Diptera - Science topic
An order of the class Insecta. Wings, when present, number two and distinguish Diptera from other so-called flies, while the halteres, or reduced hindwings, separate Diptera from other insects with one pair of wings. The order includes the families Calliphoridae, Oestridae, Phoridae, SARCOPHAGIDAE, Scatophagidae, Sciaridae, Simuliidae, Tabanidae, Therevidae, Trypetidae, Ceratopogonidae; Chirnomidae; Culicidae; Drosophilidae; Glossinidae; Muscidae; Tephriridae; and Psychodidae. The larval form of Diptera species are called maggots (see larva).
Questions related to Diptera
I recently analyzed the video data of climbing assay in drosophila using Fiji. I used a valid plug-in for tracking flies. At the end of the analysis, Fiji converts the results to excel documents (.xls). I suppose to have sequentially decreasing data in the excel files, but I get a series of crazy numbers instead. Besides, some of the data (numbers) are changed to dates. Please find the attached file to have a look at.
When I try to analyse the videos on another computer (Imac), there is no problem with the excel files. I can get the proper results without mistakes. Something seems wrong with my computer (Mac) or program. Everything is updated. Has anybody had a similar problem before, or do you have any idea what's wrong with Fiji on my computer?
I appreciate your help.
The stomach content belong to Lepomis gibossus also known by the pumpkimseed sunfish who lives in fresh water . So please I want to know the ID of the this diptera ( family, genus... ).
I'm an undergrad biology student from Denmark, and i work on a project with D. melanogaster. We're having a problem we can't figure out, and therefore i've created this account, hoping some of you have had the same experience with the CAFE and knows the reason.
We're feeding D. melanogaster with the Capillary Feeder Assay (CAFE) with 5 μL capillary tubes. Our problem is that after we've been feeding the flies for 24 hours or so, an air pocket starts to form in the bottom of the capillary tubes (green arrow, see attached picture), therefore making the liquid food inaccessible to the flies. The liquid food should "fall down" after the flies drinks from the capillary tubes, but instead this air pocket forms. This happens to at least 9/10 capillary tubes. The red ring (see attached picture) is how the capillary tubes should look like with liquid food and no air pocket in the bottom.
We're feeding them with 5 % sucrose and tap water in both 20 and 23 degrees Celsius.
Capillary tubes are 23 mm long and made of glass. Unknown inner or outer diameter. The capillary tubes we use: https://www.sigmaaldrich.com/DK/en/product/sigma/p1799
I hope the problem is clear and that i've provided all the necessary information
I knocked in a fragment of about 1000 bp to the fly genome. In order to validate that the knock in worked, I designed primers roughly 250 bp upstream of the insert, and 250 bp downstream of the insert, expecting that any fly not containing the insert would show a 500 bp product and any fly with the insert would show a 1500 bp product, using the Pfu enzyme. However, when I run the PCR, I only get the expected 500 bp band on yw flies that didn't undergo the knockin at all, and no bands in my experimental flies. I tried doing a touchdown PCR, but that also came up blank.
I am trying to rear D. hydei on Carolina Instant Medium (in an incubator at 25C with 12:12 L:D). I have not been adding yeast. The flies are surviving but not thriving. I’m looking for recommendations to improve things.
Ultimately, I want to observe oviposition behavior. I’ve read females begin courting much earlier than males. At what age have others tested egg-laying? Males and females same age? How long to allow them to lay eggs? Since they re-mate often, should I keep the males in the oviposition chamber? With D. melanogaster, I use 4-day old females that have been kept continuously with males but I put only females in the oviposition chamber (overnight for ~15 hours). Wondering how I might need to modify this protocol for best results with hydei.
Hello, can someone help me with an estimate of known species for Diptera, Coleoptera, Lepidoptera and Hymenoptera in these two territories? I have searched without much success, it seems as if all the papers I have reviewed shy away from offering a number of species.
Thanks in advance!
I am trying to express a short RNA sequences in the nucleus in Drosophila. Are there any UAS-based expression vectors in flies that do not have PAS in the 3'? Or on the other hand, is it possible to just express a transcription termination sequence at the 3' of my transcript so that my transcript terminates before reaching the PAS? I don't want to have to mess with the vector backbone itself (removing PAS...etc.). I am quite new to molecular work regarding vectors, so if I am missing something important, under a misunderstanding of sorts, or if you have good suggestions, please let me know. Thank you!
