Help needed with DNA barcoding of European tachinids

Tachinid Times #26 has some very interesting articles on a few projects around the world to compile DNA phylogenies of tachinids. One such is by my good friend Jaakko Pohjoismäki, who is looking at European tachinids. He has a few genera and species that he is having trouble locating so I am listing them here in case anyone can help:

Exoristinae

Admontia seria (Meigen)

Bessa parallela (Meigen)

Carcelia puberula Mesnil, C. tibialis (R.-D.)

Chetogena apart from C. tschorsnigi Ziegler

Drino atropivora (R.-D.), D. bohemica Mesnil, D. gilva (Hartig)

Erycia apart from E. fatua (Meigen)

All Exorista sg. Adenia apart from E. rustica (Fallén) Exorista fasciata (Fallén)

All Istocheta spp.

Ligeriella aristata (Villeneuve)

Medina luctuosa (Meigen)

Myxexoristops arctica (Zetterstedt), M. bonsdorffi (Zetterstedt)

Oswaldia eggeri (B. & B.), O. reducta (Villeneuve)

Phebellia clavellariae (B. & B.)

Policheta unicolor (Fallén)

Senometopia confundens (Rondani), S. intermedia (Herting), S. lena(Richter)

Thecocarcelia acutangulata (Macquart)

Vibrissina turrita (Meigen)

Winthemia erythrura (Meigen), W. venusta (Meigen)

 

Tachininae

Actia infantula (Zetterstedt), A. maksymovi Mesnil

All Anthomyiopsis spp.

All Aphantorhaphopsis spp.

Ceranthia pallida Herting, C. tristella Herting, C. verneri Andersen

Ceromya dorsigera Herting, C. flaviceps (Ratzeburg), C. flaviseta (Villeneuve)

Cleonice keteli Ziegler, C. nitidiuscula (Zetterstedt)

All Germaria spp.

All Graphogaster spp.

Linnaemya haemorrhoidalis (Fallén), L. olsufjevi Zimin, L. rossica Zimin

Macroprosopa atrata (Fallén)

Panzeria laevigata (Meigen), P. vagans (Meigen)

Peleteria ferina (Zetterstedt), P. popelii (Portshinsky)

Peribaea longirostris Andersen, P. setinervis (Thomson)

Phytomyptera nigrina (Meigen), P. riedeli (Villeneuve), P. vaccinii Sintenis

Siphona grandistylum Pandellé, S. hungarica Andersen, S. immaculata Andersen, S. variata Andersen

 

Dexiinae

Billaea fortis (Rondani)

Blepharomyia piliceps (Zetterstedt)

Dexia vacua (Fallén)

All Pandelleia spp.

All Rondania spp.

Stomina tachinoides (Fallén)

Villanovia villicornis (Zetterstedt)

 

Phasiinae

Besseria melanura (Meigen)

Opesia cana (Meigen)

All Strongygaster spp.

The specimens should preferably be collected in the 2000s, but as noted earlier we welcome also older samples. Dry and ethanol-preserved material are both acceptable. Ideally, we would like to borrow the whole specimen for documentation purposes. If you do not wish to donate the specimen, then it will be returned after sampling (removing a leg) and documenting together with a label, which helps to connect the specimen with the barcode in the future. Please feel free to contact us with your thoughts and suggestions. We’re hoping to hear from you!

How to find the prosternum

The prosternum can be quite a difficult feature to find on a tachinid – partly because many British workers, used to Belshaw’s key will never have had to find it unless they have used the European or Palearctic keys. Here is a nice photo and the prosternum is the thin, dark, vertical strip of chitin in the middle of the picture, under the flies chin and between and slightly in front of the front coxae. Either side of it is a membranous area.

The keys nearly always ask whether the prosternum is hairy or bare – in the following photo the prosternum is bare but hairy examples have 1 or more fine, black hairs along the lateral edges.

Thelaira solivaga (male) showing the prosternum feature, which in this case is bare.

… and here is an example with a hairy prosternum – it is very difficult to see clearly but you should just be able to see small dark hairs towards the edge of the prosternum.

Exorista rustica (male) showing the prosternum, which in this case is hairy.

