by Piter Kehoma Boll

More a politician than a naturalist, today we remember a British explorer that was central in the mess that Europe turned Africa into, but also a important in recording Africa’s culture and biodiversity.

Henry Hamilton Johnston, more commonly known as Harry Johnston, was born on 12 June 1858 in London, the son of John Brookes Johnstone and Esther Laetitia Hamilton. He studied at Stockwell grammar school and later at the King’s College London, after which he studied painting at the Royal Academy for four years. During his studies, he traveled through Europe and visited the interior of Tunisia.

In 1882, aged 24, he traveled to southern Angola with the Earl of Mayo (which I guess was Dermot Bourke at that time). Traveling north from there, he met the Welsh explorer Henry Morton Stanley in the Congo River the following year. There, he became one of the first Europeans to see the Congo River above the Stanley Pool (currently known as Pool Malebo), a widening of the river near the cities of Kinshasa and Brazzaville. He published a book in 1884 called “The River Congo: From its Mouth to Bolobo” and, in that same year, was appointed leader of a scientific expedition to Mount Kilimanjaro, in current Tanzania. In this expedition, he managed to conclude treaties with local chiefs. The reports of this expedition were published in his 1886’s book “The Kilema-Njaro Expedition”.

In 1886, the British government appointed Johnston the vice-consul in Cameroon and the Niger River delta area. The British had claimed the area but the local leader, Jaja of Opobo, refused to give up the territory. Invited by Johnston to negotiate, Jaja was arrested and deported to London. During the following years, Johnston took part in several expeditions and diplomatic missions that helped the British Empire to dominate more and more of Africa’s territory.

In 1896, Johnston married Winifred Mary Irby, daughter of the fifth Baron Boston. That same year, afflicted by tropical fevers, he was transferred to Tunis as consul-general in order to recover. In 1899, he was sent to Uganda as special commisioner to end an ongoing war. There, he found out that a showman was abducting Pygmy inhabitants of the Congo for exhibition. Johnston helped to rescue them and the pygmies mentioned to him a creature, some sort of “unicorn donkey” previously referred to by Stanley. There were some reports about explorers seeing an animal with a zebra-like pattern moving through the bushes and the expectation was that it was some sort of forest-dwelling horse. The pygmies showed tracks of the creature to Johnston and he was surprised to find out that it was actually a cloven-hoofed beast and not a single-hoofed animal as a horse. Johnston never saw the animal, but managed to obtain pieces of the striped skin and a skull, which led the creature to be classified as Equus johnstoni in 1901. The inclusion in the genus Equus was mostly motivated by the pygmies referring to the creature as a kind of horse. Analyses of its skull, however, soon concluded that it was a relative of the giraffe. This animal is currently know as the okapi, or Okapia johnstoni.

In 1902, when Johnston was back to London, his wife gave birth to twin sons, but both died few hours later. They did not have any other children. That same year Johnston was appointed member of the Zoological Society of London. In the following years, he spent most of his time at home writing novels and accounts of his voyages through Africa. In 1925, he had two strokes that partially paralyzed him. He died two years later, on 31 July 1917, aged 69.

Johnston, as all imperialists of his time, believed that Europeans, and British especially, had superiority over Africans. Nevertheless, he was against using violence against the subjugated peoples and had a more paternalistic view. Although such views are seen as horrible today (or at least they should to any reasonable human being), he was considered some sort of radical for his time, as others had a much worse vision of the African cultures.

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References:

Wikipedia. Harry Johnston. Available at < https://en.wikipedia.org/wiki/Harry_Johnston >. Access on 11 June 2019.

Wikipedia. Okapi. Available at < https://en.wikipedia.org/wiki/Okapi >. Access on 11 June 2019.

by Piter Kehoma Boll

With today’s post, I intend to start a series of three Friday Fellows that are connected. After all, that’s what life is, right? Organisms interacting.

So to start, let’s talk about a magnificent tree today, the fig tree Ficus microcarpa, commonly known as Chinese banyan, Malayan banyan, Indian laurel, curtain fig, gajumaru and many other names. Its native range goes from China to Australia, including all the southeastern Asia and several Pacific islands in the way. However, it can be found in many other countries today as it has become a somewhat popular ornamental plant.

In its natural tropical habitat, the Chinese banyan reach a height of 30 meter or more, with a crown spreading across more than 70 meters and a trunk more than 8 m in thickness. Most trees are smaller, though, and they never reach such an astonishing size in temperate climates. Its bark has a light gray color and its leaves are smooth, entire, oblanceolate, and about 5 to 6 cm long. Its figs are considerably small, hence the name microcarpa (small-fruited). It is common for large specimens to produce aerial roots, which grow from the branches and touch the soil, forming an intricate and beautiful system.

As typical among fig trees, the Chinese banyan is pollinated by a fig wasp, in this case the species Eupristina verticillata. Outside of its native range, the tree can only produce viable seeds in the presence of the wasp, so the insect must be introduced along with it. Its fruits are very attractive to birds, who spread its seeds in their feces. After passing through a bird’s gut and reaching the outer environment again, the seeds attract ants, which spread them even further. Being quite versatile regarding the substrate to germinate, the Chinese banyan can grow on a lot of surfaces, often sprouting through crevices on walls and sidewalks and breaking them as it grows.

The Chinese banyan is used in traditional Chinese medicine to treat a variety of conditions, including pain, fever, flu, malaria, bronchitis and rheumatism. Laboratory studies have isolated anti-cancer, antioxidant and antibacterial compounds from the bark, leaves, aerial roots and fruits, as well as anti-fungal compounds from its latex. The tree has, therefore, the potential to be used for the development of many medicines.

Due to its impressive size and the intricate labyrinth formed by its network of aerial roots, the Chinese banyan tree has an important role to many religious groups in its native range, being often considered the house of spirits, either good or bad ones and its presence usually marks places of worship. Regardless of these beliefs, though, this magnificent tree deserves the admiration that it gets.

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References:

Ao C, Li A, Elzaawely AA, Xuan TD, Tawata S (2008) Evaluation of antioxidant and antibacterial activities of Ficus microcarpa L. fil. extract. Food Control 19(10): 940–948. doi: 10.1016/j.foodcont.2007.09.007

Chiang YM, Chang JY, Kuo CC, Chang CY, Kuo YH (2005) Cytotoxic triterpenes from the aerial roots of Ficus microcarpa. Phytochemistry 66(4): 495–501. doi: 10.1016/j.phytochem.2004.12.026

Kaufmann S, McKey DB, Hossaert-McKey M, Horvitz CC (1991) Adaptations for a two-phase seed dispersal system involving vertebrates and ants in a hemiepiphytic fig (Ficus microcarpa: Moraceae). American Journal of Botany 78(7): 971–977. doi: 10.1002/j.1537-2197.1991.tb14501.

Taira T, Ohdomari A, Nakama N, Shimoji M, Ishihara M (2005) Characterization and Antifungal Activity of Gazyumaru (Ficus microcarpa) Latex Chitinases: Both the Chitin-Binding and the Antifungal Activities of Class I Chitinase Are Reinforced with Increasing Ionic Strength. Bioscience, Biotechnology and Biochemistry 69(4): 811–819. doi: 10.1271/bbb.69.811

Wikipedia. Ficus microcarpa. Available at < https://en.wikipedia.org/wiki/Ficus_microcarpa >. Access on June 8, 2019.

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

by Piter Kehoma Boll

Last week I presented the magnificent Chinese banyan Ficus microcarpa. Today I’m bringing a little insect that loves it but is not loved in return, the Cuban laurel thrips, Gynaikothrips ficorum.

As its name suggests, the Cuban laurel thrips is a thrips, i.e., an insect of the order Thysanoptera. Adults of this species measure about 3 mm in length and have a black and elongate body and two pairs of thin wings that fold over the dorsum when at rest. Its mouth parts, as typical of thrips, are asymmetrical, with a reduced right mandible and a developed left mandible that it uses to cut the surface of plants in order to suck its juices. It is, therefore, a plant pest.

The Cuban laurel thrips prefers to feed on juices of fig trees, such as the Chinese banyan from last week. It’s common name, though, is a reference to another fig species, Ficus retusa, commonly known as the Cuban laurel. Both fig trees, as well as the thrips itself, are native from Southeast Asia. Other, less common host plants include Citrus trees and orchids. They prefer to feed on young, tender leaves, and cause dark, usually purplish red dots, on the leaf’s surface. It is common for the leaf to curl and become hard, eventually dying prematurely. Although most infestations do not cause serious damage to the plant’s development, the curling of the leaves can reduce a plant’s ornamental value.

The reproduction of the Cuban laurel thrips is basically constant, so that several generations occur across one year. The adults take advantage of the curled leaves produced by their feeding behavior and use them as a protection to put their eggs. The immature stages, after hatching, remain inside the shelter provided by the curled leaf. They are transparent in the first two instars and then become light yellow. Only the last, adult stage, is black.

Since the Cuban laurel thrips makes ornamental plants ugly, humans are always trying to find ways to kill them, especially by using pesticides or, sometimes, natural predators of the thrips. But the little insect can also fight back. When they accidentally fall on people’s bodies, they tend to bite, most likely by accident, but this can end up causing a serious and annoying itch. That’s the price for messing with them.

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References:

Denmark HA, Fasulo TR, Funderburk JE (2005) Cuban laurel thrips, Gynaikothrips ficorum (Marchal) (Insecta: Thysanoptera: Phlaeothripidae). DPI Entomology Circular 59

Paine TD (1992) Cuban Laurel Thrips (Thysanoptera: Phlaeothripidae) Biology in Southern California: Seasonal Abundance, Temperature Dependent Development, Leaf Suitability, and Predation. Annals of the Entomological Society of America 85(2): 164–172. doi: 10.1093/aesa/85.2.164

Piu G, Ceccio S, Garau MG, Melis S, Palomba A, Pautasso M, Pittau F, Ballero M (1992) Itchy dermatitis from Gynaikothrips ficorum March in a family group. Allergy 47(4): 441–442. doi: 10.1111/j.1398-9995.1992.tb02087.x

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

by Piter Kehoma Boll

When we think of animals changing colors to adapt to the background, we readily think of chameleons, or maybe of some extremely rapid color switchers such as cephalopods like octopuses and cuttlefish. However, many other animals have this ability too.

One example are tree frogs of the family Rhacophoridae, especially of the genus Rhacophorus.

Recently, the phenomenon was recorded for the first time for the species Rhacophorus smaragdinus in northeastern India. The animal was of a vivid green color when found but, as soon as the researchers handled it, it turned into a dull brown color in a matter of seconds, only to slowly go back to green after left alone.

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Reference:

CK D, Payra A, Tripathy B, Chandra K (2019) Observation on rapid physiological color change in Giant tree frog Rhacophorus smaragdinus (Blyth, 1852) from Namdapha Tiger Reserve, Arunachal Pradesh, India. Herpetozoa 32: 95–99. doi: 10.3897/herpetozoa.32.e36023

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

by Piter Kehoma Boll

Last week I introduced the Cuban Laurel Thrips, which feeds on several fig trees, such as the Chinese Banyan and the Cuban Laurel. Today, we will continue up the food chain and talk about a mite that is a parasite of the cuban laurel thrips. Named Adactylidium gynaikothripsi, I decided to give it the common name “Cuban-Laurel-Thrips Mite”.

The Cuban-laurel-thrips mite was described only in 2011 from Cuban laurel thrips populations in Greece. This is the fourth mite of the genus Adactylidium known to parasitize the Cuban laurel thrips, the other four being Adactylium ficorum (“fig-thrips mite”), A. brasiliensis (“Brazilian thrips mite”) and A. fletchmani (“Fletchman’s thrips mite”). As you can imagine, in order to parasitize an insect as small as the Cuban laurel thrips, these mites are even smaller, measuring about 0.1 mm in length.

The life cycle of the Cuban-laurel-thrips mite, which is basically the same for all species of Adactylidium, is very bizarre. Adult females feed on the eggs of the Cuban Laurel Thrips. They start their adult life wandering over fig leaves looking for a suitable thrips egg to attack. Once finding one, they pierce the egg’s shell with their chelicerae and attach to it like ticks and start to eat. They feed on a single egg across their entire life. If they are unable to find an egg, they may also attach to an adult thrips as a last resource, or else they die of starvation in a few hours.

Once a female starts to eat, a small group of eggs, usually between 5 to 10, begins to develop inside her. The eggs grow during the first 48 hours after the female attached to the egg, making her double in size and becoming something like a spherical egg sac. The eggs hatch around this time and the mite larvae remain inside their mother. These larvae lack mouth parts, so it is believed that they absorb nutrients from her mother directly through the body surface. About 24 hours later, the larvae turn into nymphs, which remain inactive inside the shed skin of the larva. They also lack any mouth parts.

Another 24 hours pass and the nymphs turn into adult mites. They are still inside their mother when this happens. The adults consist always of a single male and several females. This male then starts to copulate with his own sisters, still inside their mother’s abdomen, and, when copulation is finished, they start to tear their mother’s body apart to get free, killing her in the process. Once outside the body, the male dies in a few minutes, never eating anything other than his own mother. The females, on the other hand, start to look for thrips eggs on which to feed, only to be killed by her own children less than 4 days laters.

This entire life cycle may look very insane from our human perspective, but nature was never interested in following our moral rules.

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References:

Antonatos SA, Kapaxidi EV, Papadoulis, GT (2011) Adactylidium gynaikothripsi n. sp. (Acari: Acarophenacidae) associated with Gynaikothrips ficorum (Marshal) (Thysanoptera: Phlaeothripidae) from Greece. International Journal of Acarology, 37(sup1), 18–26. doi: 10.1080/01647954.2010.531763

Elbadry, EA, Tawfik, MSF (1966) Life Cycle of the Mite Adactylidium sp. (Acarina: Pyemotidae), a Predator of Thrips Eggs in the United Arab Republic. Annals of the Entomological Society of America, 59(3), 458–461. doi: 10.1093/aesa/59.3.458

by Piter Kehoma Boll

Here is a list of species described this month. It certainly does not include all described species. Most information comes from the journals Mycokeys, Phytokeys, Zookeys, Phytotaxa, Zootaxa, Mycological Progress, Journal of Eukaryotic Microbiology, International Journal of Systematic and Evolutionary Microbiology, Systematic and Applied Microbiology, Zoological Journal of the Linnean Society, PeerJ, Journal of Natural History and PLoS One, as well as several journals restricted to certain taxa.

