis Lively (1987) launched P. antipodarum as system for use in the field of evolutionary biology when he sought to answer the question of why sexual reproduction evolves and is maintained in populations over time. In the early 1990’s it was discovered that some snails contain two copies of each chromosome (diploid) and are sexual – females produce on average 50% female, 50% male. While other snails contain three copies of each chromosome (triploid) and are asexual – females produce mostly all females (Wallace, 1992). Since then, snail populations containing up to six copies of each chromosome have been discovered (Neiman et al., 2011). 

Research on the relationship between the snail and the sterilizing trematode parasite has found a strong positive association between presence of parasites and presence of males, which is a measurement of the percent of snails in a population that are sexual (Lively, 1987; Jokela, 2009; King et al., 2011). This means that where there are parasites that coevolve with their hosts, there is sexual reproduction. Sexuality is favored in environments with coevolving parasites because sexual lineages generate genetic diversity in each generation, while asexual females produce offspring that are 100% related. The sexual portions of the population are a “moving target” and the asexual lineages are a “static target” resulting in parasites more easily evolving the ability to infect asexual lineages. Parasites keep the asexual populations “in check” allowing the sexuals to remain in the population. 

Fig. 3.  Baby snail just born.

Fig. 3. Baby snail just born.

Although some biological and ecological characteristics of the snail have been documented, other aspects remain a mystery. For example, it is not known what determines that a snail embryo will develop into a male vs a female adult snail. The sexual life of this species is unique because unlike most snails that lay eggs, P. antipodarum female snails undergo “pregnancy” (internal gestation) and give live birth (see Fig. 3 of baby snail just born). Occasionally, baby snails can be born inside their gestational sac (see video below). Males can be identified by external genitalia that they use to fertilize sexual females (Fig. 1). A male can initiate mating by crawling on top of a female at which time the female can reject the male mating attempt by vigorously shaking her shell back and forth to knock the male off. Recent research has found that females can be choosey, but what females pay attention to during mating attempts and what causes females to prefer particular males also remain unanswered questions. This little snail has provided important insights to evolutionary biology, ecology, and reproductive biology, but still holds the answer to many questions that need to be explored.

Potamopyrgus antipodarum born in gestational sac.  Video Credit: Deanna Soper

About the Author

Deanna Soper is an Assistant Professor of Biology at the University of Dallas where she uses P. antipodarum to understand how parasitic selective pressure and host/parasite coevolution influences the evolution of reproductive behaviors. Her lab website can be found here.


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Maynard Smith, J. 1978. The Evolution of Sex. Cambridge University Press. Cambridge.

Neiman, M., D. Paczesniak, D. M. Soper, A. T. Baldwin and G. Hehman. 2011. Wide Variation in Ploidy level and Genome Size in a New Zealand Freshwater Snail with Coexisting Sexual and Asexual Lineages. Evolution. 65: 3202–3216.

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Winterbourn, M. J. 1972. Morphological Variation of Potamopyrgus jenkinsi (Smith) from England and a comparison with the New Zealand species, Potamopyrgus antipodarum (Gray).  Proceedings of the Malacological Society London. 40: 133.

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Daniela Vergara

Postdoctoral Fellow, University of Colorado, Boulder, Colorado


We end the year with the plant genus Cannabis, belonging to the family Cannabaceae, which also includes hops (Humulus sp.) and Hackberries (Celtis). Cannabis is most famous today for its use as a recreational drug, aka marijuana, although the legalization of this plant as a drug has been quite controversial in the United States and across the world. Despite its notoriety, however, the origins, chemical properties, reproductive strategies and dispersal of Cannabis across the globe are quite fascinating and this plant genus has been impacting human culture for ages. Cannabis is one of the oldest domesticated plants and various ancient human cultures have used it for spiritual rituals, medicinal purposes, and fiber for rope or clothing that has been extracted from hemp plants (Li 1973, 1974; Russo, 2007, 2008). The genus most likely originated in Central, South or Eastern Asia but the exact origin of Cannabis is difficult to determine because of shifts in its distribution between glacial cycles (Clarke & Merlin 2013). Humans brought this species to Europe and later to Africa and the Americas where it was cultivated and domesticated into different varieties (Clarke & Merlin 2013). 

