Friday Fellow: Blue Jacaranda

by Piter Kehoma Boll

Let’s keep the trend of last week and present again a South American species, but today’s fellow has moved way beyond its original range.

Scientifically known as Jacaranda mimosifolia, the common names of this species are blue jacaranda, fern tree or simply jacaranda. Its native range includes a considerably small area between Argentina and Bolivia, but it is grown as an ornamental tree throughout the whole world.

Flowers of a specimen in its native range in Argentina. Photo by Martin Arregui.*

An iconic tree, the blue jacaranda reaches up to 20 m in height. Its bark is smooth at first but later becomes scaly and rough as it typical of trees of the family Bignoniaceae, to which it belongs. The leaves are large, up to 45 cm long, and are bipinnately compound, i.e., the compound leaf itself consists of compound leaflets, which is likely the reason why it is sometimes called fern tree.

A blue jacaranda leaf. Photo by Wikimedia user Crusier.**

The flowers are, however, the most iconic feature of this tree, appearing in spring and early summer. They are tubular, reach about 5 cm in length, have a pale purple-indigo color and are grouped in large panicles. The fruits are dry woody pods with a somewhat oval shape and are often gathered for decoration purposes, including the decoration of Christmas trees or as body ornaments, such as the confection of earrings.

The dry woody pods of the blue Jacaranda. Photo by Wikipedia user Babbage.**

The wood of the blue jacaranda has a light color and is considerably soft, being often used for the creation of sculptures and bowls, especially when still green.

Wood of the blue Jacaranda. Photo by Wikimedia user SybillKaesedick.***

The blue jacaranda became an important cultural element in many regions of the world. It is often featured in songs, especially in Argentina and Brazil. In South Africa, the city of Pretoria is also known as the Jacaranda City due to the large number of blue jacarandas that turn the city blue in Spring. In Australia, the blue jacaranda became associated with the final exams of students in the University of Queensland, which is known for its jacarandas. The trees flower during the time the students are running to complete their assignments and study for their final exams, which give rise to the expression “purple panic”.

A blue jacaranda in Campinas, Brazil. Photo by Enio Prado.**

Despite its widespread occurrence as an ornamental plant, the blue jacaranda is considered vulnerable in its native habitat by the IUCN’s Red List. In other areas, such as South Africa and Australia, for example, the tree is sometimes an invasive species, outcompeting native trees by blocking their growth.

Blue jacaranda trees in Pretoria, South Africa. Photo by Paul Saad.***

Due to such negative impacts, planting new jacarandas in Pretoria is now forbidden. The idea of removing the adults trees, which was the original plan, was discarded due to their popularity with locals. Nevertheless, in some decades or centuries (provided that humanity will survive that much as a civilization), the Jacaranda City will eventually lose all its Jacarandas.

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References:

Prado D (1998) Jacaranda mimosifolia. The IUCN Red List of Threatened Species 1998: e.T32027A9675619. https://dx.doi.org/10.2305/IUCN.UK.1998.RLTS.T32027A9675619.en Access on 17 September 2020.

Wikipedia. Jacaranda mimosifolia. Available at < https://en.wikipedia.org/wiki/Jacaranda_mimosifolia >. Access on 17 September 2020.

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* This work is licensed under a Creative Commons Attribution-NonCommercial 4.0 International License.

** This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.

*** This work is licensed under a Creative Commons Attribution-ShareAlike 4.0 International License.

A superbeetle that couldn’t care less about cyanide poisoning

by Piter Kehoma Boll

In order to try avoiding predators, many species develop powerful toxins that would harm anyone trying to eat them, sometimes even killing them. However, predators can fight back by developing a strong resistance to the prey’s defenses, sometimes to amazing levels.

Millipedes are known for as distasteful prey that evolved a variety of toxins do deter predators. Nevertheless, some species have found ways to deal with millipede’s defenses, making the poor creatures desperate for new strategies to survive.

In North America, ground beetles of the genus Promecognathus are specialist predators of millipedes. The species Xystocheir dissecta, one of their main prey, produces cyanide as a chemical defense. Cyanide is a very toxic compound for most life forms.

The cyanide-producing millipede Xystocheir dissecta. Photo by iNaturalist user mhertel.*

In a recent study, 18 different ground beetle species were exposed to sodium cyanide (NaCN) in the lab to assess their resistance. While most species succumbed in less than 10 min when exposed to 15 mg of NaCN or less, three species did not give a damn even to quantities as high as 100 mg. These three species included Promecognathus crassus, P. laevissimus and Metrius contractus. While both Promecognathus species feed on Xystocheir dissecta, Metrius contractus does not.

Promecognathus laevissimus, the “I could have cyanide for breakfast” ground beetle. Photo by Eddie Dunbar.*

In another trial, the species were exposed to 100 mg of potassium cyanide (KCN) for up to two hours. While M. contractus ramained active during the first hour, all specimens succumbed in less than two hours, but after 120 min, some specimens of Promecognathus laevissius were still moving around as if nothing was happening.

Metrius contractus, resisting cyanide just for fun. Photo by iNaturalist user tparkeressig.*

This study is the first evidence of predators having resistance to cyanide. While this superpower in P. laevissimus is easily explained by its predatory behavior, the high resistance of M. contractus is still a mystery, as this species is not specialized in millipedes, although it is possible that it may eat them as an alternative food, especially sick or injured specimens. Both species, however, are resistant to amounts of cyanide way above the ones that they would find in any millipede. It’s a real superpower.

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Reference:

Weary BP, Will KW (2020) The Millipede-Predation Behavior of Promecognathus and Exceptional Cyanide Tolerance in Promecognathus and Metrius (Coleoptera: Carabidae). Annals of the Entomological Society of America. https://doi.org/10.1093/aesa/saaa023

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* This work is licensed under a Creative Commons Attribution-NonCommercial 4.0 International License.

Friday Fellow: Trapezoid Temnocephalan

by Piter Kehoma Boll

Although most people only know flatworms through their parasitic groups (monogeneans, trematodes and cestodes) and freshwater planarians, the diversity of this phylum is much greater. One peculiar group is that of the temnocephalids or temnocephalans, small commensal species that live attached to the surface of freshwater animals, especially crustaceans and gastropods, but also turtles and water bugs.

Some species are known to live attached to the common Brazilian river crab, including Temnocephala trapeziformis, a species that was only described in 2006. Without a common name, I decided to call it the trapezoid temnocephalan.

Three preserved specimens with their “chubby glove” look. Scale bar = 1 mm. Credits to Amato et al. (2006).*

The body of the trapezoid temnocephalan measures about 2 to 3 mm in length and has the typical shape of that of other temnocephalans. It looks like a chubby glove at first, with an oval body that has five finger-like projections at the anterior end and a sucker on the ventral side near the posterior end, through which it attaches to its host. The mouth lies a little behind the anterior end and connects to a short cylindrical pharynx. The name trapeziformis comes from the fact that this species has a trapezoid-shaped plate surrounding each nephridiopore (the excretory pores).

The trapezoid-shaped plates that surround the excretory pores (n) of the trapezoid temnocephalan give it its name. Credits to Amato et al. (2006).*

Being a commensal ectosymbiont, the trapezoid temnocephalan is not a parasite and usually does not cause much trouble to its host, although they can interfere in the host fitness if they occur in large numbers. Although the diet of the trapezoid temnocephalan was not studied, it probably feeds on a variety of unicelular and very small organisms, such as algae and small crustaceans, like other temnocephalans.

A preserved and prepared specimen showing the mouth and pharynx (oval shape above in the middle), intestine (brown), sucker (circle below), testes (two largest bluish circles on each side) and cirrus (small elongate tube below the intestine slightly to the right of the midline). Credits to Amato et al. (2006).*

Adult trapezoid temnocephalans can be found in almost every region of the crab’s surface, including the cavities of the eyes. They are hermaphrodite and mate using internal fertilization by delivering sperm through a penis-like structure called the cirrus. Fertilized eggs are attached to the host surface mostly on the fourth pair of walking legs but also at the sides of the carapace and in the eye cavities. When the young hatch, being already smaller versions of the adults, they often attach to the same crab in which they were born. As their number increase, we can imagine that the crabs starts to become annoyed and may remove them by self-grooming.

