Questions related to Chemical Ecology
I'm planning an experiment to pinpoint which chemical(s) from a group of 10 components is/are causing an observed ecological effect.
Unfortunately there is no data which chemical is more or less likely to be relevant.
(Edit: To avoid confusion, this is not a question about pollution in aquatic systems, but about odours causing a behavioural response.)
The plan is to have different mixes of the chemicals as my treatments. Each mix will contain eg 4 of the chemicals in different combinations.
The concentration of each component will be constant, the difference will just be a binary 0 or 1 whether it is present or not.
Question 1: Is there a name for this kind of procedure so that I can read up on it?
Question 2: Are there ways or best practices how to design my mixes, i.e. how many different ones and how many components in each.
Question 3: What would be the best way to analyze this? As far as I've read it could potentially be done with a logistic PCA, NMDS or SIMPER analysis, but I cannot figure out which one would be best.
I've been trying to do olfactometer experiments with Callosobruchus spp. (Coleoptera: Bruchinae) for two months, but the results are unreliable. I have two arms for beans and the other two arms for pure air, but the results are that 50% of the insects go to the beans and 50% go to the pure air. In reality the insects should prefer odour of the beans over the pure air.
I think that the light source might induce bias as I work in a control environment room where is ideal for growing plants, so the light source in the room is fluorescent tubes. My olfactometer is on a shelve, and I illuminate the olfactometer from the tubes beneath the shelve (see attached picture). I have to record videos, so I can't open the light above the olfactometer as it creates reflection on the olfactometer. I think that the light beneath the shelve scatters not evenly in the room, so the insect behaviour is influenced by the light.
What type of lamp should I use in this situation while I also can record the video of the insect?
I have calculated values using the formula for Kovats retention indices, and my question is how off does my calculation, based on my column (HP-5MS), have to be from the accepted value for a certain compound for me to be able to definitively identify a compound. I used a C8-C20 n-alkane ladder for my calculations.
Say that I have calculated and RI of 1005 for one of my peaks, and one of the possible compounds has an RI value of 1000. Is that too far off to possibly consider my compound the compound with the RI of 1000?
We would like to characterise the antennal sensilla of a moth using TEM. Does anyone have any tips about ultra thin sectioning? I have read several protocols about it, but none of them describe the process in depth. How could I recognise each type of sensillum after I section the segment? (Our moth has seven types). Some papers even mention the approximate region of the sensillum that was sectioned. How could I do do that??
Any help would be greatly appreciated!
Thesis statements were given to us in our ecology class. One of which included the possible need to terraform a non-habitable planet due to drastic effects of human manipulation in natural occurrences of the environment. Given the fact that the earth's environment, which sustains life, should be followed, is it safe to say that terraforming might be a solution to the now depleting earth? Also, are there calculations as to which we will already be able to determine if terraforming is the only option?
Need chlorophyll free sample to estimate enzyme concentration but unable to remove trace amount of chlorophyll which impairing the result..!
My standard test compound showed repellency against Aedes, Anopheles & Culex. The flight orientation results support the repellency result. But in EAG studies no response was observed in all species. While all antennaes respond well to the puff of glacial acetic acid. What could be the reason of no response in EAG studies? 50, 500, 5000 ppm concentration was used for EAG studies.
What is the chemical reaction of ammonium sulfate and ammonium chloride when each of them is dissolved in seawater or fresh water?
I am trying to detect a compound which may be present at concentrations as low as 10pg/µl via MSMS on an ion-trap GCMS. When looking at the TIC of my samples there are analytes present at concentrations many thousands of times greater than my target analyte. Is there any way that, even when running the machine in MSMS mode, these high abundance analytes could prevent the machine from detecting my target compound?
I want to start a behavioral study between Braconid parasitoid and its host, also some experiments about chemical ecology between the plant, host and parasitoid. I need some suggestions about the lab equipment and also software that needed for this kind of study.
Thank you in advance.
Does anyone know if there's any evidence of pupae being able to sense chemical cues from the environment, etc? Or to produce chemical signals? I'm mainly interested in Lepidoptera, but any other group could be interesting as well.
I'm planning to run a GC-EAD using larval antennae to find out the chemical components they can detect. Working with adult antennae has been successful, but mounting larval antennae is a challenge since, once the head region is chopped off, the antennae retracts hence no contact can be made between the electrodes. Any suggestion will be appreciated.
