Science topic

Larva - Science topic

Wormlike or grublike stage, following the egg in the life cycle of insects, worms, and other metamorphosing animals.
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I Know the adult oyster lost its foot and byssus, but what about their larvae? Does they still need byssus for adhesion?
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Thanks for your all nice reply!
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I am studying the carbon dioxide production of woodlice, maggots and germinating seeds by the hydrogen carbonate indicator solution. Among three species, which one change the color of the indicator solution most quickly when they release carbon dioxide?
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I'm sorry but I don't know about CO2 emissions by biota. However I am a chemist, and I can say that the pH buffering of the bicarbonate and the action range of the indicator may make that test relatively insensitive to CO2 emission. Titration may be more accurate. Ba(OH)2 may also be a more useful base than NaOH.
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Please anybody help me to identify which group this species belong to?
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observing the length of the buccal tentacles and especially the tentacular membranes, the specimen in the photo reminds me of a pectinariidae, an adult specimen, is already beyond the larval stage
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Hi,
I have vignettes of gastropods from multiple samples (from coast to open ocean). Is there a possibility to distinguish between gastropods larvae and holoplanktonic gastropods just from the vignettes?
Thanks in advance.
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Are tropical coastal reefs sinks or sources of mesozooplankton? A case study in a Brazilian marine protected area
  • Gleice S. Santos,
  • Lars Stemmann,
  • Fabien Lombard &
  • Ralf Schwamborn
Coral Reefs volume 38
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I took these photos long ago, now I am interested in identifying the species of this larva from these pictures. I collected these larvae specimens from seasonal valleys of Southwest Saudi Arabia, and tested their efficacy of mosquito larvae predation in the lab. I need someone, if possible, to confirm or suggest to me a certain species as they appear in the pictures. I`m sure the appearance of the larvae in the pictures is not quite clear, nevertheless, I would appreciate any effort. According to me, as I searched in the literature, I propose that they could belong to the genus Sternolophus and the species could be decens.
Please, see the attached file ( Morphological characteristic of Hydrophilini ................).
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It is a Sternolophus larva. I can´t tell the species, you should check which species are present in that area.
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Why do we care about it and raise it if its larvae are destructive to crops, and is there a relationship to environmental diversity in this?
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Yes, butterflies play an important role as a pollinators.
Not all the butterflies host on crops, some host on shrubs as well.
You may take a read in the following article:
Hope it helps!
Drop me a message if you need further discussion.
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Hi I have a problem to get the mRNA from larva of hookworm. Can anyone suggest me with the procedure?
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What have you tried so far and what was the result?
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Can anyone come across such parasitized beetle larvae on eggplant??
Thanks
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It seems to me that they are Coccinelide larvae (ladybugs) before moulting.
It is not clear if they are parasitic, these larvae are themselves predators on aphids. If they are dead, then they can also be parasitized by wasps or spiders arhnidae.
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Dear reserachers,
currently I am working with Ostrea edulis larvae. However, I can't get rid of copepods via mechanical separation as the have more or less the same size as the oyster larvae. I was thinking about using a concentrated light source as copepods are positively phototactic, what do you think? Any further ideas ? Thank you in advance!
Best regards
Dominique Noetzel
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Hi !!
copepods are a source of protein for other species, under the principles of integrated multitrophic aquaculture, adding some species that controls them would be very appropriate. Normally the first trophic level is fish, and bivalves go to the last, it could be an option, I don't know your essay and characteristics but we'd be happy to talk.
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I've thought about different reasons, but can't seem to find more specifical information
-Maybe they're harder to eat/digest due to higher chitin content, and sharper exoskeleton
-Generally bitter or bad taste, but my question here is whether it is safe to eat or not?
-I've read some beetles are vectors of diseases to chickens such as Alphitobius diaperinus, but its consumption has just been approved by the EFSA, maybe the diseases are associated with wild harvested beetles and not farm raised?
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It is all about foraging efficiency. Why would you bother looking for scarce adults when you can break open a sago palm and find lots of large larvae? Conversely, when you have thousands of grasshopper adults, or millions of Chaoborus edulis adults flies around lake Malawi you can just scoop up the adults, so why bother with larval stages?
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Is there a standard procedure in disposing brine shrimp larvae along with the selected plant extract once the experimentation is concluded?
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they can be converted to compost or biofertilizer; you can consult this book
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Dear All
We are trying to detect Wolbachia in Culex and Aedes spp (adult, larvae and pupae). Can anyone please tell what would be the preferred approach to start with considering adult moquito samples. For instance, some articles refer to use to whole body while others have isolated gDNA from organs such as ovaries. It is also mentioned that removal of head improves detection due to presence of inhibitors. It is also proposed somewhere that Wolbachia could exhibit tissue tropism. Kindly share your experience and expert advice on said query.