Bacteria

• 13 new actinobacteria: Desertihabitans aurantiacus; Bifidobacterium jacchi; Thermomonospora catenispora; Streptomyces rhizosphaericola; Streptomyces altiplanensis; Streptomyces cyaneochromogenes; Streptomyces paromomycinus; Streptomyces dangxiongensis; Glaciihabitans arcticus; Microbacterium bovistercoris; Kribbella turkmenica; Cellulomonas algicola; Amycolatopsis suaedae; Propionibacterium ruminifibrarum; Cryobacterium tepidiphilum; Cellulomonas aurantiaca;
• 10 new bacteroids: Flavobacterium stagni; Flavobacterium silvisoli; Pedobacter chinensis; Sandaracinomonas limnophila; Hymenobacter humicola; Hymenobacter oligotrophus; Ancylomarina salipaludis; Empedobacter tilapiae; Flavivirga rizhaonensis; Flavisolibacter galbus;
• 1 new chlamydia: Chlamydia buteonis;
• 2 new cyanobacteria: Myxacorys chilensis, M. californica;
• 19 new firmicutes: Lactobacillus jixianensis, Lactobacillus baoqingensis, Lactobacillus jiayinensis, Lactobacillus zhaoyuanensis, Lactobacillus lindianensis, Lactobacillus huananensis, Lactobacillus tangyuanensis, Lactobacillus fuyuanensis, Lactobacillus tongjiangensis, Lactobacillus fujinensis, Lactobacillus mulengensis; Ornithinibacillus gellani; Clostridium prolinivorans; Lactobacillus suantsaicola, Lactobacillus suantsaiihabitans; Lactobacillus huachuanensis; Enterococcus florum; Chengkuizengella marina; Staphylococcus pseudoxylosus; Dysosmobacter welbionis; Weissella cryptocerci;
• 32 new proteobacteria: Pelagibacterium sediminicola; Inhella crocodyli; Sphingorhabdus pulchriflava; Providencia huaxiensis; Neisseria weixii; Chakrabartia godavariana; Bosea caraganae; Dickeya undicola; Pseudogemmobacter bohemicus; Sphingobium terrigena; Acidimangrovimonas sediminis; Pseudomonas nitrititolerans; Hydrocarboniclastica marina; Alteromonas fortis; Erythrobacter marisflavi; Chromobacterium phragmitis; Marinicauda salina; Pseudopuniceibacterium sediminis; Aquisediminimonas profunda, Aquisediminimonas sediminicola; Croceibacterium ferulae; Tabrizicola sediminis; Parvularcula marina; Azospirillum palustre; Lujinxingia litoralis, Lujinxingia sediminis; Bradyrhizobium amphicarpaeae; Thiomicrorhabdus aquaedulcis; Paracoccus tegillarcae; Pantoea endophytica; Microvirga calopogonii; Vibrio profundi;
• 2 new spirochaetes: Leptospira yasudae, Leptospira stimsonii;

Archaeans

• 3 new species: Natronolimnobius sulfurireducens, Halalkaliarchaeum desulfuricum; Halomicrococcus hydrotolerans;

SARs

• 8 new bacillariophytes: Achnanthidium jiuzhaienis, Achnanthidium epilithica, Achnanthidium limosua, Achnanthidium subtilissimum, Kolbesia sichuanenis; Diploneis implicatus,Diploneis lacustris, Diploneis pumicosus;
• 1 new chrysophyte: Mallomonas gusakovii;
• 1 new apicomplexan: Isospora borbai;

Plants

• 2 new rhodophytes: Hypnea caraibica, Hypnea schneideri;
• 2 new marchantiophytes: Riccia sahyadrica; Cololejeunea lobulopapillata;
• 13 new ferns: Pteridrys catenata, P. cicuzzai, P. clarinervis, P. dongshiyongii, P. dorsifixa, P. malesiana, P. nonattingens, P. reticulata, P. triangularis, P. vanderwerffii , P. wusugongii; Pteris rugosifolia, Pteris subesquirolii;
• 74 new angiosperms: Senna acatlanensis; Marsdenia goromotoorum; Jasminanthes borneensis, J. hosei, J. kinabaluensis, J. sarawakensis; Polyalthia khaoyaiensis; Polystachya danielana; Ocimum sebrabergensis; Lepidagathis shrirangii; Isotrema plagiostomum; Guihaia heterosquama; Thesium ovatifolium; Onobrychis maassoumii; Hieracium jasiewiczii, H. wojcickii; Rhynchosia lewisii; Senecio stella-purpurea; Sinosenecio peltatus; Hedyotis tonggulingensis; Gynochthodes honbaensis; Puya colcaensis; Lippia diversifolia; Paphiopedilum erythroanthum; Bletilla guizhouensis; Begonia balangcodiae; B. inggitiae. B. dracopelta; Begonia pseudoheydei;  Begonia palemlemensis; Begonia ciliatifolia; Begonia zhongyangiana; Begonia yizhouensis; Begonia acutis, B. remusatifolia; B. kemiriensis; Begonia danxiaensis; Senegalia atlantica, Senegalia rafinesqueana, Senegalia cupuliformis; Piper caranoense; Gymnosporia sekhukhuniensis; Cymbidium jiangchengense; Rourea barbata, R. prostrata; Eugenia caipora; Werneria spp.; Oreocharis odontopetala; Justicia thailandica; Zeylanidium manasiae; Solanum plastisexum; Dysosma villosa; Sacoglottis perryi; Thismia domei, T. terengganuensis; Impatiens jenjittikuliae; Croton rizzinii; Griffinia albolineata; Mesosetum filgueirasii; Polylepis argentea; Dalechampia macrobractea; Cnidoscolus eglandulatus, Cnidoscolus infernidialis; Acalypha almadinensis; Microlicia longirostrata; Pennellia yalaensis; Aspidosperma huberianum; Mycetia suedixieana; Xenostegia lomamiensis; Solanum sp.; Artemisia baxoiensis; Moquiniastrum glabrum; Lepidaploa congesta, Critoniopsis canaliculata; Eryngium arenosum;

Fungi

• 21 new ascomycetes: Pseudosclerococcum golindoi; Beauveria baoshanensis, B. yunnanensis; Annabella australiensis; Deniquelata quercina; Alternaria omanensis; Arthrinium phyllostachium; Phaeosphaeria ampeli; Neoroussoella alishanense; Octospora conidiophora; Macrophomina vaccinii; Elbamycella rosea; Posidoniomyces atricolor; Saturnispora galanensis; Candida xylosifermentans; Candida yunnanensis, Candida parablackwelliae; Toxicocladosporium aquimarinum, Toxicocladosporium qatarense; Penicillium lusitanum; Valsaria ostryae;
• 12 new basidiomycetes: Dennisiomyces fibrillosus; Parasola psathyrelloides; Ganoderma shanxiense; Nia lenicarpa; Pluteus bizioi; Neofavolus yunnanensis; Amanita bweyeyensis, Amanita harkoneniana; Cacaoporus pallidicarneus, C. tenebrosus; Erythrophylloporus paucicarpus, E. suthepensis;

Poriferans

• 4 new demosponges: Exsuperantia archipelagus; Biemna lutea, Hamigera cleistochela; Coelosphaera (Coelosphaera) koltuni;
• 4 new calcareans: Achramorpha antarctica, Megapogon schiaparellii, Achramorpha ingolfi, Sarsinella karasikensis;

Cnidarians

• 1 new myxozoan: Ceratomyxa siganicola;

Flatworms

• 11 new monogeneans: Anacanthorus scyphophallus, Anacanthorus ataidei, Anacanthorus siphonocommus, Anacanthorus maratininguensis, Anacanthorus lacinimentulatus, Anacanthorus cururutuiensis, Anacanthorus circumspatulatus, Anacanthorus acrophallus; Gyrodactylus chiapaneco, Gyrodactylus guatopotei, Gyrodactylus tlaloci;
• 3 new trematodes: Rhopalias oochi; Pojmanskatrema balcanica; Cryptocotyle lata;
• 3 new cestodes: Acanthobothrium soniae, Acanthobothrium vidali; Caulobothrium pedunculatum;

Rotiferans

• 1 new species: Lecane langsenensis;

Bryozoans

• 2 new species: Parasmittina cryptoavicularia, Trematooecia persica;

Brachiopods

• 2 new species: Ospreyella mayottensis, Thecidellina leipnitzae;

Mollusks

• 11 new gastropods: “Inella” galo, “Inella” euconfio, “Inella” leucocephala, “Inella” faceta, “Inella” maculata, “Inella” vanilla, Strobiligera campista, Strobiligera santista; Olea hensoni; Aplysia ghanimii, A. hooveri;

Annelids

• 2 new polychaetes: Petta investigatoris, P. williamsonae;

Kinorhynchs

• 10 new species: Cristaphyes fortis, Higginsium mazatlanensis, Cephalorhyncha teresae, Echinoderes xalkutaat; Echinoderes kohni; Daracoderes spyro; Triodontoderes lagahoo; Gracilideres mawatarii; Condyloderes agnetis, Condyloderes clarae; Cristaphyes cornifrons, Echinoderes barbadensis;

Nematodes

• 10 new species: Daptonema papillifera, Pseudosteineria anteramphida; Spinonema cuticulatum, Spinonema spirale, Spinonema absente; Serpentirhabdias mussuranae; Rhabdias glaurungi; Xiphinema primum; Molgolaimus kaikouraensis, Aponema pseudotorosum;

Arachnids

• 13 new mites: Cosmolaelaps siberiensis; Neodicrothrix grandcaputus, Latitudo asiaticis; Gaeolaelaps lankaensis, G. setillus; Lepidocunaxoides robustus; Diplobodes thailande. Rwandabodes kayoveae; Leptogamasus (Medioperigamasus) lamelligynus, L.(M.) parcus, L.(M.) ramosus, L. (M.) sarcidanus, L.(M.) sasha;
• 30 new spiders: Otacilia acerosa, O. lata, O. palmata, O. rulinensis, O. xingdoushanensis; Tarrocanus jaffnaensis; Agroeca batangensis, A. lata, A. mainlingensis, A.nigra, A. tumida; Cicurina daweishanensis, C. kekei, C. lichuanensis, C. yuelushanensis; Caraimatta brescoviti; Lineacoelotes lifengyuanae, L. tiantaiensis, L. zhongbaensis, L. ziboensis; Thaloe maricao, Thaloe leboulet, Thaloe ebano; Pionothele gobabeb; Platythomisus xiandao; Pimoa binchuanensis, P. xinjianensis; Flexicrurum wuzhishanense, F. yangjiao, F. qishi;
• 4 new pseudoscorpions: Tenebriochernes concavus, T. mohani, T. pilosus; Cornuroncus sp.;
• 3 new harvestmen: Gonyleptellus angeloi; Protimesius lucifer, P. orcus;

Myriapods

• 33 new diplopodans: Antichiropus anguinus, A. antius, A. apricus, A. cirratus, A. confragus, A. cristatus, A. cucumeraceous, A. cunicularis Car, n. sp, A. echinus, A. filiolus, A. forcipatus, A. georginae, A. gibbus, A. hystricosus, A. julianneae, A. literulus, A. lucyae, A. nicholasi, A. nimbus, A. patriciae, A. pendiculus, A. picus, A. procerus, A. quaestionis, A. rupinus, A. salutus, A. servulus, A. simmonsi, A. sloanae, A. spathion, A. uvulus, A. verutus. A. vindicatus;
• 2 new chilopodans: Hessebius prospinosa, Lithobius (Ezembius) maqinensis;

Crustaceans

• 3 new copepods: Dendropsyllus kimi, Dimorphipodia changi, Laophontodes georgei;
• 12 new decapods: Geosesarma spectrum; Oxypleuron alisae, O. papuense, Rochinia cidaris, R. despereaux, Tunepugettia corbariae, Crocydocinus ewok, C. panglao, C. porg, C. vanuatu, Neophrys inopinata; Cristimenes brucei;
• 3 new amphipods: Metaprotella guileri, M. solitaria, Notoprotella cornuta;