Dear colleagues, I am specialist form Chironomidae, but not much familiar with the midge larvae from the other families. I have one, strange to me, from spring in mountains from Kosovo. Would you help me with the identification? Maybe you could give me at least the family or maybe genus or species?
I have a dataset (fastq files) of 15 fastq flies uploaded on Galaxy bioinformatics portal. I deleted them some time back to free some space. Now I want to retrieve these files. I can see these files as deleted, but not able to restore or download them. Is there any way to get these files back on my portal.
I'm looking for articles where I can find tests of food attractants used to monitor fruit flies. I am not looking for pheromone-based attractants.
We have already performed a series of tests where we tried to have a glue trap (containing aroma dissolved in propylene glycol) and the raw materials from which the aroma was made were used next to them (eg grapes and grape aroma). Fruit flies almost always chose the raw material itself and they were not interested in the aroma in the glue. Even after removing the raw material itself, the glue trap with aroma did not attract them.
I will be happy for any of your knowledge and articles on the topic.
I found this insect in the digestive tract content of Hypoatherina temminckii (marine fish). The sampling location in seagrass waters of Karang Congkak Island, Kepulauan Seribu (Seribu Island), Indonesia. I can't identify the insect groups, I just suppose this is part of Diptera but have never seen marine (or semi-aquatic) insects in my sampling location.
i have been working on fly of asilidea family i want to ask you how i can collect these insects specially from soil? and if there are more of them in places with herbaceous covering or not?
I'm using Ctrax to autotrack multiple flies, the videos have a good quality but when I run fixerrors script I have a lot of errors. I tried to change many parameters but I always have lots of errors (2000 and more) Does anyone can help?
Thank you in advance
We are looking for a suction tool/machine that would allow us to automatically count small insects (aphids, fruit flies) as they are sucked into a tube or container. It could be a counting with a laser cell, for example, or any other method.
To be clear: we would like to count aphids on infested plants, and one easy solution would be to use a suction/vacuum device (active sampling) so the insects would be counted as they are sucked into the device.
Do not lose focus trying to kill the flies, rather work hard to get rid of what's attracting the flies - is this the right problem-solving philosophy/mindset for successful research? How else can a research approach the fly-source problem?
Which chemical is best for preservation during the time of collection and after some months I need to identify them.
Could anyone please explain if a drosophila melanogaster line can express some white eye flies whereas its wild type is red eye. Is this fading out possible or is it a contamination?
Tachinidae flies are difficult to identification , so , I need classification key to the species which parasitoid on Lepidoptera>
I'm trying to use this DAC technology in the aviation industry. Can anyone suggest some ideas on how to do so? I too have some ideas and am willing to work on them.
My work will be designing a cost-efficient module that can be used for carbon capture. If you have seen a CC plant, there are large fans that suck the air, I thought of eliminating that energy of running that motor. As the plane already flies through a lot of air in its journey, we can almost make the plane carbon neutral.
Carrion Ecology: What is the best way the statistically analyse the correlation of bacteria community data to volatile profiles and to fly community data across 3 time points using ordination techniques in R? I have data of carrion-associated bacteria (snout swabs, NGS) for timepoint 1,2,3 and cadaver volatiles (GC-MS data) for time points 1,2,3 together with blow fly, flesh fly and true fly abundance and species data for time points 2 and 3 derived from 75 stillborn piglet cadavers. Cadavers were exposed in differently managed forests in 3 study regions (75 sites). I want to correlate the bacteria matrices with volatile matrices and fly matrices to see if there is a temporal change of bacteria, volatiles and flies and want to look for indicator species. I want to use ordination methods not to loose species information. I read about coinerta analysis but I have more than one matrix to correlate with traits. Do you have any fancy idea how to analyse the data?
Please, delete Marco Bonelli from co-aouthorship of the paper Revision of Carexomyza Roháček with descriptions of three new Nearctic species (Diptera: Anthomyzidae) !
He has been generated as co-author without any my action. Terrible!
Scientists have invented a panacea which may expand the life span of human being. A group of researchers of University of Southern California Dornsife College of Letters, Arts and Sciences has revealed a report in this regard. Journal of Gerontology: Biological Science has published this report. The report shows that the new medicine Mifepristone may bring unprecedented success for the human being and other species. Applying this drug on the body of flies it has been found that the life span of female flies that interact with male flies has been increased. At that time their fertility decreased while their immunity has been increased. Researchers hope that same thing will happen in case of human being after applying this medicine upon them. If this endeavor becomes success, are the human beings going to live long or be immortal?