Looking forward
Thank you
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maybe you can start by MLST?
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if I have survival(mortality) data of larvae of one butterfly species on different plant species for one month, how can it be analysed?
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You can consult this:
Estimating arthropod survival probability from field counts: a case study with monarch butterflies
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Hello everyone,
here a few pictures of an anophthalmic aquatic insect, I am unable to identify.
It was found in a small brook and lives in fastflowing hygropetric environment. Region: Jura mountains, Switzerland, 860 m asl.
Dorso-ventrically flattened, no eyes, no antennae, six short legs, white patches on the edges of the dorsal segments, long thorns laterally on the ventral segments. Could be some coleoptera larva?
In advance many thanks for any suggestion.
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Nothing to add more than colleagues' opinion.
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Hi! So I had a question, which I am hoping someone who perhaps has worked with honeybee tissue before, can give me some much needed help. I am working with honeybee larvae cryosections, cutting them up to 20 μm thick and staining afterwards with alpha-bungarotoxin for visualization of acetylcholin receptors. The problem is, the tissue keeps breaking and it's way too brittle. It has holes in the middle too. I have tried two methods: the first being fixating the larvae O/N with 4% PFA at 4°C, then immersing O/N again in 15 % sucrose solution, and finally embedd in OCT medium in dry ice. The second is freezing them directly in OCT in dry ice and fixating after cryosectioning. Any advice for preserving the integrity of the tissue? Thank you.
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passage it to 30 % sucrose solution after 15% sucrose immersion for 2-3 days then passage it to cold (-80) methyl butane solution for remove bubble in fast freezing prosses then embed in OCT medium in dry ice.
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As a Biomarkers, I have protein concentration, protein carbonyl content, and GST activity in tadpoles. From this data result, I prepared plots and now I want to explain the relation between them. Does anyone know how they are related to each other?
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Thank you Azadeh Eshraghi for your answer.
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For the LEIA technique, we test different concentrations of forage plants (e.g.) in contact with larvae of gastrointestinal parasites (e.g. Haemonchus contortus).
These larvae are in the L3 stage.
We artificially induce larval exsheathment to obtain after 1 hour 100% exsheathment.
We count the unsheathed and sheathed larvae every 20 minutes for one hour.
If an inhibitory effect of the larval unsheathing is detected at the highest concentration (i.e. at 100% for example) and the negative control gives 0% inhibition.
If the lower concentrations give less inhibition between 0% and 100%, we can determine an effective concentration at 50% effect a.k.a. EC50.
Is it therefore necessary to carry out the intermediate counts at times 20min and 40min to calculate an EC50 afterwards?
------------------------------------------------------------------------------------------------------------------------------------------------French version
Les valeurs d'EC50 calculées peuvent-elles variées significativement en fonction des mesures prises dans le temps ?
Pour la technique du LEIA, nous testons différentes concentrations de plantes fourragères (par exemple) mises en contact avec des larves de parasites gastro-intestinaux (Ex. Haemonchus contortus).
Ces larves sont au stage L3.
Nous provoquons artificiellement le dégainement larvaire pour obtenir au bout de 1H environ 100% de dégainement.
Nous comptons les larves dégainées et engainées tous les 20 minutes pendant une heure.
Si un effet inhibiteur du dégainement larvaire est décelée à la concentration la plus élevée (i.e. à 100% par exemple) et que le témoin négatif donne bien 0% d'inhibition.
Si les concentrations plus faibles donnent une inhibition moindre comprise entre 0% et 100 %, nous pouvons déterminer une concentration efficace à 50% d'effet a.k.a. EC50.
Est-il donc nécessaire de réaliser les dénombrements intermédiaires aux temps 20min et 40min pour calculer une EC50 ensuite ?
