Asiloid Flies

Field-work in South Africa and Namibia Sep 2019

Before we turn into 2020, I am taking this moment to summarize my field trip to South Africa and Namibia with #TeamDiptera member Allan Cabrero in September 2019.

This trip included a visit to the amazing Diptera collection at the KwaZulu-Natal Museum (NMSA) and conduct field-work in coastal areas of eastern South Africa and the Namib Desert in Namibia at the Ameib Private Nature Reserve (web-site) and the Gobabeb Namib Research Institute (gobabeb.org).

I would like to thank Jason Londt and John Midgley from the NMSA for allowing us to collect under their Ezemvelo KZN Wildlife permits in KwaZulu-Natal and Tricia Pillay for access to the collection at the NMSA. In Namibia, I would like to thank the Namibian National Commission on Research, Science and Technology (NCRST) and the Ministry of Environment and Tourism (MET) for issuing collecting and export permits. Furthermore, the staff at the Gobabeb Namib Research Institute was very helpful as always and in particular, Gillian Maggs-Kölling, Leena Kapulwa, and Eugene Marais assisted in many ways.

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Here are some of the habitats we visited:

1. Coastal forest at Umlalazi Nature Reserve (KwaZulu-Natal, South Africa) where Ommatius sp. (Asilidae) was collected.

2. Leeward side of vegetated coastal dunes at Umlalazi Nature Reserve where Euscelidia natalensis Dikow, 2003 (Asilidae) was collected.

3. Dry, open Acacia bushveld at Ameib Private Nature Reserve (Erongo, Namibia) where Anasillomos chrysopos Londt, 1983, Sisyrnodytes sp. (see photo below), Afroscleropogon sp., and Empodiodes namibiensis Londt, 2012 (see photo below) (all Asilidae) were collected.

4. Dry, open Acacia bushland at ≠Gaingu Conservancy with Grosse Spitzkoppe in the back (Erongo, Namibia) where Anasillomos chrysopos Londt, 1983 and Remotomyia brunales Londt, 1983 (see photo below) were collected. The ≠ symbol in the reserve name denotes a click sound in the Khoekhoegowab language.

5. Small dunes along dry Kuiseb riverbed in Namib-Naukluft National Park at Gobabeb Namib Research Institute (Erongo, Namibia) where Acnephalomyia sp. (Asilidae) and an undescribed genus of Mydidae were collected.

6. Hope Mine Wash in Namib-Naukluft National Park near Gobabeb were Anasillomos chrysopos Londt, 1983, Corymyia sp., and an unidentified genus of Willistonininae (all Asilidae) were collected. This is the 1st record of the genus Corymyia from Namibia – all four known species occur in western South Africa. The Willistonininae specimens are likewise extremely interesting as they do not fit into the currently known Afrotropical genera – more work needs to be done on these specimens!

7–8. Kahani Dunes in Namib-Naukluft National Park near Gobabeb were Stichopogon hermanni Bezzi, 1910, Sporadothrix sp., Torasilus sp., and an undescribed genus of Mydidae were collected. Photo 8 shows a dry leaf of Stipagrostis sabulicola (Poaceae), which was used as a perch by Sporadothrix sp. If you look closely, you can see two impressions from the rim of my collecting net on the sand – I caught one fly and walked on the dune for a few minutes before I circled back to this specific site. A 2nd fly was now sitting on the same perch and was collected.

9. An unnamed dune near Gobabeb with Allan catching flies. No asiloid flies were collected here, but a number of other Diptera including large flesh flies (Sarcophagidae, see photo below).

10. Dry, very wide Kuiseb riverbed near Swartbank in Namib-Naukluft National Park north of Gobabeb where an undescribed genus of Mydidae and a species of Therevidae were collected.

11. Dry, wide Kuiseb riverbed and small sand dunes at a field-site 20 km N of Gobabeb in the Namib-Naukluft National Park where an undescribed genus of Mydidae was collected (see photo below).

12. For the 4th time, I visited this site in the Namib Desert at the eastern-most edge of the Namib sand sea on the C14 north of Solitaire (Khomas, Namibia) in the Namib-Naukluft National Park and collected Stichopogon hermanni Bezzi, 1910, Anypodetus sp., and Afroholopgon sp. (all Asilidae). The Afroholopogon species is especially interesting and was collected for the 1st time here and likely is a new species.

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I was able to collect the following genera / species (I still need to identify some taxa to the species level):

Apioceridae: –
Asilidae: Acnephalomyia sp., Afroholopgon sp., Afroscleropogon sp., Anasillomos chrysopos Londt, 1983, Anypodetus sp., Corymyia sp., Empodiodes namibiensis Londt, 2012, Euscelidia natalensis Dikow, 2003, Ommatius sp., Remotomyia brunales Londt, 1983, Sisyrnodytes sp., Sporadothrix sp., Stichopogon hermanni Bezzi, 1910, and Torasilus sp.
Mydidae: undescribed genus.

I was also able to collect several species of bee flies (Bombyliidae), micro bee flies (Mythicomyiidae), hover flies (Syrphidae), shore flies (Ephydridae), quasimodo flies (Curtonotidae), house flies (Muscidae), flesh flies (Sarcophagidae), stiletto flies (Therevidae), and horse flies (Tabanidae).

Below are some of the photographs of flies in nature taken with an Olympus OM-D EM-5 Mark II and a f/2.8 60 mm macro-lens (= 120 mm focal length in 35 mm photography):

13. Crane-fly species Maekistocera filipes filipes (Fabricius, 1805) (Tipulidae) basking on a tree trunk in Umlalazi Nature Reserve (KwaZulu-Natal, South Africa). See iNaturalist observation: 63658122. (identification by Jon Gelhaus.)

14. Mating pair of Ephydridae (shore flies) at Tugela River mouth (KwaZulu-Natal, South Africa). See iNaturalist observation: 63672413.

15. Sisyrnodytes sp. (Asilidae) at Ameib Private Nature Reserve (Erongo, Namibia) feeding on a plant bug. See iNaturalist observation: 63672673.

16. Unfortunately, I didn't get a better photo of this small Stenopogoninae species (Asilidae). I only saw this female fly and collected another female. After studying the specimen in the lab for a considerable time, I was able to identify it as Empodiodes namibiensis Londt, 2012. I believe this record is the southernmost record of the primarily northern Namibian species (see Londt 2019 doi:10.3897/AfrInvertebr.60.33075 Fig. 16 (/article/33075/element/2/110/)).

17–18. Remotomyia brunales Londt, 1983 (Asilidae) at ≠Gaingu Conservancy and Grosse Spitzkoppe area. These flies were very fast and I wasn't able to get very close – photo 18 is a crop of 17 to show fly more clearly in the center of the photo. This is my 1st time collecting flies of this genus on which I published in one of my 1st articles with Jason Londt in 2000 (doi:10.5281/zenodo.11581).

19. Flesh-fly species Khowaba atrox Pape, 1991 (Sarcophagidae) resting upside down on Stipagrostis sabulica (Poaceae) on high dunes near Gobabeb. See iNaturalist observation: 63291181. (identification by Thomas Pape and Daniel Whitmore.)

20–21. Undescribed genus and species of Mydidae (Syllegomydinae) at field-site at Kuiseb riverbed 20 km N Gobabeb. Photo 20 shows an overview of where the fly is resting (in center of photo) and since I was able to get closer and closer without disturbing the fly, I was able to take a close-up as well. This photo also showcases that even if you get just a few centimeters above the ground, you can rest in much cooler environments. From crouching down to photograph this (and other insects) during exceptionally hot unseasonal weather, my knees got extremely hot on the sand ...

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Other impressions and encounters in South Africa and Namibia:

22–24. The KwaZulu-Natal Museum in Pietermaritzburg (South Africa) is a great place to see wonderful exhibits including Diptera, meet up with old friends and colleagues, and study flies in the amazing collection, which is undoubtedly the most important collection of true flies in Africa. I first worked at the NMSA when I was an international student at the University of KwaZulu-Natal studying entomology and spent every free hour I had at the collection 20 years ago. This is where my taxonomic research on assassin flies really began under the tutoring of Jason Londt.

25. Jason Londt and I at Doreen Clark Nature Reserve just outside Pietermaritzburg (KwaZulu-Natal, South Africa, photo by Allan Cabrero).

26. Same as above with Allan Cabrero (photo by Jason Londt).

27. A free-living antlion larva (Neuroptera: Myrmeleontidae) on coastal dunes at Umlalazi Nature Reserve (KwaZulu-Natal, South Africa). See iNaturalist observation: 63658711.

28–30. Beautiful subtropical, coastal forest and mangrove at Umlalazi Nature Reserve (KwaZulu-Natal, South Africa).

31–32. Catching flies with giraffes (Giraffa camelopardalis Linnaeus, 1758) and seeing ancient rock paintings at Ameib Private Nature Reserve (Erongo, Namibia).

33. A Common Namib Day Gecko (Rhoptropus afer Peters, 1869, Gekkonidae) hiding in shade under a rock in dry wash near Gobabeb (Erongo, Namibia). It was extremely hot at this field-site!

34. Solifuge Solpugiba lineata (C.L.Koch, 1842) (Arachnida: Solifugae: Solpugidae) at Kahani Dunes near Gobabeb (Erongo, Namibia). See iNaturalist observation: 63673759. (identification by Dewald du Plessis.)

35. One of the highlights of this trip was to see a Namaqua Chameleon (Chamaeleo namaquensis Smith, 1831) on the dunes at the eastern-most edge of the Namib sand sea on the C14 north of Solitaire (Khomas, Namibia).