Eggs attached to the legs (above) and to the eye cavity (below) of the common Brazilian river crab. Credits to Amato et al. (2006).*

A dettached trapezoid temnocephalan can survive a good time without a host if it is able to obtain food, but its ideal home is on a common Brazilian river crab where it can not only find food without having to move around but also have access to others of the same species to reproduce.

The diversity and ecological role of temnocephalans are largely understudied. Thousands of unknown species are out there waiting to be discovered and have their relationship with their hosts better investigated.

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References:

Amato JFR, Amato SB, Seixas SA (2006) A new species of Temnocephala Blanchard (Platyhelminthes, Temnocephalida) ectosymbiont on Trichodactylus fluviatilis Latreille (Crustacea, Decapoda, Trichodactylidae) from southern Brazil. Revista Brasileira de Zoologia 23(3): 796–805. https://doi.org/10.1590/S0101-81752006000300026 

Cannon LRG, Joffe BI (2000) The Temnocephalida. In: Littlewood DTJ, Bray RA (Eds.) Interrelationships of the Platyhelminthes. CRC Press.

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* This work is licensed under a Creative Commons Attribution-NonCommercial 4.0 International License.

Friday Fellow: Common Brazilian River Crab

by Piter Kehoma Boll

Today we reach 250 friday fellows! Yay! And to celebrate one quarter of a thousand species, we will have two fellows today!

Our first species is a bit controversial because it is, in fact, a complex of several closely related species. However, for the sake of this post, we will consider them still as one species since the issue has not been settled yet. Named Trichodactylus fluviatilis, it does not seem to have a common name in English, so I decided to coin the name “common Brazilian river crab”.

A specimen in Espírito Santo, Brazil. Photo by Flávio Mendes.*

As the name implies, the common Brazilian river crab is a crab that is commonly found in Brazilian rivers and streams. It is widespread across several river basins near the Brazilian coast, mainly in areas of the Atlantic Forest. The size and shape of invididuals vary considerable even within the same population, which hinders an easy taxonomic determination of the different lineages. Adults have a carapace measuring between 15 and 40 mm in width, with females being only slightly larger than males. Males, however, have larger chelipeds (pincers) than females, while females have wider abdomens than males. The color varies from light to dark brown or even reddish brown.

A specimen on a human hand for size comparison. Photo extracted from https://www.leialab.com/.

The common Brazilian river crab lives in the leaf litter in streams and rivers together with several other invertebrates. Considered to be an omnivore, its main food consists of algae, but it also ingest decaying plant material and eventually feeds on dead animals as well.

During the reproductive period, males often fight for the females, which is why their chelipeds are larger. Females with eggs often remain hidden and, after their eggs have been fertilized, they carry them with them and even carry juvenile crabs for some time to protect them. This is the reason why they have much wider abdomens than males.

A female carrying her young in a stream in Espírito Santo, Brazl. Photo by Flávio Mendes.*

In small rural comunities, the common Brazilian river crab is an important cultural element and is often captured and consumed as food, especially by the poorest families. An ethnocarinological study conducted with inhabitants of a settlement in the state of Bahia in Brazil revealed that the population has a considerably good knowledge about the morphological, reproductive and ecological aspects of this species that are consistent with the results of research studies.

Traditional knowledge is not just a bunch of superstitions and miscoceptions. It can actually include relevant scientific data that a community acquired through observation and experimentation in their daily activities.

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References:

Costa L, Kiffer Jr. W, Casotti C, Rangel J, Moretti M (2016) Zoological Studies 55: e54. https://dx.doi.org/10.6620/ZS.2016.55-54

Costa Neto EM (2007) O caranguejo-de-água-doce, Trichodactylus fluviatilis (Latreille, 1828) (Crustacea, Decapoda, Trichodactylidae), na concepção dos moradores do povoado de Pedra Branca, Bahia, Brasil. Biotemas 20(1):59-68. https://periodicos.ufsc.br/index.php/biotemas/article/view/20781

Lima DJM, Cobo VJ, Alves DFR, Barros-Alves SP, Fransozo V (2011) Onset of sexual maturity and relative growth of the freshwater crab Trichodactylus fluviatilis (Trichodactyloidea) in south-eastern Brazil. Invertebrate Reproduction & Development 57(2):105–112. https://doi.org/10.1080/07924259.2012.689263

Pescinelli RA, Mantelatto FL, Costa RC (2020) Population features, sexual dimorphism and handedness of the primary freshwater crab Trichodactylus cf. fluviatilis (Brachyura: Trichodactylidae) from southeastern Brazil. Invetebrate Reproduction & Development 64(2):95–105. https://doi.org/10.1080/07924259.2019.1699176

Souza-Carvalho EA, Magalhães C, Mantelatto FL (2017) Molecular phylogeny of the Trichodactylus fluviatilis Latreille, 1828 (Brachyura: Trichodactylidae) species complex. Journal of Crustacean Biology 37(2):187–194. https://doi.org/10.1093/jcbiol/rux005

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* This work is licensed under a Creative Commons Attribution-NonCommercial 4.0 International License.

Friday Fellow: Lion’s Mane Jellyfish

by Piter Kehoma Boll

Today is time to talk again about a celebrity.

Although three cnidarians have been featured as Friday Fellows already, none of them was a jellyfish until now. So let’s introduce the first one and let it be a considerably famous species, the lion’s mane jellyfish Cyanea capillata.

Lion’s mane jellyfish near Newfoundland, Canada. Credits to Derek Keats.**

The lion’s mane jellyfish is a very large jellyfish, among the largest species known to date. Its bell can reach up to 2 m in diameter and its tentacles can grow up to 30 m in length, thus becoming longer than a blue whale! It is an inhabitant of the very cold waters of the Arctic and neighboring areas of the Atlantic and Pacific oceans. It cannot survive in warm waters and specimens living in the southernmost areas of its range cannot even grow to the full size.

The color of an adult lion’s mane jellyfish is pale red or pale yellow. Its genus name, Cyanea, which refers to a blue color, is due to another species of the genus, the blue jellyfish, Cyanea lamarckii. Its specific epithet, capillata as well as its common name are references to its dense mass of tentacles that resemble a lion’s mane. The jellyfish’s bell is divided into eight lobes, each lobe having from 70 to 150 tentacles. The indentations between the lobes contain a special organ, the rhopalium, that helps jellyfishes orient themselves in water.

A specimen in Norway. Credits to Arstein Rønning.*

Like all jellyfishes, the lion mane jellyfish has a complex life cycle. Adult specimens reproduce sexually in summer, with males releasing sperm into the water. The sperm swims into the body of the female, where the eggs are fertilized. The first life stage, the larva, grows inside the body of the female and is then released into the water where it attaches to a surface to become a polyp. The larvae seem to prefer rougher surfaces to attach and especially in darker places. The polyp grows during winter and reproduces asexually during spring. The asexual reproduction, called strobilation, occurs by the polyp releasing segments that become ephyrae, which are like very young jellyfish. The ephyrae grow to become adult jellyfish and restart the cycle.

The lion’s mane jellyfish feeds on a great variety of species during its life cycle, including plankton, invertebrates and even small vertebrates. Nevertheless, its huge body is also used as the habitat of several other animals that live in the cold northern waters. It also has a complicated relationship with the moon jellyfish, Aurelia aurita, with which it shares its habitat. While adult lion’s mane jellyfish prey on adult moon jellyfish, adult moon jellyfish prey on larvae and ephyrae of the lion’s mane jellyfish, and both species also compete for the same prey.