Hexapods

• 5 new collembolans: Nothobrya sertaneja; Troglaphorura gladiator; Acherontides serrasapoensis; Tetracanthella annulata, T. tardoki;
• 2 new odonates: Andaeschna occidentalis; Heteragrion demarmelsi;
• 1 new ephemeropteran: Traverella maranhensis;
• 7 new dictyopterans: Rhabdoblatta similsinuata, Rh. densimaculata, Rh. gyroflexa, Rh.chaulformis, Rh. maculata, Rh. ecarinata; Laevifacies quadrialata;
• 16 new orthopterans: Oedipoda cynthiae; Diestramima arbora, Diestramima lamina; Lebinthus sandakan; Anaxipha nigritorquis, Metiochodes gracilus, Sectus integrum; Svistella argentata; Paraxiphidium iriodes; Austrosalomona destructor, A. poecila, Salomona nori; Homogryllacris obtusitubera; Formosatettix serrifemora , Formosatettix zhejiangensis, Formosatettix guposhanensis;
• 1 new phasmatodean: Pachyphloea magnoliae;
• 13 new plecopterans: Flavoperla basomarginata; Leuctra abkhaziae; Kamimuria petasus; Chinoperla biprojecta; Flavoperla furcomaculata, F. triangulata; Acroneuria henana; Microperla qinlinga; Filchneria songi; Kiotina bilobata; Paraleuctra cuihuashana; Alloperla clarki; Anacroneuria lacandona;
• 10 new thysanopterans: Mesandrothrips austrosteensia, M. googongi, M. kurandae, M. lamingtoni, M. oleariae; Urothrips probolus; Thrips rhamni, T. shiranensis, T. unisetifer; Haplothrips aliceae;
• 37 new hemipterans: Spockia tagala; Sadoletus abathonotus, Sadoletus alphus, Sadoletus biprotuberans, Sadoletus planus, Sadoletus variabilis; Sinchocoris giupponii; Daimachus matheranensis, D. robustus, D. sirsiensis, D. sudindicus, Ulopsina (Indoulopa) himalayana; Armorseliza spp.; Selymbria boliviaensis, S. chevauxensis, S. guianensis, S. cinctifera, S. ecuadorensis, S. guatemalensis, S. iguazuensis, S. loretoensis, S. madredediosensis, S. puntarenasensis; Nakaharanus lii; Ambrysus brailovskyi, A. henryi, A. schuhi, A. sitesi; Rotundocoris stenonotum, R. obliquonotum; Bhatia longiradiata; Tsauria brevispina, T. longispina; Pediopsoides (Sispocnis) rectus, P. (S.) triangulus; Metaphycus anophococcusi;
• 93 new coleopterans: Parastasia qiului; Atheta prolixa, A. vegrandis; Gyrophaena (s.str.) aryanamensis; Eurhamphus pancinii; Termitozophilus belleae; Elathous agilis, Megathous tannourinensis; Dysiatus sp.; Tropidion flechtmanni; Pelthydrus schoenmanni; Hydaticus panguana; Hamadiana chapadensis; Shirozuella flavosemiovata, Shirozuella limbata, Shirozuella poststigmaea; Stenus wutongshanus; Euconnus (Cladoconnus) rudimentalis; Luispenaia atacamensis, L. paposo, L. paulseni; Pseudoechthistatus hei; Carlschoenherria adoradae, C. hadsallae, C. gapudi; Anthaxia (Anthaxia) tichyi; Clidicus qiuae, C. yingjiangus; Eniclases kusyi, Schizotrichalus halmaherensis; Batoctenus kawmontis, B. kociani; Gyronotus dracomontanus, Gyronotus ovalis, Gyronotus kearneyorum; Sternocampsus coriaceus; Pseudocneorhinus angustus, P. glaber, P. hlavaci, P. obliquehumeralis, P. setosicallus; Ochthebius sasakii; Aulonochares lingulatus, Aulonochares novoairensis, Aulonochares tubulus, Ephydrolithus hamadae, Ephydrolithus minor, Ephydrolithus ogmos, Ephydrolithus spiculatus, Ephydrolithus teli, Primocerus cuspidis, Primocerus gigas, Primocerus neutrum, Primocerus ocellatus, Primocerus petilus, Primocerus pijiguaense, Primocerus maipure, Primocerus semipubescens, Primocerus striatolatus; Lochmaea tsoui, L. cheni, L. jungchani; Lankaphthona yunnantarsella; Zapotecotoma sumichrasti, Clinops inexpectatus, C. perpessus; Scutellathous habenularis, S. nanlingensis, S. quadrata; Amphicrossus kabitae, Amphicrossus brunneus, Amphicrossus adustipennis; Morulaptoma canuta; Triariodes admiratio, Triariodes segonku, Triarius novoleonis, Triarius texanus; Philothalpus sp.; Miridiba (Pledina) lamellata; Styphlomerus devagiriensis, Styphlomerus striatus; Canthidium nebularum, C. chimalapense; Cryptognatha pam, C. kellie, C. hannah, C. whitney, C. karla, C. celia, C. shelia, C. gayle, C. della, C. vicki;
• 48 new hymenopterans: Grotea goianiense, G. paulista, G. amazonensis, G. surinamese; Dipogon (Stigmatodipogon) siam, D. (S.) wasbaueri; Kiwi barrattae, K. canterberus, K. earlyi, K. gronous, K. oreteus, K. ruzelus, K. waitakerus, Zealochus abominosus and Z. stepheni; Probolomyrmex cegua, P. dentinodis, P. kelleri, P. lamellatus; Charops angelicae, C. eduardoi , C. katiae, C. lucianae, C. mariae, C. mucioi, C. terezae; Probles alejandroi, P. belokobylskii, P. clypeola, P. contrerasi, P. juanitae, P. lunai, P. megasoma, P. miquihuana, P. picus, P. spectabilis, P. xalapana, P. zacapoaxtlana; Anaphes triapitsyni, A. kailashchandrai; Conura baturitei; Stenodynerus lombokensis; Rhaconotus directus, R. laevigatus, R. robustus, R. simulatus; Trachusa tenebrosa; Traumatomutilla pilkingtoni;
• 1 new mecopteran: Palaeopsylla (Palaeopsylla) aysenurae;
• 48 new dipterans: Blepharella bomolocha, B. grisescens, B. yaeyamana; Megaselia falsum; Bryopharsos uncinatum, Bryopharsos paulistensis, Bryopharsos amazonensis; Hirtodrosophila minshanensis, H. lambda, H. zhangae, H. zouae, H. nigrispina; Grzegorzekia kerri, G. quercus; Hyadina breva, Parahyadina angusta, P. atra, P. bifurcata, P. bulla, P. debilis, P. edmistoni, P. hennigi, P. irwini, P. latistylis; Ophiomyia falcifera, Phytomyza varronivora; Hyphantrophaga adrianguadamuzi, Hyphantrophaga albopilosa, Hyphantrophaga anacordobae, Hyphantrophaga calixtomoragai, Hyphantrophaga calva, Hyphantrophaga ciriloumanai, Hyphantrophaga danausophaga, Hyphantrophaga diniamartinezae, Hyphantrophaga duniagarciae, Hyphantrophaga edwinapui, Hyphantrophaga eldaarayae, Hyphantrophaga eliethcantillanoe, Hyphantrophaga gilberthampiei, Hyphantrophaga guillermopereirai, Hyphantrophaga hazelcambroneroae, Hyphantrophaga luciariosae, Hyphantrophaga manuelriosi, Hyphantrophaga morphophaga, Hyphantrophaga nigricauda, Hyphantrophaga osvaldoespinozai, Hyphantrophaga pabloumanai, Hyphantrophaga similis;
• 9 new trichopterans: Helicopsyche bendego, H. daome, H. dinoprata, H. luziae, H. petri, H. shaamunensu; Smicridea (S.) lata, S. (S.) spatulata, S. (S.) dividua;
• 62 new lepidopterans: Artines panama, Artines solange, Artines delfos, Artines litoralis, Artines liege, Artines bamba, Artines angelica, Artines cofus; Cimeliomorpha jarujini, C. inflata, C. perspinosa; Barsine amoenissima, B. selene, B. cao, B. speideli, B. mesomene, B. pandeia, B. euryphaessa, B. ivanovamariae, B. eos, B. mene, B. syntypicoida, B. dubatolovi, B. visaya, B. nemea, B. kishidai; Deltoplastis acutangulata, D. anatoliana, D. multidentalis, D. similihoristis, D. aculeata, D. spatuliunca, D. ovidiscalis; Idiopteryx jansei, Lecithocera minyodes, Protolychnis natalensis; Alphaea alfreda; Andronymus magma, A. fenestra; Taikona ikedai; Mustilizans zolotuhini; Eretmocera yingjiangensis, E. octopunctata, E. artemisiae; Gnorimoschema pamira, G. brachyptera, G. altaica, G. tabazhok, G. yakovlevi, G. kozlovi; Athaumasta dzhungarica, A. kurchuma, A. arida, A. etugen, A. tarbagata, A. kuchinichi; Eraina chelifera; Michaelophorus salensis, Oidaematophorus androsensis, Hellinsia bahamensis, Hellinsia lucayana;

Chondrichthyans

• 1 new squaliform: Mollisquama mississippiensis;
• 1 new myliobatiform: Potamotrygon marquesi;

Actinopterygians

• 1 new perciform: Opsarius putaoensis;
• 1 new cypriniform: Garra paratrilobata;
• 1 new aulopiform: Lestrolepis luxiocula;
• 1 new anabantiform: Channa brunnea;
• 1 new gobiiform: Psilotris vantasselli;
• 1 new syngnathiform: Leptonotus vincentae;

Amphibians

• 9 new anurans: Gracixalus yunnanensis; Megophrys dongguanensis, M. jiulianensis, M. mufumontana, M. nankunensis, M. nanlingensis, M. wugongensis; Pristimantis andinogigas; Micryletta aishani;

Reptiles

• 15 new squamates: Lerista anyara; Opisthotropis haihaensis; Cyrtodactylus muluensis, C. limajalur; Cyrtodactylus mombergi; Hemidactylus kolliensis, Hemidactylus chikhaldaraensis, Hemidactylus sankariensis; Oligosoma salmo, O. albornense, O. auroraensis; Sinonatrix yapingi; Atractus dapsilis, A. trefauti, A. aboiporu;

Mammals

• 2 new rodents: Rhynchomys labo, R. mingan;

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

by Piter Kehoma Boll

I recently presented a thrips in the Friday Fellow section, in that case a thrips that infects mostly fig trees. This group of insects, which make up the insect order Thysanoptera, is poorly known by the general public, but is certainly known by gardeners and farmers, as they can be a serious nuisance for many plant types.

We could imagine thrips as being kind of the mosquitoes of plants. They pierce the surface of plants and suck their juices just like mosquitoes do with vertebrates. And we all know that a mosquito bite may lead to much more than a small blood loss and local irritation of the skin. Many parasites use mosquitoes as vectors to travel from host to host, including protists such as Plasmodium falciparum, which causes malaria, and many types of virus, such as those of the genus Flavivirus, which cause the yellow, dengue and zika fevers.

A similar thing happens in the association of thrips with plants. A special genus of virus, called Tospovirus, infects many plant species and uses thrips as a vector. Inside the thrips bodies, the viruses reproduce after infecting the epithelial cells of the gut and, from there, travel via blood to the salivary glands and, when a thrips perforates a plant, the virus is injected in it. The cycle is basically the same used by Flavivirus in mosquitoes and ticks to infect vertebrates. Isn’t it amazing how a virus such as Tospovirus can infect both an animal and a plant? But what exactly is the disease caused by these viruses?

One of the most common Tospovirus is the so-called Tomato spotted wilt virus (TSWV), which is considered one of the most economically devastating plant viruses in the world. It can infect many crops, such as tomato, tobacco, bellpepper, peanut and basil. The symptoms vary from plant to plant, but usually include stunting, poorly developed fruits, commonly with ring spots on the surface, and necrosis of the leaves. It is transmitted to plants by thrips of the genus Frankliniella, mainly the western flower thrips Frankliniella occidentalis. Although the virus usually needs several hours to be able to reinfect a plant after infecting a thrips, in ideal conditions the time can e as short as five minutes.

But why would a thrips feed on an obviously sick plant, all ugly and full of spots? They would certainly prefer a healthy plant, but that would prevent the virus to spread. As a result, the virus developed several strategies to attract the thrips. The TSWV is able to increase the amount of free aminoacids in infected plants, and these are essential nutrients for egg production in thrips. As a consequence, infected plants become more nutritious and attract more thrips. Feeding on infected plants, the thrips will certainly get infected and at the same time ingest more nutrients than non-infected thrips. Thus, a sick thrips actually has an increased fitness and usually lays more eggs. The plants would certainly get effing scared if they were able to have emotions.

The Soybean vein necrosis virus (SVNV) is another Tospovirus of economic concern. As it names suggests, it attacks mainly soy plants, and its main vector is the soybean thrips Neohydatothrips variabilis. Infected soybean thrips produce significantly more offspring than non-infected ones, although heavily infected individuals lay few viable eggs. How do thrips bypass this problem? It’s simple! Once they are infected, they stop feeding on infected plants and prefer non-infected ones, which increases their reproductive success by avoiding becoming heavily infected and at the same time they spread the virus further to non-infected plants. A nightmare for the plants once more.

A recent study investigated the relationship of another Tospovirus-thrips pair, this time of the iris yellow spot virus (IYSV), which commonly attacks garlic and onion plants, and its main vector, the onion thrips, Thrips tabaci. Infected thrips did not show an increased daily fecundity but had an increased lifespan, allowing them to lay more eggs simply because they lived longer.

But the effect of Tospovirus on thrips can go further. For example, although plants infected by the TSWV release more aminoacids that attract and increase the fecundity of thrips, the infections still seems to have some deleterious effects on the insect. Infected males of Frankliniella occidentalis increase their consumption of food juices and increase the transmission of the virus. Females, on the other hand, seem to need nutrients that cannot be found in plants. As a result, they increase the consumption of eggs of the two-spotted spider mite Tetranychus urticae, with which they often coexist. Although primarily herbivorous as most thrips, the western flower thrips eventually feeds on mite eggs, and being infected by TSWV makes females become more eager to eat eggs. This is certainly not a strategy of the virus itself as the other ones, since a female that is feeding on mite eggs does not contribute for the virus’ reproductive success. Nevertheless, this is an interesting phenomenon that show us how the interactions in a trophic web can be dynamic, changing, for example, due to an uninentional side effect of a virus trying to survive.

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References:

Keough S, Han J, Shuman T, Wise K, Nachappa P (2016) Effects of Soybean Vein Necrosis Virus on Life History and Host Preference of Its Vector, Neohydatothrips variabilis , and Evaluation of Vector Status of Frankliniella tritici and Frankliniella fusca. Journal of Economic Entomology 109(5): 1979–1987. doi: 10.1093/jee/tow145

Leach A, Fuchs M, Harding R, Nault BA (2019) Iris Yellow Spot Virus Prolongs the Adult Lifespan of Its Primary Vector, Onion Thrips (Thrips tabaci) (Thysanoptera: Thripidae). Journal of Insect Science 19(3): 8. doi: 10.1093/jisesa/iez041

Shrestha A, Srinivasan R, Riley DG, Culbreath AK (2012) Direct and indirect effects of a thrips‐transmitted Tospovirus on the preference and fitness of its vector, Frankliniella fusca. Entomologia Experimentalis et Applicata 145(3): 260–271. doi: 10.1111/eea.12011

Stafford-Banks CA, Yang LH, McMunn MS, Ullman DE (2014) Virus infection alters the predatory behavior of an omnivorous vector. Oikos 123(11): 1384–1390. doi: 10.1111/oik.01148

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by Piter Kehoma Boll

During the past three weeks, I presented a fig tree, the Chinese Banyan, a thrips that parasitizes it, the Cuban Laurel Thrips, and a mite that parasitizes the thrips, the Cuban-Laurel-Thrips Mite. However, I haven’t wrote yet about one of the most interesting creatures that interacts with a fig tree: its pollinator.

In the case of the Chinese Banyan, its pollinator is the fig wasp Eupristina verticillata, which I named the Chinese Banyan Wasp. As all fig wasps, this species is very small and completely adapted to live with figs. They cannot survive without the exact fig species with which they interact and the fig species cannot reproduce without that exact wasp. How does this works?

Let’s start our story with an adult female Chinese banyan wasp. The females are black and very small, measuring around 1 to 1.2 mm in length only. This female is flying around looking for a young fig which will serve as her nest and her grave.

A fig, in case you don’t know, is not a real fruit in the botanical sense. It is actually a special kind of inflorescence called a syconium that is basically a flower-filled sack. The inner walls of a fig have many tiny male and female flowers and the only way to get to them is through a tiny hole at the fig’s appex. And this hole is only open during the initial stages of the fig’s development.

When the female Chinese Banyan fig wasps is flying around, she is looking for a fig that is at this exactly stage of development. Once she finds one, she crawls inside the fig through that tiny hole. She usually loses her wings while doing that because the passage is too narrow. She evens needs to use her especially adapted mandible to help her go through. Once inside the fig, she looks for the female flowers, which are located at the base of the fig, away from the entrance. The male flowers, located right at the entrance, are not mature yet. However, the female wasps arrived with pollen that she gathered elsewhere (you will learn about that soon). When she reaches the female flowers, she introduces her ovopositor (the long structure at the end of her abdomen that is used to lay eggs) inside the female flower and lays one egg inside the flower’s ovary. Her ovopositor needs to have the exact size to reach the ovary to lay the egg. If it is too short, she is unable to complete her task. And while she is moving from flower to flower to lay eggs, she ends up pollinating them. After she has finished, she dies still inside the fig.

The ovaries that received an egg start to grow into a gall (a “plant tumor”) by influence of the insect and serve as food and shelter for the larvae that hatch from the eggs. A larva grows, pupates and turns into an adult inside a single gall. When the wasps have finally reached their adult stage, they leave the gall in which they were born. This happens when the fig reached its mature stage.

Males are the first ones to emerge. They are even smaller than the females and have a yellow to light-brown color. They gnaw their way through the gall and, once outside it (but still inside the fig) they start to look desperately for female wasps to inseminate. They do that by tearing other galls apart and, when a female is found trapped inside, they inseminate her. After that, the males dig a hole through the fig to the outside and die soon after, never experienced the external world.

Female wasps then leave their galls and move towards the hole opened by the male. While doing that, they move over the now mature male flowers and become covered in polen. After leaving the fig, they search for another fig that is in its early stage of development, restarting the cycle.