36. Myself and beautiful Euphorbia sp. at Hope Mine Wash in central Namib Desert (photo by Allan Cabrero).

37–38. It's always great to be back at the Gobabeb Namib Research Intitute in the Namib Desert.

39. Before heading back home, it is advised to have German-style cheesecake (here with Rooibos tea and freshly squeezed juice mix) in Windhoek.

Posted by Torsten Dikow.

Update 1 2021-03-30: Observations on iNaturalist and some identifications were added.

Update 2 2021-07-18: Some identifications were added.

Update 3 2021-10-05: Some identifications were added.

Ollie Flint 1931 – 2019

This post is dedicated to my colleague and long-time NMNH curator of Aquatic Insects and Neuropterida Ollie Flint who passed away on 18 May 2019. He will be sorely missed by his family, his colleagues in the Department of Entomology, and the entomological community at large.

Ollie had an amazing career dedicated to the study of aquatic insects (Odonata, Ephemeroptera, Plecoptera) and in particular Trichoptera as well as Neuropterida and Mecoptera and the curation of these important collections at the Smithsonian NMNH. He described more than 1,200 species of caddisflies and included countless more in his many research articles.

Below are the short obituary written by the Department of Entomology and shared by Sant Director Kirk Johnson with NMNH staff on 24 May 2019 as well as photographs and presentation slides of the celebration of Ollie's career at the NMNH held on 7 October 2019.

A comprehensive obituary has now been published by Steve Roble in Banisteria and can be downloaded here ( Download Roble_2019). Another obituary was written by Ollie's caddisfly colleagues Ralph Holzenthal and Joaquin Bueno in Braueria including a list of all publications, caddisfly species named by Ollie, and species named after Ollie and his wife Carol Flint (thanks to Dr Simon Vitecek for providing a scanned copy, Download Holzenthal_bueno_2020).

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Photographs of Ollie Flint by Flint family, Bill Murphy, Dave Ruiter, and Paul Frandsen.

Ollie collected insects world-wide including Antarctica as can be seen from the below map (also available here), which is based on collecting events in the NMNH EMu database as of September 2019 (we are working on locating the Antarctica specimens). These specimens can also be explored at Bionomia (bionomia.net/Q21283508) based on data from the Global Biodiversity Information Facility (GBIF, gbif.org).

Ollie also identified specimens in the Smithsonian USNM and numerous other museums. An overview can be accessed at Bionomia. These data will change over time as more and more specimens are being databased and made available to GBIF (www.gbif.org).

One measure to appreciate the impact Ollie had is to list the species named after him and his wife Carol. As of January 2020, three genera and 99 species in 11 orders have been named after Ollie and Carol by 86 authors (source John Morse, Clemson University, and Steve Roble, Virginia Department of Conservation and Recreation).

List of patronyms compiled by John Morse and Steve Roble ( Download Oliver-Carol_Flint_Patronyms_4Jan2020_Morse+Roble).

 

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Short obituary shared with NMNH staff on 24 May 2019

Dear Friends,

It is with great sadness that I write to tell you that our dear colleague and friend Oliver "Ollie" S. Flint, Jr., Emeritus Curator of Neuropteroids in the Department of Entomology, passed away at age 87 this past Saturday, May 18th. Ollie was a passionate and energetic presence in our museum community for almost 60 years and was active in the collection until very recently. His constant smile and devotion to his work and our museum’s mission will be sorely missed.

Ollie grew up in Amherst, MA just down the street from renowned crane fly expert C.P. Alexander, who became his mentor in all things entomological. Ollie received Bachelor’s and Master’s degrees from the University of Massachusetts before earning his Ph.D. in Entomology from Cornell University in 1960. He joined the NMNH Department of Entomology in 1961 as Curator of Neuropteroids, serving in this role for 35 years before becoming an Emeritus Curator in 1996. During his long career in the museum, Ollie curated a number of aquatic insect orders including Trichoptera (caddisflies), Megaloptera (fishflies), Ephemeroptera (mayflies), Plecoptera (stoneflies), and Odonata (dragonflies) as well as Neuroptera (lacewings), Mecoptera (scorpionflies), and Raphidioptera (snakeflies) – a collection of more than 12,000 species and almost 400,000 specimens. Ollie was a knowledgeable and dedicated curator for these groups, with our important dragonfly collection notably becoming the best curated group within all of Entomology.

Ollie authored more than 230 papers and books on aquatic insects. This remarkable scientific output spanned almost 70 years; his first paper was in 1951 and publications are in press. The majority of his research output focused on the systematics of Neotropical caddisflies, a group for which he was a world-recognized authority. Ollie described more than 1,200 species, 21 genera, and one family of caddisflies during his career. Ralph Holzenthal (University of Minnesota) relates that his own dissertation research and those of nine of his Ph.D. students depended almost entirely on specimens collected by Ollie. His and his wife Carol’s generosity in helping students and hosting visiting researchers was unlimited. Ollie loved fieldwork as probably only few people do, and his collecting and research resulted in the most comprehensive and best curated caddisfly collection in the world, particularly for Neotropical species. He also collected and deposited specimens of at least 13 other insect orders throughout the world, including Antarctica, adding more than 26,000 specimens of dragonflies and damselflies to the USNM collection as well as untold thousands of caddisflies. His local field work in collaboration with retired USDA scientist David Smith involved intensive malaise trapping that resulted in numerous new state records for several insect groups. More generally, Ollie was an avid hiker and an expert on the natural history and entomology of the greater Washington, DC area. Throughout his career, Ollie held key positions and received significant honors from organizations including the Entomological Society of America, the North American Benthological Society, and the Virginia Museum of Natural History Foundation (another local organization, like ours, particularly dear to his heart). Ollie was honored for his expertise in caddisflies at the 12th International Symposium on Trichoptera in 2006 in Mexico City, as can be seen on the “greater than life size mural” hanging on the 6th floor of the East Court opposite his office. Three genera and more than 60 species from six insect orders have been named in Ollie's honor to recognize his extensive contributions to the field.

Please join me in expressing our deepest sympathies to Ollie’s wife Carol, his daughters Cathy, Lisa, and Maria, and their mother Ruth.

Sant Director Kirk Johnson

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Celebration of the career of Ollie Flint at NMNH on 7 October 2019

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Program of event (Download Flinto_celebration_2019-10-07_program).

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Welcome by NMNH Deputy Director Ian Owens.

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Introduction by Department of Entomology Chair Seán Brady.

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Reflections by John Morse (Clemson University, retired).

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Presentation slides by John Morse (Download MorseJ_07Oct2019_small).

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Reflections by Ralph Holzenthal (University of Minnesota).

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Presentation slides by Ralph Holzenthal (Download 02_holzenthalr_small).

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Reflections by Desiree Robertson (National Park Service).

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Presentation slides by Desiree Robertson (Download 03_RobertsonD_small).

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Reflections by Joaquin Bueno (Universidad Nacional Autónoma de Mexico, retired).

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Presentation slides by Joaquin Bueno (Download 04_buenoj).

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Reflections by Lourdes Chamorro (USDA Systematic Entomology Laboratory).

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Presentation slides by Lourdes Chamorro (Download 05_chamorro_small).

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Reflections by Paul Frandsen (Brigham Young University).

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Presentation slides by Paul Frandsen (Download 06_frandsenp_small).

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Reflections by Atilano Contreras-Ramos (Universidad Nacional Autónoma de Mexico).

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Presentation slides by Atilano Contreras-Ramos (Download 7_Contreras_small).

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Reflections by Larry Stevens (Museum of Northern Arizona).

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Reflections by Kyle DeMarr (UC Berkeley & Woods Hole Oceanographic Institution).

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Reflections by Steven Roble (Virginia Department of Conservation and Recreation).

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Presentation slides by Steven Roble ( Download 10_RobleS).

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Reflections by Chris Thompson (USDA Systematic Entomology Laboratory, retired).

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Reflections by Wayne Mathis (NMNH Department of Entomology, retired).

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Reflections by Richard Greene (Smithsonian Institution Libraries).

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Presentation slides by Torsten Dikow (NMNH Department of Entomology, Download Dikowt_presentation) highlighting the collection, number of holotypes described by Ollie at USNM, and specimens of Odonata collected by him.

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Closing by the Flint family – Carol Flint (2^nd from left) and Ollie's daughters Lisa, Cathy, and Maria (from left to right).

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Impressions of the social gathering to mingle and share stories about Ollie over light snacks and beverages.

Updates:
2020-03-03 - a newly published obituary by Steve Roble was added.
2021-03-30 - another obituary by Ralph Holzenthal and Joaquin Bueno was added.

Field work with #TeamDiptera in Namibia Nov 2018

I seem to always take a long time to finally post a summary of my field-work in photos. Here is a summary of a trip I undertook with #TeamDiptera members Charlotte Alberts, Allan Cabrero, and Brandon Claridge in November 2018 prior to the 9th International Congress of Diptera in Windhoek, Namibia.

This trip included two main field-sites: habitats around Rooisand Desert Lodge near the Gamsberg (rooisand.com) and the Gobabeb Research & Training Centre in the heart of the Namib Desert (gobabebtrc.org).

I would like to thank the Namibian National Commission on Research, Science and Technology (NCRST) and the Ministry of Environment and Tourism (MET) for issuing collecting and export permits. Furthermore, the staff at the Gobabeb Research & Training Centre was very helpful as always and in particular Gillian Maggs-Kölling, Leena Kapulwa, Gregory Golando, and Eugene Marais assisted in many ways.

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Here are some of the habitats we visited:

1. Rooisand Desert Lodge habitat where Anasillomos juergeni Dikow, 2015 (image below) and Leptogaster aganniphe Janssens, 1957 (both Asilidae) were collected.