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#LEIA #EHIA #gastrointestinal #nematods #tanins #condensed #sainfoin #invitro #biosassays #test #method #statistical #EC50 #IC50 #Haemonchus contortus #Trichostrongylus #design #experimental #tools #software
  • from "Age of Haemonchus contortus third stage infective larvae is a factor influencing the in vitro assessment of anthelmintic properties of tannin containing plant extracts" G.S. Castañeda-Ramírez
=> 2.4. Larvae exsheathment inhibition assay (LEIA)
The assays were performed once every week for seven consecutive weeks. Moisture, temperature and general conditions in the laboratory were kept homogenous along the entire experimental period. The only condition that varied was the age of larvae (from 1 to 7 weeks). The LEIA were conducted following the procedure described by Jackson and Hoste (2010). The negative controls used were larvae not treated with extract and only exposed to the PBS. The different concentrations ap plied to evaluate the AH effect of A. pennatula acetone:water extract were 1200, 600, 400, 200, 100, 40 μg/mL. Stock solution (5000 μg/mL) of acetone:water extract were made in PBS prepared with purified water. One tube was used as negative control containing 1000 μL of PBS without extract. Finally, 1000 μL of infective larvae solution (L3 ∼ 1000/mL) were added to each tube to obtain the final extract concentrations (1200, 600, 400, 200, 100, 40, 0 μg/mL PBS). Infective larvae were incubated with the plant extract for 3 h at 24 °C. After incubation, larvae were centrifuged for 3 min at 168g and washed 3 times with PBS solution. Then, aliquots of each larvae solution were placed in eppendorf vials (200 μL each). Four re petitions were performed for each concentration and PBS control. The process of exsheathment was artificially induced by contact with Milton® (Laboratoire Rivadis, France), which is a solution of sodium hypochlorite (2.0%) and sodium chloride (16.5%) diluted in PBS. The quantity of Milton® solution to use for each assay was determined every week by testing different concentrations (25, 30, 35 and 40 μL/6 mL PBS). During the first two weeks the concentration used for the bioas says was 30 μL/6 mL PBS and it was changed to 25 μL/6 mL PBS for the following weeks. The exsheathment kinetic was observed with a mi croscope using the 10× objective and recorded at 0, 20, 40 and 60 min. (https://oatao.univ-toulouse.fr/25140/1/Castaneda-Ramirez_25140.pdf)
#pIC50
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*Before determining any EC50, are the observed differences and effects interpreted correctly?
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French version
*Avant de déterminer un EC50 quelconque, est-ce que les différences et les effets observés sont interprétés correctement ?
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Spanish version
*Antes de determinar cualquier EC50, ¿se interpretan correctamente las diferencias y los efectos observados?
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Dear community,
I am a post doc at ULiège in Belgium, and in my research group we are looking to buy a microbalance to weight the dry weight of parasitoids of drosophila larvae. These insects are super tiny (dw ~ 250 µg).
The company we started to discuss with, proposed us a Sartorius microbalance (model MCE3.6P-2S00-M). In the past I (and other colleague from the team) only worked with Metler Toledo microbalances, but MT ones are far more expensive (the price is almost twice higher). So the point is that we never used Sartorius balances, and we therefore don’t know the quality of this equipment. Does anyone have feedbacks on Sartorius microbalance to weight such small individuals? Thank you! Best regards, Thomas
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Dear Thomas: I have used both MT and Sart. balances, and MT is so accurate, however, sartoruis is so good, and you can use it for these small and minute masses. You need a well levelled plane or bench and a quite room to get excellent results. Regards.
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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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I have a container of greater wax worms (larvae of Galleria mellonella) and I noticed that about half of them have black dots on them. They almost look like specks that can be removed, but they adhere to their skin. Some of them their color have turned into black as they appeared in the picture. Could this be a disease or fungus of some sort?
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I am not an expert on this topic but according to what I know these dark spots (melanized areas) are the reaction of the larva when it detects certain enzymes secreted by a parasitic fungus. This reaction consists of the segregation of various substances and among them the melanin that surrounds and isolates the fungus infection, stopping it.
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Normally, fish and larval stages are preserved in 5-10% buffered formalin prior to morphological identification. If we are going to use, molecular level identification, we have to preserved them in 75% ethanol. However, in most of the recently published papers, 75% ethanol is used, even for morphological identification.
1. what is your recommendation of using ethanol for this preservation if we are going for morphological identificaton?
2. If you have literature on preservation technics of fish eggs and larvae can you share them with me?
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Interesting question.
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Our lab has been trying to establish zebrafish larvae culture. We purchase and introduce 2 dpf zebrafish into our culture and it always crashes on 8 or 9 dpf even though the fish seem just fine until 7 dpf.
Our culture is a closed system with no filtration but the water quality is closely monitored and all the parameters (ammonia, pH, temp, etc.) stay in the suitable/tolerable ranges. We feed rotifers in the morning and 50% water change in the evening. We have not tried feeding brine shrimp yet due to some experimental restrictions.
I, as a novice, was assuming that it would be either the amount/frequency/quality/type of feed that is causing the mass mortality at the same timing. But would that be really the likely case or should I be looking into something else first? If anyone could weigh in I would be extremely grateful.
Thank you for your help in advance!
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Filippo Del Bene Thank you all so much for your advice! I will try them.
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Does it act as a visual deterrent for predators or serves a purpose in the body's physiological processes?
Also, is there any similarity between the Sphingidae caterpillars and the larvae of Trilocha varians (Bombycidae) which also has a horn-like structure in the larval stage ?
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The horn is absolutely not for defense. I’ve tried super hard to get hurt on the horn but the horn does nothing at all. It’s too floppy to do anything defense-related. I’m guessing it has something to do with sensory functions or tricking predators. This would be a cool capstone research project for anyone interested!