A lion’s mane jellyfish (top-right) capturing a moon jellyfish (bottom-left). Photo by W. Carter.

Like all cnidarians, the lion’s mane jellyfish stings. The contat with a single tentacle in humans usually does not cause much complication except for those with some sort of allergy or sensitivity. However, if you are unfortunate enough to end up swimming directy into the tentacle mass of a specimen, becoming covered by that stinging nightmare, you may end up having to be taken to a hospital quickly. Despite the low risk of killing a human, one lion’s mane jellyfish became famous as the assassin in one of Sherlock Holmes’ cases. The victim was an unfortunate guy with a heart condition, though.

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More cnidarians:

Friday Fellow: Deep-Sea Marr (on 22 April 2016)

Friday Fellow: Portuguese Man o’ War (on 7 July 2017)

Friday Fellow: Blue Coral (on 18 May 2018)

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References:

Brewer RH (1976) Larval settling behavior of Cyanea capillata (Cnidaria: Scyphozoa). The Biological Bulletin 150(2). https://doi.org/10.2307/1540467

Gröndahl F, Hernroth L (1987) Release and growth of Cyanea capillata (L.) ephyrae in the Gullmar Fjord, western Sweden. Journal of Experimental Marine Biology and Ecology 106(1):91–101. https://doi.org/10.1016/0022-0981(87)90149-3

Gröndahl F (1988) A comparative ecological study on the scyphozoans Aurelia aurita, Cyanea capillata and C. lamarckii in the Gullmar Fjord, western Sweden, 1982 to 1986. Marine Biology 97: 541–550. https://doi.org/10.1007/BF00391050

Wikipedia. Lion’s mane jellyfish. Available at < https://en.wikipedia.org/wiki/Lion%27s_mane_jellyfish >. Access on 3 September, 2020.

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* This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.

** This work is licensed under a Creative Commons Attribution-Share Alike 2.0 Generic License.

New Species: August 2020

by Piter Kehoma Boll

Here is a list of species described this month. It certainly does not include all described species. You can see the list of Journals used in the survey of new species here.

Tichowtungia aerotolerans is a new bacterium of the small phylum Kiritimatiellaeota. Credits to Mu et al. (2020).*

Bacteria

• 16 new actinobacteria: Agrococcus sediminis; Nonomuraea nitratireducens; Nonomuraea antri; Agromyces humi; Streptomyces apocyni; Streptomyces lycii; Streptomyces harenosi; Trebonia kvetii; Bifidobacterium canis; Gulosibacter macacae; Brevibacterium rongguiense; Planctomonas psychrotolerans; Arthrobacter yangruifuii, Arthrobacter zhaoguopingii; Actinomadura rubteroloni, Actinomadura macrotermitis;
• 8 new bacteroids: Hymenobacter artigasi; Hymenobacter fodinae, Hymenobacter metallicola; Algibacter marinivivus; Flavobacterium ichthyis; Agriterribacter humi; Salinimicrobium nanhaiense, Salinimicrobium oceani;
• 2 new cyanobacteria: Affixifilum floridanum, Neolyngbya biscaynensis;
• 3 new firmicutess: Alkalibaculum sporogenes; Paenibacillus algicola; Metabacillus sediminilitoris;
• 1 new hadobacterium: Deinococcus terrestris;
• 1 new kiritimatiellaeote: Tichowtungia aerotolerans;
• 2 new planctomycetes: Symmachiella dynata, Symmachiella macrocystis;
• 28 new proteobacteria: Intestinirhabdus alba; Ideonella livida; Terricaulis silvestris; Ketobacter nezhaii; Pseudomonas viciae; Pseudomonas karstica, Pseudomonas spelaei; Rhizobium rhizophilum; Rhizobium dioscoreae; Bradyrhizobium cosmicum; Paraburkholderia solitsugae, Paraburkholderia elongata; Rhodovarius crocodyli; Vibrio algicola; Nocardia macrotermitis, Nocardia aurantia; Halomonas rituensis, Halomonas zhuhanensis; Aquitalea aquatilis; Zoogloea dura; Amaricoccus solimangrovi; Yersinia artesiana, Yersinia proxima, Yersinia alsatic, Yersina vastinensis, Yersinia thracica, Yersinia occitanica; Kangsaoukella pontilimi;

Nonomuraea nitratireducens is a new actinobacterium isolated from the rhizosphere of the plant Suaeda sutralis in China. Credits to Ou et al. (2020).*

Trebonia kvetii is a new genus of actinobacteria foun in the Czech Republic. Credits to Rapoport et al. (2020).*

Archaeans

• 1 new euryarchaeote: Halomicroarcula amylolytica;

Excavates

• 5 new metamonads: Cononympha koidzumii, Cononympha monstrummogolloni, Cononympha skunkapei, Holomastigotoides bigfooti, Holomastigotoides batututi;

SARs

• 4 new ciliates: Prototapirella sabyinyo, P. sulcata, P. curiosa; Pseudokeronopsis parasongi;
• 1 new apicomplexan: Acroeimeria grismeri;
• 6 new ochrophytes: Nupela eremica; Brachysira aristidesii; Gyrosigma centropunctatum, G. dongzhaiense; Fistulifera alcalina; Aulacoseira umanai;
• 1 new oomycete: Periplasma isogametum;

Sonerila cardamomensis is a new melastomatacean from Cambodia. Credits to Shin et al. (2020).*

Plants

• 1 new marchantiophyte: Frullania amazonica;
• 1 new bryophyte: Brachymenium ayangannensis;
• 3 new pteridophytes: Hypodematium ryukyuense; Antrophyum nambanense; Hymenophyllum sp.;
• 1 new cycadophyte: Ceratozamia leptoceras;
• 4 new magnoliids: Magnolia poqomchi; Magnolia ottoi; Peperomia pasionana, Peperomia tancitaroana;
• 18 new monocots: Campylocentrum alvesii; Eremitis limae; Aloe vatovavensis, Aloe rakotonasoloi; Dendrochilum ignisiflorum; Lophiaris quichensis; Tillandsia itatiensis; Paepalanthus campanulatus; Peliosanthes nagalandensis, P. tobuensis; Anthurium siapidaarae; Laobambos calcareus; Cryptanthus cinereus; Hechtia elegans, H. ensifolia, H. medusae, H. platyphylla, H. pycnostachya;
• 52 new eudicots: Nepenthes putaiguneung; Microlicia windischii; Lippia krenakiana; Ditaxis grazielae; Phyllanthus pedicellatus; Begonia beijnenii; Cephalaria kurdistanica; Indigofera yuanjiangensis; Lactuca anatolica; Helosciadium milfontinum; Meriania microflora; Meriania ardyae, Meriania zunacensis; Stylosanthes minima, S. mitzii; Firmiana daweishanensis; Pinguicula tejedensis, P. casperiana; Schizanthus carlomunozii; Aglaia monocaula, Aglaia nyaruensis; Primula sunhangii; Geum sunhangii; Sonerila cardamomensis; Oreocharis eriocarpa, O. wenshanensis, O. fulva, O. lacerata; Oreocharis flavovirens; Oreocharis jasminina; Oreocharis wumengensis; Petrocosmea nanchuanensis; Didymocarpus lobulatus; Allocheilos maguanensis, A. rubroglandulosus; Petrocodon luteoflorus; Petrocodon rubiginosus; Petrocodon wenshanensis; Deinostigma fasciculatum; Paraboea myriantha, P. brevipedunculata; Neptunia windleriana; Cuscuta difficilis; Siphocampylus nebularis; Eugenia flavicarpa; Dactylaena boliviensis, D. zmarztyae; Plukenetia brevistyla, Plukenetia megastyla, Plukenetia chocoensis, Plukenetia sylvestris; Ipomoea bonsai; Randia heteromera;

Schizanthus carlomunozii is a new solanacean from Chile. Credits to Morales-Fierro et al. (2020).*