Carolus Linnaeus, the founder of the binomial species system, was the first person to classify the Cannabis genus in 1753, and only identified a single species, Cannabis sativa L. However, Jean Baptiste Lamarck (yes, the one who came up with a first theory of adaptation, now known and disregarded as “Lamarckian evolution”) described a second species, Cannabis indica, in 1785 (Watts, 2006). While the validity of the second species is debated, the groupings “sativa” and “indica” are still commonly used. Interestingly, Cannabis has an unusual amount of genetic diversity when compared to other plant groups (Sawler et al. 2015; Lynch et al. 2016; Vergara et al. 2016). Recent scientific research has found that there are genetic clusters, and thus it is possible that several other species will be described. However, these do not seem to reflect Lamarck’s classification of C. sativa and C. indica (Sawler et al. 2015; Lynch et al. 2016; Vergara et al. 2016). 

What molecular properties and processes make this plant so popular as a recreational drug? Cannabis produces cannabinoids, which interact with our own endocannabinoid system within the brain and nervous system (Gertsch 2008). The endocannabinoid system is involved in regulating multiple physiological processes, including sleep and hunger. One of the primary and most widely known cannabinoids produced by the Cannabis plant is Δ-9-tetrahydrocannabinolic acid (THCA), which is converted to the neutral form Δ-9-tetrahydrocannabinol (THC) once heated. This neutral form interacts with the endocannabinoid system producing a psychoactive effect (gets us “high”). THC also seems to have important medical uses potentially serving as treatment for Parkinson’s disease (Carrol et al. 2012), dementia (Walther et al. 2006), and autoimmune disorders (Lyman et al. 1989). The other well-known cannabinoid in Cannabis is cannabidiolic acid (CBDA), which produces cannabidiol (CBD) when heated. Data suggest CBD, which is not psychoactive, may mitigate some of the negative effects of THC (such as anxiety and paranoia) and has potential uses in treating cancer (Soliman et al. 2015) and epilepsy (Mechoulam et al. 2002; Devinsky et al. 2014). Besides THC and CBD, Cannabis produces around 74 different cannabinoids (ElSohly et al. 2005; Radwan et al. 2008; ), which are present at varying potencies and ratios across particular cultivars and may also have medical importance, including cannabigerol (CBG) Borelli et al., 2014), cannabichrome (CBC) (Izzo et al. 2012) and Δ-9-tetrahydocannabivarin (THCV) (McPartland et al. 2015).

Cannabis also has interesting reproductive strategies. It can be either dioecious, meaning that there are male and female plants, similar to what we see in humans and other animals (Soltis et al. 2005; Bell et al. 2015). However some Cannabis varieties are monoecious, and thus produce both males and female flowers in the same plant. To make this even more confusing, when environmentally stressed, some male plants can produce female flowers and some female plants can produce male flowers. Additionally, sex is determined by two chromosomes, X and Y (thus males are XY and females are XX), but the hermaphrodites have undifferentiated chromosomes (Hirata 1929; Yamado 1943; Sakomoto et al. 1998). Interestingly, males, females and hermaphrodites can cross with each other and produce fertile offspring.

About the Author

Dr. Daniela Vergara is post-doctoral researcher at the University of Colorado, Boulder, where she is working in Dr. Nolan Kane’s lab Cannabis Genomic Research Initiative. Specifically, Daniela has been exploring the cannabinoid genes in the genome and understanding the how these genes relate to the chemotypes. Daniela also founded and is the director of a non-profit organization The Agricultural Genomics Foundation that holds a 501(C)(3) status (AGF; and aims in becoming a genomic repository (“library of genomes”) helping CGRI perform their research. AGF also educates the public about science, Cannabis, evolutionary biology, and genomics, through public talks.