When a female leaves a mature fruit, she needs to find an immature one soon after that because she will die in a couple of days. In other words, the only way for this to work is if there are figs in the right stage all year around, and that is what happens. Differently from most plant species, which produce flowers in a specific time of the year, fig trees are always flowering. Well, not exactly. One individual fig tree produces figs only in a specific period of the year. All the figs of that tree ripen at the same time, i.e., a fig tree has an intra-individual synchrony of flower maturation. However, other trees of the same species have different moments to produce flowers, i.e., there is an inter-individual asynchrony of flower maturation. This assures that a wasp will always find a fig at the suitable maturation stage when there are enough fig trees around and also assures that a fig tree will not be fertilized by its own pollen.

As I mentioned when I presented the Chinese Banyan, this tree can only produce viable figs when the wasp is present, so that populations introduced outside of their native range will only reproduce if the waps is introduced as well. However, the wasp will be unable to survive if there are not enough fig trees to provide it with figs all year round. It is a delicate relationship between a tiny, fragile and short-lived insect and a huge, resistant and long-lived tree. And they need each other to survive.

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References:

Cook J, Rasplus J-Y (2003) Mutualists with attitude: coevolving fig wasps and figs. TRENDS in Ecology and Evolution 18(5): 241–248.

Kjellberg F, Jousselin E, Hossaert-McKey M, Rasplus J-Y (2005) Biology, Ecology, and Evolution of Fig-pollinating Wasps (Chalcidoidea, Agaonidae). In Raman A, Schaefer CW, Withers TM (Eds.) Biology, ecology and evolution of gall-inducing arthropods. v.2. New Hampshire, Science, p.539-572.

McPherson JR (2005) A Recent Expansion of its Queensland Range by Eupristina verticillata, Waterston (Hymenoptera, Agaonidae, Agaoninae), the Pollinator of Ficus microcarpa l.f. (Moraceae). Proceedings of the Linnean Society of New South Wales: 126: 197–201.

Weiblen DG (2002) How to be a fig wasp. Annual Review of Entomology 47: 299–330.

Wiebes JT (1992) Agaonidae (Hymenoptera, Chalcidoidea) and Ficus (Moraceae): fig waps and their figs, VIII (Eupristina s.l.). Proceedings of the Koninklijke Nederlandse Akademie van Wetenschappen 95(1): 109–125.

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by Piter Kehoma Boll

It’s been a very long time since the last time I presented a lepidopteran here, so today I decided to go back to this amazing group of insects. The species I chose for today is quite popular, maybe the most popular moth in the world. Its name is Actias luna, commonly known as the luna moth.

The luna moth is native from Canada and the United States. It is a quite large moth, with a wingspan of about 8 to 12 cm, although some individuals can be as big as 18 cm. Its wings, covered with scales as usual in lepidopterans, have a light green color. The forewigs have a brown anterior border that connects to two eyespots (one on each wing) by a stalk. The hindwings also have one eyespot each, but they are not connected by a stalk to the border. The hindwings also have a long tail that is characteristic of the genus Actias and somewhat resembles the similar (but shorter) tails in some butterflies, such as those of the family Papilionidae. Males and females are very similar and can be often distinguished by the size of the abdomen, which is much thicker in females.

In colder climates, such as in Canada, the luna moth has one generation per year, but southern populations, in places where the climate is warmer, can have up to three. The females lay eggs on suitable plants to serve as food for the larvae. There are several identified tree species that are used as food, including birches, walnuts, hickories and persimmons. The larvae feeding on a tree never, or very rarely, reach a number that can cause significant damage to the plant.

The eggs are brown and laid in irregular clusters on the underside of the leaves. They usually hatch one to two weeks after being laid and originate small, green larvae. The larvae are green in all instars and pass through five of them during a period of about 7 weeks. The fifth and final instar then descends the tree in which it lives to reach the ground. There, it starts to spin a silk coccoon and, after finishing it, turns into a pupa. In warmer regions, the pupa takes about two weeks to become an adult, but in colder regions it enters into diapause over winter, taking about nine months to complete the cycle.

When females become adults, they search for a suitable tree of its preferred species (usually the same species in which it was born) and emits pheromones to attract males. Adults lack mouth parts and, therefore, do not eat, living only enough to mate and lay eggs. The nice long tails on the hindwings, more than just beautiful, seem to decrease the ability of bats to detect them using their echolocation.

The luna moth is one of the most popular insects in North America. In fact, it was the first insect ever to be described from the continent, being named Phalaena plumata caudata by James Petiver in 1700. When Linnaeus started the binomial nomenclature for animals in 1758, he renamed it Phalaena luna as a reference to the Roman goddess of the moon.

Although not considered a vulnerable species at the moment, the luna moth faces some threats caused by human interference, such as habitat loss and damage caused by invasive species. Fortunately, due to its popularity, it is likely to have considerable support from the public for its conservation when that time comes.

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References:

Lindroth RL (1989) Chemical ecology of the luna moth: Effects of host plant on detoxification enzyme activity. Journal of Chemical Ecology 15(7): 2019–2029.

Millar JG, Haynes KF, Dossey AT, McElfresh JS, Allison JD (2016) Sex Attractant Pheromone of the Luna Moth, Actias luna (Linnaeus). Journal of Chemical Ecology 42(9): 869–876.

Wikipedia. Luna moth. Available at < https://en.wikipedia.org/wiki/Luna_moth >. Access on 11 July 2019.

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by Piter Kehoma Boll

Finding efficient ways to deal with agricultural pests in crops is a challenging work. Currently, as we all known, the main strategy to control such pests is the use of chemical pesticides. However, this approach only serves the interests of those seeking profit over well-being, as we all know that such pesticides increase the risk of several health issues in those consuming the crops. More than that, chemical pesticides not only kill the targeted pest but many other life forms, causing a devastating effect on ecosystems.

Fortunately, there has been an increasing interest in finding alternative, healthier ways to deal with the problem. One way is the production of genetically modified organisms (GMOs) that are naturally resistant to pests. There are, however, two main problems with this approach. The first one is that the population in general has an irrational fear of GMOs, apparently believing that they can be more harmful than the poisonous chemical pesticides, which is completely absurd. The second problem with GMOs is that the technology to create them is dominated by the same companies that produce most pesticides and, as all big companies, only seek profit and do not give a damn about the people and the environment.

A third strategy is the use of natural enemies of the pests to control them in organic farms. Although many natural enemies are great doing their job, they may also cause negative impacts by interfering with the surrounding ecosystems. Many crop pests are not native from the area where they are pests, i.e., they are invasive species and, in order to control them efficiently, a predator from its native area must be introduced as well, and this predator may end up becoming a threat to other species that it elects as food.

Fortunately, some nice strategies have been recently developed. One of them includes the use of additional plants in the fields that change the way that pests behave without posing a threat to surrounding areas. These additional plants consists of two types: trap crops and insectary plants.

A trap crop, as the name suggests, is an additional crop that is not intended to be commercially exploited, but serves as a trap for the pests. Instead of attacking the main crop (called the ‘cash crop’), the pests are attracted to the trap crop, reducing their density in the cash crop. This system is more efficient if the trap crop is similar to the cash crop, such as another plant of the same genus, or another variety of the same species, because it must be as attractive to the pest as the cash crop, or perhaps even more attractive.

Insectary plants, on the other hand, are intended to attract other insects to the plantation, especially predatory insects that prey on the agricultural pest. Insectary plants should produce flowers in abundance, thus attracting many insect species, which will increase the interest of predators in the area. However, when used alone, insectary plants will only provide predators to control the pest in crop plants that are near the insectary plants and, as they are usually planted in an area surrounding the plantation, they would not protect the plants that are near the center of the plantation.

In a recent study, Shrestha et al. (see references) decided to combine trap crops and insectary plants together with the cash crops in a strategy that they called a ‘botanical triad’. The cash crap was organic cabbage (Brassica oleracea var. capitata) planted in the eastern United States; the trap crops were three other crops of the genus Brassica: mighty mustard (Brassica juncea), kale (Brassica oleracea var. acephala) and collard (Brassica oleracea var. italica); and the insectary plants were buckwheat (Fagopyrum esculentum) and sweet alyssum (Lobularia maritima).

As a result, the number of herbivores (i.e., crop pests) was larger in the trap crops than in the cash crop. The trap crops were, therefore, more attractive than the cash crops for the pests. The presence of insectary plants increased the number of predatory and parasitoid insects, such as lady beetles and parasitoid wasps, in the trap crops when compared to treatments without insectary plants. The number of parasitized pests also increased in the presence of insectary plants.

In general, the “team work” of trap crops and insectary plants greatly reduced the influence of agricultural pests on the cash crops. The trap crops attracted the pests to an area close to the insectary plants, allowing the predators to reach them.

Efficient ways to raise crops organically are possible. We just have to focus on a healthy ecosystem and not on money. If we work together, we can defeat the “Big 6” corporations that dominate the food production in the world. They are the real pests.

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Reference:

Shrestha B, Finke DL, Piñero JC (2019) The ‘Botanical Triad’: The Presence of Insectary Plants Enhances Natural Enemy Abundance on Trap Crop Plants in an Organic Cabbage Agro-Ecosystem. Insects 10(6): 181. doi: 10.3390/insects10060181

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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 4.0 International License.

by Piter Kehoma Boll

It’s time to talk about an ostracode, or seed shrimp, again and, as usual, this is a difficult time due to the little information easily accessible regarding any particular species of this group. But there is, indeed, one that is considerably well studied. Being one of the most common ostracodes in North America and Eurasia, its scientific name is Cypridopsis vidua, to which I coined the common name “stonewort seed shrimp”.

The stonewort seed shrimp is a freshwater crustacean with the typical ostracode appearance, looking like a tiny bivalve measuring about 0.5 mm in length. Its valves have a distinctive light and dark pattern.

A relatively mobile species, the stonewort seed shrimp lives at the bottom of water bodies, over the sediment, and is common in areas that are densely vegetated by stoneworts (genus Chara). This association with stoneworts gives the stonewort seed shrimp both protection from predators, which are mostly fish, and a good food source.

The main food of the stonewort seed shrimp are microscopic algae that grow on the stems of stoneworts. While foraging, the stonewort seed shrimp swims from one stonewort stem to another using its first pair of antennae and clings on the stems using the second pair of antennae and the first pair of thoracic legs. Once realocated, it starts to scrape the microscopic algae using its mandibles.

The stonewort seed shrimp is one more of those species in which males do not exist, not even in small quantities. During the warm months of summer, females produce the so-called subitaneous eggs, which develop immediately into new females. However, when winter is approaching, they produce another type of eggs, the so-called diapausing eggs, which remain dormant in the substrate during winter. The adult animals all die during this season and, when spring arrives, a new population appears from the hatching eggs. Since not all eggs hatch in the spring, some of them may remain in the substrate for years before hatching, which usually increases the genetic diversity every year, as it not only depends of the daughters of the last generation.

But how does genetic diversity appear if there are no males and, as a result, the daughters are always clones of the mothers? This mystery is not yet fully solved. Genetic recombination during parthenogenesis, by exchanging alleles between chromosomes, does not seem to be very common. It is possible that different populations are genetically different and that they colonize new areas very often, mixing with each other. Since males are known in closely related species, it is still possible that, some day, we will find, somewhere, some hidden males of the stonewort seed shrimp. It is also possible that, somehow, males went all extinct in the recent past, like in the last glaciation, for example. If so, only time can tell what is the destiny of the stonewort seed shrimp.

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More Ostracods:

Friday Fellow: Sharp-Toothed Venus Seed Shrimp (on 22 June 2018)

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References:

Cywinska A, Hebert PDN (2002) Origins of clonal diversity in the hypervariable asexual ostracode Cypridopsis vidua. Journal of Evolutionary Biology 15: 134–145. doi: 10.1046/j.1420-9101.2002.00362.x

Roca JR, Baltanas A, Uiblein F (1993) Adaptive responses in Cypridopsis vidua (Crustacea: Ostracoda) to food and shelter offered by a macrophyte (Chara fragilis). Hydrobiologia 262: 121–131.

Uiblein F, Roca JP, Danielpool DL (1994) Experimental observations on the behavior of the ostracode Cypridopsis vidua. Internationale Vereinigung für Theoretische und Angewandte Limnologie: Verhandlungen 25: 2418–2420.

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

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

by Piter Kehoma Boll

Millipedes, which make up the class Diplopoda, are very cute arthropods in my opinion and include amazing species, such as the animal with the largest number of legs in the world. Many species are not well studied, though. However, one that is very well known is the Portuguese Millipede Ommatoiulus moreleti.

As its common name suggests, the Portuguese millipede is native from Portugal, more precisely from Southern Portugal and nearby areas in Spain, living in the soil of pine and oak forests. Its body, measuring about 4 cm as adults, has the typical cylindrical and elongate shape seen in most millipedes and is very dark, almost black, with legs that have a light color, usually whitish, but sometimes purplish.

Despite its relatively small size, the Portuguese millipede takes more than a year to reach maturity and grow for about three years. The mating period is usually during Autumn, and after having its eggs fertilized, the female lays from 60 to 80 of them in a chamber about 2 cm deep in the soil. When the eggs hatch, the first stage is a small, pupoid legless animal that remains inside a membrane until it molts into a small six-legged larva. During the first year, the juvenile molts about 8 times and the number of legs increases at each new stage. At about stage 10, they are sexually mature, but continue to molt and gaining more legs until reaching about 90 legs at the 14th stage. Males have an interesting reproductive strategy called periodomorphism, in which mature individuals molt into a “castrated” form, with reduced sexual organs, and becomes sexually mature again in the next molt, only to return to the immature form again in the next molt and so on.

The Portuguese millipede became famous after its accidental introduction in southeastern Australia, apparently in the 1950s. It soon became a very abundant species and, as a consequence, a nuisance for humans. As most millipedes, the Portuguese millipede is mainly detritivorous, feeding on dead plant material, such as rotten wood and dead leaves, so its introduction is not that much an ecological catastrophe, although it can have some negative impacts by competing with native millipede species.

The main problems caused by the introduction of the Portuguese millipede in Australia affect mostly humans. They are attracted to weak light sources, such as those emitted by houses at night, and, as a result, end up invading residences, sometimes hundreds of them at a time. When threatened, the Portuguese millipede emits a pungent yellow secretion that can irritate the eyes and, in contact with clothes, mark them with a permanent stain. Addtionally, the Portuguese millipede sometimes can feed on some crops, especially fruits.

In Portugal, the populations of the Portuguese millipede are controlled by native predators, such as the European hedgehog Erinaceus europaeus and the beetle Ocypus olens. Released from these natural enemies, the millipede spread quickly through southeastern Australia. However, about 30 years later, its population in Australia started to decrease. Apparently some nematode parasites that infect native millipedes adapted to parasitize this invasive species as well, helping to contain its population size. Some other native Australian predators have also observed feeding on the Portuguese millipede, including the blue garden flatworm, Caenoplana coerulea.

Other than Australia, the Portuguese millipede was also introduced in several Atlantic Islands, such as the Macaronesian Islands, Bermuda and the UK, as well as in South Africa. However, it does not seem to be that much of a nuisance there.