Hello everybody, I am not a specialist and I want test the effect of some substances on mosquitoes and flies but I have no idea how can I get them and how to keep them alive for treatment?
Thanks in advance
I would like to visually identify and quantify the pollen morphotypes carried by insects which have been stored in 70% ethanol. The insects are Diptera (mainly Syrphids), and Hymenoptera (mainly wild bees, bumblebees and honeybees). To facilitate pollen identification, we have collected the anthers of all flowering plants species encountered on the studied sites, and also stored them in 70% ethanol.
Firstly, is it possible to visually identify pollen after it has remained for several months in alcohol, because I read about pollen hydration that could cause damage to the exine? If it is, what is the best way to collect the "free" pollen from ethanol and the pollen that remains on the insect's body? Then, is there a specific method for preparing pollen that has been stored in 70% ethanol for visual identification under the microscope?
Thank you very much for taking the time to read and answer!
I understand that light is one of the strongest external cues to affect our clock system. I am curious about other possible factors that are less discussed. I am finding a great amount of literature of EMFs on insects such as fruit flies, but when it comes to human studies the results are always contradictory.
I would be open to any suggestions on researchers and the latest studies in the field.
I have some Thaumaleidae larvae from Bosnia and Herzegovina. Using key of Disney (1999) I identified them as Thaumalea testacea and Thaumalea verralli. Disney lists in his key only T. verralli, T. testacea and T. truncata whereas Fauna Europaea give a checklist of few dozens of them for Europe. Are there any other keys for Thaumaleidae larvae which I could use for my material identification, and what is current list of taxa recorded from Bosnia and Herzegovina?
Thank you for any help..
Does anyone know a fast and simple method (apart of using tweezers) to sample ectoparasites (especially bat flies, Nycteribiidae, but also mites and ticks) on small bats (15 g)? We release the bats after sampling.
The hexagonal boron nitride powder synthesised via the tube furnace flies off from the crucible during the synthesis and gathers at the cooler end of the furnace.
Can somebody suggest a possible solution for the same as all my synthesised products are gathering at one end of the tube furnace
Does insect such as flies and mosquitoes transmit the COVID-19 virus as a mechanical transmission?
Virus transmission take place by insect legs, mouth parts and other body parts.
When crossing grey body flies with normal wings, 25% of their progeny had a black body. Approximately 25% of all daughter individuals did not have fully developed wings. What traits dominate? What genotypes do parents have?
I have a problem confirming some specimens which identified as B. neocognata using the Dorsalis CD-ROM however the costal bands of my specimens are confluent R+3. In many references such as Dorsalis CD-ROM, Drew & Hancock, 1994, and Drew & Romig, 2013, it's written that B. neocognata costal band is overlapping or slightly overlapping R2+3. Is there any chance of B. neocognata with costal band confluent to R+3?
Thank you very much.
I am trying to do a 3 point bend test for a lattice structure. The beam is already latticed and is a nodal structure. The pins are discrete rigid shells.
3 interactions were created where the surface of the pins were made Master and the slave being a nodal set consisting of just the top most nodes of the latticed beam. This can be seen in the image. The friction defined is 0.2.
The simulation runs but in the animation, the beam flies out instead of bending. I have attached the video file here.
Any advice on improving this would be very helpful. Is there something missing in the interactions or the way I am defining it?
Thank you for your inputs!
I spent all day reading about stats, and I remember exactly none of it. Basically, here's the problem:
I have 2 different morphological trait that are continuous(ish) variables that could, if necessary, be treated as categorical. I want to see if they co-occur significantly.
The issue is my supervisor wants me to take this and compare it to the phylogenetic tree of Syrphidae using "statistics". I... do not know how to do that?? There's so many papers on this topic, with so many different possible routes to take, but a lot of them seem unnecessarily convoluted and complicated. Where is a good place to start?
I am doing a research on fruit flies (Bactrocera zonata) and i have tried to rear it many times in a cage under laboratory conditions but i didn't get any egg and even there were no mating. The flies well lived for more than a month but without mating and eggs.
Please give me your suggestions and advises that help to induce mating and egg laying.