2. View from partly vegetated sand dune at Rooisand Desert Lodge towards towering Gamsberg in the far distance, which is with 2,347 m the 3rd highest mountain in Namibia. On this sand dune Anasillomos juergeni (photo below), Anypodetus sp., and Microstylum sp. (all Asilidae) were collected.

3. For the 3rd time, I visited this site in the Namib Desert at the eastern-most edge of the Namib sand sea on the C14 north of Solitaire (Khomas, Namibia). #TeamDiptera at work (Brandon (white shirt) and Allan on the left and Charlotte on the right). We collected the following Asilidae on the dunes and interdune valleys: Anasillomos juergeni, Anypodetus sp. (different species from Rooisand), Neolophonotus sp., Astiptomyia bikos Londt, 2010, Microstylum sp., and Rhabdogaster sp. A stiletto fly in the genus Orthactia (Therevidae) was also flying on the dunes.

4. Hope Mine wash in Namib-Naukluft National Park near Gobabeb. It was very windy at this site when we visited in the afternoon and collecting wasn't quite as good as we hoped. However, we were able to collect Trichardis picta Hermann, 1906, Microphontes sp., and Neolophonotus sp. (all Asilidae).

The single female Microphontes specimen we collected represents an undescribed species. This was a great discovery on the one hand, but on the other hand the timing could not have been worse as a few days after we visited this site the revision of the genus Microphontes including the description of four new species was published (see 10.3897/afrinvertebr.59.30684).

5. With information from our Polish colleagues Krzysztof Szpila, Marcin Piwczynski, and Andrzej Grzywacz, who were also visiting Gobabeb, we found this site some 20 km north of the research station. The dry riverbed of the Kuiseb River is much wider here and made for excellent collecting. We collected Namibimydas psamminos Dikow, 2012 (image below), an undescribed genus of Mydidae (image below), Eremohaplomydas sp., and Krzysztof Szpila also collected a species of Afroleptomydas (all Mydidae). In terms of Asilidae, we collected Prytanomyia kochi (Lindner, 1973), Neolophonotus sp., Trichardis picta, and Stichopogon hermanni Bezzi, 1910.

6. Collecting in the various habitats near the Gobabeb station itself was very productive. We collected the following species: Anasillomos juergeni, Prytanomyia kochi, Stichopogon hermanni, Stichopogon punctum Loew, 1851, Laphyctis eremia Londt & Dikow, 2018, Astiptomyia bikos, and Torasilus solus Londt, 2005 (all Asilidae), an undescribed genus of Mydidae, Syllegomydas sp., and Eremohaplomydas sp. (all Mydidae), and a few Therevidae species. Krzysztof Szpila also collected a species of Rhipidocephala (Asilidae), which represents a new record for Gobabeb.

The most amazing specimen was collected by Charlotte – she collected a female of one of the largest Afrotropical Mydidae species in the genus Syllegomydas in the dry Kuiseb River bed, which represents the 6th specimen of this yet undescribed species. The Eremohaplomydas specimens also represent an undescribed species new to science. The Torasilus solus specimens are, so far, the only assassin flies known to fly on the high dunes (middle photo above).

7. On our drive back to Windhoek, we stopped at this roadside near the Gamsberg Pass in the Khomas Hochland (highland) with typical Acacia savannah. We collected Sisyrnodytes sp., Stichopogon punctum, Stichopogon caffer Hermann, 1907, Afroholopgon sp., and Rhabdogaster sp. (all Asilidae). To collect five species of assassin flies in this dry habitat was somewhat of a surprise, but shows how these flies are adapted to live in arid environments.

 

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Here are some of the photographs of flies in nature taken with an Olympus OM-D EM-5 Mark II and a f/2.8 60 mm macro-lens (= 120 mm focal length in 35 mm photography):

8. Anasillomos juergeni Dikow, 2015 (Asilidae) at Rooisand Desert Lodge. See iNaturalist observation: 64122992.

9. Stichopogon hermanni Bezzi, 1910 (Asilidae) on small sand dunes at Gobabeb Research & Training Centre. See iNaturalist observation: 64174876.

10. Trichardis picta Hermann, 1906 (Asilidae) mating (female on the right) in dry Kuiseb River bed at Gobabeb. See iNaturalist observation: 64175823.

11. Namibimydas psamminos Dikow, 2012 (Mydidae) at Kuiseb River bed 20 km north Gobabeb. See iNaturalist observation: 64176614.

12. Undescribed genus and species of Mydidae (Syllegomydinae) at Gobabeb.

13. Namibiotes argentata Barraclough, 2000 (Ulidiidae) on !nara (Acanthosicyos horridus, Curcubitaceae). This fly is so far only known to live on the !nara plant which is endemic to the Namib Sand sea within the Namib Desert. See iNaturalist observation: 64177007.

 

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Other impressions of Namibia:

14. Male Welwitschia mirabilis (Welwitschiaceae) – enigmatic plant of the Namib Desert – at Hope Mine wash.

15. A sight I have never seen on my four visits to the Namib – the desert can be green.

16. Springbok (Antidorcas marsupialis Sundevall, 1847, Bovidae) grazing the Namib gravel plains.

17. Lesser flamingoes (Phoenicoparrus minor Geoffroy Saint-Hilaire, 1798, Phoenicopteridae) near Walvis Bay.

18. A praying mantis (Chroicopteridae, ID via iNaturalist) at Hope Mine wash. See iNaturalist observation: 64175314.

19. Fruit of Phaeoptilum spinosum (Nyctaginaceae) at roadside near Gamsberg Pass. See iNaturalist observation: 64177671.

 

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TeamDiptera:

20. #TeamDiptera (Allan Cabrero, Charlotte Alberts, Brandon Claridge, Torsten Dikow from right to left) in front of huge !nara plant.

21. #TeamDiptera equipment.

Posted by Torsten Dikow.

Update 2021-04-13: Identifications or links to observations at iNaturalist were added.

Field work in South Africa and Namibia Sep 2017

Finally, several months delayed is a post about my recent field work in South Africa and Namibia that took place in September 2017.

My field work took me from Cape Town, South Africa to Windhoek, Namibia – a distance of some 1,500 km (930 miles) if you drive straight, which I didn't. I had planned to visit several habitats in the Western and Northern Cape of South Africa and Karas in Namibia to collect spring-active assassin flies (Asilidae), mydas flies (Mydidae), and other Asiloidea and pollinators such as Nemestrinidae (tangle-veined flies) for a project on the evolutionary relationships of Asiloidea and Nemestrinoidea based on morphological data. Because of a severe drought in this part of southern Africa, my collecting wasn't very successful with a few exceptional collecting localities though. Further north, I was planning to visit the Namib Desert again and in particular the Gobabeb Research & Training Centre (www.gobabebtrc.org). Once I arrived in the true desert, collecting was wonderful as the flies occurring here are not reliant on winter rainfall such as further south.

I would like to thank the government agencies in the Western Cape (CapeNature), Northern Cape (Permit Section), and Namibia (Ministry of Environment and Tourism) for issuing collecting permits. Furthermore, the staff at the Gobabeb Research & Training Centre was very helpful as always and in particular Ritha Kapitango, Leena Kapulwa, Angi Storbeck, and Theo Wassenaar assisted in many ways.

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Here are some of the habitats I visited:

1. Rocherpan Nature Reserve (Western Cape, South Africa) where Acnephalomyia dorsalis (Macquart, 1838), Haroldia trivialis (Oldroyd, 1974), Hypenetes sp., Corymyia melas Londt, 1994, and Neolophonotus sp. (Asilidae), Neotabuda lanigera Lyneborg, 1980 (Therevidae, stiletto flies), Corsomyza sp., Megapalpus capensis Hesse, 1938, and Tomomyza sp. (Bombyliidae, bee flies) were collected. Haroldia trivialis (see image below) was collected in abundance perching on the sand of the coastal dune (foreground) as was Corymyia melas, but the other species were collected near the pan (in the distance).

2–3. Gifberg Pass (Western Cape, South Africa) where Neolophonotus sp., Rhipidocephala sp., and Damalis conspicua Curran, 1934 (Asilidae), Neotabuda sp. (Therevidae), and Apatomyza sp. (Bombyliidae) were collected. This fynbos habitat was amazing (even on a cloudy day) and the Rhipidocephala species (see images below) surprised me by perching mostly on little rocks and not on vegetation.

 

4. A special treat was driving through the /Ai-/Ais Richtersveld Transfrontier Park (Karas, Namibia) along the "mighty" Gariep or Orange River, which is the largest river in western South Africa. In the adjacent sandy habitat, I was able to collect Stichopogon punctum Loew, 1851 (Asilidae) and Orthactia sp. (Therevidae).

5. A rocky hillside at Klein-Aus Vista (Karas, Namibia) where Neolophonotus sp. and Anasillomos chrysopos Londt, 1983 (Asilidae) were collected. This is the northern-most area where the winter rainfall should bring out spring flowers, but the last proper rain fell here in 2014.

6. For the 2nd time I visited this site in the Namib Desert at the eastern-most edge of the Namib sand sea on the C14 north of Solitaire (Khomas, Namibia) where Anypodetus sp., Microphontes gaiophanes Markee & Dikow, 2018, and Stichopogon hermanni Bezzi, 1910 (Asilidae) and Orthactia sp. (Therevidae) were collected. I have never seen Microphontes (see images below) before and the collected specimens represented an undescribed species for which I was able to record some peculiar flight behavior (see publication at 10.3897/AfrInvertebr.59.30684). The Anypodetus species likewise appears to be undescribed.

 7. The Gobabeb Research & Training Centre (Erongo, Namibia) is an amazing place to visit in the heart of the Namib Desert. This time, I was unable to collect any assassin flies on the high dunes (as shown here), which are situated west and south of the research station (visible in the distance to the right of the wooded dry Kuiseb riverbed). I was able to collect Xeramoeba sp. (Bombylliidae) on these high dunes though.