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More millipedes:

Friday Fellow: Leggiest Millipede (on 12 February 2016)

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References:

Baker GH (1985) Predators of Ommatoiulus moreletii (Lucas) (Diplopoda: Iulidae) in Portugal and Australia. Australian Journal of Entomology 24(4): 247–252. doi: 10.1111/j.1440-6055.1985.tb00237.x

Baker GH (1978) The post-embryonic development and life history of the millipede, Ommatoiulus moreletii (Diplopoda: Iulidae), introduced in south-eastern Australia. Journal of Zoology 186: 209–228. doi: 10.1111/j.1469-7998.1978.tb03366.x

Gregory SJ, Owen C, Jones G, Williams E (2018) Ommatoiulus moreleti (Lucas) and Cylindroiulus pyrenaicus (Brölemann) new to the UK (Diplopoda, Julida: Julidae) and a new host for Rickia laboulbenioides (Laboulsbeniales). Bulletin of the British Myriapod & Isopod Group 30: 48–60.

McKillup SC, Allen PG, Skewes MA (1988) The natural decline of an introduced species following its initial increase in abundance: an explanation for Ommatoiulus moreletii in Australia. Oecologia 77:339–342. doi: 10.1007/BF00378039

Terrace TE, Baker GH (1994) The blue land planarian, Caenoplana coerulea Moseley (Tricladida: Geoplanidae), a predator of Ommatoiulus moreleti (Lucas) (Diplopoda: Julidae) in southern Australia. Australian Journal of Entomology 33(4): 371–372.

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

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

by Piter Kehoma Boll

Here is a list of species described this month. It certainly does not include all described species. Most information comes from the journals Mycokeys, Phytokeys, Zookeys, Phytotaxa, Zootaxa, Mycological Progress, Journal of Eukaryotic Microbiology, International Journal of Systematic and Evolutionary Microbiology, Systematic and Applied Microbiology, Zoological Journal of the Linnean Society, PeerJ, Journal of Natural History and PLoS One, as well as several journals restricted to certain taxa.

Bacteria

• 18 new actinobacteria: Galactobacter caseinivorans, Galactobacter valiniphilus; Actinomadura logoneensis; Actinomadura roseirufa; Marinitenerispora sediminis; Corynebacterium choanae, Corynebacterium pseudopelargi, Corynebacterium gerontici; Micromonospora radicis; Arthrobacter celericrescens; Gandjariella thermophila; Fudania jinshanensis; Nocardia yunnanensis; Agromyces tardus; Cryobacterium melibiosiphilum; Rhodococcus subtropicus; Labedella phragmitis, Labedella populi;
• 5 new bacteroids: Aquirufa antheringensis, Aquirufa nivalisilvae; Hymenobacter edaphi; Muricauda amoyensis; Mucilaginibacter terrenus;
• 1 new cyanobacterium: Sinocapsa zengkensis;
• 28 new firmicutes: Anaerobacillus isosaccharinicus; Indiicoccus explosivorum; Streptococcus hillyeri; Thermoflavimicrobium daqui; Lactobacillus yilanensis, Lactobacillus bayanensis, Lactobacillus keshanensis., Lactobacillus kedongensis, Lactobacillus baiquanensis, Lactobacillus jidongensis, Lactobacillus hulinensis, Lactobacillus mishanensis. Lactobacillus zhongbaensis; Lactobacillus pingfangensis, Lactobacillus daoliensis, Lactobacillus nangangensis, Lactobacillus daowaiensis, Lactobacillus dongliensis, Lactobacillus songbeiensis, Lactobacillus kaifaensis; Enterococcus pingfangensis, Enterococcus dongliensis, Enterococcus hulanensis, Enterococcus nangangensis; Enterococcus songbeiensis; Aquibacillus sediminis; Oceanobacillus piezotolerans, Bacillus piezotolerans;
• 31 new proteobacteria: Oceaniglobus ichthyenteri; Acidithiobacillus sulfuriphilus; Schlegelella brevitalea; Ruegeria lutea; Sphingosinithalassobacter portus; Iodobacter ciconiae; Paenimaribius caenipelagi; Colwellia ponticola; Ferruginivarius sediminum; Sphingomonas flavalba; Paracoccus nototheniae; Roseovarius amoyensis; Pseudomonas edaphica; Rhodoferax lacus; Aliishimia ponticola; Cohaesibacter intestini; Parashewanella tropica; Sphingomonas ginkgonis; Pectobacterium odoriferum, Pectobacterium brasiliense, Pectobacterium actinidiae, Pectobacterium versatile; Sulfitobacter sabulilitoris; Erythrobacter suaedae; Pseudomonas huaxiensis; Acidibrevibacterium fodinaquatile; Kosakonia quasisacchari; Poseidonibacter antarcticus; Bradyrhizobium nanningense, Bradyrhizobium guangzhouense, Bradyrhizobium zhanjiangense;

SARs

• 3 new apicomplexans: Amoebogregarina taeniopoda, Quadruspinospora mexicana; Eimeria woyliei;
• 2 new bacillariophytes: Aulacoseira lauquenensis, Aulacoseira liucoensis;

Plants

• 1 new bryophyte: Macromitrium maolanense;
• 56 new angiosperms: Isostigma resupinatum; Muscari pamiryigidii; Petrocodon longitubus; Tagetes imbricata; Ledebouria weberi; Cremanthodium magnificum; Convolvulus hormodensis; Salvia gavilanensis; Begonia zunyiensis; Begonia saolaensis; Sphyrospermum grandiflorum; Minuartiella serpentinicola; Puccinia guassuensis; Desmodium amplistipulaceum; Crotalaria kanchiana; Lycianthes breedlovei, L. fredyclaudiae; Cornus sunhangii; Ilex montebellensis; Glandularia rupestris , G. sessilifolia; Barleria sahyadrica; Pilosocereus jamaicensis; Allium dönmezii; Zehneria tuberifera; Primulina lianchengensis; Primulina purpureokylin, P. persica, P. cerina, P. niveolanosa, P. leiyyi; Aiphanes decipiens, Aiphanes gloria, Aiphanes tatama; Coccothrinax viridescens; Hedysarum wangii; Aconitum tawangense; Stachytarpheta grandiflora; Liparis popaensis; Annona caput-medusae, A. oleifolia, Guatteria aliciae, G. attenuata, G. kamakusensis, G. pseudoferruginea, G. pseudorotundata, G. rubiginosa, G. turrialbana, Klarobelia rocioae, Tetrameranthus trichocarpus, Xylopia longicaudata; Cynometra cerebriformis, Cynometra dwyerii, C. tumbesiana, C. steyermarkii; Lachemilla rothmaleriana, Lachemilla argentea, Lachemilla cyanea; Tashiroea villosa; Rhaptopetalum rabiense; Clematis guniuensis; Pseuderanthemum melanesicum;

Fungi

• 40 new ascomycetes: Savoryella sarushimana; Cryptothecia panchganiensis; Asterina cerradensis, Asterina malvacearum, Lembosia matogrossensis, L. miconiphylla; Micarea fennica; Sphaeropezia leucocheila; Akanthomyces araneicola; Cladosporium passiflorae, C. passifloricola; Wicklowia submersa; Pachyphlodes cinnabarina, P. wulushanensis; Stemphylium dianthi; Ophiosphaerella taiwanica; Conioscypha tenebrosa; Dicephalospora albolutea, D. shennongjiana, D. yunnanica; Rugonectria microconidia and Thelonectria guangdongensis; Conlarium baiseense, C. nanningense, C. sacchari; Neothyronectria citri; Coryneum ilicis, C.songshanense; Ophiocordyceps asiatica, O. brunneirubra, O.khokpasiensis, O. mosingtoensis, O. pseudocommunis, O. pseudorhizoidea, O. termiticola; Micarea aeruginoprasina, M. azorica, M. isidioprasina, M. microsorediata, M. nigra, M. pauli; Aspergillus olivimuriae; Aspergillus takadae; Metschnikowia baotianmanensis; Phalangispora sinensis; Wickerhamiella shivajii;
• 13 new basidiomycetes: Laetiporus lobatus; Volvariella niveosulcata, V. guttulosa, V. ptilotricha, V. pulla; Stropharia atroferruginea; Tubaria squamata; Amanita mansehraensis; Amanita ahmadii; Leccinellum alborufescens, L. fujianense; Hygrophorus alboflavescens, Hygrophorus scabrellus; Heterocephalacria mucosa; Spencerozyma acididurans;

Poriferans

• 9 new demosponges: Penares mollis, P. aureus, P. vermiculatus, P. kermadecensis, P. turmericolor, P. deformis, P. okokewae, P. orbis, P. astronavis;

Rotiferans

• 6 new acanthocephalans: Plagiorhynchus (Plagiorhynchus) aznari; Acanthogyrus (Acanthosentis) fusiformis; Pachysentis lauroi; Rhadinorhynchus circumspinus, Rhadinorhynchus pacificus, Rhadinorhynchus multispinosus;

Flatworms

• 1 new polyclad: Prosthiostomum torquatum;
• 15 new rhabdocoels: Cheliplana gibarenha, C. santiaguera, C. spuriaseminalis, C. subproximalis, C. verrucosa, Carcharodorhynchus smilodon, C. papillaris, Carcharodorhynchus spiniformis, C. nativus, Schizochilus favus, S. bueycabonensis, S. atlanticus, S. espinosai, S. banesensis, S. maximus;
• 3 new monogeneans: Lamellodiscus occiduus, Lamellodiscus vesperus; Neoheterocotyle quadrispinata;
• 3 new trematodes: Atamatam amazonensis, Paratamatam sp.; Metadelphis tkachi;
• 1 new cestode: Dieffluvium nidulorum;

Annelids

• 17 new polychaetes: Aphelochaeta abyssalis, A. clarionensis, A. clippertonensis, A. spargosis, A. tanyperistomia, A. wilsoni, Caulleriella bathytata, Chaetozone akaina, C. grasslei, C. truebloodi, Tharyx hessleri; Fauveliopsis levensteinae, F. magalhaesi, Laubieriopsis blakei, Riseriopsis santosae; Amphiglena seaverae, Amphiglena joyceae;
• 1 new clitellate: Branchellion spindolaorum;

Mollusks

• 3 new gastropods: Sinochloritis lii, Bradybaena linjun; Chamalycaeus rarus;

Kinorhynchs

• 4 new species: Cristaphyes retractilis, Fujuriphyes dalii, Echinoderes brevipes, Echinoderes parahorni;

Nematodes

• 10 new species: Ascarophis morronei; Pterygodermatites (Pterygodermatites) atlanticaensis; Acanthostrongylus secundus; Malenchus gilanensis; Sabatieria chukchensis, Sabatieria parvamphis, Sabatieria major, Sabatieria multisupplementia; Physaloptera amazonica; Gyrinicola dehradunensis;

Tardigrades

• 2 new species: Meplitumen aluna; Hansenarctus toyoshiomaruae;

Arachnids

• 3 new mites: Ololaelaps elongatus; Knopkirie flascus, Kuzinellus rarotonga;
• 1 new scorpion: Centruroides romeroi;
• 9 new pseudoscorpions: Garypus latens, G. malgaryungu, G. necopinus, G. postlei, G. ranalliorum, G. weipa, G. dissitus, G. reong, G. yeni;
• 23 new spiders: Actinopus coboi, A. fernandezi, A. simoi, A. uruguayense; Macrophyes pacoti; Anicius chiapanecus, Anicius cielito, Anicius faunus, Anicius grisae, Anicius maddisoni; Herbiphantes acutalis, Labullinyphia furcata; Epiceraticelus mandyae; Cryptachaea pilar; Drassodes mylonasi , Echemus kaltsasi, Minosiella apolakia, Phaeocedus vankeeri, Turkozelotes attavirus; Draconarius baibaensis, Draconarius budanlaensis, Draconarius yigongensis, Draconarius yingbinensis;

Myriapods

• 6 new diplopods: Hyleoglomeris rukouqu, Hyleoglomeris xuxiakei, Hylomus yuani, Eutrichodesmus jianjia, Trichopeltis liangfengdong, Glyphiulus maocun;

Crustaceans

• 1 new ostracod: Chelonocytherois omutai;
• 4 new copepods: Monsmeteoris wiesheuorum, M. reductus, Cylindropsyllus valentini, C. flexibilis;
• 9 new amphipods: Propejanice lagamarensis; Colomastix iemanja, Colomastix tubulosa, Colomastix marielle; Gazia gazi, Talorchestia anakao; Aciconula tinggiensis; Sarothrogammarus yiiruae; Hyalella puna;
• 1 new isopod: Caecidotea camaxtli;
• 1 new tanaid: Xenosinelobus balanocolus; Hexapleomera sasuke;
• 7 new decapods: Nennalpheus gabonensis; Synalpheus amintae; Spongicola liosomatus; Xiphocaridinella sp.; Geosesarma mirum; Barathricola thermophilus; Macrobrachium laevis;