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It is the agri entomology experiment conducted in two different contrasting locations for two years and two seasons per year. The data collected from the experiment was number of eggs, larvae, pupa, adult insect
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I would suggests you can use multivariate analysis, correlation analysis , anova using sas software accordingly you can see the intraction effect.
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I selected three essential oils that their larvividal effects were proofed in the articles, and I used dipping method to study their effects on house fly larvae (put larvae for 60 second in diluted essential oils, I used 50 % ethanol as solvent after that I put larvae in a plate with food but they were survived after 24, 48 hours. Could you help me what is the problem? solvent, or method? In most paper acetone used as solvent but dipping larvae in pure aceton killed all them but this was not happened with ethanol so I choose ethanol as solvent.
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Dear Maryam bagheri Varzaneh
From literature point view there are few solvents which can be used in this dilution process such as
1. DMSO
2. n Hexane
3. Acetonitrile
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Can anybody help me with the protein extraction from the gut of adults and Larvae of Zebrafish?
I have a problem: I can't get a good result in Western Blot with adult zebrafish gut or with zebrafish larvae (5 dpf). When I extract the proteins, the bands on SDS-page appear not very concentrated and after the transfer, when I paint the membrane with ponceau red, the result is not clear and there aren't defined bands , but everything is very blurred as if the loaded samples were dirty. The protocol we use provides for the homogenization of the sample with lysis buffer
(SDS 1%, EDTA pH 8.0 10 mM, Tris-HCl pH 8.1 50 mM, PMSF 1 mM, protease inibitor) or Ripa buffer (Sodium chloride 150 mM, Tris-HCl pH 8.0 50 mM, Nonidet P-40 1%, Sodium deoxycholate 0.5%, SDS 0.1%, protease inhibitor).  Then I heat the samples for 5 minutes and centrifuge them at 12,000g for 15 minutes at 4°C.  After, I load from 20 to 50 ug of protein in each well in a 10% or 15% gel.  I do either dry or wet transfer, but either way it doesn't work well. What could I do to improve my homogenization of samples and transfer?
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Make a fresh loading buffer and ensure that the protein samples are quickly mixed in loading buffer and the vials are kept in boiling water for 7-10 min. And reduce the total protein/lane. Hope this will resolve your issue.
Best
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Hey,
I am working on nematodes and my interest is in strongyloides stercoralis. I have obtained larvae from fecal samples of goats and sheep by kato katz culturing method aka charcoal culture method and extracted larvae on 3rd-20th day. Now I am facing difficulty in differentiation of strongyloides stercoralis larvae and other hook worms larvae. I am trying to confirm them by microscopy and focusing on tail morphology of larvae i.e bifurcation in the tail of the larvae but it’s quite difficult and time taking process as for tail ive to see all the larvae at 100X (oil immersion). I am sharing some pictures (10X, 40X and 100X) of the larvae in hope to find some suggestions and materials for morphological identification.
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Dear Nadir,
This paper can be useful for you.
Please consider the possibility you are dealing with other species from goats, such as Strongyloides papillosus.
Hudson
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Can anyone help me in identifying larvae based on Video clip and photographs?
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It is most likely a species of the family Yponomeutidae, perhaps of the genus Yponomeuta. Knowing the name of the host plant should be easy to ascertain the name of the specie.
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Dear all,
We are trying to quantify different proteins in 6 dpf zebrafish larvae. After exposure we collect the larvae in PBS which is then replaced by RIPA buffer containing PMSF and a cocktail of inhibitors. Following homogeneization and centrifugation, we quantify protein by BCA method and apply 50 ug in a 4/12% SDS gel. After running the gel and performing the transfer, we proceed with the common blocking (5% milk), primary antibody incubation (in 5% BSA) overnight at 4 ºC, secondary antibody incubation (in 5% milk) and proceed with the detection using a chromogenic substrate. Yet, and after deeply optimize every step in the western blot procedure, we are still not getting protein signals except for the GAPDH antibody.
We have tried this same procedure with adult zebrafish brain and we get bands for some of the antibodies that we're testing but the larvae do not show any band (an example can be found attached).
Does anybody have a possible explanation for not getting any signal from larval samples? Is this possibly related to antibodies specificity for each organ?
Thanks in advance
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Dear @Elisabeth E LeClair
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Hello,
I need to stain in a complete way all the cell membrane of a zebrafish larvae around 6dpf.
I approached bodipy, but this staining doesn't work in fixed larvae.
Some of you have some ab, or staining usable?
Thankyou
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Some proteins against human proteins thats stain cell membrane also cross react with zebra fish, its worth to try some of those.
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Compliment of the season.
A project was given to my team members and I to study the aforementioned topic but we have being unable to get any articles to review... could you be of help please
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Thanks Nasrin.
If I may ask what's your research area?