 

8. The edge of the high dunes adjacent to the wooded dry Kuiseb riverbed at Gobabeb proofed very interesting though where Sporadothrix sp. and Prytanomyia kochi (Lindner, 1973) (Asilidae, see images below) as well as Parectyphus namibiensis Hesse, 1972 and an undescribed genus and species of Mydidae (see image below) were collected.

9. Just as in my previous visit to Gobabeb, Orthactia gobabensis Lyneborg, 1988 (Therevidae), Trichardis picta Hermann, 1906 (see image below), and Stichopogon punctum Loew, 1851 (Asilidae) were collected in the dry Kuiseb riverbed.

 

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Here are some of the photographs of flies in nature taken with an Olympus OM-D EM-5 Mark II and a f/2.8 60 mm macro-lens (= 120 mm focal length in 35 mm photography):

10. A female stiletto fly of Caenophthalmus gracilis Lyneborg, 1976 (ID via iNaturalist Shaun Winterton, Therevidae: Therevinae) ovipositing at Cape Columbine Nature Reserve (Western Cape, South Africa). See iNaturalist observation: 63854583.

11. Acnephalomyia dorsalis (Macquart, 1838) (Asilidae) perching on sand at Rocherpan Nature Reserve (Western Cape, South Africa). See iNaturalist observation: 63854661.

12. Female Haroldia trivialis (Oldroyd, 1974) (Asilidae) perching on sand dune at Rocherpan Nature Reserve (Western Cape, South Africa). See iNaturalist observation: 63855130.

13. A male Neotabuda lanigera Lyneborg, 1980 stiletto fly (Id via iNaturalist Shaun Winterton, Therevidae) resting on sand at Rocherpan Nature Reserve (Western Cape, South Africa). See iNaturalist observation: 63854969.

14. Male Neolophonotus sp. (Asilidae) feeding on an aphid on a rock at Gifberg Pass (Western Cape, South Africa). See iNaturalist observation: 63855347.

15–16. Rhipidocephala sp. (Asilidae) perching on rock and my collecting net at Gifberg Pass (Western Cape, South Africa). See iNaturalist observation: 63855466.

 

17. Female Stichopogon punctum (Asilidae) feeding on a hemipteran at Gariep (Orange) River in /Ai-/Ais Richtersveld Transfrontier Park (Karas, Namibia). See iNaturalist observation: 63855586.

18–19. Microphontes gaiophanes Markee & Dikow, 2018 (Asilidae) perching on sand dune (18, male) and dry twigs (19, female) at Namib-Naukluft National Park near Solitaire (Erongo, Namibia). See iNaturalist observations: 63855855 + 63855886.

20–21. Female Sporadothrix sp. (Asilidae) perching on sand dune (20) and ovipositing in sand (21) at Gobabeb in the Namib-Naukluft National Park (Erongo, Namibia). See iNaturalist observation: 63856014.

22. Female Trichardis picta (Asilidae) perching on sand in dry Kuiseb riverbed at Gobabeb in the Namib-Naukluft National Park (Erongo, Namibia).

23. Female Prytanomyia kochi (Asilidae) perching on dung on small sand dune at Gobabeb in the Namib-Naukluft National Park (Erongo, Namibia). See iNaturalist observation: 63856269.

24. Female mydas fly of an undescribed species and genus (Mydidae) resting on dry leave on small sand dune at Gobabeb in the Namib-Naukluft National Park (Erongo, Namibia). This species was one of the target taxa on my list to collect and so I was extremely happy to collect and photograph it in nature.
 

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Some impressions of southern Africa:

25. Whenever I can, I stop by botanical gardens and Kirstenbosch National Botanical Garden (www.sanbi.org/gardens/kirstenbosch) in Cape Town is one of the world's most beautiful gardens.

26. I also visited the National Botanic Garden (www.nbri.org.na/sections/botanic-garden) in Windhoek.

The flower diversity in western South Africa always amazes me and here are just some impressions.

27. Baboon root (genus Babiana, Iridaceae) at Cape Columbine Nature Reserve (Western Cape, South Africa). See iNaturalist observation: 63856269.

28. Capebells (genus Wahlenbergia, Campanulaceae, ID via iNaturalist) at Gifbeg Pass (Western Cape, South Africa). See iNaturalist observation: 63855308.

29. King Kalkoentjie (Gladiolus alatus, Iridaceae, ID via iNaturalist) at Gifbeg Pass (Western Cape, South Africa). See iNaturalist observation: 63855241.

Namibia is a place of huge distances between towns.

30. Getting ready for the drive through /Ai-/Ais Richtersveld Transfrontier Park along Gariep (Orange) River (Karas, Namibia).

31. Near Rosh Pinah (Karas, Namibia).

32. Near Aus (Karas, Namibia).

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Other wildlife:

33. Four-striped grass mouse (Rhabdomys pumilio (Sparman, 1784)) at Rocherpan Nature Reserve (Western Cape, South Africa). ID by Molly McDonough.

34. Friendly tortoise at Rocherpan Nature Reserve (Western Cape, South Africa).

35. Chacma baboons (Papio ursinus Kerr, 1792) in /Ai-/Ais Richtersveld Transfrontier Park (Karas, Namibia) watching what I am doing with my collecting net.

36. The ever so beautiful Kokerboom (Aloidendron dichotomum (Masson) Klopper & Gideon F.Sm.) just outside /Ai-/Ais Richtersveld Transfrontier Park (Karas, Namibia).

37. Tree Locust (Anacridium moestum (Serville, 1838), Orthoptera: Acrididae) hiding among dry vegetation at Klein-Aus Vista (Karas, Namibia). See iNaturalist observation: 63855760.

38. Hartmann's Mountain Zebra (Equus zebra hartmannae (Matschie, 1898)) at Namib-Naukluft National Park near Solitaire (Erongo, Namibia).

39. Blister beetle (Meloidae) on dune plant at Namib-Naukluft National Park near Solitaire (Erongo, Namibia).
 

40. Thread-winged lacewing Nemopterella sp. (Nemopteridae, ID via iNaturalist) at Namib-Naukluft National Park near Solitaire (Erongo, Namibia). This thread-winged lacewing was very pale and I assume it had recently emerged from the pupa and had not fully hardened/colored. See iNaturalist observation: 63855936.

41. Dune lark (Calendulauda erythrochlamys (Strickland, 1853)) on high dunes at Gobabeb, Namibia-Naukluft National Park (Erongo, Namibia) – Namibia's only true endemic bird species. See iNaturalist observation: 63856209.

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42. My companions – Olympus and collecting net.

43. Cheese cake (Quarkkuchen) is always good and the German heritage can still be found in Namibia.

Posted by Torsten Dikow.

Updates: 2018-03-08 - Molly McDonough kindly identified the grass mouse photograph.

2018-03-12: Several species identifications were added.

2021-04-13: IDs and links to iNaturalist observations added.

Photographing asiloid flies in amber

Having imaged a number of amber fossils for an upcoming publication, I thought it might be good to summarize the workflow I have been using with my equipment in a blog post. Here it is in images and short captions.

• Stereo microscope: Zeiss SteREO Discovery.V12 (Zeiss web-site), PlanApo S 1x objective, Zeiss VisiLED illumination, magnification varies.
• Camera: Micro 4/3 Olympus OM-D E-M5 (see dpreview.com for details) - I have also used an Olympus E-PL5 with this stereo microscope.
• Settings camera: S program, shutter time = 1/125, RAW image, low ISO (= 100), auto White Balance.
• Settings VisiLED lightsource: combined/mixed Bright Field (light from above) and Dark Field (light from side) - in some instances also Transillumination (light from below).
• Image stacking: manual, 15–30 slices/images per specimen.
• Software: Adobe Lightroom 5, HeliconFocus Pro 6. The workflow below is entirely RAW-based - RAW-in-DNG-out.

1. Specimen, here †Protoloewinella keilbachi Schumann, 1984 (Asilidae: Laphriinae: Atomosiini, K-4739), placed on stage.

2. VisiLED combined Dark and Bright Field lights being lowered.

3. Look into the "light chamber" that will go over the specimen. The lower ring of LED lights (closest here) will provide the Dark Field (lateral) light and the 2nd set of LED lights (two rings, further away surrounding the objective opening) will provide the Bright Field (dorsal) light.

4. The specimen is covered/surrounded by light in a "light chamber" and we are ready to take images.

5. View of the camera display with a preview of the image and camera settings

Since the Zeiss SteREO Discovery.V12 doesn't have a motor drive to move the objective/camera down to take several images at different focal plains (the V20 does), I trigger the camera with a remote control and manually move down the objective/camera before taking the next image. I try to always move it the same distance, but that's not always possible. I take between 15–30 slices/images per specimen. For higher magnification of certain parts of the specimen, the number can be smaller.

Once all of the images are taken, I use the same workflow with Lightroom and Helicon Focus as with other images I take of pinned flies. You can read more in this blog post by starting at number 17. Differences are:

• no need to rotate the images
• keywords will be the magnification setting at the stereo microscope
• no scale line (although one could add the scale line here)

A few general comments:

• the Dark Field LED lights were set to be much brighter than the Bright Field lights
• it is good to play with the transillumination - sometimes it enhanced the image and sometimes it didn't make much of a difference
• of course, one always sees more looking through the stereo microsocope than can be captured in the image, but the images provide access to these specimens and will all be uploaded and made available through Morphbank for future dipterists at publication of the manuscript.

Here are a few final images.

†Protoloewinella keilbachi (K-4739) in dorsal view. 10x magnification.