A combined image, zoomed in to show the hairs or lack of. Try to ignore the bristles on the coxae – under a microscope it is a little easier because you have a greater perception of 3-dimensions and the ability to move the specimen to the best orientation:

One of the big impediments to viewing the prosternum is usually poor pinning/mounting, which allows the head to drop or the front coxae to close together and block the prosternum from view. I have noticed this most often when specimens are pinned directly and dorsoventrally. If you pin a specimen laterally the head is less likely to drop down and if, in the rare event that it isn’t in a good position, you can easily manipulate it and use micro-pins to hold the specimen in position while it dries.

Distinguishing species of Thelaira

In my experience a lot of people have trouble with this genus because the traditional features used in keys are quite variable. In particular it is difficult to separate the common Thelaira nigripes from the rare solivaga.

All keys use the size of the outer-vertical bristles and the anterodorsal bristles on the middle tibia, which are a bit variable, and Belshaw (1993) uses the colour of the abdomen, which is one of the worst features to rely on – it is a confirmatory feature at best. This leads to a lot of Thelaira nigripes being wrongly classified as Thelaira solivaga – the 2 species are very similar but they can be distinguished if the correct features are used.

Belshaw is a little ambiguous when he talks of the middle leg  only having “2 long bristles on its anterodorsal surface”. What constitutes “long”? All Thelaira have both long and short bristles on the ad surface of the mid tibia but I prefer to ignore the smaller ones at the top and bottom of the tibia and focus on the rest, in the middle. Here are some photos of Thelaira nigripes tibiae:

Here is a photo of a Thelaira solivaga mid tibia:

Also, a very useful approach is to look at the male genitalia – here is a nice set of figures from an article I saw a while ago:

43Thelaira leucozona (Panzer, 1809); 44Thelaira solivaga (Harris, 1780); 45Thelaira nigripes (Fabricius, 1794); 46Phenicellia haematodes (Meigen, 1824) (not British).

Pay particular attention to the segment just before the genitalia themselves and the way that the surstylus curves. The distinction between nigripes and solivaga is still very fine but in conjunction with the other features you should be able to make a more confident determination. Later I will add some closeup photos of the male genitalia.

Eurithia – female sternite 6

These are some figures from Tschorsnig & Herting (1994), showing the sternite 6 of some female Eurithia spp.:

AEurithia consobrina (furrow along entire length)

BEurithia connivens (furrow on only anterior half)

CEurithia vivida & intermedia (anterior indentation)

Unfigured (sternite domed) – Eurithia caesia & anthophila (slightly)

Big Nature Day

Today Matt & I attended “Big Nature Day” at the Natural History Museum, in London – a day for wildlife organisations to show what they do. The weather was  hot & sunny and the event was very colourful and well-attended, with lots of events for children.

Many thanks to Lucy Carter and all those that helped organise the event and make it so successful.

Some name changes

Belshaw’s 1993 handbook was a great improvement over the nomenclature used in van Emden but since then many of the names have changed. This is pretty boring stuff, so I apologise in advance, but here are the changes and explanations, as far as I know them:

  • Timavia Robineau-Desvoidy 1863 is a junior synonym of Smidtia Robineau-Desvoidy 1830 so Timavia amoena becomes Smidtia amoena Meigen 1824.
  • Erycilla Mesnil 1957 is a junior synonym of Allophorocera Hendel 1901 so Erycilla ferruginea becomes Allophorocera ferruginea (Meigen 1824).
  • Chrysocosmius Bezzi, 1907 not a valid genus name by ICZN rules (complicated story) so Chrysosomopsis Townsend, 1916 is used instead, which means that Chrysocosmius aurata becomes Chrysosomopsis aurata (Fallen 1820).
  • Microsoma exigua Meigen 1824 is corrected to Microsoma exiguum Meigen 1824.
  • Actia nudibasis Stein 1924 is a junior synonym of Actia resinellae Schrank 1781, which replaces it.
  • Peribaea fissicornis Strobl 1909 is a junior synonym of Peribaea setinervis Thomson 1869, which replaces it.
  • Siphona mesnili Andersen 1982 is a junior synonym of Siphona confusa Mesnil 1961, which replaces it.
  • Ernestia Robineau-Desvoidy 1830 is a junior synonym of Panzeria Robineau-Desvoidy, 1830 – this has quite a far-reaching effect because Ernestia puparumrudis, vagans & laevigata all move to Panzeria
  • Cyrtophleba Rondani, 1856 is corrected to Cyrtophloeba Rondani, 1856 following the rule of the first reviser. See explanation in Botria (below).