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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 encountered some problems during the western blotting of protein extracted from zebrafish larvae. I use the chloroform: methanol=1:3 configuration to remove the yolk solution, but after centrifugation, an insoluble precipitate is formed, resulting in poor protein quality. I would like to ask how to remove the yolk efficiently without affecting the quality of the protein. thank you!
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Deyolking embryo homogenates for running on gels is a perennial problem. We've used everything from the old-school titurating with a p200
to the heptane/MeOH method published by Schnable
Especially for later stages, the best method is to mechanically deyolk by drawing the embryos through a glass transfer pipette that has been drawn out in a flame and broken open such that the opening is narrower than the yolk ball but wider than the embryo's head.
If you come up with a better approach, you should publish it.
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I study larvicidal effects of essential oils on house fly larvae in the lab. I use dipping method and put larvae into essential oils for 60 second after that put them on a plate with some feed for 24 hour then count number of live larvae. My problem is that I can not find death larvae between feeds particles. I used feed ingredients based on literature review. I tried different materials such as alfalfa, soy powder, yeast, grape syrup and barley powder. My question is that which feed is suitable for this experiment?
another question is that my essential oils dont kill all larvae in high concentration ( 1000 ppm was tested for Thymus vulgaric, lavendur officialis, neem oil lemon grass,...) larvicidal effect of this compouns were reported by many papers before.
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Colleague
You can use beer bagasse mix with bovine blood and/or cheese whey. Put attention in the quantity of oils when you propose scale up your recommended dosage.
Regards,
Redimio
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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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Dear all,
I am new to the world of Artemia and I am running into some troubles I hope someone might be able to help with :) I grow A. franciscana nauplii in the lab (starting salinity : 45g.l) in glass tanks, at 23 degrees C and feed them with dry spirulina (1g diluted in 200 ml of RO water, and then I dispense ~1-2 drops of this solution per individual). I use Red Sea Salt diluted in RO water but I do not proceeded at any water change between the moment they are naupli instar 1 and to the moment they form pairs. I incrementally increase the salinity by 5% every other day when the larvae are 10 days old to reach a salinity of 90g/l. I noticed from the literature that my Artemia are growing slowly: ~4weeks to reach adulthood. Not sure why, could be density, but this is not my biggest issue. See below...
When pairs form in the tanks, I isolate them into individual containers (so far, I have tried plastic cups, plastic drosophila vials, and glass beakers) in a volume of 150 ml of freshly made 90g/l salt water, give the pairs a drop of food. Within 48-72 hours, all of these isolated pairs die whereas those in the main tank, at the same salinity, with the same diet, exposed to the same photoperiod regime and the same air, are just fine. Does any one have any idea as to why the isolated pairs die when the others are fine?
Worth noting: in the isolated pairs, males develop very quickly some black spots on their legs are claspers. I do not know what this is, I can't really find any info on this potential (?) issue, but suggestions on what that might be are welcome :)
Many thanks in advance for your help and your time
Clementine
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Optimum salinity around 36 psu is reqd., but NH3 and DO level to be considered
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i need help regarding designing oral feeding bioassay for adult beetle (scotinae, coleoptera). my study involves profiling the effect of a toxin on midgut of the insect through transcriptomics. i have tried some semi artificial diet based protocol. but it did not work and i am unable to find much literature on it. please suggest any other way to feed the insect so as the chemical reaches the gut.
i have been working on adult beetles. can i also use lower developmental stages (larva or pupa) for such a study.
thank you :)
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Dear Aisha,
Good job. This seems really an interesting topic for me to work on and therefore, I got inspired to have a quick survey in Google scholar about it. Interestingly I came across bunch of papers describing the transcriptome analysis profiling the effect of toxins like Bacillus thuringiensis based biopesticides on mainly lepidopterans. Just few of them worked on Coleopterans. So, your work may be a great job to find new achievements in this case. Hereby, I send you two related papers published recently one on lepidopterans and one on coleopterans. First read the method part and then check the references you may have the chance to find new approaches in the case of your research. Good luck.
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Adults, larvae and pupae are Larger than D. melanogaster.
Longer life cycle.
Pictures (on millimetric swuared paper).
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Hi, Celine, thank you. I have no answer, from anybody so far. Can you identfy the specimen?
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I am Kodai Fukada, a member of the Chemistry Department at UNIVERSITY OF TOYAMA FACULTY OF HUMAN DEVELOPMENT ATTACHED SCHOOL. I can secure beetle larvae, but I don't have HPLC, so I can't purify them. Please answer me!