†Protoloewinella keilbachi (K-4739) in ventral view. 10x magnification.

†Protoloewinella keilbachi (K-4739) male terminalia detail with Transillumination. 60x magnification.

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I have previously used the same stereo microscope, lighting set-up, and post-processing worklfow to image male and female terminalia of extant flies. See Dikow 2015.

Anasillomos juergeni Dikow, 2015 (USNMENT00832201) male terminalia dorsal. 50x magnification, Morphbank 850965.

Anasillomos juergeni (USNMENT00832201) male terminalia lateral. 50x magnification, Morphbank 850967.

Anasillomos juergeni (USNMENT00832201) male terminalia ventral. 50x magnification, Morphbank 850969.

Field work in South Africa in Dec 2015

Finally, several months delayed is a post about my recent field-work in South Africa that took place in December 2015.

This field trip was funded by the Global Genome Initiative (GGI) with the purpose of increasing the holdings of flies (Diptera) available for genome or genomic sequencing in the NMNH Biorepository. For that purpose, I sent a liquid nitrogen dry shipper to South Africa and back via FedEx. During the field work I kept the dry shipper in the back of my field vehicle.

The field vehicle in Addo Elephant National Park near Port Elizabeth, Eastern Cape (note: I did not collect in the national park).

The liquid nitrogen dry shipper secured in the back of the bakkie (a.k.a. pickup truck).

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I proposed this field work in order to visit a particularly diverse place in the Nama Karoo of South Africa called Willowmore. This small town and its surroundings in the south-eastern Nama Karoo (Eastern Cape Province of South Africa, 33°17'00''S 023°29'00''E) are particularly diverse for asiloid flies with 1 species of apiocerid flies, 62 assassin flies, and 11 mydas flies known to date. The article by Londt (1998, download PDF) nicely summarizes the Asilidae species that were collected by Dr. J. Brauns in and around Willowmore in the first quarter of the 20th century. Other collections also house specimens from J. Brauns and a single new genus and species to this list, Ammodaimon sp. (USNMENT00995300, see record), was discovered in the USNM collection. So, this area is a great place to collect asiloid flies (and other Diptera).

During this field trip I visited several different habitats from coastal dunes and riverine habitats to several kinds of Karoo veld and habitats as can be seen from the following habitat photographs.

1. Coastal dunes and dune vegetation at Riet River Mouth near Port Alfred, Eastern Cape, where Afroleptomydas sp. and Ectyphus pingius Gerstaecker, 1868 (Mydidae) and a Neolophonotus species (Asilidae) were collected.

2. Riverine habitat of the Groot Rivier at Meiringspoort, Western Cape, where Stichopogon hermanni Bezzi, 1910 (Asilidae) was collected perching on the rocks.

3. Nama Karoo north of Olifantskop Pass, Eastern Cape, where on a roadside Acnephalomyia, Neolophonotus, and Trichardis species (Asilidae) were collected.

4. Nama Karoo mountain side near Graaff-Reinet, Eastern Cape, where a Neolophonotus and a Afroscleropogon species (Asilidae) were collected.

5. Nama Karoo at Aberdeen Nature Reserve outside of Aberdeen, Eastern Cape, where Apiocera braunsi Melander, 1907 (Apioceridae) and Anypodetus fascipennis Engel, 1924, Habropogon, Microstylum, Neolophonotus, Promachus, Smeryngolaphria, and Trichardis species (Asilidae) were collected.

6. Nama Karoo at Aberdeen Nature Reserve outside of Aberdeen, Eastern Cape. This is one of the pans at which I caught Apiocera braunsi Melander, 1907 running around the margins near the vegetation.

7. Nama Karoo on the De Zeekoe Guest Farm near Oudtshoorn, Western Cape, where an undescribed genus of Mydidae as well as Afroscleropogon, Daspletis setithoracicus (Ricardo, 1925), Leptogaster aganniphe Janssens, 1957, Microstylum, Nusa infumata (Loew, 1851), Synolcus acrobaptus (Wiedemann, 1828), Sisyrnodytes, Trichardis, and Trichoura tankwa Londt, 1994 (Asilidae) were collected.

8. Vegetated dune ridge in Goukamma Nature Reserve near Knysna, Western Cape, where an Afroleptomydas species (Mydidae) as well as Damalis, Pegesimallus, and Sisyrnodytes species (Asilidae) were collected.

9. Coastal forest in Goukamma Nature Reserve near Knysna, Western Cape, where Oligopogon penicillatus Loew, 1858 (Asilidae) was collected. In my phylogenetic work on Asilidae, I was unable to place this genus with morphological data (it is currently incertae sedis) and now we can utilize molecular data as I collected a few specimens in liquid nitrogen.

In the end, I didn't even make it to Willowmore as I found many other suitable collecting localities and time was limited on this 10-day trip.

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I was able to collect the following genera / species (I still need to identify several taxa to the species level):

Apioceridae: Apiocera braunsi Melander, 1907
Asilidae: Acnephalomyia sp., Afroscleropogon sp., Anypodetus fascipennis Engel, 1924, Damalis annulata Loew, 1858, Daspletis setithoracicus (Ricardo, 1925), Habropogon sp., Hyperechia nigripennis (Wiedemann, 1830), Leptogaster aganniphe Janssens, 1957, Microphontes ericfisheri Markee and Dikow, 2018, Microstylum sp. (3 species), Neolophonotus sp., Nusa infumata (Loew, 1851), Oligopogon penicillatus Loew, 1858, Pegesimallus sp., Promachus sp., Sisyrnodytes sp., Smeryngolaphria sp., Stichopogon hermanni Bezzi, 1910, Synolcus acrobaptus (Wiedemann, 1828), Trichardis sp., and Trichoura tankwa Londt, 1994.
Mydidae: Afroleptomydas sp., Ectyphus pinguis Gerstaecker, 1868, and an undescribed genus.

For the majority of these species, specimens were preserved both in ethyl-acetate and pinned for morphological work and proper identification as well as in liquid nitrogen for genome and genomic sequencing in the future. An example record of a pinned fly can be viewed on the USNM collection web-site (USNMENT01115072) and a liquid nitrogen-preserved fly on the Global Genome Biodiversity Network (GGBN) data portal (AA8IQ31).

I was also able to collect several species of bee flies (Bombyliidae), hover flies (Syrphidae), and stiletto flies (Therevidae) as well as a horse fly (Tabanidae) and a tachinid fly (Tachinidae) species.

Below are some photographs I was able to take of flies in the field.

 

1. Leptogaster aganniphe Janssens, 1957 (Asilidae: Leptogastrinae). See iNaturalist observation: 63361814.

2. Microstylum sp. (Asilidae: Stenopogoninae). See iNaturalist observation: 63361220.

3. Microstylum sp. (Asilidae: Stenopogoninae). See iNaturalist observation: 63361579.

4. Oligopogon penicillatus Loew, 1858 (Asilidae: incertae sedis). See iNaturalist observation: 63362338.

5. Synolcus acrobaptus (Wiedemann, 1828) (Asilidae: Asilinae). See iNaturalist observation: 63362567.

6. Anypodetus fascipennis Engel, 1924 (Asilidae: Laphriinae). See iNaturalist observation: 63361137.

7. Trichoura tankwa Londt, 1994 (Asilidae: Willistonininae). Se iNaturalist observation: 63361460.

8. Daspletis setithoracicus (Ricardo, 1925) (Asilidae: Stenopogoninae). See iNaturalist observation: 63361712.

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South Africa is a beautiful place to visit and the Nama Karoo especially. I was fortunate to be able to see Meerkats for the first time through Meerkat Adventures at the De Zeekoe Guest Farm near Oudtshoorn, Western Cape.

The last afternoon of my stay in South Africa I spent in the Addo Elephant National Park where I saw plenty of Loxodonta africana (Blumenbach, 1797) and other mammals.

Did you know that the Nama Karoo has an incredible diversity of tortoises? Here is just one of them, the Angulate Tortoise Chersina angulata (Schweigger, 1812).

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Posted by Torsten Dikow.

Thanks to the Eastern and Western Cape authorities for collecting permits. Thanks also to Dolf Grobler and J.D. Glinister for helping me with field work on the De Zeekoe Guest Farm property.

Updates: 2016-11-05 - the identification of a few species has been changed/updated.

2021-04-14: Several IDs and iNaturalist observations added.

Photographing asiloid flies

I have been thinking about a blog post to showcase the macro-photo equipment I use for photographing asiloid flies for a while. Here it is in images and short captions.

• Camera + lens: Micro 4/3 Olympus OM-D E-M5 (see dpreview.com for details), 60 mm f2.8 Macro (see Olympus web-site, 120 mm in 35 mm photography).
• Stand: base, stand, and StackShot from Visionary Digital Passport II system.
• Dome light: Falcon FLDM-i250-W.
• Settings camera + lens: manual focus, manual program, f/8, RAW image, low ISO (= 100), auto White Balance, depending on size of fly 1:2, 1:1.3, or 1:1 magnification.
• Settings StackShot: for dorsal and lateral view 15 steps, for head in anterior view 7 steps, automatic step mode.
• Software: Adobe Lightroom 5, HeliconFocus Pro 6. The workflow below is entirely RAW-based - RAW-in-DNG-out.

1. Simple specimen holders made of plastozote and cut DGK grey card (DGK Color Tools Digital Grey Kard), which are not glued together.