Fungi

• 1 new zygomycete: Cunninghamella guizhouensis;
• 16 new ascomycetes: Hypoxylon wujiangensis; Epicoccum endophytica; Aspergillus telluris; Arcopilus amazonicus; Muyocopron thailandica; Scheffersomyces jinghongensis, S. paraergatensis, S. anoplophorae; Dictyoarthrinium musae; Stylonectria qilianshanensis, Trichoderma gansuanum; Corylicola italica; Neoascochyta fuci, Paraconiothyrium salinum; Elaphomyces marmoratus, Elaphomyces fuscus;
• 25 new basidiomycetes: Lepiota condylospora; Lepiota cholistanensis; Rhodophana guandishanense; Ganoderma bambusicola; Amanita lacerosquamosa; Hebeloma sagarae; Fuscoporia ambigua; Junghuhnia austrosinensis, J. nandinae, J. subcollabens; Athelia termitophila; Hyphoderma fissuratum, H. mopanshanense; Colacogloea armeniacae; Phylloporia alyxiae, P. mori, P. murrayae, P. rubiacearum; Fomitiporia nubicola, F. conyana, F. murrilli; Phlebia fuscotuberculata, Ph. tomentopileata, Ph. tongxiniana; Clitopilus abprunulus;

Scheffersomyces jinghongensis is a new yeast from rotten wood in China. Credits to Jia et al. (2020).*

Junghuhnia subcollabens is a new crust fungus from China. Credits to Du et al. (2020).*

Poriferans

• 5 new demosponges: Radiospongilla fungosa; Hyattella sulfurea, H. pedunculata, Amphimedon palmata, Phoriospongia mozambiquensis;

Cnidarians

• 2 new anthozoans: Blastopathes medusa; Anemonactis tohrui;
• 10 new hydrozoans: Zygophylax dispersa, Z. encarnae, Z. laertesi, Z. medeae, Z. niobae, Z. pseudoabietinella; Actinopluma mirifica, Schizoplumularia helicoidalis, Corhiza patula, Thamnopteros uniserius;
• 3 new myxozoans: Myxobolus vetuschicanus, Myxobolus mineirus; Sphaeromyxa scorpaena;

Flatworms

• 3 new triclads: Obama apiguara, Paraba tata, Piima ata;
• 8 new monogeneans: Microcotyle kasago; Urocleidoides digitabulum, Urocleidoides solarivaginatus, Urocleidoides falxus, Urocleidoides sapucaiensis, Urocleidoides tenuis, Urocleidoides sinus, Urocleidoides uncinus;
• 5 new trematodes: Gaevskajatrema balistes; Aenigmatrema undecimtentaculatum, A. inopinatum, A. grandiovum; Retroporomonorchis pansho;
• 1 new cestode: Polypocephalus barsami;

Paraba tata is a new land planarian from Brazil. Credits to Oliveira et al. (2020).*

Mollusks

• 28 new gastropods: Caecum lightfootanum, C. draperi, C. shaskyi, C. galapagoense, C. spiculum; Amalda miriky, A. cacao; Kartvelobia sinuata, K. kinchkha, K. shishaensis, Imeretiopsis prometheus, I. gorgoleti, I. nakeralaensis, I. cameroni, I. iazoni, Caucasopsis letsurtsume, Caucasopsis olsavskyi, Caucasopsis egrisi, Caucasogeyeria gloeri, Caucasogeyeria ignidona, C. colchis, C. pseudocolchis, C. chrysomallos, Pontohoratia vinarskii, P. pichkhaiai, P. mapeli, Hausdorfenia shareula.

Nemerteans

• 2 new species: Euborlasia maycoli; Proamphiporus kaimeiae;

Annelids

• 14 new polychaetes: Gesaia csiro, Phalacrostemma timoharai; Syllis cambuk, Syllis hampirmenyatu, S. maganda; Aricidea (Acmira) filamentosa, Aricidea (Acmira) pearti, Aricidea (Acmira) bbkingi; Mauretanaspis longichaeta; Aricidea (Acmira) katzmanni, Aricidea (Acmira) pseudoassimilis; Acromegalomma sumbense, Claviramus olivager, Notaulax montiporicola;
• 2 new clitellates: Cataladrilus porquerollensis, Scherotheca portcrosana;

Bryozoans

• 1 new species: Smittoidea complexa;

Kinorhynchs

• 1 new species: Echinoderes vulcanicus;

Nematodes

• 7 new chromadoreans: Cobbionema brevispicula, Cobbionema acuminata; Rhabdochona iriomotensis, Spinitectus mirabilis; Contracaecum jorgei; Rhabdias kiri;

Tardigrades

• 1 new species: Crenubiotus revelator;

Arachnids

• 11 new mites: Cornigamasus allotritosternus; Eutegaeus parapapuaensis, Eutegaeus paralagrecai; Transeius conyzoides, Typhlodromus (Anthoseius) huangjiaensis; Cheladonta afshari, Neotrombicula tehranensis; Torrenticola pseudosiamis, T. anziensis, Monatractides sichuanensis; Mideopsis milankovici;
• 6 new harvestmen: Troglosiro dogny, T. pin, T. pseudojuberthiei, T. sharmai; Abaddon despoliator, Turonychus fadriquei;
• 2 new pseudoscorpions: Parobisium laevigatum, P.muchonggouense;
• 57 new spiders: Macrothele undata, M. arcuata, M. sanheensis; Spinirta jinyunshanensis, S. forcipata, S. sparsula, S. aurita, S. aviforma, S. quadrata, S. leigongshanensis, S. qizimeiensis, S. rugosa; Loxosceles tolantongo; Trilacuna changzi, T. hponkanrazi, T. triseta, T. zhigangi; Merizocera baoshan, M. betong, M. colombo, M. galle, M. hponkanrazi, M. kachin, M. kandy, M. mandai, M. krabi, M. kurunegala, M. lincang, M. mainling, M. nyingchi, M. peraderiya, M. phuket, M. putao, M. ranong, M. ratnapura, M. salawa, M. tak, M. tanintharyi, M. tengchong, M. thenna, M. uva, M. wenshan, M. wui, M. yala, M. yuxi; Heteroonops scapula, H. jurassicus, H. aylinalegreae, H. verruca, H. renebarbai, H. yuma, H. carlosviquezi, H. gabrielsantosi, H. solanllycarreroae, H. constanza; Ocrepeira klamt; Cybaeus daimonji;

Crustaceans

• 1 new anostracan: Phallocryptus fahimii;
• 1 new copepod: Rhinergasilus unguilongus;
• 3 new cumaceans: Cyclaspis goytacazes, Cyclaspis lilianae, Cyclaspis tesauna;
• 2 new amphipods: Carpentaria tropicalis; Pseudocrangonyx wonkimi;
• 5 new isopods: Baeticoniscus bullonorum; Fissarcturus argentinensis; Elthusa fistularia, Elthusa pseudorhombus, Elthusa uranoscopus;
• 16 new decapods: Arcotheres ocularius; Turleania rubriguttatus; Zoukaris festivus; Badistemon fulvum; Salmoneus rashedi; Caridina barakoma, C. choiseul, C. intermedia, C. maeana, C. nana, C. piokerai, C. pisuku, C. paratypus, C. poarae, C. sikipozo, C. turipi;

Holocerus devriesei is a new grasshopper from Madagascar. Credits to Skejo et al. (2020).*

