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Insect - Science topic

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Other than the dead heart?
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Thanks much
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Hello everyone!
Recently, i have started working with SF9 cell line from thermo (Cat No. B82501) for which i was able to culture and grow the newly received vial in complete Grace's insect media at 27 degree Celsius in a non-humidified incubator as per the instructions (bright field images attached for reference). But i am not able to witness the viable cells after revival of frozen cells from previous batch. Approximately 10*7 cells/ml cells were frozen with 80% complete Grace's insect media, 10% heat inactivated FBS, and 10% DMSO as per instructions. Even though the doubling time is 24-30 hrs, there is slow growth of cells even after 7days.
Seeking expert advice regarding the possible reason for such slow growth of cells and how to enhance the culture conditions.
Thanks
Kanan
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Wen Liu is it required to use medium supplemented with 20% FBS when subculturing the culture or we use it only in the first step?
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Cameron (2014) reports that the mitogenomes of 28 insect orders have been sequenced so far. Is there a recent publication that presents similar data for different taxonomic levels?
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Agreed with Dong. Just search in NCBI and count it.
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I want to research the effect of black aphids inside the plant
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The thickness and structure of plant cell wall may affect the percentage of aphid attack.
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    My research topic is to explore the biogeograpgic patterns of species richness of insects. I have the regional richness data of all insects and different orders from many locations. It's well known that insects include c. 30 orders with different numbers of species and phylogenies. I want to group different insect orders into several groups, and make a clear description of their diversity patterns. The problem is in grouping different insect orders into several groups.
    I'm also looking for someone interested in this project. Please contact me if you want to join me. 
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By taxonomic characters for example the order of Lepidoptera includes butterflies
The diptera order include flies
The hymenoptera order includes bees, wasps, hornets and ants
Coleoptera includes bettles
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I am involved in a project on biological control of the Comstock mealybug Pseudococcus comstocki in Switzerland. As part of this project, we are doing host specificity tests of a parasitoid and, besides P. comstocki, have tested so far the following non-target species: Pseudococcus longispinus, Planococcus citri and Phenacoccus aceris. We would like to test more species of the family Pseudococcidae and are looking for someone in Europe who could give us an identified starting colony for this purpose.
Thank you for your help!
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Dear Jinan, thanks for your answer, that is good to know! Which species are those? Just in general whatever is attacking ornamental plants? Are there some dominant species? Greetings, Lukas
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I am working on my Tesis, and I need papers related with morphology and morphometry of cockroaches or any related insect (opthopteroids).
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thank you!
Best wishes
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We purchased a new CG-MS and the provider suggested to use intrastent grade helium (to be cleaned with filters) instead of analytical grade He. I have always used analytical grade before and I am unsure of the impact of the swap on the quality of my samples (I mostly analyse plant and insect volatile compounds, some of which are present in minute amounts). I would be thankful for any technical advice you can provide.
With kind regards,
Andrea
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As friends and professors have said, one of the best ways is to use alternative gas, where hydrogen and nitrogen can be the best choices.These articles can help you in this way
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Being from the same field of insect breeding, can I have a video of this project?
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I recently deployed some insect pitfall traps and in several of them I unfortunately caught mice. How can I avoid this? I have seen suggestions of using shallow traps or small ladders, what is the best way to obtain quality insect samples and minimize risk to small mammals?
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Great question. I have gone through the same issue while laying baited pitfall traps and have sadly captured reptiles in the trap, along with my study group of beetles. Please do checkout the discussion thread here for some ideas : https://www.researchgate.net/post/How-to-prevent-that-small-vertebrata-get-caught-in-regular-soil-pitfall-traps
I would also recommend you to go through the methodology followed by where their intent was to capture small vertebrates but exclude larger predators like mammals? Might be useful.
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I have recently deployed some insect pit traps in the field, but I am concerned they may flood and overflow during rainstorms. Does anyone have recommendations for solving this problem?
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If the container is made of plastic, drill a small hole in the the upper part such that the liquid will flow out of the container but the insect will be retained. Alternatively, install a "roof" (we used painted plywood) such that the rain will be deflected and will not enter the container. The roof will also prevent the liquid to evaporate quickly.
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Dear Everyone!
This insect inclusion is sitting a Cretaceous amber from the Carpathian Basin. I am looking for ideas on what this insect could be.
The dorsal side (?wings) might bear some scale-like structures. Any ideas are welcome!
Sincerely;
Márton
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Hi Márton,
It looks very similar to Alienopterix smidovae Hinkelman, 2021, this genus is tentatively included in Umenocoleidae, a enigmatic family within Dictyoptera.
Further reading:
Vršanský P, Sendi H, Hinkelman J, Hain M. 2021. Alienopterix Mlynský et al., 2018 complex in North Myanmar amber supports Umenocoleoidea/ae status. Biologia
Luo C, Beutel RG, Engel MS, Liang K, Li L, et al. 2022. Life history and evolution of the enigmatic Cretaceous–Eocene Alienopteridae: A critical review. Earth-Science Reviews 225: 103914
Luo C-H, Beutel RG, Thomson UR, Zheng D-R, Li J-H, et al. 2021. Beetle or roach: systematic position of the enigmatic Umenocoleidae based on new material from Zhonggou Formation in Jiuquan, Northwest China, and a morphocladistic analysis. Palaeoworld 31: 121–30
Hope it helps.
