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Hey,
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.
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It is fine to use continuous variables in the Y matrix. I mentioned a binary response because of the way you framed your question. Both the X and Y matrices could be continuous, and for statistical reasons it is better that they are not binary. So use concentrations in X instead of presence/absence, and use your growth parameters in Y. Y can contain one or more variables. I am sure that there are good published tutorials on PLS-DA that would explain further.
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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?
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Hello Ussawit: Mr Holloway has it right. On the other hand, you might try to null experiment to determine if the beetles' behavior is influenced by the light. Do it without beans. A random distribution is predicted. That's quicker than rearranging the whole apparatus. Best regards, Jim Des Lauriers
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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. 
For example:
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?
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Hello, You need a table next to the mass spectra to confirm the compounds (attached as an attached file)
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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!
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Do not know about antennal sensilla, but speaking in general, if you can see place of interest in optical microscope you can mount your specimen in a way that allow to cut through it. I prefer flat molds for such task, it's easy to use a microscope to orient a specimen in a proper way in a flat mold. After polymerization a semi-thin sections may be needed for precise location of place of interest.
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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?
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Terraforming, especially for a planet like Mars, would take millions of years but is possible when the time comes that the Earth cannot supply the resources we need. It would start with manned missions to the planet (which we haven't even done yet and wouls cost billions of dollars). It would also involve changing the planet's atmosphere to alleviate the planet's current harsh atmosphere. Creating domes or structures that people could live should also be done before having people stay in the planet. Moreover, studies show that even after alleviating Mars' atmosphere, people may still need to wear body suits in order to breathe efficiently in the said planet. In conclusion, humans proliferating Mars is possible yet would entail sufficient preparation and funds.
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Need chlorophyll free sample to estimate enzyme concentration but unable to remove trace amount of chlorophyll which impairing the result..!
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You may try to use GCB (Graphitized Carbon Black) and or PSA (Primary Secondary Amine) to remove chlorophyll and other pigment also.
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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.
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 Thank you Dr. Andreas Reinecke for your valuable discussion and suggestion. 
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What is the chemical reaction of ammonium sulfate and ammonium chloride when each of them is dissolved in seawater or fresh water?
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thank you
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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?
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Ion traps in general use a mechanism called AGC (Automatic Gain Control) to prevent the trap from overloading. The ion injection time is set to maintain the optimum quantity of ions for every scan. AGC consists of a prescan and the following analytical scan. During the prescan the intesity of incoming ions is measured and used to determine the ion sampling time for the analytical scan. This leeds to an influence of coeluting analytes especially for in context with the LOD.
If the target analyte and interfering analyte have different retention times there should be no influence at all.
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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.
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Three essential sets are required,the models may vary.
1. Air entrapping device or head space collection : for violates collection
2. GC and HPLC ,GC/MS : For characterisation of volatiles.
3. Olfactometer or wind tunnel.
Your insect rearing should lab should quite sound to maintain pure cultures for good results.
EAG etc may also be essential sometimes.
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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. 
Thank you. 
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I think that a secondary part of your question (pupal production of chemical signals, received by conspecific individuals or individuals of other insect species) can find answers by exploring for instance two very wide fields of entomological research, i. e., a) communication among social insects inside their colonies and, b) relations parasitoid-host and predator-prey (as regards entomoparasitic and entomophagous insects looking for pupae). 
I can add that, for some insects, intraspecific communication by pharate adults (adults still into the pupal cuticle) is known or hypothesized, having the function to synchronize (together with other factors) adult emergence of more individuals.
Best regards,
Rinaldo Nicoli
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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.
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Charles,
Which larva you are using for your study? Does they contain olfactory sensilla to perceive airborne volatiles in the GC effluent?
This paper on Spodoptera litura by FM. Poll et al. 2014 may help.
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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?
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if you consider Wolbachia as an endosymbiont this might be one candidate as it interferes with sexual differentiation of several insect species and infects ovaries. But not sure about how it works, i.e. if it really influences ovary development or uses other mechanisms.
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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!
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Use a Random Forest classifier. This approach will give you, among many other things, descriptive variables (in your case volatiles) for each class (here treated vs non-treated). Random forest can be found in the R package. For a nice description of the merits of this approach in analysing volatiles, see Ranganathan & Borges "Reducing the babel in plant volatile communication: using the forest to see the trees" Plant Biology 2009. 