Hexapods

• 4 new collembolans: Hymenaphorura urbana, H. kalindera; Hylaeanura emiliae; Hypogastrura herrerosvelai;
• 3 new archaeognathans: Machilinus petrophilus, M. obscurus, Charimachilis rostoviensis;
• 2 new odonates: Coeliccia tongbiguan, Coeliccia yunnanensis;
• 2 new dictyopterans: Microphotina panguanensis; Rugitermes tinto;
• 9 new orthopterans: Sinohaplotropis amurensis; Tagasta mizoramensis; Boroseiyla tupeba, Caauara pinima, Capiguara trimaculata, Hyalipenna tetralineata, Tuaia panacarica; Gigagryllus omayrae; Holocerus devriesei;
• 2 new plecopterans: Sinacroneuria huzhengkuni, S. lii;
• 4 new thysanopterans: Podothrips etsukoae; Parabaliothrips betulaceae, P. brevisetosus, P. robustus;
• 87 new hemipterans: Pachymetopius anamikus, Yaontogonia unipuncta; Usanus xochipalensis, U. igualaensis; Spaltumtettix coloradus, Pseudonapo waorani, P. huanucensis, Goiattus reyesi; Acrogymnia alagoas, A. blanki, A. brevis, A. chapada, A. clypea, A. deulufeuti, A. fascia, A. krylita, A. listoni, A. pulla, D. australis, D. fernandezi, D. taegeri; Macropsis aselae; Calyria hyperochelabrys, Carineta trinidadensis, Herrera grammosticta; Pachyjassus alatus, Pachyjassus basifurcatus, Pachyjassus ranomafanensis, Petalojassus ochrescens, Phaiojassus acutus, Phaiojassus bispinosus, Phaiojassus constrictus, Phaiojassus grandis, Phaiojassus spatulatus, Phaiojassus undulatus, Phaiojassus unispinosus, Pictojassus kirindiensis, Pictojassus productus, Pictojassus tulearensis, Platyjassella ancora, Platyjassella ohahelensis, Platyjassella attenuata, Platyjassella cormorana, Platyjassella emarginata, Platyjassella immaculata, Platyjassula cyclura, Platyjassula heterofurca, Platyjassula isofurca, Platyjassula mahajangensis, Platyjassus acutus, Platyjassus asymmetricus, Platyjassus fisheri, Platyjassus griswoldi, Platyjassus harinhalai, Platyjassus irwini, Platyjassus pedistylus, Platyjassus pennyi, Platyjassus pictipennis, Platyjassus symmetricus, Platyjassus vestigius, Plerujassus brunnescens, Plexijassus caliginosus, Pseudocurtara minima, Pseudocurtara nigripicta, Pseudocurtara quadrata, Pseudocyrta hyalina, Pseudomarganana olivacea, Pseudomarganana rosea, Pulchrijassus anjozorobensis, Pulchrijassus eunsunae, Pulchrijassus pallescens, Pulchrijassus roseus, Pulchrijassus rubrilineatus, Pulchrijassus sindhuae, Pulchrijassus talatakelyensis, Pulchrijassus toamasinensis, Punctijassus circularis, Punctijassus compressus, Punctijassus ivohibensis; Betacixius latissimus, B. maguanensis; Indolipa fugongensis, I. longlingensis; Trioza turouguei; Mitjaevia shibingensis, Mitjaevia dworakowskae; Vittaliana reticulata;
• 122 new coleopterans: Anthobium formosanum, A. lackneri, A. kabateki, A. marci, A. convexior, A. auritum, A. kabaki, A. semenovi, A. inopinatum, A. bengalicum; Batriscenellus suthepensis, B. vietnamicus; Cephalocyclus iohannisgabrielis; Nogunius sokani, N. aogashimanus, N. kerri, N. fukuuzanus; Carpophilus (Myothorax) brunneus; Laccophilus leguyaderi, L. bergsteni, L. makay, L. rakouthae; Beckhoplia bicolor, B. caliginosa, B. castanea, B. dolichiocnemis, B. elkeae, B. fusca, B. gifbergensis, B. nigra, B. nigrofasciata, B. nigrosetosa, B. pallidibrunnea, B. pulchra, B. pumilla, B. setosa, B. suturalis; Strongylium purcharti; Accolabass monteithi, Alpestriscyphon bartlefrere, Al. spurgeon, Anocyphon lepus, Copiacyphon brindaleensis, C. cardinalis, C. dytikos, Pumiliocara peneparva, Ruborcara saintae, Saltuscyphon montanus, Sal. teraniaensis, Saprocyphon bithongensis, Sisyracyphon brisbanensis, S. bulburinensis, Spilotocyphon occidentalis, Sp. orientalis, Sp. zwicki, Vadumcyphon centralis, V. rugosus; Bubastes barkeri, B. deserta, B. dichroa, B. flavocaerulea, B. hasenpuschi, B. iridiventris, B. iris, B. macmillani, B. magnifica, B. michaelpowelli, B. pilbarensis, B. remota, B. viridiaurea; Leptopeltus trogloxenus; Elacatophora jambiana, E. kerinciana, E. thailandiana, E. indica; Dichotomius valoisae; Tapinomorphus angolanus, T. kudrnai, T. latipennis, T. verunkae; Zarcosia barlayi, Zarcosia bedosae, Zarcosia brunnea, Zarcosia deharvengi, Zarcosia gerstmeieri, Zarcosia ilonae, Zarcosia intermedia, Zarcosia lemairei, Zarcosia schawalleri, Zarcosia sinuata, Zarcosia spinifemur Zarcosia srilankaensis, Zarcosia sumatrensis, Zarcosia weigeli; Australonotha monteithi; Xylodromus schweiki; Ancystrocerus lueliangi, A. philippinus; Blakealtica fusca; Exocelina athesphatos, E. tsinga; Ivalia antennata, I. caligulata, Cangshanaltica mindanaoensis, C. luzonica; Nephus apolonia; Agabus anguluverpus, Agabus austellus, Agabus riberae, Agabus agulhas; Metapogonia snizeki; Anchylorhynchus chrysomeloides, Anchylorhynchus goiano, Anchylorhynchus imitator, Anchylorhynchus latipes, Anchylorhynchus multisquamis, Anchylorhynchus rectus; Liodessus altoperuensis, Liodessus caxamarca; Apatrobus hekosanus; Ceratanisus diyarbakiricus; Dilychnia dumasi;
• 3 new neuropterans: Austrochrysa lihongyui, Austrochrysa angusta; Nusalala peruana;
• 68 new hymenopterans: Trieces confusus; Synergus aurofacies; Odontomutilla sairandhriensis, O. fletcheri; Aivalykus microaciculatus; Cestrus itatiensis; Alastor (A.) brevicornis, Alastor (A.) carens, Alastor (A.) gestroides, Alastor (A.) moody, Alastor (A.) namibiensis, Alastor (A.) parilis, Alastor (A.) sabrinae, Alastor (A.) temperatus, Alastor (Alastorellus) planipunctus; Aulacus davidi, A. shizukii; Gynecaptera albicrus, Macroocula madinaensis; Dicamptus curvus; Lymaenon funiculus, L. ledongus, L. longitus, L. naiquanlini, L. radiculus, L. zhui; Megischus sp., Foenatopus spp., Parastephanellus spp.; Microstenus rufithorax; Norbanus persicus, Stenomalina delvarei; Aleiodes andinus, A. angustus, A. asenjoi, A. bahiensis, A. barrosi, A. brevicarina, A. coariensis, A. goiasensis, A. gonodontivorus, A. hyalinus, A. inga, A. joaquimi, A. lidiae, A. mabelae, A. maculosus, A. ovatus, A. santarosensis, A. taurus; Periclistus orientalis; Pheidole antsahabe, Pheidole atsirakambiaty, Pheidole clara, Pheidole flammea, Pheidole flavodepressa, Pheidole mantadioflava, Pheidole maro, Pheidole ovalinoda, Pheidole similis, Pheidole tenebrovulgaris, Pheidole uranus, Pheidole voreios, Pheidole zirafy; Coccobius islandicus; Sphinctus pereponicus;
• 117 new dipterans: Sarcophaga (Hoa) membranijuxta; Tanap cinar; Winnertzia acutistylus, W. angustistylus, W. arctostylus, W. bicolor, W. brachytarsus, W. dentata, W. egregia, W. ekdalensis, W. fraxinophila, W. grytsjoenensis, W. hamatula, W. hemisphaerica, W. imbecilla, W. incisa, W. inornata, W. lapponica, W. lobata, W. longicoxa, W. normalis, W. oelandica, W. ombergensis, W. parvidens, W. pilosistylus, W. pratensis, W. pustulatula, W. quercinophila, W. rickebasta, W. ruliki, W. serri, W. setosa, W. silvestris, W. smalandensis, W. sundini, W. tumidoides, W. upplandensis, W. panguan, W. warraensis; Coniceromyia spp.; Condylostylus comorensis, C. friedmani, C. gavryushini, C. kaplini, C. madagascarensis; Cheilosia candida, Paragus thracus, Psilota aegeae; Macropelopia pergrandis; L. anaphlecter, L. apoecus, L. asilomar, L. bitumineus, L. canningsi, L. condylophorus, L. esau, L. karli, L. nelsoni, L. odontotus, L. sierra, L. tumulicola, L. wilcoxi; Dasythereva penai, D. patagonia; Tephritis kutuki; Epiphragma amamiense; Atelestus turcicus, Atelestus ibericus;  Massalongia nakamuratetsui; Colobaea deemingi; Oxysarcodexia alectoris, O. angulosa, O. ariozanoi, O. graminifolia, O. maiae, O. rimata; Schizomyia maesae;
• 55 new trichopterans: Cernotina longa; Agmina tuberosa, A. semiovale, A. rocheta, A. tenuisa, A. multidentata, A. cornuta, A. sagittata, A. circulata, A. digitata, A. longispina, A. magnahamata, A. longicordata, A. campanula, A. semicampanula, A. cunicula, A. cerritula, A. monstrosa, A. rectangulata, A. chela, A. piscaria, A. amplexa, A. caraffa,, A. rostrata, A. dathioensis, A. rougensis, A. viklundi, A. lata, A. falx, A. guttula, A. amieuensis, A. spina, A. complexa, A. dognyensis, A. mana, A. anterohamata, A. curvatacua, A. recurvata, A. taoensis, A. triangulata, A. bleuensis, A. touhoensis, A. wardi, A. parallela, A. christinae, A. brevis, A. ninguana, A. scopula; Ecnomus petchanaae, E. boonsawaengae, Polyplectropus proukaewi, Psychomyia srichanai; Machairocentron chorotegae, M. eugeniarguedasae, M. kalinae;
• 84 new lepidopterans: Dichomeris fascialobella, D. flavimaculata, D. infundibularis, D. lancea, D. longivalvata, D. marginalis, D. myriaodontosa, D. ochriella, D. quadratella, D. sclerospinula, D. spinosella, D. unilobata, Helcystogramma albimacularis; Abantiades cephalocorvus, Abantiades tembyi; Ypsolopha admirandella; Yelenka calciferella; Panca puri, Panca xavante; Lophocnema mackeyi, Oligophlebia insurgia, Micrecia capillaria, Micrecia kuukuyau, Micrecia hawkei, Lamellisphecia sanguinea, Specodoptera hiltoni, Aegerosphecia rufea, Melittia flanneryi, Melittia pyroptella, Archaeosesia milleri, Nokona kungkay, Nokona bipora, Pseudosesia muyuae, Pseudosesia sumoensis, Pseudosesia sepikensis, Paranthrenella terminalia, Paranthrenella brandti, Paranthrenella lelatensis; Epicrocis datianensis, E. maculata, E. flabelloformis; Leucinodella banthaensis; Exoasota pursatensis; Metallolophia taleensis; Meleonoma anisodonta, M. catenata, M. dilativalva, M. dorsibacilliformis, M. lanceolata, M. longaedeaga, M. longiprocessa, M. lunulata, M. papillata, M. parvissima, M. rugulosa, M. tenuiuncata, M. tianmushana, M. triacantha, M. ventrospinosa; Stathmopoda atrifusca, S. cellifaria, S. cornuta, S. digitiprocessa, S. dolichantha, S. falsistimulata, S. guangxiensis, S. ingena, S. liberata, S. purpurata, S. serrifasciaria, S. similatridorsalis, S. spinicornuta; Eidophasia assmanni; Epiparbattia multispinalis; Sacada dzonguensis, S. umtasorensis; Megacraspedus cottiensis; Nagiella bispina; Catoptria samnitica; Nokona vietnamica, Nokona cucphuongae; Phragmataecia effendii; Pseudodebis nakamurai, P. pieti;