2. The macro-photography set-up with specimen labels ready to be photographed.

3. Labels just photographed.

4. Fly, here Apiocera braunsi Melander, 1907, ready to be photographed in dorsal view.

5. Fly positioned with evenly grey background.

6. Fly photographed.

7. Fly positioned for lateral view (1).

8. Fly positioned for lateral view (2).

9. One of the resulting images on the camera screen.

10. Lens set to photograph at 1:1.3 magnification (fly would not fit on screen when 1:1 would be selected).

11. Other side of lens where different distance settings can be pre-selected.

12. Camera writing one of the many images to the card.

13. StackShot triggering the camera to take image 10 of 15.

14. Fly positioned for photographing head in anterior view (1).

15. Fly positioned for photographing head in anterior view (2).

16. Look inside the currently switched off, but super bright Falcon dome light.

17. Images get imported into Adobe Lightroom 5 as native .orf RAW files. A secondary copy is made as a back-up to my RAID.

18. The original images get rotated, meta-data adjusted (e.g., the copyright holder changed), and renamed.

19. Keywords get allocated that indicate the magnification at which the images were taken. Important if one would want to re-do the scale bar.

20. Images get exported in native .orf RAW to HeliconFocus for stacking.

21. In HeliconFocus, Method C is used for stacking into a single Digital Negative .dng file.

22. HeliconFocus stacking in progress.

23. HeliconFocus stacking finished - left = one of the original images, right = final stacked image.

24. Scale line of 5 mm is applied to image (here 23% of the automatically inserted scale). I calculated the scale line following the guidelines provided by HeliconFocus. Basically taking an image with the macro-photography set-up of a microscopic slide with a scale.

25. Stacked image saved as Digital Negative (.dng) file and added to Lightroom catalog. White Balance is always correct as images are taken with the same Falcon Dome Light and grey background.

26. In Lightroom, image can be edited just as any other .dng file and I adjust the exposure here slightly.

27. The final 4 .dng images. A new keyword (scale=5mm) is assigned so that I remember that this scale is 5 mm long. From here, the images can be exported for different uses such as Twitter, Morphbank, or our institutional database KE EMu.

Here are the 4 final images of Apiocera braunsi:

Posted by Torsten Dikow

update 2016-03-16: The wrong grey card was listed above for the specimen holder. Instead of WhiBal G7 grey card it should read DGK Color Tools Digital Grey Kard, which is very sturdy and has an almost smooth surface.

Collecting asiloid flies for genomic research

The assassin fly Scleropogon duncani photographed in the field near Parker, AZ feeding on a Heteroptera species.

I have collected flies for my research for almost 20 years now and over the past 15 preserved specimens for different research applications. The standard for morphological taxonomic and phylogenetic research on asiloid flies is to hand-net specimens, preserve them in ethyl acetate, and pin them in the field. For internal morphological study, I have preserved specimens in Kahle's Fluid (see information here) and for molecular phylogenetic studies in 95% ethanol. It has always been my aim to collect a series of specimens of a particular species at the same site, pin several of them for proper species identification, and preserve a few in 95% ethanol and Kahle's fluid to utilize the specimens in combined morphological and molecular phylogenetic analyses. In our genomics age, my collecting efforts have changed slightly in that I strive to preserve specimens in liquid nitrogen to achieve the highest quality of preservation of RNA and DNA for transcriptome and genome sequencing or other next-generation sequencing applications in addition to specimens for morphological study.

Below is an outline of a workflow developed during my first trip to southern Arizona and California, in which I used a liquid nitrogen dry shipper and databased every single specimen in the field, conducted in April 2015. This trip was a good test scenario because I visited collecting sites I had visited in previous years and had an idea which genera and species of asiloid flies I could expect. The field sites were mostly inland sand dunes or sandy habitats such as shown here.

Vegetated sand dunes east of Parker, AZ.

Vegetated sand dunes and interdune valleys east of Parker, AZ.

The Smithsonian Institution's National Museum of Natural History (NMNH) has recently established a liquid nitrogen repository called Biorepository that is a new "type of collection" not restricted to taxonomic groups and which will store vials with tissues or whole organisms of disparate taxa adjacent in a liquid nitrogen tank. The Biorepository is integrated in the Global Genome Initiative (GGI) and the Global Genome Biodiversity Network (GGBN), which are both headquartered at the NMNH.

A few words about specimen data

Collecting specimens today involves keeping track of associated data right from the beginning, i.e., directly in the field. In order to associate the pinned fly, which will serve as an exemplar in the NMNH Diptera collection for future verification of identification or morphological study, with the specimen(s) of the same species preserved in liquid nitrogen, it needs to have a unique specimen identifier assigned to it right away. Likewise, each specimen preserved in liquid nitrogen will need a unique identifier. In the case of the NMNH Biorepository, a second identifier is necessary to track the vial in the liquid nitrogen tanks, which is captured in a FreezerPro database. Once the record of the pinned fly is in the NMNH institutional KE Emu database, both records can be associated and the information forwarded to the GGBN data portal. At the NMNH, the field data can either be captured in a Field Information Management System (FIMS) Excel spreadsheet or, as in my case, in a personal research database from where they will be exported following the FIMS template.

Prior to field work

I prepared the vials by affixing two specimen identifiers (UNMENT and Bioreposiory identifier) to each of them so that I won't have to do this step in the field. (I brought a hand-held label scanner into the field to scan the identifier labels and enter them into the correct field to avoid typing errors.) I also sent the liquid nitrogen dry shipper, such as the one below, to the FedEx office at the Phoenix airport from where I picked it up and had it filled with liquid nitrogen at a welding supply store.

In the field

I am collecting asiloid flies (chiefly my research taxa Apioceridae, Asilidae, and Mydidae) with a hand-net by collecting one specimen at a time. These flies are big enough so that they can be identified to at least the genus in the field. Since I needed to make a decision of which specimens will be preserved in liquid nitrogen and which one in ethyl acetate in the field, I decided to keep individual flies at a specific site alive in separate vials. Once I had collected several specimens of the same species, I preserved the next few in ethyl acetate for pinning. The flies destined for liquid nitrogen were placed in a small cooler with ice packs and later transferred to a fridge in the hotel room before processing. This had the added advantage that the flies slowed down and could be moved easily to one of the pre-labeled vials.

Databasing specimen records in the field

In the evening back at the hotel, specimens were processed. The specimens preserved in ethyl acetate were pinned, received a unique specimen identifier (USNMENTXXXXXXXX), and were added to my specimen-level database.

Part of the pinned specimens collected with unique specimen identifiers assigned.

Screen shot of custom research database (in FileMaker Pro) capturing occurrence data for pinned specimen.

Next, the genomic sample specimen that was kept alive and which is vouchered by the above pinned specimen was placed in a pre-labeled vial with both the USNMENT unique identifier and the unique Biorepository identifier (such as AA8IQ05) and added to the database.

Screen shot of research database capturing occurrence data for genomic sample specimen.

These two records need to be connected, which is done by adding the USNMENT identifier of the exemplar and genomic sample to the respective records. A pinned exemplar can have several genomic samples, but each genomic sample refers to only one pinned exemplar in this case.

Screen shot of research database showing relationship of pinned exemplar specimen to genomic sample specimens (one-to-many relationship).

Screen shot of research database showing relationship of genomic sample specimen to pinned exemplar specimen (one-to-one relationship).

Specimens to be preserved in liquid nitrogen were placed in the vials, wrapped in aluminum foil (to prevent the labels getting disassociated form the vial), and then dropped into the dry shipper containing liquid nitrogen.

Pre-labeled cryovials with two identifiers (USNMENT and Biorepository).

Cryovials wrapped in aluminum foil before being dropped in liquid nitrogen.

From each pinned exemplar and genomic sample specimen, a hind leg was placed in a buffer plate so that a genetic barcode (mitochondrial COI) can be sequenced to provide a second means of identification of the specimens.

96-well plate with buffer for preservation of DNA from leg for sequencing of COI gene.

This leg gets it's own record and unique specimen identifier and likewise is connected to the voucher specimen (exemplar or genomic sample).

Screen shot of research database showing record for leg removed for sequencing COI gene.

After the field trip

Since I sent the liquid nitrogen dry shipper back to the Biorepository from Phoenix via FedEx, the GGI team moved the specimens from the dry shipper to the liquid nitrogen tanks and added the vial numbers to FreezerPro. The data of all specimens I had collected and the legs for COI sequencing were exported from my database to the FIMS spreadsheet and uploaded into EMu by the Department of Entomology data management team.

The pinned specimens needed to be labeled and I use my database to generate the locality and identification labels. I added a script that would include a label indicating whether a pinned specimen is an exemplar for a specimen in the Biorepository and whether a leg has been removed for COI barcoding.

Example labels for a pinned exemplar specimen.

Below is the set of pinned Scleropogon duncani specimens with two of them being exemplars for specimens in the Biorepository as well as vouchers for a leg removed for COI sequencing.

 

Unit tray with assassin fly Scleropogon duncani specimens labeled and ready to go into main USNM Diptera collection.

Today, the data of the flies I collected are publicly accessible. For example, the exemplar specimen can be found on the NMNH Entomology collection database: USNMENT01115108. The associated specimen in the Biorepository can't be found in the same database yet (NMNH doesn't publish genomic samples to the public directly yet), but it is accessible through the GGBN record identified by the unique NMNH Biorepository number AA8IQ05. 

The assassin fly Albibarbefferia albibarbis photographed in the field near Portal, AZ feeding on a Heteroptera species and ovipositing in crevices in the bark.

Now I am all set for a GGI-funded field trip to the Eastern Cape in South Africa to collect flies for genomics research in December 2015.