 
 
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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.
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You should also read the follow paper, and get more informations.
Schettgen, C., Schmidt, A. and Butte, W. 1999: Variation of accumulation and clearance of the peredioxin 5-chloro-2-(2,4-dichlorophenoxy)-phenol (Irgasan DP 300, triclosan) with the pH of water. Organohalogen Compounds 43, 49-52.
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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?
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no idea about this
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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.
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I think, I read in one of the research papers prefer to normal body temperature 37 C
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I am monitoring Sciothrips cardamomi in the field, and i want to study the dynamic population using pheromones
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Thank you very much.
We have been evaluating some pheromones on the market, but none have worked against the species of thrips in cardamom
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I am a researcher and I want to make bioassays on insect chemo-ecology, in our laboratory
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Be prepared on 1 year delivery time from Syntech, Peter Ochenfels is very very busy.
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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.
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I would like to elaborate on this question:
Let’s take the Mediterranean fruit fly, Ceratitis capitata, (Medfly) as an example. Usually in nature, one does not see this fly walk for more than a few centimeters on the fruit or the leaf surface. The majority of its movement is airborne. Its flying range may be from centimeters (inside that tree canopy) to several kilometers (like when crossing the Sea of Galilee, 12 Km across).
For me it seems wrong to consider same set of chemical cues for both activities. I think when the fly walks it responds to low volatile molecules with short range of activity or/and to contact cues. These cues become irrelevant a few meters above the top of the canopies.
If I am right, then it is wrong to use a glass Y- olfactometer, with inner diameter of several centimeters that forces the fly to walk, or to move about with short range jumps or flights, to detect the cues that this fly uses in a long distance flight.
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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?
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Hi Seyed, what I do is to run a GLM with a binomial function (landing/no landing) considering each insect a replicate (1 or 0). You can do this rather straight forwardly using  the software "R" for example. Good Luck, Marco
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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?
Thanks!
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Release rate is important because if the rate change the amount of total volatile also change as a result the attraction or replancy also change .i.e. 1ml/min and 1ml/30 sec is not same amount of volatiles pushing in the wind tunnel.
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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?
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Because it is possible to measure many parameters from the 2D tracks that are recorded with Syntech tracksphere there is not a single response.The first point to consider is what kind of data you will extract from the raw tracks.
You  might be more interested in directionality, final vector length and vector angle,  or mean angles that  require circular statitstics.  But in case you need to compare two treatments that appears to show some attractivity, other parameters that measure a response "intensity" may be calculated directly or with R scripts working from the csv files: total walked distance, speed, both can be treated by standard statistics.  And you can also make distribution stats on the numbers of animals that for instance walk a minimal distance or show positive attraction. 
I agree with Sven that 5 replicates is far from enough.  Due to variability in insect behaviour, 30 replicates per treatment are recommandable. You can find examples of application in our paper Party et al  2013 PlosOne (http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0052897)
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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.
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In terms of actual chemical characterization, none have been described (to my knowledge). However, Fraker et al 2009 (Horm Behav) showed a nice neuroendocrine response from a skin-released compound in larval amphibians. Fish have been well described (e.g., ostariophysan alarm pheromone). The Fraker paper has a nice summary of what is known as well. I doubt my answer helps, but I would also love to know if you have any updates on this question.
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I will be happy to see what you think.
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I would analyze their headspace volatiles or SPME based extraction of their cuticular profile and after identification of the compounds would test them for ant behavior
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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 [800] compared to control [370]). 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.
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Hi Vivek, Thank you for this wonderful question. I believe that B. mori's preference to increase egg laying on paper with EAD volatiles is something akin to the C. maculatus (cow pea weevil) preferring to oviposit on black eyed beans. I work with them. After splitting the primary population (lab based) onto different other bean types for over 3 years (e.g. mung, urid, nigerian honey, moth beans), there was a strong preference to the black eyed beans (personal observation) that could be maternally influenced. So, yes it could be (heritable) maternal effects. Look up some of Charles Fox's papers: adaptive maternal effects. Find some of his interesting articles attached with this answer. Hope this helps.
