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 : www.cricketfarm.org or my more frequently updated instagram : www.instagram.com/bkcrickets or email me at : brooklyncrickets@gmail.com.

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: www.adamchadbrody.com.

References

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. https://doi.org/10.1665/034.025.0208

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”).

Acknowledgements

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.

References

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.

Desportes, I. 1984. The Paramyxea Levine 1979: An original example of 10volution towards multicellularity. Origin of Life 13: 343–352.

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.

Ford, S.E., N.A. Stokes, E.M. Burreson, E. Scarpa, R.B. Carnegie, J.N. Kraeuter & D. Bushek. 2009. Minchinia mercenariae n. sp. (Haplosporidia) in the hard clam Mercenaria mercenaria: implications of a rare parasite in a commercially important host. J. Euk. Microbiol. 56: 542–551.

Janouškovec, J., D.V. Tikhonenkov, F. Burki, A.T. Howe, F.L. Rohwer, A.P. Mylnikov & P.J. Keeling. 2017. A New Lineage of Eukaryotes Illuminates Early Mitochondrial Genome Reduction. Curr. Biol. 27: p3717–3724.

Larsson, J.I.R. & M. Køie. 2005. Ultrastructural study and description of Paramyxoides nephtys gen. n., sp. n. a parasite of Nephtys caeca (Fabricius, 1780) (Polychaeta: Nephtyidae). Acta Protozool. 44: 175–187.

Lester, R.J.G. & P.M. Hine. 2017. Paramyxida. in Handbook of the Protists (eds. Archibald J., Simpson AGB., Slamovits C.).

Seenivasan, R., N. Sausen, L.K. Medlin & M. Melkonian. 2013. Picomonas judraskeda Gen. et sp. Nov: The first identified member of the Picozoa Phylum Nov., a widespread group of picoeukaryotes, formerly known as ‘Picobiliphytes’. PLoS ONE 8: e59565.

Simpson, A.G.B. &. Y. Eglit. 2016. Protist diversification. Encyclopedia of Evolutionary Biology 3: 344–360.

Stentiford, G.D., A. Ramilo, E. Abollo & R. Kerr. 2017. Hyperspora aquatica n.gn., n.sp. (Microsporidia), hyperparasitic in Marteilia cochillia (Paramyxida), is closely related to crustacean-infecting microspordian taxa. Parasitol. 144: 186–199.

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 (http://www.wec.ufl.edu/faculty/olim/olilabs.html#Elise). 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.

References

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 https://www.hbw.com/node/56351 on 22 January 2018).

BirdLife International. 2016. Pseudocalyptomena graueri. The IUCN Red List of Threatened Species 2016: e.T22698719A93699841.

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.

Chapin, R.T. 1978. Brief accounts of some central African birds based on the journals of James Chapin. Rev. Zool. Afr. 92: 805–836.

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.

Janzen, D.H. 1967. Why mountain passes are higher in the tropics. The American Naturalist 101: 233–249.

Lowe, P.R. 1931. On the anatomy of Pseudocalyptomena and the occurrence of broadbills (Eurylaimidae) in Africa. Proc. Zool. Soc. London 1931: 445–461.

Lindsell, J.A. 2001. A breeding record and behavioural observation of African Green Broadbill Pseudocalyptomena graueri in south-western Uganda. Scopus 22: 68–70.

MacArthur, R.H. & MacArthur, J.W. 1961. On bird species diversity. Ecology, 42, 594–598.

McCain, C.M. 2009. Vertebrate range sizes indicate that mountains may be ‘higher’ in the tropics. Ecology letters, 12: 550–560.

Phillipps, P. G. & Seimon, A. 2010. Potential climate change impacts in conservation landscapes of the Albertine Rift. WCS Albertine Rift Climate Assessment. whitepaper report.

Plumptre, A.J., Davenport, T.R.B., Behangana, M., Kityo, R., Eilu, G., Ssegawa, P., Ewango, C., Meirte, D., Kahindo, C., Herremans, M., Peterhans, J.K., Pilgrim, J.D., Wilson, M., Languy, M. & Moyer, D. 2007. The biodiversity of the Albertine Rift. Biol. Conserv. 134: 178–194.

Rahbek, C. 1995. The elevational gradient of species richness: a uniform pattern? Ecography 18: 200–205.

Rockefeller, J.S. & Murphy, C.B.G. 1933. The rediscovery of Pseudocalyptomena. Auk 50: 23–29.

Terborgh, J. (1977). Bird species diversity on an Andean elevational gradient. Ecology 58: 1007–1019.

Terborgh, J. 1990. Mixed flocks and polyspecific associations: costs and benefits of mixed groups to birds and monkeys. American Journal of Primatology 21: 87–100.

Wilson, J.R. & Catsis, M.C. 1990. A preliminary survey of the forest of the 'Itombwe' mountains and the Kahuzi-Biega National Park extension, east Zaire, July-September 1989. Unpublished report for the World Wide Fund for Nature, London.

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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.

References

Darwin, C. R. On the Origin of Species by Means of Natural Selection, or the Preservation of Favoured Races in the Struggle for Life. London: John Murray. (1st edition).