Hexapods

• 2 new collembolans: Aethiopella ricardoi; Megalothorax dobsinensis;
• 2 new ephemeropteran: Thraulus thiagarajani; Potamocloeon edentatum;
• 1 new dictyopteran: Brachylatindia xui;
• 6 new orthopterans: Phyllotrella hainanensis, P. transversa, P. fumingi; Criotettix longispinus, Criotettix undatifemurus; Indigryllus kudremu;
• 3 new plecopterans: Cerconychia trilobata; Capnia s.l. bilobata; Perlesta sublobata;
• 1 new thysanopteran: Heliothrips longisensibilis;
• 23 new hemipterans: Orius hibiscus; Stictotettix cleyerae, S. morishimai; Limnocoris chaetocarinatus, L. major, L. nanus, L. zacki; Physatocheila nigrintegerrima; Myittana (Benglebra) biflaka, M. (Benglebra) curvata; Maiestas borealis, M. maritima, M. peninsularis; Phytocoris (Compsocerocoris) amardus, Phytocoris (Compsocerocoris) hyrcaniaensis; Megacanthaspis guiyangensis; Ceratogergithus brachyspinus, Neohemisphaerius clavatus; Malaxa hamuliferum, M. tricuspis; Bambusana longispina; Esakia borneensis, E. mazzoldii;
• 87 new coleopterans: Vitacinis luziae, Nealcidion antonkozlovi, Onalcidion antonkozlovi, Leptocometes antonkozlovi; Derisemias tsutsumiuchii, Derisemias kojimai; Agrilus acacivorus, A. albizivorus, A. coco, A. monadikos, A. occultus; Sathytes borneoensis, S. liuyei, S. larifuga, S. shihongliangi; Tropanisopodus antonkozlovi, Paranisopodus antonkozlovi; Notomicrus meizon. N. teramnus; Cephennomicrus yunnanicus, C. andreasi; Scybalocanthon acrianus, S. adisi, S. arnaudi, S. chamorroi, S. federicoescobari, S. haroldi, S. martinezi, S. papaxibe; Xalpirta mauryi; Sichuanortia zhouchaoi; Barypalpus chinensis, B. holzschuhi, B. shibatai; Cionus armeniacus, C. boroveci, C. colonnellii, C. dodeki , C. harani, C. himalayensis, C. khorasanicus, C. laibalei, C. maurus, C. negevicola, C. neglectus, C. osmanlis, C. rossicus, C. rufescens, C. winkelmanni, C. yunnanensis; Yara oyaguei; Agelosus auricomus, Apecholinus imitator, Apecholinus canifer; Zeugophora cupka; Oxytrechus chioriae, O. cayambeensis; Tmesiphorus kinomurai, T. okinawensis; Rosalba contracta, R. venusta, R. skelleyi, Tethystola alboangulata, Blabicentrus similis, Pygmaeopsis apicalis, Pseudestola maculata; Esemephe peba; Hyalonthophagus pulcher; Peripus brunkei, P. didontus, P. madrededios, P. monodontus; Linan qiniangmontis; Isacanthon mariaelinae; Pharetis thayerae, Spherita newtoni; Canthon (P.) arriagadai, Canthon (P.) bonaerensis, Canthon (P.) vidaurrei, Canthon (P.) ziggy; Cymatodera acuminata, Cymatodera unica, Cymatodera parva, Cymatodera magdalena;
• 11 new neuropterans: Afroptera acuta, A. alba, A. brinkmani, A. balli, A. cylindrata , A. folia, A. koranna, A. maraisi, Nemopterella kabas, N.cedrus; Nothochrysa ehrenbergi;
• 110 new hymenopterans: Dicranorhina dinesani, D. georgei, D. sreeramani; Orthostigma (Patrisaspilota) enduwaense; Brachygaster gussakovskiji; Passaloecus bisulcatus, P. profundesulcatus, P. tuberangustus, P. tuberculiformis; Anteon ambrense, Anteon beankanum, Anteon elongatum, Anteon hoekense, Anteon mabibiense, Anteon majunganum, Anteon malagasy, Anteon musmani, Anteon nigropictum, Anteon nimbense, Anteon pseudohova, Anteon sakalavense, Anteon tulearense, Aphelopus sequeirai, Apoaphelopus fisheri, Apoaphelopus wallacei, Bocchus forestalis, Bocchus granulatus, Bocchus harinhalai, Bocchus nigroflavus, Bocchus parkeri, Bocchus ruvidus, Conganteon hawleyi, Conganteon sensitivum, Crovettia afra, Deinodryinus ambrensis, Deinodryinus granulatus, Deinodryinus nigropictus, Deinodryinus piceus, Dryinus bellicosus, Dryinus dentatiforceps, Dryinus erenianus, Dryinus milleri, Dryinus mobotensis, Dryinus nigrithorax, Dryinus teres, Dryinus tulearensis, Dryinus whittleorum, Gonatopus avontuurensis, Gonatopus bellicosus, Gonatopus comorensis, Gonatopus costalis, Gonatopus flavotestaceus, Gonatopus gumovskyi, Gonatopus hantamensis, Gonatopus harinhalai, Gonatopus karooensis, Gonatopus koebergensis, Gonatopus marojejyanus, Gonatopus minutus, Gonatopus nigropictus, Gonatopus ranomafanensis, Gonatopus robertsoni, Gonatopus rugithorax, Gonatopus scholtzi, Gonatopus wikstrandae, Lonchodryinus madagascolus, Madecadryinus ranomafanensis, Neodryinus bimaculatus, Neodryinus keleboensis; Acrogymnidia catalina, Ptenos amazonicus, Heteroperreyia andina, Derecyrta risaraldensis; Xorides nigrithorax, X. rufofacialis, X. simoni; Eniacomorpha hermetiae; Ecclitura shawi, Marshiella tupi, Streblocera brasiliana; Liphanthus jenamro, L. sapos, L. domeykoi, L. discolor, L. centralis, L. molavi, L. abotorabi, L. cochabambensis, L. fritzi, L. amblayensis, L. ancashensis, L. tregualemensis, L. yrigoyeni, L. sparsipunctus, L. aliavenus; Trachusa (Paraanthidium) pingdaensis, T. (P.) staabi, T. (P.) wuae; Telostholus celebes, T. rinjani, T. sulawesi; Acmaeodera (Acmaeodera) strumiai, A. (Acmaeotethya) dhofarica; Alexeter beijingensis; Microplitis bomiensis, M. paizhensis; Hoplocryptus qingdaoensis; Ponera tudigong; Brachymyrmex bahamensis, Brachymyrmex bicolor, Brachymyrmex iridescens, and Brachymyrmex sosai;
• 50 new dipterans: Drapetis abrollensis; Pieza ankh, P. aurislepus, P. bittencourti, P. parakake, P. parnasecon, P. rafaeli, P. silvanae, P. yeatesi; Pseudacteon trapeziformis; Medinodexia japonica; Ormiophasia guimaraesi, Ormiophasia seguyi, Ormiophasia crassivena, Ormiophasia manguinhos, Ormiophasia tavaresi, Ormiophasia chapulini, Ormiophasia buoculus, Ormiophasia townsendi; Anabarhynchus aurantilateralis, A. halmaturinus, A. venabrunneis; Ammophilomima amamiensis, A. rikioi; Riethia azeylandica, Riethia hamodivisa, Riethia paluma, Riethia phengari, Riethia queenslandensis, Riethia donedwardi, Riethia noongar, Riethia wazeylandica, Riethia kakadu, Riethia neocaledonica; Psectrosciara ahuatla, Psectrosciara otumba; Ptychoptera emeica ,P. circinans P. lucida, P. separata; Domodon caxiuana, D. inaculeatus, D. sensibilis; Eumerus grallator, E. tenuitarsis; Neuratelia altoandina, N. colombiana; Neophyllomyza clavipalpis, N. motuoensis, N. obtusa; Pseudasphondylia tominagai;
• 1 new trichopteran: Hydroptila nago;
• 33 new lepidopterans: Bucculatrix yingjingensis, B. liubaensis; Diduga bayartogtokhi, D. nigridentata, D. quinquicornuta, D. hanoiensis; Pleurota variocolor, P. azrouensis, P. ternaria; Zographetus rienki; Astrotischeria ochrimaculosa, A. colombiana; Phlogophora butvili; Sarbanissa pseudassimilis; Synanthedon auritinctaoidis, Paranthrenella helvola; Sabulopteryx botanica; Herpetogramma biconvexa, H. longispina, H. brachyacantha; Neosphecia cecidogena; Thraumata peruviensia; Zaphanta acuta, Z. anas, Z. bahiana, Z. beckeri, Z. elephanta, Z. elephanticula, Z. machaera, Z. rawlinsi, Z. stiletto; Antispila longcangensis, A. emeishanensis;

Actinopterygians

• 1 new anabantiform: Aenigmachanna mahabali;
• 1 new cypriniform: Paraschistura kermanensis;
• 2 new siluriforms: Parotocinclus pentakelis; Mystus prabini;
• 1 new perciform: Scolopsis lacrima;
• 1 new labriform: Cirrhilabrus wakanda;
• 1 new cyprinodontiform: Leptopanchax sanguineus;
• 2 new characiforms: Hemigrammus amacayacu; Knodus nuptialis;

Amphibians

• 9 new anurans: Microhyla fanjingshanensis; Dendrophryniscus davori, D. haddadi, D. izecksohni, D. jureia; Allobates sp.; Paratelmatobius sp.; Pristimantis ferwerdai; Nidirana yaoica; Nidirana leishanensis;

Reptiles

• 15 new squamates: Cyrtodactylus pinlaungensis; Glaucomastix itabaianensis; Ptychozoon cicakterbang, P. kabkaebin, P. tokehos; Calotes zolaiking; Hemidactylus varadgirii; Asthenodipsas jamilinaisi, A. stuebingi; Dipsadoboa montisilva; Scincella badenensis; Microgecko varaviensis; Cnemaspis tarutaoensis, Cnemaspis adangrawi; Proahaetulla antiqua;

Mammals

• 1 new primate: Mico mundukuru;

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by Piter Kehoma Boll

Look at this thing:

It is so beautifully red like a strawberry that I feel my mouth salivating and an urge to bite it. But instead of a juicy sweet fruit like a strawberry, this is a hard salty seashell belonging to the species Clanculus puniceus that has the appropriate common name of strawberry top shell.

This species is found in the Indian Ocean along the eastern coast of Africa, from the Red Sea to Cape Agulhas, including nearby islands such as Madagascar and the Mascarenes. It belongs to the family Trochidae, commonly known as top shells or top snails because their shell resembles a spinning top.

The shell of the strawberry top snail measures, in the adult, at least 15 mm in diameter, reaching up to 23 mm, and has a beautiful red color, caused by uroporphyrins, that can vary from orange-red to crimson. The spiral of the shell, when seen from above, has a line formed by black dots, caused by melanin, intercalated by two or three white dots. When seen from below, there are two additional lines with this pattern that run side by side near the shell opening.

As usual among top snails, the strawberry top snail lives in intertidal and subtidal zones and feeds on algae that it scrapes from rocks using its toothed tongue (the radula). They are dioecious, i.e., there are male and female individuals, as in most sea snails, but there is no sexual dimorphism.

Due to its beauty, the shell of the strawberry top snail is highly desired by shell collectors. However, little is known about the natural history of this particular species. I wasn’t even able to find a photograph in which the snail itself is visible.

If you work with this species or at least has a photograph of a living specimen showing the snail inside the shell, please share it! We need more available information on the wonderful creatures that share this planet with us.

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More marine snails:

Friday Fellow: Ornate Limpet (on 3 May 2019)

Friday Fellow: Tulip Cone (on 29 December 2017)

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References:

Herbert DG (1993) Revision of the Trochinae, tribe Trochini (Gastropoda: Trochidae) of southern Africa. Annals of the Natal Museum 34(2): 239–308.

Wikipedia. Trochidae. Available at < https://en.wikipedia.org/wiki/Trochidae >. Access on 29 July 2019.

Williams ST, Ito S, Wakamatsu K, Goral T, Edwards NP, Wogelius RA, Henkel T, Oliveira LFC, Maia LF, Strekopytov S, Jeffries T, Speiser DI, Marsden JT (2016) Identification of Shell Colour Pigments in Marine Snails Clanculus pharaonius and C. margaritarius (Trochoidea; Gastropoda). PLoS ONE 11(7): e0156664. doi: 10.1371/journal.pone.0156664

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by Piter Kehoma Boll

It’s time to introduce a new insect order here and, again, this is a complicated taxon. The order Trichoptera consists of small moth-like insects known as caddisflies. They are closely related to moths and butterflies, the order Lepidoptera, being a sister-group of them. Having 10 times fewer species than the order Lepidoptera, the order Trichoptera is less common and much less popular, so that it is hard to find species that are well studied to present here.

The species I picked is called Glyphotaelius pellucidus and popularly known as the mottled caddisfly or mottled sedge. It lives in middle and northern Europe and has the typical life cycle of any caddisfly.

The larva of the mottled caddisfly inhabits still and slow-running waters that are overgrown by trees, especially alders, oaks and beeches, in areas of lower altitude. As usual among caddisflies, the larva of the mottled caddisfly builds a silk case (a “caddis”) in which it lives and attaches pieces of debris, especially leaf fragments of the trees mentioned above, to make it stronger. In this species, the fragments that are attached make the case very large and characteristic. To the sides of the case, the larva attaches small and irregular leaf fragments, while to the dorsal and ventral sides, it attaches large, circular sections that are much wider than the larva’s body.

The larva lives several months, from about October to April, and feeds on leaf fragments, the same material with which it builds its case. In April, the larva turns into a pupa which, usually during summer (around June or July), turns into an adult. The adult is not aquatic as the larva and the pupa. Thus, the pupa swims to the surface before breaking and releasing the adult. During this moment, the adult is very vulnerable to predators, especially fish. This is why fake adult caddisflies are commonly employed as fishing baits.

If the adult menages to leave the water alive, it still has to spend some time waiting for its wings to dry, which is another very vulnerable moment. The color of the adult is brown and the wings have a mottled pattern of dark and light marks that makes it resemble a fragment of dried leaf.

Adult caddisflies in general rarely eat and this is not different with the mottled caddisfly. The only purpose of adults is to mate and lay eggs. After mating, the female lays the eggs in a mass on the surface of leaves hanging over a water body. One female may lay up to six egg masses, which decrease in size from the first to the last, and then dies. When the eggs hatch, the larvae fall into the water, restaring the cycle.

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References:

Crichton MI (1987) A study of egg masses of Glyphotaelius pellucidus (Retzius), (Trichoptera: Limnephilidae). In: Bournaud M., Tachet H. (eds) Proceedings of the Fifth International Symposium on Trichoptera. Series Entomologica, vol 39. Springer, Dordrecht. doi: 10.1007/978-94-009-4043-7_30

Gullefors B (2010) Seasonal decline in clutch size of the caddisfly Glyphotaelius pellucidus (Retzius) (Trichoptera: Limnephilidae). Denisia 29: 125–131.

Kiauta B, Lankhorst L (1969) The chromosomes of the caddis-fly, Glyphotaelius pellucidus (Retzius, 1783) (Trichoptera: Limnephilidae, Limnephilinae). Genetica 40: 1–6.

Otto C (1983) Behavioural and Physiological Adaptations to a Variable Habitat in Two Species of Case-Making Caddis Larvae Using Different Food. Oikos 41(2): 188–194. doi: 10.2307/3544262

Rowlands MLJ, Hansell MH (1987) Case design, construction and ontogeny of building in Glyphotaelius pellucidus caddisfly larvae. Journal of Zoology 211(2): 329–356. doi: 10.1111/j.1469-7998.1987.tb01538.x

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by Piter Kehoma Boll

If you walk through eucalyptus forests in eastern Australia, you may find today’s fellow in its natural environment. Its name is Acacia pycnantha, commonly known as the golden wattle and, as obvious by its scientific name, is a species of acacia.

The golden wattle is a peculiar tree. It reaches a height of about 8 m, although most individuals grow only up to 6 m. As common among Australian species of the genus Acacia, the golden wattle does not have true leaves. Instead, it has modified leaf stems, called phyllodes, that are widened to look and function like leaves. The phyllodes have a lanceolate and falcate shape, i.e., they look like a typical leaf that is slightly curved to one side, like a sickle. The outer side of this “sickle” has an extra-floral nectary, a structure that produces nectar and attracts insects and birds that feed on it.

The plant produces flower buds all year round but only those produced between November and May will develop further and open between July and November of the next year. The flowers occur in inflorescences and have a strong yellow color and the typical fluffy aspect of acacia flowers caused by the very long stamens.

Despite the huge amount of flowers that a single tree produces, this species is self-incompatible, meaning that it cannot fertilize itself and needs its pollen to be taken to the flowers of another individual of the same species. It has been shown that birds are very important pollinators of the golden wattle and the tree uses the extra-floral nectaries to aid that. When a bird visits the tree, it feeds on the nectar from the extra-floral nectaries and, in the process, brushed against the flowers, becoming covered with pollen. When the birds visit the next tree and brush against its flowers, part of the pollen of the first plant passes to the flowers of the second one.