Best wishes,
Cihang
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Hi, I am looking for help emulating the process depicted in the image below. Essentially I am looking for a method to extract the juices out of feeder insects such as crickets, roaches, mealworms, etc in large quantities that ideally doesn't require any costly tools, without altering the juice in any physical (heating/desiccating) or chemical way. In the image below it was done without the use of a centrifuge, so I would like to avoid needing to purchase one. I'm aware that there are procedures for extracting hemolymph alone, but I need all liquid contents of the insects, not just the hemolymph. I tried using a juicer machine but all that did was sadly turn the insects into a paste. Any advice is more than appreciated, thanks!
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I want to do research on some insects organs association of functional morphology, there I want steps and procedure to make research. Please help me. Here I have attached pictures for reference
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Dear Babu Kes
The software product will suit you Aidos-X. It is written for entomologists. Many articles are devoted to ground beetles. If you have any questions write.
Regards, Sergey
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Would the dominance of one of the microbes in the gut change if we give different foods to insects or larvae? Suppose the insect is supplemented with amylase-producing bacteria, fed with carbohydrates such as rice, corn, and other sources.
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Kindly check the following RG link in which a representative strain collection of dominant aerobic bacteria from black soldier fly larvae (Hermetia illucens, BSFL) has been established:
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We are running qPCR experiments with cDNA samples derived from insect tissues using a QuantStudio Real-Time-PCR-System from Applied biosystems. Since a few weeks, we are facing severe problems with S-shaped or sigmoid amplification curves, leading to extremely decreased CT values for the relevant PCR samples. Intriguingly, comparing two technical replicates running in two adjacent wells, one sample is associated with this problem while the other one is normal. This is demonstrated in the attached figure that shows the amplification curves of two technical replicates: the one on the right hand is normal whereas the other is sigmoid.
The problem is not linked to a specific primer pair since it occurs with different primer pairs that have been used in the past without any problems.
Does anyone have a good idea how to avoid this problem in the future?
Best regards,
Joerg
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The problem comes from difficulties of the software to recognize the correct exponential phase. This point needs to be estimated to determine the range of the baseline fluorescence, which is subtracted from the curve. In your case, the software seems to wrongly interpret some early signal increase as he exponential phase. You can solve this problem by adjusting the settings for background correction. You should at least be able to manually define the range of cycles from which the baseline fluorescence should be estimated (should be about 3-19 or 12-19 to exclude the "problematic" region in the early cycles).
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The targeted insects include fall armyworm and locusta migratoria. Need proteins study for further information. Suggest me proteins, database, or any research paper. Just share the specific insects proteins which are not involved in humans.
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For the DNA encoding any locust candidate protein of interest, you can perform a BLASTn search in NCBI to see if any part of it has homology to human genes: Nucleotide BLAST: Search nucleotide databases using a nucleotide query (nih.gov)
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Hello insect cell experts,
I am currently culturing Sf9 insect cells for a protein expression and noticed some tadpole-like cells (red circled in the picture). They seem to reduce/disappear when cells are close to confluence. Are they just unhappy Sf9 cells, or could it be a contamination?
These cells are recovered from a cryo-stock and not yet transfected.
For the culturing, I use supplemented Grace's medium with 10% FBS and Pen/Strep.
I would appreciate if anyone with insect cell experience could tell what they are.
Thanks in advance!
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The elongation and spreading are the properties of normal and healthy cells under tissue culture conditions. Since you have recovered these cells from a cryo-stock and supplemented Grace's medium with 10% FBS and Pen/Strep, the cells of your concern are happily growing in the culture flask. I could see some other cells that are spreading, widening and getting attached to the flask and must be having focal adhesion points on the bottom of the flask. These functions of the cells are markers of normal and healthy culture conditions indicating the availability of sufficient amount of nutrients and growth factors in the culture medium.
One the other hand, if there is shortage of nutrients and growth factors in the culture medium due to large number of cells in confluent conditions, cells feel stressed and then loos spreading-elongation & adhesion properties thats why you do not see these features in cells under confluent conditions. Cells, in order to protect from the stress insult and increase the possibility of survival, they tend to become rounded in order to minimize the surface area and thereby stress insult.
Hence, it is neither contamination not your cells are unhappy. These are the sign of healthy cells getting nutrients and growth factors in the culture medium and are really happy. So be happy.
For more details, you can see our original research publication in Journal of Cellular Biochemistry. The link is given below:
Modulation of a5b1 Integrin Functions by the Phospholipid and Cholesterol Contents of Cell Membranes. .J Cell Biochem 2000 Apr;77(4):517-28.
Best
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I need some transparent medium for mounting small insects. I need a medium that a bit dissolves soft tissues and not needs complicated chemical procedure with mounting.
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Thanks Chen - Digging further, I did get another possible place:
I looked into my ancient records: I got mine from Lonza Inc. It's called DANTOIN 739 there, and a search for that might give you some more paths to follow.
Best wishes, Owen.