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McPartland, J. M., M. Duncan, V. Di Marzo & R. G. Pertwee. 2015. Are cannabidiol and Δ9‐tetrahydrocannabivarin negative modulators of the endocannabinoid system? A systematic review. British journal of pharmacology 172: 737–753.

Mechoulam, R., L.A. Parker & R. Gallily. 2002. Cannabidiol: an overview of some pharmacological aspects. The Journal of Clinical Pharmacology 42: 11S-19S.

Radwan, M. M., S. A. Ross, D. Slade, S. A. Ahmed, F. Zulfiqar & M. A. ElSohly. 2008. Isolation and characterization of new Cannabis constituents from a high potency variety. Planta medica 74: 267–272.

Russo, E. B. 2007. History of Cannabis and its preparations in saga, science, and sobriquet. Chemistry & Biodiversity 4: 1614–1648.

Russo, E. B., H. E. Jiang, X. Li, A. Sutton, A. Carboni, F. Del Bianco, G. Mandolino, D. J. Potter, Y. X. Zhao, S. Bera & Y. B. Zhang. 2008. Phytochemical and genetic analyses of ancient cannabis from Central Asia. Journal of Experimental Botany 59: 4171–4182.

Sakamoto, K., Y. Akiyama, K. Fukui, H. Kamada & S. Satoh 1998. Characterization; Genome Sizes and Morphology of Sex Chromosomes in Hemp (Cannabis sativa L.). Cytologia 63: 459–464.

Sawler, J., J. M. Stout, K. M. Gardner, D. Hudson, J. Vidmar, L. Butler, J. E. Page & S. Myles. 2015. The Genetic Structure of Marijuana and Hemp. PloS One 10: e0133292.

Solinas, M., V. Cinquina & D. Parolaro. 2015. Cannabidiol and Cancer—An Overview of the Preclinical Data. In Molecular Considerations and Evolving Surgical Management Issues in the Treatment of Patients with a Brain Tumor. InTech.

Soltis, D. E., P. S. Soltis, P. K. Endress & M. W. Chase. 2005. Phylogeny and evolution of angiosperms. Sinauer Associates Incorporated.

Walther, S., R. Mahlberg, U. Eichmann & D. Kunz. 2006. Delta-9-tetrahydrocannabinol for nighttime agitation in severe dementia. Psychopharmacology 185: 524–528.

Watts, G. 2006. Science commentary: Cannabis confusions. BMJ: British Medical Journal 332: 175.

Vergara, D., H. Baker, K. Clancy, K. G. Keepers, J. P. Mendieta, C. S. Pauli, S. B. Tittes, K. H. White & N. C. Kane. 2016. Genetic and Genomic Tools for Cannabis sativa. Critical Reviews in Plant Sciences 35: 364–377.

Yamada, I. 1943. The sex chromosome of Cannabis sativa L. Seiken Ziho2: 64–68.


Jonathan Foox

Postdoctoral Associate, Institute for Computational Biomedicine, Weill Cornell Medicine, New York, New York


This month's featured taxon is Myxozoa: a bizarre, poorly understood group of microscopic, obligate parasites. Members of this taxon are typically found parasitizing teleost fish and annelid worms, though they have been observed in a wide spectrum of hosts including amphibians, birds, bryozoans, cephalopods, reptiles, shrews, and waterfowl. These parasites are globally distributed in marine and freshwater aquatic environments (though some are exclusively terrestrial), and have been found in nearly all tissue and organ types. Myxozoa is an extremely diverse group not only in distribution but in species richness, comprising over 2,200 described species distributed among over 60 genera (Lom and Dyková, 2006) – which likely represents a small fraction of the total diversity, with some estimates of 16,000 species in the Neotropics alone (Naldonia et al., 2011).

Although most myxozoan infections are innocuous, some species are well known pathogens that cause fatal diseases that can have significant economic impact, particularly on fish farms (Kent et al., 2001). One especially nasty example is Tetracapsuloides bryosalmonae, the causative agent of Proliferative Kidney Disease, which can wipe out 90% of infected salmonid populations, and even caused authorities to shut down a 183-mile stretch of Yellowstone River last summer (Young, 2016).