The bark of the golden wattle produces large quantities of tannins, more than any other Australian acacia, which led to its cultivation for this purpose. When stressed, the trunk exudes a gum (resin) that is similar to the gum arabic produced by African species of acacia.

The golden wattle has been introduced in several other countries, especially in Europe and Africa, for ornamental or economic purposes. In South Africa, its cultivation for tannin production made it spread quickly through the native ecosystems, becoming invasive. And now, as always, we have to deal with the consequences of our irrational acts and run to solve this problem.

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References:

Hoffmann JH, Impson FAC, Moran VC, Donnelly D (2002) Biological control of invasive golden wattle trees (Acacia pycnantha) by a gall wasp, Trichilogaster sp. (Hymenoptera: Pteromalidae), in South Africa. Biological Control 25(1): 64–73. 10.1016/S1049-9644(02)00039-7

Vanstone VA, Paton DC (1988) Extrafloral Nectaries and Pollination of Acacia pycnanthaBenth by Birds. Australian Journal of Botany 36(5): 519–531. doi: 10.1071/BT9880519

Wikipedia. Acacia pycnantha. Available at < https://en.wikipedia.org/wiki/Acacia_pycnantha >. Access on 9 August 2019.

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by Piter Kehoma Boll

(First of all, I wish it were Bolsonaro, that piece of diarrhea-shaped cancer, who were dying by fire instead of the Amazon forest.)

(Now let’s go to the post itself:)

The São Francisco de Paula National Forest (FLONA-SFP) is a protected area for sustainable use in southern Brazil. Its was originally covered by Araucaria forest but currently is composed of a mosaic of the native forest and plantations of Araucaria, Pinus and Eucalyptus trees. This protection area is one of the main study areas of Unisinos’ Planarian Research Institute, where I conducted my undergradate, Master’s and PhD studies.

After studying the land planarian community of FLONA-SFP for many years, we conclude that it includes a fairly large number of species. Take a look at some of them and how cool they are:

Land planarians live in the leaf litter of forest soils and prey on other invertebrates. The 22 species shown above are the ones found in FLONA-SFP that are formally described but there are still some awaiting description. We could say that there are at least 30 different species coexisting in this protected area.

How can they all persist together? Isn’t there any sort of competition for food? Thinking of that, I conducted my master’s research investigating the diet of those and other land planarians. My results suggest that, although some species share many food items, most of them have a preferred food or an exclusive food item that could be considered what Reynoldson and Pierce (1979) called a “food refuge”.

Here is what we know about the FLONA-SFP’s species until now:

• Obama ficki feeds on slugs and snails and seems to prefer large slugs;
• Obama ladislavii feeds on slugs and snails and seems to prefer snails;
• Obama maculipunctata feeds on slugs and snails with unknown preference;
• Obama anthropophila feeds on slugs, snails and other land planarians, especially of the genus Luteostriata, and prefers the latter;
• Obama josefi apparently feeds on other land planarians only;
• All species of Luteostriata feed exclusively on woodlice;
• Species of Choeradoplana apparently feed on woodlice and harvestmen;
• Cephaloflexa araucariana apparently feeds on harvestmen;

The diet of the remaining species is still completely unknown but, based on other species of the same genera, it is likely that species of Pasipha feed on millipedes, species of Paraba feed on slugs and planarians, and Imbira guaiana feeds on earthworms.

There are plenty of different invertebrate groups that share the leaf litter with land planarians. Despite the apparently simple anatomy of these flatworms, they were able to adapt to feed on different types of prey and have muscular and pharyngeal adaptations for that. And attempt to relate anatomical adaptations to the diet of land planarians was part of my PhD research. As soon as it is published, I’ll make a post about it. There are some nice results!

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More on land planarians:

Friday Fellow: Abundant Yellow Striped Flatworm

Friday Fellow: Ladislau’s Flatworm

Darwin’s Planaria elegans: Hidden, extinct or misidentified?

How are little flatworms colored? A Geoplana vaginuloides analysis

Obama invades Europe: “Yes, we can!“

The fabulous taxonomic adventure of the genus Geoplana

The hammerhead Flatworms: Once a mess, now even messier

The New Guinea flatworm visits France: a menace

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References:

Boll PK & Leal-Zanchet AM 2015. Predation on invasive land gastropods by a Neotropical land planarian. J. Nat. Hist. 49: 983–994.

Boll PK & Leal-Zanchet AM 2016. Preference for different prey allows the coexistence of several land planarians in areas of the Atlantic Forest. Zoology 119: 162–168.

Leal-Zanchet AM & Carbayo F 2000. Fauna de Planárias Terrestres (Platyhelminthes, Tricladida, Terricola) da Floresta Nacional de São Francisco de Paula, RS, Brasil: uma análise preliminar. Acta Biologica Leopoldensia 22: 19–25.

Oliveira SM, Boll PK, Baptista V dos A, & Leal-Zanchet AM 2014. Effects of pine invasion on land planarian communities in an area covered by Araucaria moist forest. Zool. Stud. 53: 19.

Reynoldson TB & Piearce B 1979. Predation on snails by three species of triclad and its bearing on the distribution of Planaria torva in Britain. Journal of Zoology 189: 459–484.

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by Piter Kehoma Boll

Last week I presented a nice Australian acacia, the golden wattle, so today I decided to present a small creature lives on its branches. Called Sextius virescens, this insect is commonly known as the wattle horned treehopper, acaciia horned treehopper or simply green trehopper. It is a member of the order Hemiptera and the family Membracidae, commonly known as treehoppers, which are closely related to cicadas and leafhoppers.

The body of the wattle horned treehopper measures about 1 cm in length and is mostly green, but the legs are brown. There are also two horn-like projections on the thorax that have a brown to black color and another long extension of the thorax that lies over the abdomen. It lives in groups on the branches of acacia trees, with the individuals usually aligned on the branches.

As all treehoppers, the wattle horned treehopper feeds on the sap of the plants in which it lives, sucking it with its adapted mouth parts. They excrete a sweet liquid called honeydew that atracts ants. Such ants usually feed on the nectar produced by the acacia’s extra-floral nectaries and defend the tree against other herbivores. However, the wattle horned treehoppers make ants turn their attention to them instead of the plant. Delighted by the honeydew, the ants stop defending the tree and start defending the treehoppers, which is not at all good for the plant.

But that is nature. One creature always trying to explore the relations between other creatures to take the best to itself.

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References:

Buckley R (1983) Interaction between ants and membracid bugs decreases growth and seed set of host plant bearing extrafloral nectaries. Oecologia 58: 132–136.

Museums Victoria Sciences Staff (2017) Sextius virescens Green Treehopper in Museums Victoria Collections Available at <https://collections.museumvictoria.com.au/species/8561>. Access on 10 August 2019.

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by Piter Kehoma Boll

Spiders, mites, harvestmen and scorpions are the best known arachnids among the general public. However, another group that has a lot of species, even more than scorpions, is that of the pseudoscorpions. There is a very good chance that some of them are living very close to you, especially if we think of Chelifer cancroides, the house pseudoscorpion.

The name pseudoscorpion comes from the fact that these arachnids resemble scorpions, except for the lack of the tail. They are also much smaller. The house pseudoscorpion is brown and measures only about 0.5 cm in length and, as its name suggests, likes to live in human residences.

Male house pseudoscorpions defend a small territory with a radius of only a few centimeters. They allow females to enter their territory and, during the mating period, begin the courtship behavior, by which they initiate a dance and lead the female to a sperm sac (spermatophore) deposited on the ground. The female picks the spermatophore with her genital orifice and use the sperm to ferilize her eggs.

When the eggs are laid, they remain attached to the female genital pore and are covered collectively by a membrane. When the young hatch from the eggs, they are still larvae and remain inside the sac formed by the membrane covering the eggs. The female then secretes a milk-like substance from her uterus and the larvae feed on it. After molting for the first time, the larvae, now first-instar nymphs, leave the mother and, after three more moltings, reach the adult state.

Although it can pass unnoticed most of the time, the house pseudoscorpion is a cosmopolitan and common species living near and insie houses. Its pedipalps, which resemble those of scorpions, are very long and can reach almost 1 cm in length when extended. As most arachnids, they are predators, and their presence in human dwellings can be quite useful as they feed on smaller, annoying creatures, such as mites, bed bugs and booklice.

If you ever find one in your house, be kind and thank them for their service.

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References:

Harvey MS (2014) A review and redescription of the cosmopolitan pseudoscorpion Chelifer cancroides (Pseudoscorpiones: Cheliferidae). Journal of Arachnology 42: 86–104.

Levi HW (1948) Notes on the life history of the pseudoscorpion Chelifer cancroides (Linn.) (Chelonethida). Transactions of the American Microscopical Society 67(3): 290–298. doi: 10.2307/3223197

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

by Piter Kehoma Boll

Here is a list of species described this month. It certainly does not include all described species. Most information comes from the journals Mycokeys, Phytokeys, Zookeys, Phytotaxa, Zootaxa, Mycological Progress, Journal of Eukaryotic Microbiology, International Journal of Systematic and Evolutionary Microbiology, Systematic and Applied Microbiology, Zoological Journal of the Linnean Society, PeerJ, Journal of Natural History and PLoS One, as well as several journals restricted to certain taxa.

Bacteria

• 6 new actinobacteria: Micromonoscpora acroterricola; Alloscardovia theropitheci; Kribbella jiaozuonensis; Geodermatophilus marinus; Amycolatopsis eburnea; Actinomyces lilanjuaniae;
• 8 new bacteroids: Emticicia agri; Chryseobacterium mulctrae; Confluentibacter sediminis; Allopseudarcicella aquatilis; Flavobacterium cerinum; Pontibacter arcticus; Apibacter muscae; Pedobacter helvus;
• 1 new deinococcus-thermus: Deinococcus arcticus;
• 2 new firmicutes: Lentibacillus lipolyticus; Hungateiclostridium mesophilum;
• 28 new proteobacteria: Pseudomonas juntendi; Pseudomonas nosocomialis; Pseudomonas daroniae, P. dryadis; Pseudomonas mangiferae; Shimia thalassica; Marinomonas flavescens; Thermomonas aquatica; Tabrizicola alkalilacus; Pararhodobacter marinus; Undibacterium piscinae; Actinobacillus vicugnae; Rhodosalinus halophilus; Sphingomonas gilva; Pseudomethylobacillus aquaticus; Pleionea sediminis; Bradyrhizobium niftali; Hwanghaeeella grinnelliae; Ramlibacter humi; Steroidobacter soli; Dyella amyloliquefaciens; Facilibium subflavum, Cysteiniphilum halobium; Sphingorhabdus lutea; Mycolicibacterium stellerae; Paraburkholderia pallida, P. silviterrae; Thauera lacus;

Archaeans

• 2 new species: Halostella limicola; Halorientalis pallida;

SARs

• 1 new ciliate: Neobakuella aenigmatica;
• 4 new diatoms: Catenula exigua; Prestauroneis genkalii; Cocconeis carinata; Olifantiella onnuria;

Plants

• 1 new rhodophyte: Melanothamnus maniticola;
• 4 new chlorophytes: Coelastrella thermophila, C. yingshanensis, C. tenuitheca; Microrhizoidea pickettheapsiorum;
• 1 new charophyte: Nitella megacephala;
• 1 new pteridophyte: Asplenium simaoense;
• 1 new conifer: Amentotaxus hekouensis;
• 84 new angiosperms: Gastrodia amamiana; Strobilanthes tricostata; Chamaecrista falcata; Justicia longipetiolata; Portulaca almeviae; Gastrochilus yunlongense; Polysphaeria ribauensis, P. harrisii; Echeandia jaliscensis; Prosopis andicola; Hechtia ibugana; Rhodocodon petrae, Rhodocodon viridans, Rhodocodon rubescens, Rhodocodon perrieri, R. siederi; Hemiboea thanhhoensis; Dendrobium annulatum; Euphorbia rimireptans; Polycarpaea rangaiahiana; Myrciaria alta; Astragalus admirabilus; Myrcia psammophila; Tashiroea dayaoshanensis; Petunia correntina; Brachyscome lucens, Cardamine magnifica, Ranunculus callianthus, Geranium socolateum; Connarus aureus, C. tomentosus; Calamus spp.; Gluta laosensis; Isotrema sanyaense; Bulbophyllum reflexipetalum; Ceropegia jinshaensis; Disporum nanchuanense; Gentianella macrosperma; Lysimachia fanii; Marsdenia yarlungzangboensis; Aristolochia pseudoutriformis, A. yangii; Cylindrolobus motuoensis, C. glabriflorus; Yushania tongpeii; Dendrocalamus menghanensis; Henckelia nanxiheensis, H. multinervia; Gastrochilus prionophyllus; Primula dongchuanensis; Petrocosmea rhombifolia, Petrocosmea tsaii, Didymocarpus brevipedunculatus, Henckelia xinpingensis; Pedicularis multicaulis; Spiradiclis tubiflora; Cranichis callejasii, C. roldanii, C. schultesii, C. barkleyi, C. killipii, C. juajibioyi, C. neglecta, C. pennellii, C. popayanensis, C. pseudomuscosa; Inga kursarii; Rhynchospora chapadensis, Rhynchospora harleyi; Erythroxylum niziae; Fritzschia cordifolia; Serjania rosalindae, Serjania crucensis; Waltheria saundersiae; Tarenaya longicarpa; Varronia xinguana; Justicia birae, Justicia distichophylla, Justicia mcdadeana; Oxylobus coyulensis;

Amoebozoans

• 3 new mixomycetes: Tubifera glareata, T. tomentosa, T. vanderheuliae;
• 2 new discoseans: Paramoeba aparasomata, P. karteshi;

Fungi

• 8 new ascomycetes: Plectosphaerella guizhouensis, Plectosphaerella nauculaspora; Diaporthe millettiae, Diaporthe osmanthi; Memnoniella sinensis; Kazachstania jinghongensis, Kazachstania menglunensis; Verrucoconiothyrium ambiguum;
• 18 new basidiomycetes: Marasmius albisubiculosus, M. diversus, M. elaeocephaliformis, M. laranja, M. leptocephalus, M. paratrichotus, M. segregatus; Phellodon subconfluens; Lactarius viridinigrellus; Crassisporus imbricatus, C. leucoporus, C. macroporus, C.microsporus; Camarophyllopsis olivaceogrisea, Hodophilus glaberripes; Sanghuangporus toxicodendri; Sporobolomyces agrorum, Sporobolomyces sucorum;

Poriferans

• 7 new demosponges: Hamacantha (Vomerula) mamoi, Hamacantha (Vomerula) umisachii; Megaciella cardenasi, Crella (Crella) hennequinae, Hymedesmia (Hymedesmia) noramaloneae, Clathria (Clathria) priestleyae; Racekiela cresciscrystae;

Cnidarians

• 1 new myxozoan: Myxobolus parakoi;
• 6 new anthozoan: Edwardsia alternobomen; Bunga payung, Laeta waheedae, Phenganax marumi, P. subtilis, P. stokvisi;