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We found cotton fields of Nalgonda District, Telangana State, India
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Rakesh Davella , Oplodontha viridula doesn't occur in India. According to morphology and distribution, this is a female of Oplodontha rubrithorax, as I mentioned above. Please check the following paper:
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I will use it for insect DNA extraction. Can we use sterile distilled water to dissolve?
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Is morphological distinctiveness enough for separating a genus from another?
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Also check please the following useful RG link:
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Hello,
I am looking for a lab that can process insect DNA samples for me. More precisely, I would like to do hyRAD on my samples (they are not good enough for ddRAD). I am struggling to find one.
Thank you!
Sophie
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Tyler Chafin thank you very much !
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I am currently conducting an experimental study. I don't have any methods on how to count the generations that might be exhibited by the insect that I mass reared.
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Number of generation depend on the life cycle start from egg. the time period of one generation starts till the mortality of the insect.It mean one generation include egg, different larval period and adult longevity.
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In our case it is impossible to traced from where is originated (species is clearly different from all others and belongs to group with limited distribution in Middle East).
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Tamara: And if Ruben's suggestions have not helped, describe the species (leaving "type locality unknown") to make other students aware of its existence: maybe it turns out to be already present in some collection but either unidentified or misidentified; or hopefully somebody, knowing what to look for, will find and colect it.
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I'm trying to get some good microscopic photographs of insect genitalia for my research work but I'm having trouble removing the air bubbles that comes inside the genitalia, no matter how cautious I'm.
Suggestions are appreciated.
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How to make a model any one have experience? About insect sustainable waste management, economics value and market demand
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No newly described order as such. Why?
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Emana: It is quite simple: there really exist few tens of taxonomic groups deserving the rank of order, and these few tens have already been discovered and described - at most, perhaps, some yet unknown will be found among fossils; on theother hand there are much-much more existing species and relatively few taxonomists able to recognize, distinguish, and properly describe them, so we have already done much work on this field (some 1 000 000 already described insect species) but much more remains still o be done!
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Is there a written protocol or reference that explains how to isolate the fat body from insect larva (specifically BSF)?
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kindly see:
ORIGINAL RESEARCH article
Front. Insect Sci., 16 June 2021 | https://doi.org/10.3389/finsc.2021.693168
Fat and Happy: Profiling Mosquito Fat Body Lipid Storage and Composition Post-blood Meal
📷Matthew Pinch1*, 📷Soumi Mitra1, 📷Stacy D. Rodriguez1, 📷Yiyi Li2, 📷Yashoda Kandel1, 📷Barry Dungan3, 📷F. Omar Holguin3, 📷Geoffrey M. Attardo4 and 📷Immo A. Hansen1
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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.
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In the upper picture, in my opinion, a diptera from the family Syrphidae
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Hello!
I'm trying to know where the holotype of Psammotettix confinis (described by Dahlbom in 1850) is conserved.
I searched informations on GBIF, INPN, EOL and internet, unfortunately, I didn't find anything.
Can you help me to know how can I find where the holotype is conserved?
Thank you
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Hello everyone!
Thank you very much for all your answers!
I found an answer on EOL (Encyclopedia of Life) : Dahlbom didn't create a holotype when he described the species in 1850.
According to EOL, in 1937, Ossiannilsson decided to create the holotype for Psammotettix confinis (conserved at Carices I Rohne, in Sweden). The type is a male.
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I'd like to call your attention to browse and read a recent collection of papers on insect extinction, just published in Ecological Entomology: https://onlinelibrary.wiley.com/doi/toc/10.1111/(ISSN)1365-2311.insect-extinctions
Thanks
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Thank you for interesting thoughts that will stimulate research on this difficult topic.
Regards, Sergey
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I want to assess the similarity of insect communities within and between two groups of plants: a monophyletic clade and a paraphyletic grade.
Would you recommend calculating Bray-Curtis dissimilarities + Kruskal Wallis H test or a multivariate test such as MRPP or PERMANOVA?
Thank you for your help
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You are most welcome dear
Wish you the best always.
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How long will it take for arthropod ingredients to appear on our menu?
Recently, Nestle has released food for dogs and cats, in which, in addition to the usual chicken, they added chopped fly larvae. And no, the global corporation does not save on cats. Livestock is one of the drivers of climate change, and replacing cows with insects can reduce its turnover. Some insect products have been on the market for a long time. Tell us who you can try and what sensations to expect?
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Depends on the country and culture. Unlikely in Brazil. Most likely in China.
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I need to preserve termite samples for future quantification of juvenile hormone in their bodies. Or extract the Juvenile Hormone and keep that samples stored for future quantification.
I may need to keep the samples stored for up to two months and they also need to "survive" an international trip.
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Follow the procedure given by Brent & Dolezal, 2009
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Expert comments required to id the insect (Image Attached)
Location : J&K, India
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Dear Amit,
What can be seen from the photo seems to be a dune or desert cricket from the family schizodactylidae, the genus Schizodactylus. They mainly inhabit arid sandy areas.
Best, Elaheh
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Can any one help in the identification of tree hopper found on Guava plant?
Your expertise will be highly appreciated..
Thanks in advance..
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For sure, this is a tree hopper with taxonomic tree as below:
Order: Hemiptera, suborder: Auchenorrhyncha, superfamily: Cicadelloidea, family: Membracidae
But, I am still in doubt about confirming its ID as Leptotocentrus taurus, because the morphological characters of this species do not fit the sample in the picture. It needs to be more investigated.