Each individual myxozoan has a fantastically complex life cycle that involves radical physiological transformations. Upon penetration of a host, an individual amoeboid-like reproductive body will undergo complex rounds of cellular fusion and division, before ultimately producing a reproductive spore that will eventually emerge from its host into the water column in search of its next host. These spores exhibit a stunningly diverse array of morphologies, including spherical, fusiform, pyriform, floral, round, ovoid, flattened, elongated, with or without caudal appendages, and all variations exhibit a wide variety of variation in orientation and number of constituent parts. The image gives just a taste of the incredibly morphological diversity of this taxon. In rather dramatic fashion, these spores harbor a complex organelle known as a polar capsule, which contains a coiled up filament that, upon stimulation, will rapidly evert from the capsule like the finger of a glove. The sticky filament flies through the water and latches onto the integument of the target animal like a grappling hook, allowing the spore to wriggle its way into its next host and beginning the parasitic cycle anew.

But perhaps the most impressive thing about Myxozoa is its position within the tree of life. These microscopic, morphologically simplistic parasites are members of the phylum Cnidaria, the lineage containing animals such as jellyfish, sea anemones, and corals. Indeed, myxozoans are extremely divergent, incredibly reduced, highly derived evolutionary cousins of these commonly known creatures. And this relationship of myxozoans to its cnidarian allies renders the group one of the most dramatically degenerate parasitic radiations known to biology. Myxozoans have neither tentacles, nor gastrovascular cavities, nor even tissue layers – and yet, they are cnidarians, by virtue of their polar capsules, which are homologous to cnidocytes (the stinging organelles only found within Cnidaria).

To put it into perspective: the size difference between an individual myxozoan spore and the common moon jelly is equivalent to the size difference between a human and Mt. Everest. Quite the difference.

And yet, this evolutionary relationship was not understood for nearly two hundred years. After first discovery in the early 19th century, myxozoans were categorized as various protistan lineages. Upon their confirmation as cnidarians not much more than 20 years ago (Siddall et al., 1995), biologists realized that myxozoans are not only incredibly derived cnidarians, but that they are the smallest and perhaps simplest animals in existence. Having lost nearly all diagnostic features known to animals (cellular structures such as centrioles and cilia), myxozoans stretch the very limit of what we understand to be "animals". It is only fitting that we save for the end of the year a taxon that stretches the limits of our biological imagination.


Kent, M.L., K.B. Andree, J.L. Bartholomew, M. El-Matbouli, S.S. Desser, R.H. Devlin, S.W. Feist, P.P. Hedrick, R.W. Hoffmann, J. Khattra, S.L. Hallett, R.J.G. Lester, M. Longshaw, O. Palenzeula, M.E. Siddall ME & C.X. Xiao. 2001. Recent advances in our knowledge of the Myxozoa. J Eukaryot Microbiol  48: 395–413.

Lom, J., & I. Dykova. 2006. Myxozoan genera: definition and notes on taxonomy, life-cycle terminology and pathogenic species. Folia Parasitology 53: 1-36. London, pp. 115–154.

Naldonia, J., S. Aranab, A.A.M. Maiac, M.R.M. Silvac, M.M. Carrieroc, P.S. Ceccarellid, L.E.R. Tavarese & E.A. Adrianof. 2011. Host–parasite–environment relationship, morphology and molecular analyses of Henneguya eirasi n. sp. parasite of two wild Pseudoplatystoma spp. in Pantanal Wetland, Brazil. Veterinary Parasitology 177: 247–255.

Siddall, M.E., D.S. Martin, D. Bridge, S.S. Desser & D.K. Cone. 1995. The demise of a phylum of protists: phylogeny of Myxozoa and other parasitic Cnidaria. Journal of Parasitology 81: 961–967.

Young, Ed.  2016.  A Tiny Jellyfish Relative Just Shut Down Yellowstone River.  The Atlantic: .