Jokela, J., M. F. Dybdahl and C. M. Lively. 2009. The Maintenance of Sex, Clonal Dynamics, and Host-Parasite Coevolution in a Mixed Population of Sexual and Asexual Snails. 174: S43–S53.

King, K. C., L. F. Delph, J. Jokela and C. M. Lively. 2011. Coevolutionary hotspots and coldspots for host sex and parasite local adaptation in a snail-trematode interaction. Oikos. 120: 1335–1340.

Lively, C. M. 1987. Evidence from a New Zealand snail for the maintenance of sex by parasitism. Nature. 328: 519–521.

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.

Wallace, C. 1992. Parthenogenesis, Sex and Chromosomes in Potamopyrgus. Journal of Molluscan Studies. 58: 93–107. 

Winterbourn, M. J. 1970.  Population Studies on the New Zealand Freshwater Gastropod, Potamopyrgus antipodarum (Gray).  Journal of Molluscan Studies39: 139–149.

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.

Winterbourn, M. J. 1974.  Larval Trematoda Parasitising the New Zealand Species of Potamopyrgus (Grstropodoa: Hydrobiidae). Mauri Ora. 2: 17–30.

Cannabis

Daniela Vergara

Postdoctoral Fellow, University of Colorado, Boulder, Colorado

IMG_7824589.jpg

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; AgriculturalGenomics.org) 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.

References

Bell, C. D., D. E. Soltis & P. S. Soltis. 2010. The age and diversification of the angiosperms re-revisited. American Journal of Botany 97: 1296–1303.

Borrelli, F., E. Pagano, B. Romano, S. Panzera, F. Maiello, D. Coppola, L. De Petrocellis, L. Buono, P. Orlando & A. A. Izzo. 2014. Colon carcinogenesis is inhibited by the TRPM8 antagonist cannabigerol, a Cannabis-derived non-psychotropic cannabinoid. Carcinogenesis 35: bgu205.

Carroll, C. B., M‐L. Zeissler, C. O. Hanemann & J. P. Zajicek. 2012. Δ9‐tetrahydrocannabinol (Δ9‐THC) exerts a direct neuroprotective effect in a human cell culture model of Parkinson's disease. Neuropathology and Applied Neurobiology 38: 535–547.

Clarke, R. C. & M. D. Merlin. 2013. Cannabis: evolution and ethnobotany. University of California Press.

Devinsky, O., M. R. Cilio, H. Cross, J. Fernandez‐Ruiz, J. French, C. Hill, R. Katz, V. Di Marzo, D. Jutras‐Aswad, W. G. Notcutt & J. Martinez‐Orgado. 2014. Cannabidiol: pharmacology and potential therapeutic role in epilepsy and other neuropsychiatric disorders. Epilepsia 55: 791–802.

ElSohly, M.A. & D. Slade. 2005. Chemical constituents of marijuana: the complex mixture of natural cannabinoids. Life sciences 78: 539–548.

Gertsch, J., M. Leonti, S. Raduner, I. Racz, J.-Z. Chen, X.-Q. Xie, K.-H. Altmann, M. Karsak & A. Zimmer. 2008. Beta-caryophyllene is a dietary cannabinoid. Proceedings of the National Academy of Sciences 105: 9099–9104.

Hirata, K. 1929: Cytological basis of the sex determination in Cannabis sativa. Idengaku Zasshi. 4: 198–201.

Izzo, A. A., R. Capasso, G. Aviello, F. Borrelli, B. Romano, F. Piscitelli, L. Gallo, F. Capasso, P. Orlando & V. Di Marzo. 2012. Inhibitory effect of cannabichromene, a major non‐psychotropic cannabinoid extracted from Cannabis sativa, on inflammation‐induced hypermotility in mice. British journal of Pharmacology 166: 1444–1460.

Li, H. L. 1973. An archaeological and historical account of cannabis in China. Economic Botany 28: 437–448.

Li, H. L. 1974. Origin and use of Cannabis in Eastern Asia; Linguistic-cultural implications. Economic Botany 28: 293–301.

Lyman, W. D., J. R. Sonett, C. F. Brosnan, R. Elkin & M. B. Bornstein. 1989. Δ 9-tetrahydrocannabinol: a novel treatment for experimental autoimmune encephalomyelitis. Journal of neuroimmunology 23: 73–81.

Lynch, R. C., D. Vergara, S. Tittes, K. White, C. J. Schwartz, M. J. Gibbs, T. C. Ruthenburg, K. deCesare, D. P. Land & N. C. Kane. 2016. Genomic and Chemical Diversity in Cannabis. Critical Reviews in Plant Sciences 35: 349–363.

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.

Myxozoa

Jonathan Foox

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

Taxonofthemonth_myxozoan_diversity.png

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.

References

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: https://www.theatlantic.com/science/archive/2016/08/the-parasite-that-just-shut-down-a-montana-river-has-an-unbelievable-origin/496817/ .

Philcoxia

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).

References

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. 

Enterobacteriaceae

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.

References

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 Audobon, and researchers survey horseshoe crabs populations each year during the breeding season. This year, the MSNH will team up again with NYC Audubon on Friday, June 9 to participate 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.

References

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.