We are currently planning to do experiments where we want to study differential gene expression in both tephritid fruit flies and fly symbionts at the same time. We are however a bit unsure about how to do the rRNA depletion step because several more commonly used universal depletion kits do not appear to be produced. We are aware that recently some new kits that can be applicable to this have been made such as Ribopools and Zymobiomics universal rRNA depletion kits, but given their recent arrival we are unsure about them. Are there people here who have experiences with these kits? In addition, we read in literature also about some alternative approaches: Terminator™ 5′-PhosphateDependent Exonuclease and DSN. We do not have much experience in transcriptomics at the moment though so we are curious about what others think about these approaches. I do know that DSN can also reduce very common mRNA transcripts but i thought that it might be an advantage in this case as this would lead to more symbiont transcripts being recovered.
I have become interested in potentially doing a small study on non-bee pollinators by looking at what pollen are present on their bodies. I am unsure whether or not this would be a useful contribution, since while there appear to be many studies on the role of non-bee pollinators, there doesn't seem to be many studies indicating the state of the literature.
That being said I did find this paper which identifies some gaps in the literature but it is now somewhat out of date and only focuses on flies and not other pollinators.
I would appreciate any insights into the state of the literature on non-bee pollinators and their contribution(s), as well as any interesting papers etc.
In particular is it known:
- Which plants each pollinator is responsible for pollinating and in what proportions
- Whether species labelled as pollinators actually pollinate or simply visit plants?
- Are there any species in which it is contested whether or not they act as pollinators?
- Are there any obvious gaps in the literature?
Thanks in advance! I look forward to hearing your thoughts!
I'm trying to understand the protocol to make the CRISPR mutant stock. I have transgenic Vasa cas 9 (X chromosome) and gRNA (2nd chromosme) flies. I start by crossing cas 9 males with gRNA female flies and then I'm lost. Can anyone help me out on how to go ahead and get a balanced stock?
A high rate of modifications in wing venation in a small population of a species X (Diptera: Tipoloidea) could be the result of low genetic elasticity? Would this fact support our hypothesis that it is a relict and isolated population?
I have added some Background and Facts, which could be useful.
Background: In a study in hypogeal environments we have found a species of cranefly that is apparently isolated (trapped) in the cave (outside the cave conditions are not suitable for survival). We think that it is an eutroglophile species, of which only this population of the cave has survived, losing the population of the external environment in the past.
Fact: We found anomalies in the wing venation of almost half of the specimens. Although the wing venation in the cranflies may be surprisingly prone to individual anomalies (including the addition of crossveins), I do not think it is sufficient to explain such a high rate of anomalies as those found.
I ran an experiment to evaluate fly attraction to a number of bacterial cultures (n=50). Briefly what I did is: 20 male flies were exposed to two odour sources at a time-one containing a bacterial culture and other containing control; number of flies that showed attraction to the culture and control were counted after a certain time. The test was repeated 10 times for one bacterium. Similarly female flies attraction was also tested for the same bacterium. Thus for one bacterium I had 20 replicates in total (10 for males and 10 for females). Now, I want to analyse the data to evaluate which bacterium showed attraction at what extent and how they differ gender-wise. I am not certainly sure which statistical test will be suitable for this purpose. I was thinking about Kruskal-Wallis test but not sure if its the correct one. I am using SPSS. Could you please suggest which statistical test will be suitable to analyze this data set?
Thanks in advance.
The wound myiasis causing flies of animals are potential human myiasis agents. Monitoring the animal myiasis helps to more precisely identify the dominant and aggressive species in the province.
Dear all, we are now working evaluating the effect of a plan in a fly-diabetes type II model. This is the first time we are using this model, therefore, I would like to ask you: why are flies being fed since they are pupae? Is it more logical to feed them once they are adults? Thanks in advance for your help.
I am trying to amplify intronic region of miR-1017 from Drosophila genomic DNA. Have done this several times before with no issue. Currently I am screening flies for mutation and for the purpose I am trying to PCR amplify the region. But the same working conditions isn't working anymore. I have tried with fresh primers, different set of primers, proteinase K treatment of template. It works for few round and then I start getting blank gel picture only with the ladder.
Any suggestions on this?
I am trying to find out how the mating staus (virgin or non-virgin) of female flies affect their response to heat stress i.e. an exposure to ~39 degrees. Does virgin females take longer to be knocked down as compared to the non-virgin females.
I have performed a TAG assay on adult male and female Drosophila,
however, I seem to end up with other values that that are reported in literature (40-50µg per mg fly).