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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).
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A few suggestions:
Poropak Q is a good material for collection of plant volatiles.
Regarding impurities: - connect your 'air' source to an activated charcoal column and confirm the eluting 'air' or any gas you tend to use is clean of impurities (you chould check this with a blank run in GC-MS of the elluting gas).
It is a good idea to treat the (inner surface of) glassware with trimethylsillyl chloride to block the -OH groups on the inner surfaces of your glass apparatus to avoid loosing the plant volatiles (which tend to stick to the Si-OH surface of the glass).
Ideally, remove the poropak Q or Hayesep-Q and do a head space sampling instead of using a solvent, since solvents also also carry some impurities. Otherwise, run a solvent blank to confirm which impurities come from the solvent.
Good luck in your efforts
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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.
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Hello,
It is becoming more and more clear that the microbiota plays an astonishing role in animal and insects in particular. Usually, pheromones are produced by specific secretory cells. However, it would not be impossible that a primer released from such glands in a pouch could be finally transformed by microorganisms living in the pouch into the actual pheromone. An interesting side-question would be an the mechanism of inheritance of these micro-organims.
Best of luck with your experiments !
JLH
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I am looking for different attractants for females.
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Thanks a lot Vivek.
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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?
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There you go!
/m
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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?
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Martin, first of all I would search for special purposed software. It could be easier than coding it yourself, maybe someone else here will recommend a specific program. For example, I know two papers that describe protein spectra registration: http://arxiv.org/pdf/1311.2105.pdf and http://membres-timc.imag.fr/Sophie.Lambert/papier/Spectrometry.pdf and the software SpecAlign http://bioinformatics.oxfordjournals.org/content/21/9/2088.long It is for mass spectrometry, but may work for other kinds of spectra as well.
As for your specific question:
If there was a peak or a number of peaks that were always present in both samples, an initial estimate for a and b could be obtained by linear regression. Otherwise they could be limited by reasonable ranges. Say if the drift was not expected to be more than 10%, parameter a would fall between 0.9-1.1 and b between 0.9*tmax - 1.1*tmax. The step by which these values could then be changed in a numerical optimization depends on typical peak width. If it is, let's say 0.1% of the chromatogram width, then a and b could be incremented or decremented by a number slightly smaller than 1/1000th (or 1/1000th of tmax for parameter b). In a brute force approach this would mean a little bit more than 100*100 = 10000 chromatogram similarity calculations for every combination of a and b, to find the best combination of a and b among them. I am quite sure this is not the most efficient apprach, though. However, because the peaks are discrete, one probably cannot do things like gradient search to get the optimum.
As a fallback solution, you could also just save the chromatograms as images and try to do registration with appropriate image registration software, probably using more sofisticated methods than the one above, e.g. like the MATLAB code here: http://www.softpedia.com/get/Science-CAD/Image-Registration-Technique-for-Recovering-Rotation-Scale-and-Translation.shtml or the code here http://www.doc.ic.ac.uk/~dr/software/
Of course, the image-based approach will necessarily be inefficient, because the spectra are really just curves, one-dimensinal objects and we are forcing them into a two-dimensional analysis. Best results should be obtained with solving the "non-rigid curve registration problem", as is the case in some of the papers in the first paragraph of this message.
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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?
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The fridge is good, but I recommend getting your seeds into the extraction solvent to get the extraction started. If you don't plan on germinating any of them, but only want to preserve them, then they will preserve better in an alcohol solution anyway. This way, when you are ready to begin analyses, you just draw off a small aliquot and run it. If you don't see much, then let the samples sit at room temp for a month and try again. We've got a Geno-grinder which is awesome at pulverizing tissue for extraction. If your seeds have a thick seed coat, opening them up will speed the process. The other main question is how do you plan to analyze this chemistry? LC-MS is great for this, if you have access to one. Some prefer to use Thin layer chromatography but your options will be rather limited. It would be helpful if you tell us what types of equipment you have available.
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I recently reached Analytical Research Systems through the web but they do not respond to my enquiries.