Genome sequencing

Did we succeed in preserving RNA and DNA properly? Yes! The largest assassin fly I collected on this trip was Proctacanthus coquilletti Hine, 1911 and we are now sequencing the genome of this species based on a single specimen (see record at GGBN) using the HiSeq 2500 platform and DISCOVAR de novo assembly. We have also sequenced the transcriptomes of the imago of the apiocerid fly Apiocera parkeri Cazier, 1941, the mydas fly Messiasia californica (Cole, 1969), and the assassin flies Laphystia limatula Coquillett, 1904 and Scleropogon duncani Bromley, 1937 on the HiSeq 2000 platform in a single lane. The results are good and watch this space for updates on the progress of these projects in particular the novel genome.

The assassin fly Proctacanthus coquilletti photographed in the field.

Thanks

I need to thank Patricia Gentile-Poole and Michael Lloyd from the Department of Entomology for help with uploading the specimen data from the FIMS into EMu. Chris Huddleston and his GGI team at the Biorepository helped in preparing this trip and took care of the vials shipped to them.

Posted by Torsten Dikow

update 2016-03-09: Erroneously, the location of some of the images was given as Portal, AZ whereas it is Parker, AZ.

Prey of assassin flies - data from the USNM Diptera collection - part 1

The data presented in this blog stem from a project that was part of a summer internship of three undergraduate students conducted in 2015.

 

A Comparison of Prey Items of Asilidae subfamilies

in the USNM and NMSA Collections

Jonathan Chen | University of Maryland

Karine Camacho | Universidade Federal da Integração Latino-Americana UNILA

Suma Cheru | Wellesley College

 

Introduction

In this study, we compared the different prey items of Asilidae subfamilies in the USNM (National Museum of Natural History) collection with a data-set based on the NMSA collection (KwaZulu-Natal Museum, Pietermaritzburg, South Africa) as published by Londt (2006, http://hdl.handle.net/10520/EJC32691 ). The five Asilidae subfamilies that were compared in this study include Asilinae, Dasypogoninae, Laphriinae, Stenopogoninae, and Stichopogoninae. While the species were exclusively from the Afrotropical Region in Londt’s publication, the species in the USNM were not sorted based on region, and may have come from any of the zoogeographic regions. The purpose of this study was to observe how the prey orders compared between the exclusive Afrotropical Asilidae species with the Asilidae species in the USNM.

Methods

We first went through the USNM Asilidae collection, which is organized by subfamily, and moved any specimens collected with a prey item and placed them in a cabinet adjacent to main assassin-fly collection. We were then able to assign a unique specimen identifier (USNMENTXXXXXXXX) to each assassin-fly specimen. For each of these specimens, we identified the prey item to order and recorded the data in an Excel spreadsheet. Therefore for each subfamily, we were able to come up with a list of the different prey orders that we found. We were then able to create bar graphs and compare the number and percentages of each prey order for each Asilidae subfamily and compare our results to the data provided by Londt (2006) based on Afrotropical species. In both data-sets, about 2,000 assassin flies with prey items are included.

Results

1. Asilinae

Observations Asilinae

Diptera prey comprises 28.4% of the total prey items of the Asilinae in the USNM and similarly comprises 26.6% of the total prey items of the Afrotropical Asilinae in the NMSA. Londt reports that the Diptera is the dominant prey item for the Asilinae (Londt 2006); however in the USNM, Hymenoptera clearly is the dominant prey item, comprising 36.2% of all prey items found with Asilinae in the collection.

The dominant prey items for Asilinae in the USNM are clearly Hymenoptera and Diptera while the dominant prey items and in the NMSA are Diptera, Hymenoptera, and Orthoptera.

There is a discrepancy between the dominance of Orthoptera as a prey item between the two collections: while it comprises only 8.2% of total prey items for the USNM Asilinae, it comprises 20% of total prey items of Afrotropical Asilinae in the NMSA.

Hemiptera comprises 8.0% of the total prey items of the Asilinae in the USNM and similarly comprises 8.5% of the total prey items of the Afrotropical Asilinae in the NMSA.

Lepidoptera comprises 11.8% of the total prey items of the Asilinae in the USNM and similarly comprises 11.7% of the total prey items of the Afrotropical Asilinae in the NMSA.

Araneae comprises 0.14% of the total prey items of the Asilinae in the USNM and comprises 0.32% of the total prey items of the Afrotropical Asilinae in the NMSA.

Summary data Asilinae

Comparison of the percentages of prey items (sorted to prey order) for the USNM Asilinae* and for the NMSA Afrotropical Asilinae.

Prey Order	USNM Asilidae 3	NMSA Afrotropical Asilidae
Blattodea1	0.09 %	0.0 %
Coleoptera	6.6 %	14.7 %
Diptera	28.6 %	22.0 %
Hemiptera	7.8 %	9.6 %
Hymenoptera	40.2 %	26.4 %
Lepidoptera	8.7 %	8.2 %
Neuroptera	0.33 %	1.3 %
Odonata	1.4 %	0.2 %
Orthoptera	5.8 %	14.0 %
Araneae	0.47 %	0.8 %
Archaeognatha2	0.0 %	0.1 %
Ephemeroptera2	0.0 %	0.1 %
Isoptera2	0.0 %	1.2 %
Mantodea2	0.0 %	0.1 %
Mecoptera2	0.0 %	0.2 %
Psocoptera2	0.0 %	0.3 %
Thysanoptera2	0.0 %	0.1 %
Trichoptera2	0.0 %	0.1 %
Undetermined2	0.0 %	0.3 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Asilinae in the NMSA: Blattodea
² No data for the following prey items of the Asilinae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined.

³ Specimen USNMENT01115713 (caterpillar) in the USNM Asilinae collection.
* Apocleinae (as used in Londt 2006) combined with Asilinae (see Dikow 2009, http://hdl.handle.net/2246/5949)

 

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2. Dasypogoninae

Observations Dasypogoninae

Hymenoptera is the most dominant prey item in both the USNM and in the NMSA Afrotropical Dasypogoninae collections, comprising 68.1% and 40.8% of the total prey items, respectively. Diptera is the second-most dominant prey item in both Dasypogoninae collections, comprising 14.4% of total prey items in the USNM and 25.5% in the NMSA Afrotropical Dasypogoninae collection.

Hemiptera comprises 6.9% of the total prey items of the Dasypogoninae in the USNM and similarly comprises 7.1% of the total prey items of the Afrotropical Dasypogoninae in the NMSA.

Lepidoptera comprises 2.5% of the total prey items of the Dasypogoninae in the USNM and similarly comprises 3.1% of the total prey items of the Afrotropical Dasypogoninae in the NMSA. 

Summary data Dasypogoninae

Comparison of the percentages of prey items (sorted to prey order) for the USNM Dasypogoninae and for the NMSA Afrotropical Dasypogoninae*.

Prey Order	USNM Dasypogoninae	Afrotropical Dasypogoninae*
Blattodea1	0.0 %	0.0 %
Coleoptera	5.6 %	16.3 %
Diptera	14.4 %	25.5 %
Hemiptera	6.9 %	7.1 %
Hymenoptera	68.1 %	40.8 %
Lepidoptera	2.5 %	3.1 %
Neuroptera	0.28 %	0.0 %
Odonata	0.56 %	0.0 %
Orthoptera	1.1 %	0.0 %
Araneae	0.56 %	3.1 %
Archaeognatha2	0.0 %	0.0 %
Ephemeroptera2	0.0 %	0.0 %
Isoptera2	0.0 %	3.1 %
Mantodea2	0.0 %	0.0 %
Mecoptera2	0.0 %	1.0 %
Psocoptera2	0.0 %	0.0 %
Thysanoptera2	0.0 %	0.0 %
Trichoptera2	0.0 %	0.0 %
Undetermined2	0.0 %	0.0 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Dasypogoninae in the NMSA: Blattodea
² No data for the following prey items of the Dasypogoninae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined.
* Prey counts not precise as the NMSA Afrotropical Dasypogoninae collection includes the Brachyrhopalinae within the Dasypogoninae.

 

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 3. Laphriinae

Observations Laphriinae

Coleoptera comprises 21.9% of the total prey items of the Laphriinae in the USNM and similarly comprises 22.9% of the total prey items of the Afrotropical Laphriinae in the NMSA.

Londt reports that the dominant prey item for Afrotropical Laphriinae is Hymenoptera (Londt 2006). In contrast, the dominant prey item for the Laphriinae in the USNM is Diptera, comprising 32.8% of total prey items. However, in the USNM, Hymenoptera still makes up the second-most dominant prey item, comprising 31.3% of the total prey items.

Londt reports that Coleoptera comprises 81% of prey items for the Laphystiinae (Londt 2006). Combining the Laphystiinae with Laphriinae, it is still evident that Coleoptera is the second-most dominant prey item for Afrotropical Laphriinae, comprising 22.9% of total prey items. Diptera is the third-most dominant prey item for Afrotropical Laphriinae, comprising 19.4% of total prey items.

Summary data Laphriinae

Comparison of the percentages of prey items (sorted to prey order) for the USNM Laphriinae* and for the NMSA Afrotropical Laphriinae.

Prey Order	USNM Laphriinae	NMSA Afrotropical Laphriinae
Blattodea1	1.5 %	0.0 %
Coleoptera	21.9 %	22.9 %
Diptera	32.8 %	19.4 %
Hemiptera	12.4 %	7.6 %
Hymenoptera	31.3 %	42.3 %
Lepidoptera	0.0 %	2.8 %
Neuroptera	0.0 %	0.0 %
Odonata	0.0 %	0.0 %
Orthoptera	0.0 %	4.9 %
Araneae	0.0 %	0.0 %
Archaeognatha2	0.0 %	0.0 %
Ephemeroptera2	0.0 %	0.0 %
Isoptera2	0.0 %	0.0 %
Mantodea2	0.0 %	0.0 %
Mecoptera2	0.0 %	0.0 %
Psocoptera2	0.0 %	0.0 %
Thysanoptera2	0.0 %	0.0 %
Trichoptera2	0.0 %	0.0 %
Undetermined2	0.0 %	0.0 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Laphriinae in the NMSA: Blattodea
² No data for the following prey items of the Laphriinae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined.
* Laphystiinae (as used in Londt 2006) combined with Laphriinae (see Dikow 2009, http://hdl.handle.net/2246/5949)

 

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 4. Stenopogoninae

 Observations Stenopogoninae

Neuroptera comprises 0.89% of the total prey items of the Asilinae in the USNM and similarly comprises 0.33% of the total prey items of the Afrotropical Asilinae in the NMSA.