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Further disturbing data were published on the dramatic decline in the number of bees.
Now, in the media there was information that about 40 percent. Bees in the US did not survive the winter of 2018-2019.
Similar data is also found in many other countries.
This is very disturbing.
Is mankind able to solve this problem in time?
Will technological development solve this problem?
Apparently, a significant part of the bee population is killed not only in winter but also in other, warmer seasons. Also in the spring and summer, when large-scale spraying of crops with pesticides is used in agriculture, also used during insect feeding periods on flowers. Then many insects are poisoned and die.
How to solve the problem of a drastic drop in the population of bees and other pollinating insects?
Please reply
I invite you to the discussion
Thank you very much
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Dear Ashutosh Saini,
This is very positive news.
Thank you, Regards,
Dariusz Prokopowicz
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Hello everyone
I have a question regarding mounting mediums for insect haemocyte in microscope slide preparations.
Can I use canada balsam instead of DPX?
Thank you in advance
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You can use both. But in this case, DPX is more convenient. Since when using Canadian balm, a strict protocol must be followed. Otherwise, hemocytes "dry out"
Regards, Sergey
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Case studies like real ones where the testimony from forensic entomologists was presented in court cases. I have searched but found the research case studies on the insect fauna associated with human cadavers like in the case studies below article for instance:
Anika Sharma, Madhu Bala, Neha Singh. Five Case Studies Associated with Forensically Important Entomofauna Recovered from Human Corpses from Punjab, India. J Forensic Sci & Criminal Invest. 2018; 7(5): 555721. DOI: 10.19080/JFSCI.2018.07.555721
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As a prospect for the development of your research, I am sending you an article
Regards, Sergey
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I have been trying to dissolve TC-100 media powder obtained from US Biological in autoclaved distilled water to prepare 2X media for plaque assay.The solution is only clear at pH 4.5, as soon I make the pH to the desired pH (6.5) it starts to precipitate. I am using 1N KOH for adjusting the pH. Has anyone encountered a similar problem?? Can anybody suggest how the precipitation can be avoided?
Thanks in advance.
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May be attached file can help you thank you.
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I am doing Iodine Test of insect's egg in flour for the first time and I just want to validate my results if what I have seen in the microscope is really the weevil's eggs.
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sorry i thought the question was asked 3 hours ago but it is 3 years ago
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The availability of relatively large number of disease resistant varieties compared to insect resistant cultivars suggest that incorporating disease resistance is relatively easy. This is also evident from the list of registered donors and available literature. Should it be understood that breeding for disease resistance is easier than that for insect resistance?
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Thank you Dr Manal Hadi Kanaan for posting your thought at this thread. Due to polyphagous nature of most insect pests, it becomes difficult to create "no choice" condition while scoring for resistance. Thank you once again for your valued participation.
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I have constructed a small hut with the support of bamboo sticks and were attacked by insects( mostly in black color)which made small holes all over the bamboos resulted in powder as in the attached pictures. Daily I see a lot of bamboo poweder and form dust throughout the room. Any remedy for this. Please refer attached pics. Thanks in advance.
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Cover the hut with plastic and leave it under the sun for several hours. All insects will die from overheating with zero insecticides.
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Found in Turkey on peony flower buds.
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Yes, is Tropinota squalida
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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.
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Insect pest monitoring is typically performed in agriculture and forestry to assess the pest status in given locations (i.e., greenhouse, field, orchard/vineyard, forest) by collecting information about the target pest presence, abundance, and distribution. Within the integrated pest management programs in agriculture, the final goal of insect pest monitoring is to provide growers with a practical decision-making tool. Typically, an automatic trap equipped with a camera involves two modules: the hardware and the software. The hardware is typically composed of the trap structure containing the bait and retaining the trapped insects, an electronic box including the camera, a data transmission modem, a battery, and eventually an external power supply, such as a solar panel. The software is composed of the online repository in which the capture data pictures are stored and accessed plus optional image analysis algorithms to automatically identify and count the captures. Trap design may vary according to the target pest to be monitored, as detailed in this section.
Some to be considered: An automatic trap prototype modifying a commercial trap (Pomotrap®, currently Carpo® by Isagro S.p.A., Milan, Italy) with data acquisition and data transfer systems to monitor the codling moth Cydia pomonella L. (Lepidoptera: Tortricidae) in apple orchards;
‘Jackson trap’ was equipped with a camera device for the automatic monitoring of the Mediterranean fruit fly Ceratitis capitata Wiedemann (Diptera: Tephritidae)
Bucket traps are typically adopted to monitor fruit flies where a camera-based electronic McPhail trap was used to monitor by remote the olive fruit fly Bactrocera oleae Gmelin (Diptera: Tephritidae).
Agriculture operators are now facing the ‘Big Data Analysis’ prospect: organize, aggregate and interpret the massive sample size of available digital data with sophisticated algorithms to drive decisions based on data interpretation, prediction, and inference potentially on a global scale.
Camera-based insect monitoring can be exploited not only for pest monitoring but also for early detection and survey, allowing a prompt reaction especially for invasive species. There is a potential perspective to interconnect traps among sites and create a network at local, regional, country, continental, and global scales.
