Asiloid Flies

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.