Flatworms

• 4 new polyclads: Stylostomum mixtomaculatum; Pseudoceros agattiensis, Pseuodoceros stellans; Anthoplana antipathellae;
• 10 new proseriates: Duplominona aduncospina, D. terdigitata, D. pusilla, D. bocasana, D. dissimilispina, D. chicomendesi, D. macrocirrus, D. diademata, D. puertoricana; Pseudominona cancan;
• 1 new triclad: Matuxia tymbyra;

Annelids

• 6 new polychaetes: Trypanosyllis mercedesae; Dendronereis chipolini, Neanthes hsinchuensis; Heteromastus namhaensis, Heteromastus gusipoensis, Heteromastus koreanus;
• 4 new clitellates: Amynthas luridus, Amynthas ruiyenensis; Decimodrilus diverticulatus, D. globulatus;

Mollusks

• 1 new bivalve: Montacutona sigalionidcola;
• 3 new gastropods: Moridilla jobeli; Sinoxychilus melanoleucus; Ganesella halabalah;

Nematodes

• 10 new species: Ditylenchus paraparvus; Aphanolaimus strilliae, Makatinus africanus; Metarhabditis giennensis; Thalassironus filiformis; Paratylencholaimus sanshaensis, Tylencholaimus zhongshanensis, Dorylaimoides shapotouensis; Philometra ostorhinchi, Philometra tenuis;

Arachnids

• 14 new mites: Lorryia meliponarum, Melissotydeus bipunctata; Kokoppia lagunaensis, Setoppia parrillarensis, Graptoppia (Stenoppia) magallanesensis; Haplozetes bayartogtokhi, Magyaria leonilae; Geckobia andina, Geckobia circumdata; Unionicola (Unionicola) maolanensis, Unionicola (Unionicola) xishuiensis, Unionicola (Unionicola) suiyangensis; Ixodes barkeri; Laelaps schatzi;
• 1 new palpigrade: Eukoenenia ibitipoca;
• 1 new scorpion: Tityus guane;
• 2 new pseudoscorpions: Parobisium magangensis, P. yuantongi;
• 27 new spiders: Buitinga qingyuani, B. wamitii, Smeringopus voi; Pholcus mixiaoqii; Onocolus ankeri; Artoria weiwei; Ochyrocera tinocoi, Ochyrocera garayae, Ochyrocera itatinga; Mago brimodes; Thaiderces shuzi, T. peterjaegeri, T. ganlan, T. ngalauindahensis, T. yangcong, T.zuichun, T. miantiao, T. jiazi, T.tuoyuan, T. fengniao, T. haima, T. chujiao, T. thamphadaengensis, T. thamprikensis; Falcileptoneta baegunsanensis, F. odaesanensis, F. umyeonsanensis;

Myriapods

• 1 new chilopod: Geophilus serbicus;
• 6 new diplopods: Hyleoglomeris hoanglien, H. fanxipan; Coloradesmus hopkinsae, C. manitou, C. beckleyi, C. warneri;

Crustaceans

• 2 new ostracods: Ilyocypris incus; Xylocythere sarrazinae;
• 1 new branchiopod: Lynceus huentelauquensis;
• 5 new copepods: Panaietis bobocephala, Panaietis flavellata; Leptotachidia senaria, L. apousia; Duoergasilus basilongus;
• 1 new cirripede: Membranobalanus porphyrophilus;
• 5 new tanaids: Hamatipeda prolata, Meromonakantha mauri, Paratyphlotanais apletos, P. bessai; Hargeria chetumalensis;
• 1 new amphipod: Ceradocus kiiensis;
• 12 new isopods: Alpioniscus lossinii, A. drazinai, A. mandalinae; Alpioniscus hirci, Alpioniscus velebiticus; Ligia dante, L. eleluensis, L. honu, L. kamehameha, L. mauinuiensis, L. pele, L. rolliensis;
• 10 new decapods: Cambarus fetzneri; Salmoneus venustus; Salmoneus durisi; Salmoneus tiburon; Alpheus perlas; Caridina malanda; Paralebbeus pegasus; Axianassa planioculus; Mediapotamon liboense;

Hexapods

• 4 new collembolans: Oligaphorura wanglangensis, O. ussurica, O. kedroviensis; Sinhomidia uniseta;
• 2 new archaeognathans: Coryphophthalmus obscurus, C. serbicus;
• 3 new odonates: Gomphidia podhigai; Dactylobasis demarmelsi; Cyclogomphus flavoannulatus;
• 3 new ephemeropterans: Camelobaetidius metae; Oligoneuriella tuberculata, Oligoneuriopsis villosus;
• 2 new dictyopterans: Quadrihormetica onorei, Lucihormetica yasuniana;
• 21 new orthopterans: Alloteratura (Alloteratura) sagittala, Alloteratura (Alloteratura) flabellata, Alloteratura (Meconemopsis) pentadactyla; Kuzicus (Kuzicus) bicurvus; Xizicus (Eoxizicus) lobicercus; Anisophya una, Xenicola taroba, Xenicola xukrixi; Criotettix zhejiangensis; Xestophrys namtseringa; Tachycines (Tachycines) huaxi; Mimicogryllus splendens, Pteroplistes bruneiensis, Tembelingiola belaitensis, Vescelia sepilokensis; Rhytidaspis arfak, R. camela, R. genyem, R. nigropunctata, R. ornata, R. variata;
• 10 new plecopterans: Perlodinella mazehaoi; Isoperla qinlinga; Hemacroneuria elongata, H. ovalis; Neoperla shiwandashana; Peltoperlopsis mengmanensis; Phanoperla zwicki; Soyedina sheldoni; Indonemoura bifida; Chrysoperla duellii;
• 1 new thysanopteran: Stictothrips farsi;
• 71 new hemipterans: Mesovelia brevia, M. dilatata, M. occulta, Mesovelia andamana, M. bispinosa, M. isiasi, M. tenuia; Rubrocuneocoris vietnamensis; Empoasca giusana, E. palgongsana; Bagauda atypicus; Calodia deergha, C. keralica, C. kumari, C. neofusca, C. periyari, C. tridenta, Glaberana acuta, G. purva, Olidiana lanceolata, O. flectheri, O. umroensis, O. unidenta, Singillatus parapectitus, S. serratispatulatus, Trinoridia dialata, T. ochrocephala, T. piperica, T. ramamurthyi, T. saraikela, T. timlivana, W. andamana, W. burmanica, Zhangolidia weicongi; Tympanoterpes virgulata, Cracenpsaltria nana, Guyalna dasyeia, Guyalna fasciata, Guyalna polypaga, Parnisa santacruzensis, Carineta ensifera, Carineta hamata, Carineta pictilis, Carineta uncinata, Herrera concolor, Herrera freiae, Herrera melanomesocranon, Herrera phyllodes, Herrera signifera; Aphis ingeborgae, Aphis conspicua, Aphis fuentesi; Tynacantha cuprea, T. umeridenigrata; Naratettix cheondungsanus; Melaniphax suffusculus; Kosmiomiris carvalhoi; Mariomella singularis; Bambusiphaga unispina; Hoplonannus bifidus, Hoplonannus robustus; Cyrta webbi; Sinonissus daozhenensis, S. hamulatus, S. longicaudus; Saissetia kunmingensis; Malingeus tumidus, Yaramayahus rufescens; Cladonota erwini; Hebetica sylviae; Calopsaltria luteotorquata;
• 47 new coleopterans: Cylloepus bispoi, C. segurae; Agonotrechus spinangulus, Sinotrechiama yunnanus; Oodescelis (Planoodescelis) lii; Tychus meggiolaroi; Incoltorrida benesculpta, I. galoko, I. magna, I. marojejy, I. quintacostata, I. zahamena, Hydroscapha andringitra; Metallactus abditus, M. chamorroi, M. dicaprioi, M. madefactus, M. praetorius, M. viator; Aenigmoronia echinodes, Australycra similis, Austronitidula multimaculata, Cychramus splendidus, Lasiodites howensis, Temnelytron nigrum; Pteroplatus luculentus; Adelopsis diabolica; Brachysomus (Hippomias) tannourinensis; Clidicus forceps, C. interfector, C. kalis, C. occisor, C. shavrini; Copelatus ankaratra, Copelatus kely, Copelatus pseudostriatus, Copelatus safiotra, Copelatus vokoka; Copelatus amphibius, Copelatus betampona, Copelatus zanatanensis, Madaglymbus kelimaso, Madaglymbus menalamba, Madaglymbus semifactus; Psiloibidion boteroi,Tropidion neisi; Xenocona antonkozlovi; Ceratochodaeus darlingi;
• 1 new megalopteran: Neochauliodes flinti;
• 43 new hymenopterans: Eucera dafnii, E. wattsi; Strumigenys caiman, S. economoi, S. hubbewatyorum, S. zemi; Kudakrumia malaenglek, Kudakrumia loaibonho, Krombeinella brothersi; Bathanthidium fengkaiense; Coryptilus circalatus, Coryptilus longicervix; Mischocyttarus nazgul, M. asahi, M. kallindusfloren; Austerocardiochiles mellosus, A. simulatus; Nylanderia bibadia, N. caerula, N. coveri, N. disatra, N. esperanza, N. fuscaspecula, N. lucayana, N. metacista, N. pini, N. semitincta, N. sierra, N. wardi, N. xestonota, N. zaminyops; Clistopyga albovittata, C. lapacensis and C. speculata; Dryinus georgianus; Bombus (Pyrobombus) interacti; Epigeiobracon perplexus; Ephutomorpha tyla; Neptihormius dipterophagus, Neptihormius whakapapa, N. herbaspicatum; Diodontus polytylus, Diodontus guichardi;
• 94 new dipterans: Leucotabanus fairchildi; Simulium (Gomphostilbia) marosense; Curtonotum abrelatas, C. irksum, C. notatum, C. prolixum, C. transitus; Maruina (Aculcina) roraimensis, Maruina (Maruina) kallyntrona, M. (M.) mystax, M. (A.) pila; Pseudoaerumnosa acinacea, P. ampliata, P. annae, P. awanensis, P. banari, P. clavidactyla, P. clivicola, P. collicola, P. consuota, P. cryptoloba, P. curvifalx, P. eminula, P. exacuta, P. filispicata, P. formosa, P. fragilis, P. impensa, P. inviolata Rudzinski, 2006, P. junciseta, P. obovata, P. pilicaudata, P. quadriquetra, P. saginata, P. tenuidens, P. tkoci; Gloma pyricornis; Chaetonerius serratus; Leinendera carrerai, Leinendera mnrj; Laneella fuscosquamata, Laneella purpurea, Mesembrinella bullata, Mesembrinella chantryi, Mesembrinella epandrioaurantia, Mesembrinella guaramacalensis, Mesembrinella longicercus, Mesembrinella mexicana, Mesembrinella nigrocoerulea, Mesembrinella serrata, Mesembrinella velasquezae, Mesembrinella violacea, Mesembrinella woodorum, Mesembrinella zurquiensis; Agromyza princei, Melanagromyza vanderlindeni, Ophiomyia antennariae, O. osmorhizae, Calycomyza smallanthi, Liriomyza euphorbiella, L. garryae, L. phloxiphaga, Phytomyza nemophilae, P. salviarum; Rhabdomastix borroloola, R. dobrotworskyi, R. dooragana, R. hirsuta, R. nivalis, R. ponticulus, R. collessiana, R. rosae; Antocha (Antocha) bella; Mycodiplosis puccinivora; Xylota hauseri, X. orientiflorum, X. xanthotarsis; Cyrtopogon hollandi, C. martini, Stenopogon graminis; Tipula (Vestiplex) butvilai; Molophilus (Molophilus) rohaceki, M. (M.) soldani; Claraeola bousynterga, C. heidiae, C. khuzestanensis, C. mantisphalliga; Cladochaeta amorimi; Polletella polleti, Polletella norrbomi, Polletella inca; Aedes (Ochlerotatus) amateuri, Aedes (Protomacleaya) lewnielseni; Neolasioptera imprimata;
• 7 new trichopterans: Rhyacophila longicaudata, Rhyacophila aksornkoaei, R. limsakuli, Rhyacophila kengtungensis; Ascotrichia adirecta, A. hystricosa, A. simoma;
• 42 new lepidopterans: Organopoda acutula, O. deltaformis, O. megiste; Herminia yuksam, H. borneo, H. amamioshima; Toccolosida nigraregina; Argyrophorus idealis, A. rubrostriata; Ancylosis larissae; Cyana lada, C. titovi; Eupoecilia jakarana, E. gedui, Lumaria phuntschona, Borneogena trashiyana,Bactra cophinana, Penthostola subnigrantis, M. brunnofasciana, Peridaedala nigrifasciana, Epiblema albulusana; Homaloxestis arcuatus, H. lactizonalis; Fujimacia cornutiprocera, F. dayaoensis, F. gracilenta, F. longispinosa; Cnephasia razowskii; Meleonoma foliiformis, M. projecta; Calisto gundlachi, Calisto siguanensis, Calisto disjunctus, Calisto sharkeyae, Calisto lastrai; Grapholita chytranthusi; Tentamen atrivirgulatum, Longistrebula argalata, Tectosquamae furvalata, Tectosquamae poguei, Tectosquamae solisae; Sufetula anania;

Echinoderms

• 2 new holothuroids: Sclerothyone reichi, Sclerothyone oloughlini;

Actinopterygians

• 2 new anguilliforms: Gymnothorax smithi; Gymnothorax andamanensis;
• 1 new aulopiform: Arctozenus australis;
• 1 new characiform: Pristella ariporo;
• 3 new cypriniforms: Garra simbalbaraensis; Neobola kinondo; Cabdio crassus;
• 2 new gobiiforms: Gymnesigobius medits; Luciogobius yubai;
• 1 new gymnotifom: Eigenmannia sirius;
• 3 new perciforms: Trachinotus macrospilus; Johnius taiwanensis; Branchiostegus biendong;
• 5 new siluriforms: Leiocassis rudicula; Chiloglanis mongoensis; Glyptothorax gopii; Synodontis denticulatus; Trichomycterus astromycterus;

Anurans

• 17 new anurans: Cophyla fortuna; Spinomantis mirus; Ansonia kyaiktiyoensis; Pristimantis nelsongalloi; P. altillo, P. chomskyi, P. gloria, P. jimenezi, P. lutzae, P. multicolor, P. nangaritza, P. teslai, P. torresi, P. totoroi, P. verrucolatus; Noblella naturetrekii; Scinax sp.;
• 3 new caudates: Hynobius guttatus, H. tsurugiensis, H. kuishiensis;

Reptiles

• 15 new squamates: Lepidodactylus agnanus, L. dialeukos, L. zweifeli, L. mitchelli, L. kwasnickae; Helicops boitata; Cnemaspis amba, C. koynaensis; Cnemaspis aaronbaueri; Cnemaspis tanintharyi, Cnemaspis thayawthadangyi; Bothrops monsignifer; Larutia kecil; Takydromus yunkaiensis; Epictia rioignis;

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