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Hello,
I have a problem with plasma membrane isolation from Sf9 cells, because after cells disruption and pelleting unbroken cells, cell debris and mitochondria the clear supernatant containing plasma membranes slowly becomes turbid. Something of white color is precipitating and makes difficult further steps. Does anybody know what is the origin of such precipitate and how to prevent its development?
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Not too many people dare to answer. Did you solve the problem? If not, sorry, I have no clue, but probably your cells are loaded with a high Ca2+ concentration. I think sf9 intake calcium upon stress, or other stimuli, and then if your lysis buffer contain phosphate... Did you try to change the composition of that buffer?
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I am developing a method to detect insect DNA in food samples via DNA metabarcoding.
I already designed primers (1 forward and 1 revers) for an amplicon of ca. 200 bp length that bind to mitochondrial insect DNA. Right now I am testing those primers in PCR to find the right temperature and conditions. I consider amplification curve and melting temperature from PCR and also bands on an agarose gel of all DNA-samples. All insect samples work well, but i have a quite unusual problem with honey bees:
They have a band at 200 bp on agarose gel, and there also is a melting peak at about 78°C. This is as expected and also like all other insect samples. But there is a difference: There are no amplification curves of honey bees in PCR.
I already tried cleaning the DNA extract with magnetic beads, that didn't help.
Additional information: I use EvaGreen as a flourescence dye in PCR.
Does anyone have an idea what could be the issue or what i could try to solve it?
I am happy to give more information about the conditions, if needed.
Thank you in advance.
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I would call tech help at bio-rad. Are the bees all run on their own assay or do you have any results from a plate of some bees and some other amplimer showing this result or are all of the negative results on bee only plates?. If you have a bee pcr band amplified then I agree with Katie A Burnette then this looks like a technical problem possibly something has changed the exes on the graphing so the curves cannot be seen for your bee assay or you have found a way of hiding the display for these samples so the data may be there but just not showing
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I had been updating the old collection list from our museum and I found that there is some clash between family taxon for those three genera, some sources put them under Family Lonchodidae while some under Family Diapheromeridae.
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Hello,
According to the current taxonomy, these 3 genera, Marmessoidea, Lopaphus (please check spelling), Carausius (please check spelling, too) belong to the family Lonchodidae. Necrosciinae and Lonchodinae, formerly subfamilies of Diapheromeridae, are now subfamilies of Lonchodidae (Robertson et al. 2018).
Regards
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Does anybody have an idea on how to successfully mass-rear Nesi bug? In full details, please! Any suggestions, thank you in advance.
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I rear Nesidiocoris tenuis on tobaco plants in cages/ Feed with Sitotroga cerealella eggs and bee collected polen.
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the objectives of insect sampling in forest is to determine the abundance, diversity and habitat association of insect in forest ecosystem
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The idea is t capture all seasons and flow conditions to capture all the conditions and stages of all potential insects for the region. It will also be important to consider the spatial scale (micro- and macro-habitat diversity) and sampling effort, as these are also as important as covering all the seasons.
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Hello everyone,
I am having a very interesting issue with my protein expression. Currently, I am expressing a ~55kD protein recombinantly in insect cells (Sf9 to generate high titer virus and High-5 for protein expression). I used P3 of the recombinant virus to infect High-5 cells and have already verified the expression level of the 55kD protein via ELISA, WB and IFA. It is highly expressed in the lysate (insoluble fraction) of the pellet and some low expression was observed in the media. In addition the protein was observed to be in the High-5 membrane with IFA. Now since we do not have a SEC column in the lab yet, I was using the cell media to purify with the HisTrap 5mL column by Cytiva. After purification I got a single band at 55kD (it was not thick but it was visible) in 5 x 1.5mL elution's. Since the protein is in 500mM Imidazole solution I decided to combine, concentrate and Buffer exchange with HEPES Buffered Saline Solution for interaction assays using the Pierce Concentrators on my first try and then the Amicon Concentrators on my second try. I read on other posts here that the concentrator membrane composition might have caused the protein to bind, so I wanted to check that. I followed standard protocol supplied by the manufacturers and checked protein concentration with BCA at 595 and protein plate reader after. Both gave me a concentration of 2.9mg/mL and hence I loaded 1ug of the protein onto an 12% SDS-PAGE gel and even did WB with the antigen specific pAb. I got no bands in either assays for both concentrators (not even smaller bands)! I additionally checked the flow though material and washed the membrane of the concentrators but nothing. I am really lost as to how a protein can just disappear like that especially when the plate reader and BCA both detected protein? I was thinking maybe proteases but after purification and kept on ice all the time. My next steps would be to just purify from the insoluble lysate and use the SEC for desalting and separation of contaminating proteins, but I am worried that upon concentration the same thing will happen. I would be grateful for any advice that you might have that could possibly point me into the right direction. Thanks!
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Hi Julian Ramelow . So if I understand correctly, you are not losing total protein in the concentrators, you are specifically losing *your* protein, correct?
If so, your protein must be very hydrophobic or very prone to aggregation (or both). Check if it has glommed onto the membrane by washing it with 0.1% SDS and doing a gel.