Previously I had asked a question "Are "sex pheromones" really produced by insects?" while in the quest to answer this question, we came across a very interesting observation. We developed germ free lines of insects (GFL) by antibiotic treatment, the GFLs did not lay eggs at all. When we desiccated the insect we found its ovary underdeveloped. However, the non-GFL insects that were not treated with antibiotics laid eggs. What are the possibilities of endosymbionts being involved in the ovary development?
I currently have a growing excel spreadsheet listing all of the compounds and peaks in my samples and blanks from GC-TOF (time-of-flight). What is best practice for analyzing differences between treatments or even characterizing consistent profiles within a treatment?
Thank you for your time!
Once triclosan enters into the environment, it persists and has been found to be harmful for several species of fish and algae. I want to know what is its bioaccumulation factor in species of fish and algae.
Previously, I had asked a question "Are sex pheromones produced by microorganisms?". Therefore, to test our hypothesis, we cultured the isolated bacteria attractive to B. dorslais males. The volatiles of the cultured bacteria were collected and subjected to GC-EAD and GC-MS analysis. The insect responded very sharply to a single minor peak of the FID!! Using MS library search we identified the compound and to our astonishment the compound was a Spiroacetal. Is it possible fruit flies feed on these micro-organism and harbor them in their body to help them produce these pheromones? If yes, how could insects learn such complex behaviors of harboring a specific microorganism?
I work with head lice and I would like to know if they prefer a specific temperature or a temperature combined with other parameters of the human head.
I am interested in insect chemical behavior; especially it’s response to semiochemicals. A glass Y- olfactometer is a key method which is used in many olfaction studies on flying insects. However, such devices, due to their narrow dimensions of the glass tubes, oblige the insect to walk toward the stimuli rather then to fly, assuming similar respond and I wonder how such reflects the actual responds of the insect when flying.
For a part of my work I'm testing wind tunnel responses of a moth to odor sources. For each odor source, I test 10 moths individually in 3 replicates (A total of 30 moths per odor source). Each replicate is applied in a different day. So, how can I analyse the data?
I'm working on the insect-plant chemical communication. In a part of my work I need to test responses of a moth to its host plant volatile collection. To do this, the gray rubber septum dispenser is available to me. Can I use this dispenser in wind tunnel bioassay to release odor molecules?
I have done tracksphere experiments of a pest caterpillar with Syntech Tracksphere LC 300. I have recorded the responses of the caterpillar towards 6 different plant leaf extracts, extracted in dichloromethane (uninfested and infested leaf extracts of 3 varieties). Five replicates were recorded for each extract, each time with a new caterpillar (separate recordings were done for dcm alone and for air). Can anyone please suggest a method for its statistical analysis?
Anuran tadpoles respond to chemical cues of predation reducing foraging and swimming activity. In many cases this behavior is produced by a predation event releasing different kind of cues, in particular alarm cues, coming from tadpole itself, seem to play a key role in elicit antipredatory responses, even if they often need to be associated to kairomone (from predator) to have the whole response.
We conducted oviposition preference in B. mori and found that they still prefer to lay eggs on mulberry leaves. Further we extracted volatiles from mulberry leaves and conducted a GC-EAD and found that moths responded to specific volatiles. When silkmoths were exposed to these volatiles in a oviposition-choice assay, they selected the papers treated with EAD active volatile as oviposition site. Apart, there was enhanced egg laying (Number of eggs drastically increased on treated papers  compared to control ). Is it that before domestication, the silkmoth with million years of co-evolution with mulberry may have developed "innate recognition templates" for mulberry volatiles to easily detect its host between a plethora of other plants.
We have been using Hayesep-Q for plant volatile collection by dynamic air entrainment, with variable results in terms of impurities and recovery levels. I wonder if colleagues have had better experiences with Tenax or other adsorbents (for solvent elution).
I had previously asked a question "Can bacteria mediate "mating" in insects?" for which i had received wonderful answers from my peers. Taking all answers and reference into consideration we performed certain experiments to ascertain our hypothesis. We cultured a germ-free line (GFL) of B. dorsalis females. Virgin males were given a dual-choice between GFL and Non-GFL B. dorsalis females in a customized olfactometer. We were astonished as males were significantly (P < 0.0001) attracted to non-GFL B. dorsalis females.
We had previously identified 9 bacteria from the reproductive organ of the females and each bacteria was checked for their attractiveness. Two out of nine were attractive to virgin males. Is there a possibility that these bacteria produce "sex pheromones" that attract male towards female flies? Further work is under-process.