Beetle defensins are antimicrobial peptides found in the body of beetle larvae. It kills germs that invade the body of the beetle larvae in decomposing soil with a lot of bacteria, and prevents infection. Beetle defensins were discovered in 1996 by Miyanoshita et al. of the then National Institute of Silkworm and Insect Science (now National Institute of Agrobiological Resources). The number of amino acid residues is 43, and the sequence is as follows vtcdllsfea kgfaanhslc aahclaigrr ggscergvci cre
Beetle defensins are antimicrobial peptides, and many such antimicrobial peptides and proteins have been found in invertebrates. In particular, many insect-derived peptides exhibit antibacterial activity by disrupting bacterial cell membranes, and are being studied as antibiotics that are resistant to drug-resistant bacteria such as methicillin-resistant Staphylococcus aureus. Beetle defensins are also thought to have strong basicity, which may be effective in puncturing the phospholipid membranes of drug-resistant bacteria and causing them to lyse. In order to be used in clinical medicine for humans, the number of amino acid residues that make up beetle defensins needs to be modified. This is because if the full-length beetle defensins are administered into the human body, the antigen-antibody reaction will eliminate the beetle defensins from the body. To avoid this, attempts have been made to reduce the number of amino acid residues to about 10 while maintaining the antibacterial activity. In addition, research is underway to use beetle defensins clinically as anticancer agents. Some of the amino acid modified peptides derived from beetle defensins have been reported to be selectively cytotoxic to cancer cells only [5]. In general, cancer treatment involves chemotherapy with anti-cancer drugs and radiotherapy, but these methods also destroy normal tissues such as white blood cells, resulting in so-called side effects. Depending on the specificity of the cytotoxicity shown by the beetle defensin, a new drug with fewer side effects may be born.
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This is an interesting topic. I know these substances from the genus Nicrophorus. These are promising compounds. Moreover, the imago of these species processes the substrate for the larvae - the corpse of an animal. But purifying these compounds for medical use is very difficult. In addition, getting the required volume (at least 1 ml) is also difficult. I am just as interested in you in these important points.
Regards, Sergey
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Together with David Tempelman I am preparing a key for larvae and pupae of Orthocladius for western Europe. We still miss the larvae of O. ashei, rivicola, ruffoi and rivinus. We should be plaesed with larvae of some of this species. You can sent them to my address.
H. Cuppen
Hogeweg 8
6961 LT Eerbeek
The Netherlands
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did you get ruffoi ? Bruno Rossaro should have it. Pete
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I want to isolate the microbiom from the gut of the larvae but I have problems with the dissection. I tried to use needles and a scalpel from a dissection set for biologists but I had problems to open the larvae and to find the gut. Is there any trick?
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Larvae of Lepidoptera they have well developed head capsule, 3 pairs of thoracic leg and 3-5 abdominal leg; Duster headless and legless; Coleopyeta well develped.head capsule and only three pairs of thoracic leg; Hymenopteta well developed head capsule. Three pairs of thoracic legs and greater than five pairs of abdominal leg.
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I can't seem to find a straightforward answer to this question. I just need to know how big the average cell is at this age. Any resources or papers you find helpful would be appreciated! Thanks.
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Can anyone help me with morphological identification of filarial (Brugia sp. and Wuchereria sp.) larvae especially L1 and L2 in mosquito vectors ? Is there any pictorial guides, standard keys or publications?
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Hello!
I'm a M.S. student at the University of North Carolina at Charlotte studying fiddler crabs and their embryos/larvae. We've been preserving our samples using RNAlater throughout the summer and I'm now extracting them. I've determined that the RNAqueous Micro Kit is the best one for us to use, and I'm using a NanoDrop for quantification.
I did three extractions from the same clutch, each with 50 larvae. My nanodrop concentrations were as follows: 146.7 ng/uL, 154.4 ng/uL, and 126.5 ng/uL. My 260/280 for each was 1.99, 1.96, 1.98, which from my understanding is good. However, my 260/230 were -1.04, -1.74, -1.12.
I have two questions:
1. Are these RNA concentrations high enough to be sent off for RNA sequencing? I'm new to the world of RNA sequencing so any advice is appreciated.
2. What would cause my 260/230 values to be negative? My understanding is that they should be close to 2.
Thank you!
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Hey Caitlin! I recently did a tutorial zoom call for RNA-Seq with Tecam and they mentioned your ND concentrations should be 300 ng/uL or better. That could be for their specific kit, but I would imagine higher numbers are better, given that you will lose quite a bit of sample during all of the wash steps. Also, your 260/230 values can show negative if the instrument wasn't cleaned properly by the previous user/between samples. Alternatively, you could have a bubble in your sample. In the past if I have had weird reads, I load 1.5 uL instead of 1 uL and really watch for bubbles, but that's totally up to you. Good luck!
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Can anyone provide me some reference paper where image stitching is done with a fiduciary marker
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Elizabeth E LeClair Thanks a lot for your answer. It was very useful to us
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I can't find the appropriate handbook to identification.
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Привет! Спасибо.