If it is aggregated, then you should include stabilizers in subsequent steps to prevent aggregation. Common stabilizers include EDTA, urea, arginine, glycerol, detergents, etc. Composition depends on what you want to do with your protein.
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what method can be done for the extraction of plant/insect sample?
what method can done for separation and quantification?
for cyanogenic glucosides
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Dear Anaswara k Sreedharan a typical separation and identification method for cyanogenic clycosides (CG) has been outlined in the "Materials and Methods" section of the following potentially useful article entitled:
Metabolism of the Cyanogenic Glucosides in Developing Flax: Metabolic Analysis, and Expression Pattern of Genes
This paper has been published Open Access and is freely available as public full text (see attached pdf file). Please also have a look at this reference:
Spatial separation of the cyanogenic β-glucosidase ZfBGD2 and cyanogenic glucosides in the haemolymph of Zygaena larvae facilitates cyanide release
(also attached)
A third interesting articles is freely available as public full text on RG:
Plant-Insect Interactions-Cyanogenic Glucosides
I also strongly suggest that you search the "Publications" section of RG for more relevant information about this topic. Just search e.g. for the term "Cyanogenic Glucosides" and then click on "Publications" to receive a long list of relevant references:
I hope this helps. Good luck with your work!
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Hi all
I have a dataset of insect species and their abundances across sites (sites as rows, species as columns). I would like to determine compositional turnover between site-level assemblages (ie overlap) using the Horn similarity index. To do so, I would like to first calculate the beta diversity across sites using Shannon based on Hill numbers. Does anyone know of a package (along with a code) I can use to do this in RStudio? I have tried betadiver in vegan, but these do not give Shannon diversity indices.
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Have a look at phyloseq tutorial.
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I am wondering if there is a type of microscopy technique that people use in the field of insect physiology that allows muscle to be seen in living or prepared specimens.
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Hi Tristan -- I'm taking a look into this -- I bet you already found some answers on your own since a year has passed, but I'll look into it right now out of curiosity.
The best I could find from a quick search was a mice study here ( ) -- I particularly liked Figure 1 here -- and I am taking a look at Reference # 26, 27 ("A few studies previously reported in vivo microscopy of skeletal muscle" [I know you are not looking at skeletal muscle per se of course, since we are talking about insect muscles.]) . . .
Summarizing possible methods for what I think you are looking for:
From the 1st paper I linked: "Multilodal microscopy combining SHG and fluorescence detection" where SHG stands for second harmonic generation.
The methods from the three papers here seems to be called "intravital imagining of muscles" as a whole.
From the third paper (Ref 27) -- it mentions confocal, intravital, and superresolution microscopy . . .
The second paper (Ref 26) mentions intravital microscopy...
A quick search yielded this result for intravital microscopy:
Interestingly, for insects, when I tried to find "intravital microscopy and insects" -- I found this instead: https://www.microphotonics.com/x-ray-microscopic-inspection-of-insect-flight-muscles/ --the title is a bit strange -- but specifically, I am looking at the Micro-CT imaging part, as it can allow you to visualize the internal and external features of your insect without having to kill it. You may want to anesthetize the insects like what Dr. Ellis does with her flies, perhaps.
I think that can be a good start for now... I know this is an old post but this was an interesting and useful question nonetheless.
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Is it possible to run out BINs analyses on BOLD based on 16S sequences (wild bees)?
My fear is that BINs analysis is only possible for large dataset (such as CO1 sequences) but would not be relevant for 16S as insect species are not well represented on BOLD for this marker.
Any advice for diversity analysis of 16S for approx 100 species (up to 3 replicates each)?
I was thinking about looking at K2P distances, compare 16S and CO1 trees to validate our new barcodes.
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I think, it is possible. So, you can try to do it as early as possible Dr @Mélodie Ollivier
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I specifically look for taking digi photo from mandibles
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I agree with the technical details offered by everyone here. I too put emphasis of focus stacking. Its an "art" that needs practice. In addition, if you're using a camera the quality of the lens is critical. You want (usually a macro lens) excellent glass and a lens that offers a low f-stop. Both contribute to resolving very fine details. Add a very stable camera support in a room free of any vibrations.
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Hi everybody , I hope all is well.
I've got a question
Could you suggest me some suitable pesticides for insect larvae in spirulina farm?
Thanks
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It Is smaller than standard salinity for spirulina. The standard amount is 30 g/L which about 5 grams of this come from NaCl and the other part is from Soda, Soda ash, and other nutrients. from of my view, you didn't use enough Soda (16 g/L) and Soda ash (8g/L) which are very important for preventing predators and competitors. When evaporation occurs the salinity could rise higher and higher, in this situation you have to keep watching the salinity doesn't go more than 60 g/L and by adding freshwater keep it at 30 g/L points.
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Dear colleagues,
does any of you have some experience what is the best protease inhibitor mixtue for use in media for insect cells (S2,Sf9) please? I worked with commercial EDTA-free tablets based on inhibition of serine/ cysteine proteases and it didn´t work very properly. After FPLC I can still detect on SDS-PAGE followed by Mass spec. cleavage of my protein (small fragments of my protein). If you can recommend me some commercial mix of proteases for the insect cells media I would really appreciate it.
Good luck in your experiments and thank you!
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You can refer to the link below for the protease inhibitor cocktail form Promega.