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Hi there, ARS no longer exists. We placed an order about a year ago for $3000. They were happy to take the money and vanished after that. We came to know afterwards that the company had shut for some time already and we apparently are not the only ones having paid for items we will never receive. My advice to you is to find a local scientific glassblower. They very often are very creative people and are happy to make stuffs that are a bit different from classic orders they receive from chemistry labs.
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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?
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Thanks all for answering my question.
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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?
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So because of this overlap in the olfaction and visual transaction pathways, shouldn't these two cues that are likely to interact, be studied simultaneously?
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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.?
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Diode light. Natural spectral composition incl. UV.
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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.
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I do not understand. What is PET? polyethylene terephthalate?
I guess so, sample with those bags always contaminate plants. In case you do volatile sampling is best done in glass, no phthalates but to a lesser extent. I have a glass jar for potted plants adapted to volatile plant foliage is sampled only and is not sampled the soil. Do you want me to send a photo?
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I'm conducting behavioral study for fruit moth larval parasitoid and need to test parasitoid's antennal response to fruit derived volatiles.
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Hi Kennedy, when you get your system up and running you can view an EAG methodology video at J. Vis. Exp. 63: e3931. Good luck.
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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?
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In my opinion, both are same.
As you may know headspace (HS) volatiles can be collected by two ways 1. static HS collection 2. dynamic HS collection.
In static HS collection SPME or gas tight syringe injection is common way of volatiles collection from HS of a sample.
In case of dynamic collection, a stream of gas (mostly air) passed over a sample and HS volatiles are collected on an adsorbent packed in a tube.
In all the cases HS volatiles are collected. the second method (dynamic collection) can be called as aeration collection or extract as this in this air or some other gas use to push volatiles from the HS of a sample.
for reliable quantitative analysis, dynamic HS collection should be used if it work.
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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?
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I am on the road without access to the literature, but there is a paper form Sylvia Dorn's group in Zurich that uses a color printer to create defined reflectances. RTC
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I read that the pheromone glands of butterflies are in their wings. Is this true and how do we isolate these pheromones?
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I do not know about butterfly pheromone gland location. But the most reliable method to extract these pheromones is probably as follows: isolate the gland and crush it in a small vial with solvent. The pheromone is very likely to be hydrophobic (see literature), thus hexane would be a good bet to extract it. If it is polar, you can use methanol for example.
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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?
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Vivek,
your question on " How will a organism know to do certain things without learning" is on the very basis of ethology. This is one of the aspects of the foundational work done by Konrad Lorenz and Niko Tinbergen (their work made them Nobel prize winners together with Karl von Frisch in 1973).
In fact we assume that behaviors called innate are based on genetically tranferred information contained in a species genome (which can, of course, have variability in certain genes/behaviors).
How this sets of genes determine behavior is currently a very "hot spot" in neuroscience. This means that we still mostly base our opinions on single gene models. For a real picture to be good enough we still need to work on gene networks, which might be a better basis to understand complex outputs such as behavioral or physiological traits. Unfortunately, the whole mechanistic view is not available to answer your question properly, but this does not disqualify the opinion that an innate behavior is a genetically based set of neuronal and motor responses triggered by key stimuli recognized by specific sensory receptors which are there probably due to selection pressures favoring their role.
All the best,
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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.
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Dillman et al. 2012 PNAS... he looks at direct cues from hosts that attract EPN
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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.
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You can go through this
J. A. A. Renwick., Experientia, 15 March 1989, Volume 45, Issue 3, pp 223-228, Chemical ecology of oviposition in phytophagous insects
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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?
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As you may know, there are three importnat components to understand olfactory coding; Behavior, Physiology and Molecular Biology. For behavior in lab you need the olfactometer setups like flight windtunnel, Y-tube, T-maize, open arena etc. For physiological studies the most common techniques used are EAG, GC-EAD, SSR, GC-SSR, Calcium and Optical imaging, Intracellular, GC-MS etc. To study molecular biology the are different molecular instument and techniques are use, but all in all it depends on the availibility of fund and your interest and specialities to decide what is really needed and possible to buy it.
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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" ?
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Totally agree with Padalia Sir......Ecospecies or Ecotypes are genetically distinct geographic variety, population or race in a species which are adapted to specific environmental conditions. Whereas, Chemotypes are chemically distinct entity or species with differences in their chemical composition or secondary metabolite composition.