Ricardo Bressan Pacifico

Ph.D. Student, Plant systematics and biogeography lab - Maringá State University, Maringá, Brazil

Photo credit. A. V. Scatinga.

Photo credit. A. V. Scatinga.

With Halloween just around the corner, our taxon of the month is a recently described and unique genus of plants with unusual feeding habits, Philcoxia. This genus was first described 17 years ago, known only from three species (Taylor & Souza, 2000), although since then several additional species were recently discovered bringing the number of species in the genus to seven (Scatigna et al., 2015; 2017). All Philcoxia species are rare and are endemic to central Brazilian mountaintop grasslands, usually known as campo rupestre (Taylor & Souza, 2000). They are annual herbs, usually less than 30 cm tall, with delicate roots and stems, and small white to purple flowers measuring less than 1 cm in length. However, the most striking features of Philcoxia took more than a decade to be discovered. All species have underground leaves to which many nematodes attach and these leaves look somewhat similar to those found in carnivorous plants, a feature which caught the attention of researchers from California and Brazil, who decided to perform carnivory tests (Fritsch et al., 2007). The initial carnivory test results were negative (Fritsch et al., 2007), however, a few years later, a new and creative experiment shed light on this matter. In this experiment, radioactive nitrogen (15N) was used to feed bacteria (Escherichia coli) that were fed to to a population of nematodes (Caenorhabdtis elegans), which, in turn, were placed over the underground leaves of Philcoxia minensis for two days. The idea was to track the nutrient acquisition of Philcoxia, i. e., to see if the radioative nitrogen from nematodes would somehow be absorbed by this plant. The fast absorption of the 15N revealed by the elevated concentration of it in Philcoxia leaves strongly suggested that the nematodes were digested (instead of naturally decomposed) and absorbed by Philcoxia leaves (Pereira et al. 2012)⁠ suggesting that this genus of plants is carnivorous and feeds on nematodes. Carnivory evolved at least six times within angiosperms (flowering plants) and about 20 carnivorous genera distributed in 10 distinct families have been identified. A general cost–benefit model predicts that carnivory will be restricted to well lit, low-nutrient areas, where the major source of important nutrients such as nitrogen and phosphorus will be obtained from captured and digested invertebrates (Pereira et al. 2012).⁠ Philcoxia, like other carnivorous plants, live in nutrient-poor soils and are the only known carnivorous plants in the Plantaginaceae family (Pereira et al. 2012). The unusual new mechanism of carnivory discovered in Philcoxia caught public attention in high impact scientific journals such as Nature (Rowland, 2012).


Fritsch, P. W., F. Almeda, A. B. Martins, B. C. Cruz and D. Estes. 2007. Rediscovery and phylogenetic placement of Philcoxia minensis (Plantaginaceae), with a test of carnivory. Proceedings of the California Academy of Sciences 58: 447–467

Pereira, C. G., D. P. Almenara, C. E. Winter, P. W. Fritsch, H. Lambers and R. S. Oliveira. 2012. Underground leaves of Philcoxia trap and digest nematodes. Proc. Natl. Acad. Sci. U. S. A. 109: 1–5. doi:10.1073/pnas.1114199109.

Rowland, K. 2012. Hungry plant traps worms underground. Nature (news). doi:10.1038/nature.2012.9757

Scatigna, A. V., V. C. Souza, C. G. Pereira, M. A. Sartori, and A. O. Simoes. 2015. Philcoxia rhizomatosa (Gratioleae, Plantaginaceae): A new carnivorous species from Minas Gerais, Brazil. Phytotaxa 226: 275–280

Scatigna, A. V., Silva, N. G., Alves, R. J. V., Souza, V. C. and O. Simões. 2017. Two New Species of the Carnivorous Genus Philcoxia (Plantaginaceae) from the Brazilian Cerrado. Systematic Botany 42:351-357. doi: 10.1600/036364417X695574

Taylor, P., Souza, V. C., Giulietti, A. M. and R. M. Harley. 2000. Philcoxia: A new genus of Scrophulariaceae with three new species from eastern Brazil. Kew Bulletin 55: 155–163. 