Trioza turouguei is a new jumping plant louse from Taiwan. Credits to Tung et al. (2020).*

Echinoderms

• 4 new ophiuroids: Astrochlamys timoharai, Ophiosteira lissopatella, Ophiophyllum umbonatum, Soliophis bakeri;
• 1 new holothuroid: Elpidia soyoae;

Agnathans

• 1 new myxinid: Eptatretus albiderma;

Actinopterygians

• 1 new anabantiform: Parosphromenus barbarae;
• 1 new anguilliform: Xyrias anjaalai;
• 2 new characiforms: Bryconops sp.; Priocharax varii;
• 2 new cypriniforms: Paracobitis salihae; Rasbora adisi;
• 1 new gobiiform: Corcyrogobius pulcher;
• 3 new perciforms: Menticirrhus sp.; Upeneus floros; Etheostoma faulkneri;
• 1 new siluriform: Aspidoras azaghal;
• 1 new tetraodontiform: Canthigaster aziz;

Pristimantis chamezensis is a new frog from Colombia. Credits to Acosta-Galvis et al. (2020).*

Amphibians

• 1 new caudate: Bolitoglossa awajun;
• 12 new anurans: Platymantis sp.; Leptobrachella neangi; Pristimantis sp.; Pristimantis chamezensis; Adenomera spp.; Xenorhina salawati, X. waigeo;

Acanthosauria liui is a new agamid lizard from China. Credits to Liu et al. (2020).*

Reptiles

• 10 new squamates: Hemiphyllodactylus nahangensis; Cyrtodactylus houaphanensis, Cyrtodactylus ngoiensis; Cyrtodactylus sp.; Cyrtodactylus amphipetraeus; Cnemaspis rishivalleyensis, Hemidactylus rishivalleyensis; Atheris hetfieldi; Acanthosaura liui; Dipsas bothropoides;

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* This work is licensed under a Creative Commons Attribution 4.0 International License.

Friday Fellow: Common Pellia

by Piter Kehoma Boll

Liveworts often live in moist and shady spaces and, even if we know how to identify them as liverworts, they often look all the same. However, if we pay attention to the details, differences can often be perceived.

Pellia epiphylla, commonly known as the common pellia, is a liverwort that loves very humid places, so it often grows very close to rivers and other watercourses in North America, Europe, North Africa and some nearby areas in Asia. Its thalli are smooth and slightly fleshy, about 1 cm wide and can reach several cm in length. They like ro remain in a horizontal position, so they grow very attached to the horizontal subtrates but tend to grow away in vertical ones, acquiring a more ruffled aspect. Although usually completely green, the thalli can have a purplish or reddish tinge along the middle, especially when they grow too far from water, which can help identify this species. Otherwise it is very featureless compare to many other liverworts.

The typical aspect of the common pellia. Some thalli can be seen with a purplish tinge in the middle. Photo by Frank Vincentz.**

As with all liverworts, the thallus of the common pellia is the gametophyte, i.e., the haploid generation (with only one chromosome of each type per nucleus) and that generates the gametes. Although in many liverworts the gametophytes are either male or female, they are monoicous (i.e, hermaphrodites) in the common pelia. The male sex organs (antheridia) occur along the middle, appearing as very small light and shiny dots, while the female ones (archegonia) occur close to the tip and remain covered. Fertilization, as usually, occurs when the plant becomes wet. The antheridia absorb water to the point that they burst, releasing the sperm cells (antherozoids) that swim to the archegonia, where fertilization occurs.