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Please suggest how to isolate DNA efficiently from strongyle larvae.....I am trying it from QUIGEN STOOL KIT.....itz.not working even after freeze fracturing the larvae
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I have extracted chitin using Lehmann and White method. But I was unable to find the calculation. Can anyone tell me the calculation please
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Hello to all scientific community, I kindly request you very urgent help. Who kindly give me brief information how to differentiate the four stage of Tuta absoluta’s larvae (First instar, second instar, third instar and fourth instar)? How can we measure their lengthiness? As scientific procedure. Thank you for you cooperation
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Larvae complete 4 instars that are well-defined and are of different sizes and colors (Estay, 2000). Larvae are dorso-ventrally flattened and their color changes from creamy white to green during development. When larvae are ready to molt they stop eating and purge their stomach contents, causing their coloration to return to creamy white.
One more point that is you have to monitoring the moulting of each larvae. This could be done by keeping each larva in a single small pitre dish and try to find the exuvae
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after hatching from cyst how to enrich artemia napulii.
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Mr Senthikumaravelan....
To enrich Artemia, we can feed with vitamins or calcium or by adding an emulsion of phospholipids rich in DHA to newly hatched Artemia. The Artemia eat the emulsion. The Artemia are then fed to the fish or can then be kept refrigerated for up to 3 days. Feed the artemia minimum of 12 hours before feeding them to fishes or other organism...
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Hi,
I am working on a zebrafish model and trying to induced stress using copper sulfate. Although there is a very limited number of publications, I have found that experiments were performed at a 10uM-200uM concentration to induce stress/inflammation in larvae. However, I found that even at the lowest concentration (10um) the larvae survived only for 1 hour.
Also, the solution became turbid (Bluish).
I want to know why the solution is getting bluish turbidity? Is the concentration (10uM) is high for copper sulfate? Where I am getting wrong?
Kindly provide your suggestions or related research articles.
Thank you
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Thank you for your suggestions Jakub Grzesiak . I tried with few modifications and it worked fine.
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We are going to produce 1000 litres of 250 LE of Spodoptera NPV and 500 litres of 250 LE of Helicoverpa armigera in the laboratory. These will be distributed in different projects.
For the preparation of this NPV, how much larval population will be needed and how much time to take to produce this much quanity.
Thanking you
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Thank you sir
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Hi,
In previous field studies we observed great crested newt larvae displaying black and yellow belly pattern in the last stages before leaving water. Does anyone have an idea for how long metamorph individuals with belly pattern stay in water before terrestrial dispersal? How stable is this pattern for individualization?
Many thanks in advance!
Xavier
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The time they leave water after morphing is actually right about this time of year, meaning later half of august, but it depend on latitude.
The ventral pattern is to my knowledge the same, neither have I seen any sign of any orthogenetic variation, meaning that those who got yellow with black maculata keep that colour, and do not get orange later. But the latter statement is less certain. The number of individuals I have caught repeatedly is very limited indeed.
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I have gone through few papers that mentions LPS to induce inflammation. Can I use Hydrogen peroxide instead to induce inflammation to the larvae?
Also, I have found papers mentions copper to induce inflammation. However, I couldn't find much reference work done on the same.
Kindly provide your suggestions
Thank you
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Thank you Daniel Carneiro Moreira and Chiara A.M. Fois for your suggestions.
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I'm unable to bleach the eye and body pigments for craniofacial development. I have attached the result I got. Help me with this issue.
Protocol I followed: "Larvae at 5 dpf or older were fixed overnight in 4% phosphate-buffered paraformaldehyde and washed several times in phosphate-buffered saline with 0.1% Tween-20 (PBT). In order to enhance their optical clarity, the specimens were bleached in 30% hydrogen peroxide for 2 hours or until the eyes were sufficiently translucent. The embryos were rinsed in PBT and transferred into a filtered, Alcian blue solution (1% concentrated hydrochloric acid, 70% ethanol, 0.1% Alcian blue) and stained overnight. Specimens were cleared in acidic ethanol (5% concentrated hydrochloric acid, 70% ethanol) for 4 hours, dehydrated in an ethanol series, and stored in glycerol"
Thank you in advance!
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Dear Hari Deva Muthu .B this is a very interesting technical question. As an inorganic chemist I'm unfortunately not an expert in this field. However, you can find an excellent answer to your question when you check the following previously asked RG question:
Staining zebrafish with alcian blue?
Especially the answer provided by Melissa Chernick seems to be really good and rather comprehensive.
I hope this helps. Good luck with your experiments!
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Does anyone know of any publications that have shown Zygoptera predating on tadpole larvae?