Stephanie F. Loria

We have been pretty biased towards multicellular organisms in the Taxon of the Month posts. But this month, we are doing justice to our single-celled organism friends giving them the recognition they deserve as they are so crucial to the health of all multicellular life. For September, we focus our attention on the bacteria family, Enterobacteriaceae. Enterobacteriaceae are quite diverse and include more than 200 species in 51 genera (Octavia & Lan 2014; Janda & Abbott 2015). All Enterobacteriaceae are gram-negative, meaning that they possess a thin peptidoglycan layer in their cell walls causing them to appear pink after Gram staining (Beveridge 2001). Some well-known Enterobacteriaceae members include the medically important Escherichia coli, Salmonella and Klebsiella (Janda & Abbott 2015). E. coli is an essential human gut bacterium that can also act as a pathogen under certain conditions (Janda & Abbott 2015). Salmonella is notorious for causing illness of the human digestive system, which is sometimes fatal, and is transmitted through food and water contaminated with feces (Janda & Abbott 2015). Klebsiella species are found free-living in soil or water or in vertebrate digestive systems but are also responsible for a number of human illnesses including urinary tract infections and pneumonia.

Bacteria from the gastrointestinal tract of  Narceus americana.&nbsp; Photo credit to C. Wright.

Bacteria from the gastrointestinal tract of Narceus americana. Photo credit to C. Wright.

Many organisms rely on gut-inhabiting bacteria to assist with the digestion of various foods. For example, detritivores, organisms that eat decaying organic matter in the soil, rely on bacteria for assistance in breaking down hard-to-digest plant material, such as cellulose (Taylor 1982). Many Enterobacteriaceae inhabit animal digestive systems and are known to assist with digestion (Lauzon et al. 2003). For a class project as an undergraduate, a fellow student (C. Wright) and I agar plated the gut contents of a common detritivore, the large North American millipede, Narceus americanus. After sequencing the 16S rRNA gene of the plated bacterial colonies, we discovered several members of Enterobacteriaceae inhabiting this millipede's gut including Bacillus mycoides, Serratia sp. and Enterobacter cloacae. All three of these bacteria were previously known to inhabit animal digestive tracts. B. mycoides was previously found in both the soil (Lewis 1932) and in earthworm guts (Jensen et al. 2003). Enterobacter cloacae is known from plants and insect digestive systems (Watanabe & Sato, 1998). Serratia has been recorded in the digestive tract of flies in the genus Dacus (Lloyd et al., 1986). It is possible that these bacteria are assisting this millipede species digest its food.

Several studies have examined the diversification of Enterobacteriaceae. Research indicates that the evolution of endosymbiotic forms occurred multiple times in this family (Husník et al. 2011). Additionally, many endosymbiotic Enterobacteriaceae coevolved with their hosts (Duchaud et al. 2003; Moran et al. 2005). Given their species diversity and the wide range of hosts they inhabit, Enterobacteriaceae are great organisms to study for understanding selective pressures on symbiotic relationships.


Beveridge, T.J. 2001. Use of the gram stain in microbiology. Biotechnic & Histochemistry 76: 111–118.

Duchaud, E., C. Rusniok, L. Frangeul, C. Buchrieser, A. Givaudan, S. Taourit, S. Bocs, C. Boursaux-Eude, M. Chandler, C. Jean-Francois and E. Dassa. 2003. The genome sequence of the entomopathogenic bacterium Photorhabdus luminescensNature biotechnology 21: 1307–1313.

Husník, F., T. Chrudimský & Václav Hypša. 2011. Multiple origins of endosymbiosis within the Enterobacteriaceae (γ-Proteobacteria): convergence of complex phylogenetic approaches. BMC biology 9: 87.

Janda, J.M. & S.L. Abbott. 2015. The family Enterobacteriaceae. Practical handbook of microbiology (Goldman, Emanuel and L. H. Greens, eds) 307–319.