Young sporophytes growing from inside the archaegonia. Photo by Hermann Schachner.

The resulting zygote gives rise to the sporophyte, a diploid generation (with two chromosomes of each type per nucleus) and it grows from inside the archegonia in the form of a very long and slender whitish stalk with a dark capsule at the tip. When the capsule is mature, it bursts and releases the spores, which will germinate and originate new gametophytes. The group of sporophytes growing from the gametophyte give the set a peculiar “hairy” aspect, which also helps recognize this species.

When the sporophytes grow, they give the family a hairy look. Photo by Roger Griffith.

Being a common species across its range, the common pellia has been studied to understand physiological and reproductive characteristics of liverworts, as well as some ecological aspects. For example, it is known that, while the gametophyte absorbs water mostly through the under surface, the antheridia absorb it from the upper surface, and the lower midrib of the plant compared to the border is essential to retain water for this. While the sporophyte of many liverworts is completely dependent on its mother, the gametophyte, to receive water, that of the common pellia is much more indepenent, absorbing most of it from the environment.

Although fairly featureless, the common pellia still has its charm.

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References:

Clee A (1939) The Morphology and Anatomy of Pellia epiphylla considered in Relation to the Mechanism of Absorption and Conduction of Water. Annals of Botany 3(1): 105–111. https://doi.org/10.1093/oxfordjournals.aob.a085045

Greenwoo HE (1911) Some Stages in the Development of Pellia epiphylla. The Bryologist 14(4): 59-70. https://doi.org/10.2307/3238074

Wikipedia. Pellia epiphylla. Available at <https://en.wikipedia.org/wiki/Pellia_epiphylla >. Access on 27 August 2020.

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** This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.

Friday Fellow: Rat Acanthocephalan

by Piter Kehoma Boll

The evolution of similar traits in distantly related species is common when they have similar lifestyles, and this is particularly noticeable in some parasitic groups. Among flatworms, tapeworms have developed a complex life cycle with two hosts, larval stages and adults that live in the intestine of the definitive hosts where they absorb nutrients directly through the body surface, lacking a digestive system, and have a special structure on their head to remain attached to the host’s guts.

A very similar lifestyle and morphology evolved in a distantly related group of animals, the acanthocephalans or thorny-headed worms. For a long time, the acanthocephalans were considered a phylum of their own, Acanthocephala, but we now know that they are just a group of very specialized rotiferans, whose free-living forms are very small, so small that they are often mistaken for ciliates or other unicelular organisms.

Today’s species is an acanthocephalan that lives very close to humans, so close that they can even live inside us. Named Moniliformis moniliformis, I will call it the “rat acanthocephalan” because, well, it infects rats (and occasionally other mammals including humans).

Adults of the rat acanthocephalan are often found in the intestine of rats but other mammals can also be infected, such as dogs, cats and humans. Males reach up to 5 cm in length while females can be much longer, up to 30 cm.

Adult, probably female, specimen of the rat acanthocephalan.

The anterior end of the body has a short cylindrical proboscis covered by hooks, which the animal uses to attach to the host’s intestine. This proboscis is hollow and can be pulled back into the body. There is a septum separating the cavity of the proboscis from the cavity of the rest of the body. Like in tapeworms, the surface of acanthocephalans is covered by a syncytium, a tissue formed by cells that fused together into a single multinucleated structure. Due to the lack of a digestive system, they absorb the nutrients from the hosts intestine directly through their body surface, just like in tapeworms.

Anterior end showing the short proboscis.

After mating occurs, females release fertilized eggs into the host’s intestine and they leave the body with its feces. The eggs measure about 100 µm in length and 60 µm in width and contain the first larval stage, known as the acanthor. In the environment, the eggs are ingested by the intermediate host, usually a cockroach or sometimes a beetle, and the acanthor hatches, changing into the second-stage larva, the acanthella. After some weeks developing inside the intermediate host, the acanthella changes into the final larval stage, the cystacanth, which forms a cyst inside the intermediate host’s tissues, and there it waits.

An egg under the microscope.

For the cycle to be completed, the intermediate host needs to be eaten by the definitive host. To increase the chances of this happening, the parasite leads to behavioral changes in the intermediate hosts. Infected American cockroaches, for example, show delayed escape responses, increasing the probablity of being captured by a predator. When it happens, the cystacanths are released into the definitive host’s gut and develop into adults.

Life cycle of the rat acanthocephalan.

Humans acting as definitive hosts is a rare occurrence since it requires the ingestion of raw infected cockroaches or beetles. Most reported cases in the literature include small children, which are prone to put everything into their mouths, and the symptoms of the infection include acute abdominal pain and, in very small children, usually less than a year old, more severe symptons such as vomiting, anorexia and diarrhea can also occur. The identification of eggs in stool samples of infected humans is difficult, though, so that the actual infection rate may be much higher than thought, especially in rural areas where insect consumption is a common practice.

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References:

Moore J (1983) Altered Behavior in Cockroaches (Periplaneta americana) Infected with an Archiacanthocephalan, Moniliformis moniliformis. Parasitology 69(6):1174–1176. https://doi.org/10.2307/3280893

Salehabadi A, Mowlavi C, Sadjjadi SM (2088) Human Infection with Moniliformis moniliformis (Bremser 1811) (Travassos 1915) in Iran: Another Case Report After Three Decades. Vector-Borne and Zoonotic Diseases 8(1):101–104. http://doi.org/10.1089/vbz.2007.0150

Wikipedia. Moniliformis moniliformis. Available at < https://en.wikipedia.org/wiki/Moniliformis_moniliformis>. Access on 20 August 2020.

Friday Fellow: Dragon Tree

by Piter Kehoma Boll

When the seventh generation of pokémon was released, it introduced regional forms of previous pokémon, including an Alolan form of Exeggutor which was changed from the grass/psychic type of the traditional Exeggutor to a grass/dragon type. This led many people to become familiar with the genus Dracaena, a genus that is well-known among botanists and gardeners and includes many ornamental plants.

Alolan Exeggutor, a grass/dragon pokémon.

The name Dracaena comes from the greek word meaning dragoness, i.e., a female dragon and is given based on the type-species of the genus, Dracaena draco, or the dragon tree, which is today’s fellow.

Dragon tree in Tenerife, Canary Islands. Photo by Wikimedia user Losrealejos.es*

The genus Dracaena is closely related to the genus Asparagus and the dragon tree was intially named Asparagus draco by Linnaeus and later renamed Dracaena draco by himself based on a genus name created by the Italian naturalist Domenico Agostino Vandelli. This species is native from the African islands in the Atlantic (Canary Islands, Cape Verde and Madeira).

Closeup of a flower. Photo by Wikimedia user Philmarin.**

The dragon tree starts its life as a small unbranched stem like most ordinary species of Dracaena we see in gardens. Its growth is very slow and only after growing vertically for 10 to 15 years it will produce flowers for the first time. The flowers are white and lily-like and appear in a spike, later turning into reddish berries. After this first reprouctive cycle, the stem branches for the first time from a crown of terminal buds and then grows again for 10 to 15 years before branching again. Being a monocot, the dragon tree lacks growth rings but its age can be estimated by the number of branching points from the ground to the crown.

The fruits. Photo by Forest & Kim Starr.***

The association of this plant with dragons comes from ancient times. Not only Dracaena draco, but some other species of Dracaena as well, produce a red resin that is secreted when the leaves or the trunk are cut. A similar red resin is found in many other plants, including palm trees and crotons, and they were all collectively known as “dragon’s blood” and used for several purposes, such as dye or medicine. The ancient Romans collected dragon’s blood from the Island of Socotra, where a closely-related species, Dracaena cinnabari, the dragon’s blood tree, is found.