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Hello Clay, hopefully these papers attached may help your query. Best,
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  1. This is Entomopathogenic fungal isolates screening experiment under laboratory condition. We want to calculate feeding percentage of S. fruigiperda larval instar as we provide specific quantity of maize leaves to each instar. We count the data after 24 hrs/ 1day which goes up to 10 days. We provide 7, 12, 20, 30, 45 and 60 gram maize to 1st, 2nd , 3rd, 4th, 5th and 6th instar respectively. The maize quantity is for group of 15 larvae and we will replace it after each day. We observed that S. fruigiperda slow down feeing efficacy against some fungal isolates as compared to control. We didn't use leaf disc method because it was not convenient for us. it will be really appreciated if someone can help us to find the formula by which we can do statistical analysis.
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Thank you
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I am using probit regression to check mortality rate of larvae after exposure to various bacterial isolates and my total sample number is 16 including one control. i transformed the mortality rates to probits from finney table.
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Sorry! The table is attached here
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Dear all , I am interested in studying proteins from C elegans intestinal cells. To get rid of other cells I am planning to tag intestinal cells with GFP and then apply FACS to sort cells.
So the questions are 1. How to tag intestinal cells? Moreover,2. I want to study these proteins in all life stages of c. elegans( L1 to L4), how to do that. 
I am thinking of tagging a protein in larvae and follow-it up to L4.Any thoughts ?
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fluorescent pathogens were fed to C. elegans and fluorescence observed in the gut was considered an indicator for bacterial colonization. However, the grinder in the pharynx of these nematodes supposedly crushes the bacterial cells, and the ground material is delivered to the intestine for nutrient absorption. Therefore, it remains unclear whether intact bacteria pass through the grinder and colonize in the intestine.
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I can find many for Chironomidae larvae, but struggling to find one for pupae, but I'm sure they must exist!
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Ive just made a tagged over-expression line of my gene and I want to identify any protein-protein interactions. Does anyone have a protocol I can use, the procedure in my lab is to use Brain ring gland complex samples for a western followed by mass spectrometry. The issue is most of the time it doesn't work and takes a long time to dissect the samples. I was wondering if anyone has a protocol using whole body larvae I can use?
Thank you!
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Craig Silver by any chance do you have a drosophila protein extraction protocol I can use? Thank you so much!
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I have six lot each one contain 10 invertebrate larvae and a concentration of toxin. at the first concentration I have 0 dead individual the 2nd conceentration 1 dead, the 3rd 0 dead the 4 th 0 dead, the 5 th 1 dead and the 6th 3 dead.
how could I calculate the the median lethal dose?
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Fit a logistic-type model and derive the LDx from it. A similar example can be found here using dose.p https://www.rdocumentation.org/packages/MASS/versions/7.3-54/topics/dose.p
Note that you probably can't get a LD50 for your data as the response only reached ~30%, but you may be able to get a LD10
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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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I am conducting research of stomach content of fish that mostly larvae. However, i can't get the value of Fullness index as most fish are small so that the stomach contents can't be weighed and i have only 0.0001 precision weighing scale
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Have you tried to weight the whole fish, removing the stomach contents and then measuring the fish again? the difference of the values should be a estimation of the contents's weight.
If it isn't possible, you could pretty much use Archimedes buoyant force: the weight of dislocated water should equate the weight of the mass you put on the water... You just need a small graduated beaker filled with water. Then, weight is just density (you can get a estimation of water density on the internet) X dislocated volume (that you measure on the beaker). It is not as precise as a scale, but should do it.
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Antlions are one on soil arthropods in their early larvae stage, where they create a conical pit in the sand and wait for its prey to fall in it to devour it as a predator. So suggest the method use to count the population of these insects in the habitat.
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Hi. I hope the following website could help you:
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If yes, can you pls provide a methodology for this?
How about if L3 is straight from an animal source? Will this be much easier? How would I know if the animal has monospecific infection?
Pls forgive me, I'm not an expert and I'm just an undergrad chemistry student.
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Thank you Mr. Michael Uebel and Mr. Aissa Saidi
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Hello all,
I have a rather trivial question: what is the best way for imaging whole Drosophila larvae? I need to image control and mutant larvae to show differences in larval size.
The best I have come up with, is flash-freezing on grape juice plates, letting them thaw, transferring them to a moist slide, arranging them, wicking up the PBS so it doesn't glare, and taking a photo. While this sounds simple, the flash-freezing causes them to shrivel up, making differences in size difficult to show; and arranging them on a slide and keeping them moist so they don't stick to the slide, but also dry enough that different larvae don't all coalesce into the same puddle of PBS, is very time-consuming.
I feel there has got to be a simpler way that I am simply not thinking of, but this is one of those things that nobody describes in the methods sections of their papers, so there isn't much I can do but make protocols up until something works. But I figured I'd ask RG first.
Thanks for the help
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You can use a Stereoscope device, which equipped with a digital camera.
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Can any one help me with a step by step discription on how to Isolate Mesonephric gonad complex of catfish larva using disection microscope?
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I would like to be able to detect parasitoids in lepidopteran larvae. Mostly targeting species from Eulophidae family.
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