Jensen, G.B., B.M. Hansen, J. Eilenberg & J. Mahillon. 2003. The hidden lifestyles of Bacillus cereus and relatives. Environmental microbiology 5: 631–640.

Lauzon, C. R., T. G. Bussert, R. E. Sjogren & R. J. Prokopy. 2003. Serratia marcescens as a bacterial pathogen of Rhagoletis pomonella fies (Diptera: Tephritidae). European Journal of Entomology 100: 87–92.

Lundgren, J. G., R. Michael Lehman & J. Chee-Sanford. 2007. Bacterial communities within digestive tracts of ground beetles (Coleoptera: Carabidae). Annals of the Entomological Society of America 100: 275–282.

Lewis, I.M. 1932. Dissociation and life cycle of Bacillus mycoides. Journal of bacteriology 24: 381–421.

Llyod, A.C., R.A.I. Drew, D.S. Teakle & A.C. Hayward. 1986. Bacteria associated with some Dacus species (Diptera: Tephritidae) and their host fruit in Queensland. Australian Journal of Biological Scienes 39: 361–368.

Moran, N.A., J.A. Russell, R. Koga and T. Fukatsu. 2005. Evolutionary relationships of three new species of Enterobacteriaceae living as symbionts of aphids and other insects. Applied and Environmental Microbiology 6: 3302–3310. 

Octavia, S. & R. Lan. 2014. The family enterobacteriaceae. The Prokaryotes: Gammaproteobacteria 225–286.

Taylor, E.C. 1982. Role of aerobic microbial populations in cellulose digestion by desert millipedes. Applied and environmental microbiology 44: 281–291.

Watanabe, K. & M. Sato. 1998. Plasmid-mediated gene transfer between insect-resident bacteria, Enterobacter cloacae, and plant-epiphytic bacteria, Erwinia herbicola, in guts of silkworm larvae. Current Microbiology 37: 352–355.

Atlantic Horseshoe Crab (Limulus polyphemus)

Stephanie F. Loria

This month we honor an organism which will soon begin its breeding season in NYC - the Atlantic horseshoe crab, Limulus polyphemus. Despite their name and superficial resemblance, horseshoe crabs are not crabs. They actually belong to their own class Xiphosura in Chelicerata, an arthropod group that also includes the classes Arachnida (spiders, scorpions, ticks, etc), Eurypterida (the extinct sea scorpions and also MSNH's logo taxon), and Pycnogonida (sea spiders). The placement of Xiphosura within Chelicerata has been debated and recent research has even placed Xiphosura within Arachnida (Sharma et al. 2014). Worldwide only four extant species of horseshoe crabs exist and all species except L. polyphemus are found in the Indo-Pacific Ocean (Xia 2000). Extinct horseshoe crab species have also been described and the oldest fossil, found in Canada, dates to the Upper Ordovician, 445 million years ago (Rudkin et al. 2008)! Despite their remarkably old age, horseshoe crabs have changed little morphologically since their first appearance and are therefore often referred to as 'living fossils' in the scientific literature (Avise et al. 1994). 

The breeding season of L. polyphemus runs from March to July with peak season occurring in May and June (Rudloe 1980; Rutecki et al. 2004). During the breeding season, male and female L. polyphemus arrive on the shores of eastern North America in droves with most breeding happening at high tide on new and full moon nights (Rudloe 1980). Males typically mount females using special claspers and eggs are fertilized externally (Brockmann 1990). However, eggs may also be fertilized by satellite males which are not attached to females and surround the mating couple (Sasson et al. 2015). Eggs develop in the sand, hatching 3 to 4 weeks later and larvae disperse into the ocean (Bakker et al. 2016; Botton and Loveland, 2003; Rudloe, 1979). Horseshoe crabs live longer than dogs typically reaching 19 years of age (Rutecki et al. 2004).