Plucked dead leaves showing the red color of the dragon’s blood. Photo by Wikimedia user Sharktopus.*

The dragon tree is the official tree of Tenerife, where the largest and possibly oldest specimen is also found, the so-called “Drago Milenario”. This specimen is about 21 m tall but, despite its name (the thousand-year-old dragon), it is not actually that old and its age is most likely about 300 years or so.

The Drago milenario in Tenerife, the largest dragon tree in the world. Photo by Andrey Tenerife.**

Despite being a relatively popular species that is grown as an ornamental plant, the dragon tree is classified as vulnerable in the IUCN’s red list. It’s wild populations are close to extinction and one reason for this is likely because some of its original seed dispersers went extinct. Only two bird species have been recently recognized as effective dispersers. Due to the dragon’s tree relatively large fruit, most bird species do not eat the whole fruit and only bite off pieces of the pulp, so that seeds are not carried to new locations.

Ripe fruits. Photo by Wikimedia user Nadiatalent.*

The Guanches, the aboriginal people of the Canary Islands, used to worship a large dragon tree in Tenerife. Alexander von Humboldt apparently saw this tree when visiting the island and it was later destroyed by a storm that hit Tenerife in 1868. The Guanches were wiped out by the Spanish invaders and now their sacred tree is facing the same fate.

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References:

Bañares A et al. (1998) Dracaena draco. The IUCN Red List of Threatened Species 1998: e.T30394A9535771. https://dx.doi.org/10.2305/IUCN.UK.1998.RLTS.T30394A9535771.en. Access on 13 August 2020.

González-Castro A, Pérez-Pérez D, Romero J, Nogales M (2019) Unraveling the Seed Dispersal System of an Insular “Ghost” Dragon Tree (Dracaena draco) in the Wild. Frontiers in Ecology and Evolution 7:39. https://doi.org/10.3389/fevo.2019.00039

Wikipedia. Dracaena draco. Available at < https://en.wikipedia.org/wiki/Dracaena_draco >. Access on 13 August 2020.

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* This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.

** This work is licensed under a Creative Commons Attribution-ShareAlike 4.0 International License.

*** This work is licensed under a Creative Commons Attribution 3.0 Unported License.

Friday Fellow: Lab Dung Fungus

by Piter Kehoma Boll

In my microbiology classes as an undergraduate student, I remember seeing only two genera of ascomycetes under the microscope: Aspergillus and Penicillium. However, there is another genus that is commonly used in biology classes, Sordaria, the dung fungi, whose most popular species is Sordaria fimicola, which I decided to call the lab dung fungus.

As its name suggest, the lab dung fungus is found growing on dung, more specifically on dung of herbivorous mammals. For a long time, it was thought that this species required dung to complete its life cycle. After growing on dung, the lab dung fungus releases its spores in the environment, they adhere to the surface of plants and are ingested by grazing mammals, restarting the cycle. However, it is now known that this species can grow and reproduce on plant matter without requiring dung, although more studies are needed to understand how the presence or absence of dung affects its fitness.

Fruiting bodies (perithecia) of Sordaria fimicola growing on dead leaf tissue of the grass Bromus tectorum. Credits to Newcombe et al. (2016).*

During most of its life, the lab dung fungus exists, just like other fungi, solely as a network of hyphae, the mycelium, growing inside the medium on which it feeds, in this case decaying plant matter, especially in dung. These hyphae are haploid (n), meaning that they have only one copy of each chromosome. When two hyphae touch, they can fuse and create a cell with two nuclei, the dykarion, each nucleus coming from one of the original hyphae. The dikaryotic cells divide through mitosis without fusing their nuclei, originating a set of dikaryotic hyphae that form a fruiting body, the perithecium, that grows inside the mycelium of haploid hyphae.

A bursted perithecium with released asci. Photo by Carmelita Levin.**

The perithecium is kind of pear-shaped and, inside of it, some dikaryotic cells allow their nuclei to fuse into a single, diploid nucleus, which now has two chromosomes of each type (2n), one from each parent hypha back then when the haploid hyphae meet. This newly formed diploid cell is a zygote but instead of growing into diploid hyphae by mitosis, it immediately undergoes meiosis to originate once again a set of haploid nuclei. The four resulting nuclei from meiosis each one undergoes mitosis, resulting in eight final nuclei, which remain lined up in the elongated cell. The cell then divides into eight individuals cells, each with one of the nucleus, and they turn into spores, ascospores, and remain inside an elongated sac, the ascus. When the ascospores are mature, they are released in the environment and can germinate to create a new set of haploid hyphae.

Lineages of the lab dung fungus found in nature often have very dark ascospores and this is called the wild type. However, one laboratory lineage has lighter, often gray ascospores, and is called the tan type. The color of the spore is determined by a single gene in one of the chromosomes. Thus, if you cross the wild and the tan types, the ascus of the hybrid will have four dark and four light spores, and this is how the lab dung fungus becomes a good species to understand meiosis and chromosome crossover in biology classes.

Lab dung fungus in the lab, growing in a Petri dish. Photo by Wikimedia user Ninjatacoshell.***

Before meiosis occurs, all chromosomes in a cell are duplicated, resulting in cell with four chromosomes of each type (4n) of which two are from one parent and two are from the other. When the nucleus divides for the first time, each daughter cell will be a special case of 2n, in which the two copies of each chromosome are originally from the same parent. In the lab dung fungus, considering the chromosome with the color gene, this would create a pattern like (AA)(AA) in these two nuclei. After the second division of meiosis, the pattern becomes (A)(A)(A)(A) and, after the mitosis that leads to the eight final spores, (A)(A)(A)(A)(A)(A)(A)(A).

Asci of a hybrid showing several combinations of dark and light ascospores. Extracted from https://www.fishersci.com/.

However, if crossover occurs, one pair of chromosomes from different parents exchange pieces with each other, while the other pair remains unaffected. As a result, they exchange the gene responsible for the color and the final product, instead of being 4 of one color followed by 4 of another (4:4 patterns), shows a 2:4:2 or a 2:2:2:2 pattern.

Resulting arrangement of the ascospores in the asci when chromosome crossover occurs (below) or not (above). Extracted and adapted from http://facweb.furman.edu/.

Sometimes other weird patterns appear as well, such as 2:1:1:1:1:2 patterns, but I guess this happens because of some mechanical action where one spore can roll over another and end up outside of its original position inside the ascus, perhaps caused when they are squeezed out of the perithecium. Really stranged patterns are those in which there are not 4 spores of each color, which include very rare instances of 6:2 or 5:3 patterns, and those are explained as the result of errorSs during chromosome replication.

Unusual 2:1:1:1:1:2 pattern probably caused because the two central ascospores were swapped because of pressure applied to the asci, so that the original pattern was 2:2:2:2 as expected when crossover occurs. Photo by Wikimedia user Ninjatacoshell.***

Isn’t the lab dung fungus indeed a very cool model to use in classes? I’m sad that I haven’t had the opportunity to see this in my genetics or microbiology classes. If you studied biology, were you lucky enough to see this? Let us know!

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References:

Dotson J (2019) Life cycle of Sordaria Fimicola. Sciencing. Available at < https://sciencing.com/life-cycle-sordaria-fimicola-6909851.html >. Acccess on 6 August 2020.

Kitani Y, Olive LS, E-Ani AS (1961) Transreplication and Crossing Over in Sordaria fimicola. Science 134: 668-669. https://doi.org/10.1126/science.134.3480.668

Newcombe G, Campbell J, Griffith D, Baynes M, Launchbaugh K, Pendleton R (2016) Revisiting the Life Cycle of Dung Fungi, Including Sordaria fimicola. PLoS ONE 11(2): e0147425. https://doi.org/10.1371/journal.pone.0147425

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* This work is licensed under a Creative Commons Attribution 4.0 International License.

** This work is licensed under a Creative Commons Attribution-ShareAlike 4.0 International License.

*** This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.