My relative levels of TAG when I compare male and females have the expected fold increase in female flies.
I'm afraid I'm doing something wrong in my calculations.
Would someone be willing to share some data (that they are not using) just to check whether I'm doing something horrendously wrong?
Thanks in advance
I want to watch development of carnivorous larvae (diptera larvae) by giving a solid chemical that does not dissolve in water. How do I determine doses? how many meat do I need to put in for how many larvae? What should be the unit of doses (mg/g, mg/larvae)?
I would appreciate it if you help me design the experiment...
I'm currently doing some work with molecular identification of flies using the COI region for my masters. All the flies came from the same location. These particular three flies were identified as Peckia (sarcodexia) lambens after Genbank search. Following alignment and tree building using reference sequences obtained from Genbank, two of the three flies showed 100 percent identity with each other (where as they showed slightly lower with the third fly). On the tree, they clustered more closely together on a sub branch on their own, whereas the third fly clustered with a reference sequence. I'm not really sure how to explain this. Any ideas?
We are attempting to rear Phytomyza gymnostoma with little success. Counterparts in other areas are also struggling. Does anyone have any experience rearing this species or other Phytomyza? Recommendations of relevant literature are also appreciated.
A little about this fly in the Northeastern US:
This fly has two generations - one in spring and one in fall - with an aestivation and overwintering stage in between. The adults emerge when temperatures are ~18C and feed on plant exudates from puncture wounds made by the female flies in allium plant species. Oviposition occurs on the plant leaves and larvae feed in the leaf tissue. Pupation occurs either in the plant tissues or in the soil immediately around the base of the plant.
We are currently storing the fall generation in a refrigerator for ~3 months then transferring to 25C, 55% RH, 16:8 L:D. The eclosion rate is ~65%. We are unsure what conditions to maintain adults at and have settled on 18C, 55%RH, and 16:8 L:D. We provide water and onion plants on which flies can feed and develop. The adults only live about a week.
We are unsure about whether the spring flies will need to be stored at aestivation temps prior to eclosion. Not much is know about this species so any information is helpful.
I study the community of predators on gregarious larval web of Euphydryas aurinia butterfly. I photographed a fly belong to Odontomyia (Diptera, Stratiomyidae) and I would understand what the potential reason (diet?) of its presence on butterfly nest. Thank you very much for your help!
Good evening to you all.
I have modeled a bogie using Simpack software and i tried to connect the primary suspensions, secondary suspensions but this bogie leaves the track (flies) and after 4 seconds, it comes come back on the track and moves normally. I am confused and i dont know why this is happening? I have followed the tutorials on youtube but they are incomplete. Please help. Thank you.
I need this article to explain the relationship between hoverflies and certain plant species.
I have flies in which I have knocked out a specific (non essential) sugar transporter and I've noticed many of them turn and almost ebony colour when dead. Does anyone know why this might happen?
Thank you in advance
Does anybody know dipteran species which oviposit NEAR feeding substrate but not IN it?
So far, I could only find dipterans that oviposit directly in the feeding substrate for the developing larvae (Drosophila, blow flies, houseflies... ).
Does anyone have an idea?
There is always a lot of confusion and negative reaction when I discuss female intrasexual competition with lay people, especially women. Many feel the "female" is unnecessary and sexist or that it reduces all female disagreement down to the fight for mates. When I try to explain that the "sex" in intrasexual is a noun not a verb this rarely helps. There seems to be a taboo to even discussing female competition - even just admitting it exists. Women generally view it as negative even though competition can be healthy and is essential for cooperation. Understanding the mechanisms and anatomy of fic will also help women and girls by enabling us to target female bullying in schools and other institutions, where at the moment it flies under the radar.
I've been reading some research lately which differentiates FIC with female social competition (fsc) and discusses the pros and cons of using the different terms: fic for competition for mates and fsc for social competition. I'm not sure I see the utility as both are competition for fitness enhancing resources - BUT I also see substantial resistance by women in even discussing FIC at all.
I am searching for some genera or species in Brachycera that have hairs on wing veins. If anyone has an idea, I would be interested to read it !
Thanks in advance,
I need some good published paper about Clove (Syzygium aromaticum), Hing (Asafoetida), and Wood ash to control insect pest specially aphids, white flies and Spotted Boll worm?
if some one have these papers (Clove (Syzygium aromaticum)/, Hing/ (Asafoetida), and Wood ash). Kindly share with me.