During the breeding season, red knots (Calidris canutus rufa) feast on horseshoe crab eggs, an important food source for these birds (Niles et al. 2009). Horseshoe crabs are also harvested by humans for biomedical use as their blood contains amoebocyte lysate (ACL), a compound that can be used to detect bacterial endotoxins (Rutecki et al. 2004). Although biomedically harvested individuals are typically released once blood has been taken, mortality does occur among released individuals (Rutecki et al. 2004). Horseshoe crabs are also harvested for fishing bait and overharvesting from fishing and the biomedical industry and shoreline destruction has led to population declines (Land et al. 2015). In order to help track the health of horseshoe crab populations, nonprofit organizations such as NYC Audubon, and researchers survey horseshoe crabs populations each year during the breeding season. The MSNH teams up annually with NYC Audubon in the survey so that we can interact and help protect this fascinating and ancient species.

Horseshoe crabs ( L. polyphemus ) at Jamaica Bay. Courtesy of Maurice Chen.

Horseshoe crabs (L. polyphemus) at Jamaica Bay. Courtesy of Maurice Chen.


Avise, J. C., W. S. Nelson, and H. Sugita. 1994. A speciational history of" living fossils": molecular evolutionary patterns in horseshoe crabs. Evolution: 1986-2001.

Bakker, A. K., J. Dutton, M.  Sclafani and N. Santangelo. 2016. Environmental exposure of Atlantic horseshoe crab (Limulus polyphemus) early life stages to essential trace elements. Science of The Total Environment 572: 804-812.

Botton, M. L. and R. E. Loveland. 2003. Abundance and dispersal potential of horseshoe crab (Limulus polyphemus) larvae in the Delaware estuary. Estuaries 26: 1472-1479.

Brockmann, H. J. 1990. Mating behavior of horseshoe crabs, Limulus polyphemusBehaviour 114: 206-220.

Landi, A. A., J. C. Vokoun, P. Howell, and P. Auster. 2015. Predicting use of habitat patches by spawning horseshoe crabs (Limulus polyphemus) along a complex coastline with field surveys and geospatial analyses. Aquatic Conservation: Marine and Freshwater Ecosystems. 25: 380-395.

Niles, L. J ., J. Bart, H. P. Sitters, A. D. Dey, K. E. Clark, P. W. Atkinson, A. J. Baker, K. A. Bennett, K. S. Kalasz, N. A. Clark, and J. Clark. 2009. Effects of horseshoe crab harvest in Delaware Bay on Red Knots: are harvest restrictions working? Bioscience59: 153-164.

Rudkin, D. M., G. A.  Young, and G. S. Nowlan. 2008. The oldest horseshoe crab: a new Xiphosurid from Late Ordovician Konservat‐Lagerstätten Deposits, Manitoba, Canada. Palaeontology 51: 1-9.

Rudloe, A. 1979. Locomotor and light responses of larvae of the horseshoe crab, Limulus polyphemus (L.). The Biological Bulletin 157: 494-505.

Rudloe, A. 1980. The breeding behavior and patterns of movement of horseshoe crabs, Limulus polyphemus, in the vicinity of breeding beaches in Apalachee Bay, Florida. Estuaries and Coasts 3: 177-183.

Rutecki, D., R. H. Carmichael, and I. Valiela. 2004. Magnitude of harvest of Atlantic horseshoe crabs, Limulus polyphemus, in Pleasant Bay, Massachusetts. Estuaries and Coasts 27: 179-187.

Sasson, D. A., S. L. Johnson, and H. J. Brockmann. 2015. Reproductive tactics and mating contexts affect sperm traits in horseshoe crabs (Limulus polyphemus). Behavioral ecology and sociobiology 69: 1769-1778.

Sharma, P. P., S. T. Kaluziak, A. R. Pérez-Porro, V. L. González, G. Hormiga, W. C. Wheeler, and G. Giribet. 2014. Phylogenomic interrogation of Arachnida reveals systemic conflicts in phylogenetic signal. Molecular Biology and Evolution, p.msu235.

Xia, X. 2000. Phylogenetic relationship among horseshoe crab species: effect of substitution models on phylogenetic analyses. Systematic Biology 49: 87-100.