We recently isolated 9 cultivable bacteria from the reproductive organs of B. dorsalis. Using 4-armed olfactometer, the isolated bacteria were provided to B. dorsalis as odor source. Out of the 9 bacteria, 1 bacteria attracted males, significantly (P < 0.0001). Is there a possibility of a bacterium acting as a mediator of "Sex" in insects. Can it be a symbiotic host-microbe interaction?
The chromatograms are saved as Xcalibur files and i want to align them (e.g. via R). They actually have a large variance in peak number / concentration and i need an algorithm to make them comparable and reduce the GC-machine drift error. Any suggestions here?
I will start a study similar to Pradeep´s study. But I want identify four secondary metabolites (Tannins, Trypsin inhibitors, Saponins and Alkaloids) from seeds of 30 tropical tree species.
A I am novel in this issue, I am seeking some basic information on how to collect seeds, what exact part do I need collect and what are the conditions of storage. I will be collecting seeds in the rain forest and I do not know what I must prepare seeds and how it keeping while I bring it to the lab?
I recently reached Analytical Research Systems through the web but they do not respond to my enquiries.
Hemiptera is an order of insects most often known as the true bugs, comprising around 50,000–80,000 species of cicadas, aphids, planthoppers, leafhoppers, shield bugs, and others. How does the mechanism of feeding of this order affect the evolutionary biology and behaviour of these insects?
Many studies are addressed either to the visual stimuli (i.e. the overall attraction to colors or the spectral sensitivities of the photoreceptors) or to the olfactorial stimuli (attraction to chemical cues) however, for diurnal insects, when both systems may be stimulated simultaneously, what may be the outcome?
In other words, would an insect that in general is attracted to yellow, still prefer that even when exposed to food- or host- odors?
I am about to install two new working places in my lab for olfactometer experiments and I am thinking about the best light conditions for illuminating experiments with both y-shaped and 4-chamber olfactometers. Do you have any suggestions concerning wave length, intensity of illumination, brand of illuminant etc.?
By sampling headspace of plant volatiles by using PET oven bags we have sometimes measured contaminations by decanal and occasionally by different plant volatiles. Did you make similar experiences with such contaminations? We think the decanal is directly emitted by some charges of PET foil, while plant volatiles seem to penetrate the foil. This could be caused maybe due to quality differences of the product.
I'm conducting behavioral study for fruit moth larval parasitoid and need to test parasitoid's antennal response to fruit derived volatiles.
During the first step of feeding site selection, aphids rely on plant surface stimuli to initiate their probing. Do you know any reference mentioning that compounds produced by aphids, in honeydew for example, can induce a probing in conspecifics?
I want to choose between head space collection and aeration extract for my study, but I could not find the difference between them. Are these phrases synonymous?
I am testing the responses of my insect to visual stimuli in lab bioassays. I have an existing bioassay in which I can measure attraction to color cardboards. I would like to use the same bioassay to test how they respond to their actual host or non-host leaves. I do not want to use real leaves as they dessicate shortly,release volatiles that may affect insects' responses and also because pigment degradation affects very rapidly leaf color. I have the relfectance spectra of the leaves I would like to test. I can't reproduce similar stimuli using my color printer. Would anyone know who/which company I could contact to reproduce a stimulus based on reflectance spectra?
We work on the neuroethology of B. dorsalis. We discovered an oviposition stimulant present in mango that stimulates the flies to lay eggs. We perceived that the fly may have learnt about the cue and may have associated it to mango. However, flies that were not even exposed to this cue previously also showed similar behavior of oviposition. We concluded that this was an inborn or congential template. But, can such templates be transferred genetically ? If so, how?
I would be interested in any published information about volatiles emitted by insects used as cues by insect predators to locate they prey. Best would be studies focusing on belowground systems, but any information will be welcomed.
If so, do you have any references? There are contradictions that a blend of semiochemicals are required for the ovipositional behavior in insects. I would like to know if there is any single specific odor molecule that elicits the above response in insects. We have found a few, but I don't have any references.
I am setting up a chemical ecology lab at my department and am keen on working on Olfaction on insects. Olfactory coding seemed important and would like to know: What instruments and skills do I need prior to starting up the lab? Any suggestions?
I have some confusion about the exact definition of ecotype and chemotype, is it correct to say: If an aromatic plant continues to produce the main constituents of the essential oil in different habitat it will be defined as chemotype "because the biosynthesis has a chemical stability", while if the aromatic plant changes its main constituents of the essential oil depending on the environmental conditions it will be considered as ecotype: "very susceptible to the environmental conditions" ?