Taxa not in Belshaw:

  • Bothria Rondani, 1868 is corrected to Botria Rondani, 1856 following the rule of the first reviser. Rondani was very poor at forming linguistically correct names and used many different variations of spelling – often in the same publication, which causes the initial confusion. The next time he used one of those spellings he became the “first reviser” under ICZN rules because he chose the “correct” spelling and so it is this that we have to use. For more information on this issue see: O’Hara, Cerretti, Pape & Evenhuis (2011) Nomenclatural Studies Toward a World List of Diptera Genus-Group Names. Part II: Camillo Rondani.
This website will be changed to reflect these changes (many have been done already) but I am holding back for a little while with some until I work out the best way to do this without confusing people. In particular the changes from Ernestia to Panzeria are potentially quite confusing.

Panzeria, Eurithia, Appendicia & Fausta male genitalia

This is a reprint of the genitalia figures from van Emden’s 1957 handbook – out of print for many years. I find them quite useful for double-checking males – if I get good feedback then I might turn this into a larger article.

For reference the key to this diagram is:

PPanzeria laevigata, APanzeria rudis, BPanzeria vagans, IFausta nemorum, JAppendicia truncata, KEurithia anthophila, LEurithia caesia, QEurithia intermedia, REurithia connivens, SEurithia vivida, TEurithia consobrina

sf = superior forceps = cercus, if = inferior forceps = surstylus

Botria subalpina found in Scotland

Botria (=Bothria) subalpina is a Spring species previously thought to be found in northern & eastern Europe – I have specimens from Finland and Bavaria. But recently Murdo Macdonald sent some flies to the National Museum of Scotland and David Horsfield spotted a fly that keyed out to Bothria subalpina in the Central European key. He sent the specimen to Hans-Peter Tschorsnig in Germany who confirmed the identification!

Apparently the name has changed from Bothria to Botria due to a Rondani misspelling that has only recently come to light. Also Villeneuve wrote in his 1910 type description for subalpina that he had been sent a specimen taken near Birmingham by Wainwright. It would be interesting to track down this specimen and confirm the identification and locality.

Here is a photo of a specimen from Finland and a very bad photo of one from Germany:

Currently, in Belshaw, this species would get lost around couplet #27  because it looks like it should go to #28 (Phorocera/Parasetigena) but the basals are between 2-3x as long as the scutellum so it is weak here and of course if you go to #28 it isn’t Parasetigena because Botria has median discals and it isn’t Phorocera because the male genitalia are all wrong (smaller than Phorocera) and it has pale tibiae. I think the pale tibiae are probably the best way to split Botria out but I need to double-check all of the other alternatives from #27 onwards.

Phytomyptera nigrina seen in Lincolnshire

Had a very good record recently, sent in by Phil Porter, of Phytomyptera nigrina. Richard Davidson took the specimen at Whisby Nature Park on 22nd April. Here is a photo of the specimen – note the “disappearing” median vein and the complete lack of m-cu vein – very rare features:

The first photos through the Leica S8APO

It has taken a while to get the camera hooked up to the microscope because it is quite a complicated process, involving lots of adapters and converters – not to mention getting the optics and the extension tubes correctly arranged. The first shots really weren’t worth looking at but today I started to capture images that are getting a bit better – still not the quality I hope to achieve, but getting there.

Although the microscope is rated at 80x the actual magnification through the microscope tube is related to the size of the sensor and this works out at about 10x – 15x but this is still very good and a perfect range for what I want to do. The lighting in these photos is very rough and contrasty because I haven’t experimented with flash diffusers yet but it gives you an idea of what is possible.

This is a butterfly wing (Eunica coelina):

This is a lateral view of Dinera grisecens, showing a bit of the katepisternum; a nice row of hypopleural bristles; the hind spiracle with hairy flap; haltere and abdominal tergite 1+2:

Contrast that with the spiracle of Exorista rustica, which has a “classic” single-flap arrangement:

Exorista rustica spiracle - showing the single flap

… and the next 2 show classic polideine spiracles, with 2 equally-sized flaps:

Lypha dubia spiracle - showing the 2 equally sized flaps

Lydina aenea spiracle - showing the equally sized flaps

I hope to make a few more photos over the coming days, once I have worked out the problem with parfocality – the camera should be in focus at the same point as the main microscope eyepieces but at the moment it isn’t … a bit more experimentation needed! 😉