I'm doing a longevity assay in drosophila on a high fat diet. A JoVE article collected flies and aged them for an additional 5 days on normal food before putting on the experimental or control diet ( doi:10.3791/56029) Most other papers I've seen do not age the flies after collecting, is there a reason for the aging that JoVE did?
Hi RG people,
I've been calculating invertebrate abundance through collecting soil cores and then correcting the number of inverts against the dry mass of the core collected. I started doing this as the soils are shallow (polar) so depth variable. However, most of the inverts are closer to the surface (collembola, mites, diptera larvae).
On inspection of the data the mass correction is quite drastic, turning non-mass corrected sites from high to low abundance and vice versa.
What do you think? Mass correct, or not?
Look forward to hearing your thoughts!
I found something so strange when importing a modeling file from Catia to Ansys workbench. When I import the model file which is an impeller upon hiding one of the blade's outer surfaces (as in the attached figure), the most inner part of the blade wall which is connected to the curved guide plate is gone. I tried to hide also the same surface in Catia, it seems the same with Ansys (Please see attached file). Why Catia is doing so knowing that the curved guide plate was revolved as a unique surface but now it seems some portions are gone. Is it a sort of demonstration problem or that section is really flied away and not there?
Hi there, researchgaters...
I am trying to find descriptions of the genitalia of Hermetia illucens, the black soldier fly (Stratiomyidae), but no luck.
I identified a couple papers that seem to contain this information:
- Iide, P. & Mileti, D.I.C. (1976) Estudos Morfológicos sobre Hermetia illucens (Linnaeus, 1758) (Diptera, Stratiomyidae). Revista Brasileira de Biologia, 36 (4), 923–935.
- McCallan E. 1974. Hermetia illucens (L.) (Diptera: Stratiomyidae), a cosmopolitan American species long established in Australia and New Zealand. Entomol. Mo. Mag. 109: 232–234.
If anyone out there has PDF of these or any other paper containing photos or images of the genitalia of this species, I will be most grateful to get a copy!
Thanks in advance.
As I'm working on this group and try - with colleagues - to work on the phylogeny of this family (with subfamilies and more), is was nice to get more experience with more material from the period of that we have only very limited information.
Sincerely Rüdiger Wagner
I am looking for insight into transmission of parasites via botflies, specifically ectoparasitic Diptera.
I am studying a Dipteran fly whose larval stages are ectoparasites in birds’ nests, eating blood and tissue of nestling birds, while the adult flies are known so far to eat bird feces and fermenting fruits.
I am trying to find any information on previous literature, observations, and possibilities for ADULT flies (not the larval stage), like the species I study, to transmit blood parasites to birds. (other examples of this transmission to other animals would be welcome too).
Generally, it seems so far blood parasites need a biting vector to take a blood meal from the new host for transmission to new host. The mouthparts of my fly species of interest has a proboscis with lapping/sucking ability, but no evidence of teeth, so it does not seem suited for latching onto flesh or creating a wound. Despite this, we still have interest to ask for insights, advice on if blood feeding could be a possibility for this species/species with similar anatomy.
Any advice, thoughts, information is welcome. Thank you for your help!
There is plenty of information available about the Hemotek system working well for adult mosquitos and ticks. Does anybody have experience using it for blood-feeding Diptera larvae? We are trying to improve the rearing system that we are using to rear larvae that are ectoparasites of birds and are wondering if this is a good option Many thanks
In our case, prey might be both terrestrial and aquatic so I need primers that amplify at least:
- Diptera (including chironomidae)
... without amplifying spiders !
Of course, I have already selected LCO 1490 / HCO 2198... But maybe there are other options...
Some month ago, I met a colleague of you in Göttingen and he told about project. I'd be very much interested inamber Psychodidae. I'm retired and have now time to look for phylogenetics of that group. Probably you know some papers of that I was (co-)author to give you some idea what/who I am.
With best wishes
I'm currently undertaking a project that seeks to asses the distribution of a minute gall midge, Arthrocnodax fraxinellus, and its associated parasitoids (Aphanogmus spp.) in Europe.
I'm asking for material of ash cauliflower galls (Aceria fraxinivora) on ash (Fraxinus spp) as the gall midge feeds on the mite in the larval stage.
Material from the following countries are of interest:
Bosnia and Herzegovina
I have attached a PDF with details about the project - please have a look.
Thanks in advance!