 Londt reports that Coleoptera is the most dominant prey item for the Afrotropical Stenopogoninae, comprising 35.1% of total prey items (Londt 2006). However, in the USNM Stenopogoninae collection, Coleoptera is one of the least dominant prey item, comprising only 6.3% of total prey items found with USNM Stenopogoninae.

The most dominant prey item in the USNM Stenopogoninae is Hymenoptera, comprising 41.1% of total prey items, and Diptera is the second-most dominant prey item, comprising 27.7% of total prey items. In the NMSA Afrotropical Stenopogoninae collection, Hymenoptera follows Coleoptera as the second-most dominant prey item, comprising 21.3% of total prey items, and following is Diptera, comprising 14.4% of total prey items.

There is a discrepancy between the dominance of Coleoptera as a prey item between the two collections: while it comprises only 6.3% of total prey items for the USNM Asilinae, it comprises 35.1% of total prey items for NMSA Afrotropical Stenopogoninae. While it is clearly a dominant prey item for the Afrotropical Stenopogoninae, in the USNM, Coleoptera appears to be one of the lesser dominant prey items.

Summary data Stenopogoninae

Comparison of the percentages of prey items (sorted to prey order) for the USNM Stenopogoninae* and for the NMSA Afrotropical Stenopogoninae.

Prey Order	USNM Stenopogoninae	Afrotropical Stenopogoninae
Blattodea1	0.0 %	0.0 %
Coleoptera	6.3 %	35.1 %
Diptera	27.7 %	14.4 %
Hemiptera	5.4 %	14.1 %
Hymenoptera	41.1 %	21.3 %
Lepidoptera	8.0 %	2.3 %
Neuroptera	0.89 %	0.33 %
Odonata	10.7 %	0.66 %
Orthoptera	0.0 %	8.9 %
Araneae	0.0 %	0.7 %
Archaeognatha2	0.0 %	0.0 %
Ephemeroptera2	0.0 %	0.0 %
Isoptera2	0.0 %	1.6 %
Mantodea2	0.0 %	0.0 %
Mecoptera2	0.0 %	0.0 %
Psocoptera2	0.0 %	0.0 %
Thysanoptera2	0.0 %	0.0 %
Trichoptera2	0.0 %	0.0 %
Undetermined2	0.0 %	0.0 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Stenopogoninae in the NMSA: Blattodea
² No data for the following prey items of the Stenopogoninae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined.
* Stenopogoninae (as used in Londt 2006) do not entirely coincide with Stenopogoninae as used at USNM (see Dikow 2009, http://hdl.handle.net/2246/5949)

 

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5. Stichopogoninae

Observations Stichopogoninae

Diptera is the most dominant prey item in both the USNM Stichopogoninae collection and in the NMSA Afrotropical Stichopogoninae collection.

While the NMSA only had seven prey items corresponding to Afrotropical Stichopogoninae (Londt 2006), the USNM had 72 prey items.

While Coleoptera and Hymenoptera each comprise 14.3% of the total prey items in the NMSA, in the USNM, there are 0% Coleoptera and Hymenoptera. 

Summary data Stichopogoninae

Comparison of the percentages of prey items (sorted to prey order) for the USNM Stichopogoninae and for the NMSA Afrotropical Stichopogoninae.

Prey Order	USNM Stichopogoninae	Afrotropical Stichopogoninae
Blattodea1	0.0 %	0.0 %
Coleoptera	0.0 %	14.3 %
Diptera	86.1 %	57.1 %
Hemiptera	5.4 %	14.3 %
Hymenoptera	0.0 %	14.3 %
Lepidoptera	0.0 %	0.0 %
Neuroptera	0.0 %	0.0 %
Odonata	0.0 %	0.0 %
Orthoptera	1.4 %	0.0 %
Araneae	8.3 %	0.0 %
Archaeognatha2	0.0 %	0.0 %
Ephemeroptera2	0.0 %	0.0 %
Isoptera2	0.0 %	0.0 %
Mantodea2	0.0 %	0.0 %
Mecoptera2	0.0 %	0.0 %
Psocoptera2	0.0 %	0.0 %
Thysanoptera2	0.0 %	0.0 %
Trichoptera2	0.0 %	0.0 %
Undetermined2	0.0 %	0.0 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Stichopogoninae in the NMSA: Blattodea
² No data for the following prey items of the Stichopogoninae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined. 

 

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 6. All Asilidae

Observations Asilidae

USNM data are in accordance with Londt’s finding that Hymenoptera is the most dominant prey item (Londt 2006). In the USNM, Hymenoptera comprises 40.2% of all prey items and in the NMSA, Hymenoptera comprises 26.4% of all prey items.

There is also no discrepancy between the two collections over which is the second-most dominant prey item. Diptera comprises 28.6% of all prey items in the USNM Asilidae collection and 22% of all prey items in NMSA Afrotropical Asilidae collection. However, when it comes to the third-most dominant prey item, Lepidoptera is the third-most dominant in the USNM, comprising 8.7% of all prey items, while in NMSA Afrotropical Asilidae collection, Coleoptera is the third-most dominant prey item, comprising 14.7% of all prey items.

Summary data Asilidae

Comparison of the percentages of all prey items (sorted to prey order) for the USNM and for the NMSA Afrotropical Asilidae. 

Prey Order	USNM Asilidae 3	NMSA Afrotropical Asilidae
Blattodea1	0.09 %	0.0 %
Coleoptera	6.6 %	14.7 %
Diptera	28.6 %	22.0 %
Hemiptera	7.8 %	9.6 %
Hymenoptera	40.2 %	26.4 %
Lepidoptera	8.7 %	8.2 %
Neuroptera	0.33 %	1.3 %
Odonata	1.4 %	0.2 %
Orthoptera	5.8 %	14.0 %
Araneae	0.47 %	0.8 %
Archaeognatha2	0.0 %	0.1 %
Ephemeroptera2	0.0 %	0.1 %
Isoptera2	0.0 %	1.2 %
Mantodea2	0.0 %	0.1 %
Mecoptera2	0.0 %	0.2 %
Psocoptera2	0.0 %	0.3 %
Thysanoptera2	0.0 %	0.1 %
Trichoptera2	0.0 %	0.1 %
Undetermined2	0.0 %	0.3 %
TOTAL	100 %	100 %

¹ No data for the following prey items of the Afrotropical Dasypogoninae in the NMSA: Blattodea
² No data for the following prey items of the Dasypogoninae in the USNM: Archaeognatha, Ephemeroptera, Isoptera, Mantodea, Mecoptera, Psocoptera, Thysanoptera, Trichoptera, and Undetermined.
³ The following USNM Asilidae subfamilies had no prey items in the collection: Leptogastrinae, Ommatiinae, and Trigonomiminae.

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Total prey item numbers

Discussion

In the future, the next step in this project would be to sort through the USNM Asilidae prey item collection and sort all the Asilidae based on zoogeographical regions. We would separate subfamilies with prey items based on geography. In this way, we would then be able to compare the predatory habits/prey items between NMSA Afrotropical Asilidae flies with, for example, Nearctic or Oriental assassin flies. Furthermore, we could compare all the USNM Afrotropical Asilidae flies with the Afrotropical Asilidae flies studied by Londt to see if there are any differences.

This would allow us to determine if Asilidae subfamilies have preferential prey items. For example whether one Asilidae subfamily preferentially prey on Coleoptera while another Asilidae subfamily preferentially prey on Hemiptera.

Gauromydas recorded from Peru for the 1st time

I was handed a specimen of mydas flies (Mydidae) by my colleague Norm Woodley the other day that was collected by USNM lepidopterists in 1989 in the Amazon basin of south-eastern Peru, specifically in the Madre de Dios Region. It was tugged away in a drawer and had not been incorporated into the main collection.

The specimen seemed interesting as there are only four species of Mydidae known from Peru in the genera Messiasia Andretta, 1951, Plyomydas Wilcox and Papavero, 1971, and Stratiomydas Wilcox, Papavero and Pimentel, 1989 (see catalog of Peruvian species here). This specimen, however, is very different and belongs in the genus Gauromydas Wilcox, Papavero and Pimentel, 1989.

Gauromydas includes the largest known fly, i.e., Gauromydas heros (Perty, 1833), distributed in south-eastern Brazil (see distribution here). The entire genus is only found in north-eastern South America and north-eastern Argentina, southern Brazil, and Paraguay (see distribution here). To my knowledge, it is so far unknown from the western Amazon Basin so that this unique specimen is quite exciting. Nelson Papavero, in the most recent catalog of Neotropical Mydidae (Papavero 2009), also does not record this genus from this part of the Amazon Basin.

There are four valid species of Gauromydas known to date (see species catalog here). While the specimen is morphologically similar to Gauromydas heros and keys out to that species in the Wilcox et al. 1989 key (electronic version available here), I am hesitant to call it that species because it is so far away from other collecting localities of G. heros.

Full occurrence data soon to be available in the NMNH collections database - USNMENT01115313.

It's always surprising to see what specimens might come your way.

Torsten Dikow @TDikow #asiloidflies