OR
You can prepare the cocktail as given below:
Cocktail of protease inhibitors used in Sf9 Cell culture.
1mM benzamidine
0.2mM phenylmethysulfonyl fluoride
10ug antipapain --- add fresh
10ug leupeptin ----- add fresh
10ug aprotinin
Reference: G Protein Pathways, Part B: G Proteins and their Regulators
Inaki Azpiazu, N. Gautam, in Methods in Enzymology, 2002, vol 344, Pgs 112-125
Good Luck in your experiments too.
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I am interested in using RIP-seq or a similar protocol to identify microRNA targets in a non-model insect (Sarcophaga bullata). There is a published genome, but the annotated sequences do not include the 3' UTRs. My questions are:
How long is the fragment of mRNA that is generally pulled out using RIP-seq? Will I be able to identify the sequences I pulled out of this species using this genome?
Thanks in advance for your help!
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The length of the precipitated RNA depends on the method used. Fixation in the formaldehyde will give you the smaller fragments than without crosslinking agent.
Normally reads about 50-150 bp is enough for mapping.
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I am wanting to compare fossil insect densities with charcoal concentrations and other indexes for disturbance.
My issue is that insect samples were subsampled according to geochemical patterns in the soil and are of varying resilution (between 2 and 8 cm) and also temporal resolution (20 - 120 years). The core where the other indexes from disturbance were counted from is from the same site but was subsampled in 1-cm resolution. We have a hunch that bark beetles and charcoal densities are related, but how to test this statistically, is it even possible? To correlate these insect time slices of varying resolution, with the data points from the other core?
Example insect samples:
S1 0-4 cm 2017 - 2004
S2 4-6 cm 2004 - 1969
S3 6-8 cm 1969 - 1946
S4 8-11 cm 1946 - 1899
S5 11-15 cm 1899 -1841
S6 15-17 cm 1841 - 1806
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Finally, my colleague used some analysis in R with the libraries bincor (Polanco-Martinez, 2018), ggplot2 (Wickham, 2016) and trend (Pohlert, 2018). He used a binned correlation sapproach that allows for irregular time series (Mudelsee, 2012). In the calculation, he used the rule based on average spacing as it showed him better performance than MOnte Carlo simulations. We found a significant correlation between charcoal volumes and number of primary bark beetle fossils and are in the process of submitting our manuscript to a fitting journal. Thank you for your repplies and interest to this question. :)
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Hello!
The results of my study are leaving me a little baffled. I would appreciate any insight you could give me.
What I expect: Larger objects (living or non-living) should heat up slower than smaller objects
My results: Heating rate is increasing with size, larger beetles heat up faster than smaller beetles
I tested for: The amount of water absorbed during the rehydration process does not play a significant role.
I am wondering: Could it be related to differences in fat/protein contents?
Thank you!
Emilie
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Thanks for the interesting question. Indeed, the larger the insect, the more it heats up (but I studied the content of proteins / lipids of different insects), but there is no clear pattern. In what it really can be traced it in the development of the trachea. At the same time, insects with a low temperature threshold (those that are active at 0 degrees), for example, boreus, have a high lipid index. In general, I am also interested to know the opinion of colleagues on this issue.
Regards, Sregey
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It is very common for Taxonomists to request insect specimens, from a number of institutions and museums, to assist them in their research on specific insect groups. It can take a few years to actually make use of these specimens and reach a conclusion with published descriptions. The institutions that make these loans in good faith, find themselves having to chase researchers for the return of the loans. I have heard of loans still not returned after 30 years and this may mean the loss of these specimens due to retirement or even death of the researcher. This issue is a BIG problem and so what would be a reasonable time for these loans to be made for? What ways can these institutions encourage researchers to return loans without threatening sending in the debt collectors?
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Perhaps this is a matter of ethics. It is better to deal with specialists who are employees of large scientific centers. Then, for security reasons, it will be possible to conclude an agreement in which the period for which the samples are provided will be determined. I also once gave samples on my word of honor, but did not receive it back.
Regards, Sergey
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I collected it from my home garden.
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I do not see the image, please send it. In the mean time I have a question about another species of insect. Please ID.
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I am working on insect associated mites. The parasitism and predation of the mites on the insect is evident from the direct observation and rearing experiment. I encountered a problem in confirming the predatory/parasitic relationship of the mite on the insect in molecular level. Could you please suggest a molecular technique for analysing the predatory /parasitic relation of the mite on the insect host. If we are going for a gut content analysis of the mite, which kind of molecular tests will be more sensitive in analysing it?
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Can we use grace insect media
un-supplemented with 10 percent FBS only?
Is it necessary to add other suppliments?
I have tried to grow Sf9 with grace insect media supplemented with 10 percent FBS , antibiotics and antifungal but failed.
Please suggest me
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Is it required to add methionine and yeastolate, lactalbumin in grace medium unsupplemented?
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I have small insect samples stored in ethanol and I need to obtain dry mass. I have access to a drying oven but I have also heard you can air dry them on the bench top in a lab environment with relatively consistent humidity. I would appreciate any tips.
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I am in general agreement with the previous posts but I am concerned about 2 points. The first is that ethanol will dissolve fats from your specimens and this could make up a significant part of the specimen's weight. My solution would be to tare a piece of filter paper sized to fit in a Petri dish place and pour the sample onto the filter paper bit by bit so the ethanol does not overflow the filter paper. Otherwise the fats will stick to the glass of the Petri dish. You can let this air dry and then weigh the filter paper and specimen subtracting the tare weight of the filter paper. I would suggest using the drying oven at 35C. This comes to my second point. In northern BC you can have significant relative humidity so bench drying really is not a good idea. Your weights will vary significantly with RH. It will be important to dry the specimens on filter paper in the drying oven for accuracy and repeatability. Take the samples right out of the drying oven and weigh immediately before they absorb moisture from the air.
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I´m starting a pollination experiment. A cacti population will be selected for the study and maybe there are references to consider over an appropiate population, including some distance of human disturbance to pollinators.
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It certainly depends on the pollinators you want to observe. Birds are most sensitive, bats less so, while keeping still seems to work for most insects (although I am not sure that we would recognize a disturbed insect).
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I am planning to conduct a survey of patients who had taken an insect-derived drug given by a traditional healer in past years. I wish to know that if the medication was effective for the patients, any short or long-term side effects they know, and so that I can obtain some idea to study further on the medicinal value of that particular insect. I would be grateful if anyone can suggest to me what kind of questionnaire should I prepare, how should I analyze the data, and if anything else I should know?
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I agree with Firas Al-Zyoud.
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I am interested in the size of Lepidoptera (moths and butterflies) and would like to know what is the smallest known species of this group of insects. It is probably a Nepticulidae (pigmy moths). The species in my figure below (unidentified) measures about 4 mm with the wings spread, and its dry body weight was 0.3 micrograms (0.03 mg).
We are generally more impressed by the higher figures (the oldest tree, the heaviest vertebrate…) than by the minima. Thus for instance one can read about the largest moths (Thysannia, Attacus: http://entnemdept.ifas.ufl.edu/walker/ufbir/index.shtml). However 'smallness' has interesting biological implications (see the recent book by A. Polilov 'At the Size Limit - Effects of Miniaturization in Insects'). I have seen descriptions of other nepticulids in the same range of size as 'my' species (around 4 mm: Dooren weerd et al.: http://onlinelibrary.wiley.com/doi/10.1111/syen.12212/full). Perhaps there are slightly smaller European species (some Stigmella spp., e.g.: http://lepiforum.de/lepiwiki.pl?Stigmella_Magdalenae).
So, does anybody know of any moth smaller than 3.5 / 4.0 mm?
Thanks!
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Dear Munira Nasiruddin,
Dear colleagues,
We kindly suggest, prior to providing your answers in this discussion, to read about results of the most recent study (attached above and in the current message). Do you know anything smaller among Lepidoptera than is mentioned in the paper? Let us know please if you possess such info. We can expect that there might be many more extremely small species in the tropics and subtropics.
So far, the minimal recorded forewing length was found to be around 1.2–1.3 mm and the wingspan around 2.6–2.8 mm in two families, the Gracillaridae and Nepticulidae. Among Lepidoptera, the following species have the smallest moths globally: the European Johanssoniella acetosae (Stainton), the Peruvian Simplimorpha kailai Stonis & Diškus, the Mexican Stigmella maya Remeikis & Stonis, the Mediterranean S. diniensis (Klimesh), the Mediterranean Parafomoria liguricella (Klimesh) (Nepticulidae), the South East Asian Porphyrosela alternata Kumata, and the Central African P. desmodivora De Prins (Gracillariidae) (see Stonis et al. 2021).
Kind regards,
Andrius
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Please, l need a clear experimental Proceedure for the extraction and purification of pigments from an Insect ( Weevils and Aphids)
Please can l get the step by step extraction procedure before the use of HPLC
Solvent Extractions, l guess maybe okay , but l am not sure yet of the adequate chemicals/reagents that maybe needed.
Finally as it is a pigment( Protein) do l need to pass any of the supernatant through indirect heating by use of water bath?
I look forward to your advise, suggestions, materials, journals and practical manuals that could be of help
Thank You
Best Regards
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Dear Jerry Nwabasa many thanks for your interesting technical question. Please have a look at the following potentially useful article which could help answering your question:
Optimizing conditions for the extraction of pigments in cochineals (Dactylopius coccus Costa) using response surface methodology
Also please see this relevant article:
Isolation, Purification, and Identification of an Important Pigment, Sepiapterin, from Integument of the lemon Mutant of the Silkworm, Bombyx mori
This article is freely available as public full ext on ResearchGate
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Hello everyone,
I am getting to start a project on substitution of SBM with insect meal. The main aim of this study is to assess the effect of that replacement on performances of layers flock. I would like to know if anyone knows anything about mineral we should be careful with when using insect meal with black soldier larvae.
Thank you,
Catherine Roy, agr. M.Sc.
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Dear @ Catherine Roy I think @C.A. (Kees) Kan is right. The ingredients of both should be compared. The second aspect relates to energy use efficiency sustainability. Soya meal being product of primary producer may be more cheap and energy efficient.
Regards!
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I would like to observe/ record insects behavior under webcam setting at night. So we are looking for some suitable infrared(Red light) lamps or IR LED to do this. Can anybody recommend us some proper products?
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