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Questions regarding the rearing of the Greater Wax Moth



  • oh wow, that's a glowing review, Andrew! I better check my hives for some grubs!

  • edited November 2014


  • For those interested: I received an email stating "... one of the big Wax Worm breeders in Germany ... use a cold process at a certain Time to damage the silk gland, so they stop producing silk ... they wont tell me an accurate method ...."

    I replied the following earlier today & am posting it here what could be potential options. These are not a recommendation that one's entire colony should be experimented with until a strategy proves to work.

    At lower temperature the charge of primary compounds that can make an actual silk fiber are not able form polymers; & thus the larvae can not spin out anything. You would need to experiment to see how low the temperature would have to be in for this; I recall experiments at 4Celsius (40F) stopped silk polymerization.

    There is another approach you may want to try; it works on inhibiting the protein break down that occurs with molting before pupation. Prior to silk spinning the existing larvae has to transform key protein under a sequence of signals; it the signal sequence is wrong then the steps into moulting are not followed through.

    The chemical compound 8 hydroxy-quinoline (8HQ) is a chelator of nitrogen & oxygen. There are protein degrading enzymes (prote-ase) which are responsive to neutral metal chelation & 8HQ inhibits this in insects.

    To moult the act involves a moulting fluid & this is produced when a specific prote-ase acts on protein in the larval exterior (it lets old exo-skeleton be replaced by a new moult's exo-skeleton). Anyway, 8HQ interferes with this & then timely moulting fluid is not adequate; then chitin can't do what it naturally does (come into the right place) & the moult fails to occur.

    8HQ added to the insect diet at no more than 0.06% 8HQ (weight/weight of food) can inhibit the described proteo-lytic response by 17-37%. The larvae will be smaller in later instars than non-8HQ fed larvae of the same age. The 8HQ late instar larvae will take longer to catch up with weight (although 1st few instars they should grow as large as non-treated larvae); they will eventually pupate & can become breeding adults.

    My assumption is that you want them to hang around as larvae so that can cull them for use in a controlled manner; in some stages the 8HQ in feed will give a better growth rate than without it. 8HQ is a competitor for adenine being side tracked & this conservation of adenine lends stability to RNA/DNA; in effect it reduces the cells' trend toward senescence (aging).

    Please consider this experimental & do not expose your entire colony to 8HQ; I have not personally used this tactic. You might get better results with just 0.02% (w/w), but based on reports probably will get the better growth rate using 0.06% (w/w); "frisky" was how one researcher described mealworms fed low level 8HQ.

    This is the commonly available chelator powder that florists add to cut flowers to preserve their vase life; a Ukranian source = You may have to invent a technique for incorporating into your larval feedstock; I don't know what you are feeding; but if wax paper maybe weigh the paper, lightly scratch (or superfine sand paper) the wax paper surface & then spray it with the 8HQ solution to create the desired % coverage.

  • Hi @gringojay,

    Thanks for your attribution. I am confident that the solution lies in the cold treatment. When I put my larvae in the freezer for 5-15 minutes, depending on densities, the larvae stopped spinning silk almost completely. Putting them in the fridge (6*C) for 24 hours did stop them from spinning in the fridge, but after being put back at room temp, they started spinning again.

  • Hello I am trying to rear some G. mellonella for research experiments. I use a diet of bee´s wax and pollen but I am having some problems with fungal contamination. Does anyone know an antifungal that I could use? Thanks a lot

  • Hi victorsb, Citronella has some anti-fungal effect & although repellent to insects is not neccesarily lethal to insects (mosquitos). But I have not looked into nuance of whether insect is in larval form to inform you if it might kill them.

  • Have looked more specifically & see numerous investigations found larvae can be killed by citronella conclude that it would NOT be suitable as an anti-fungal in wax worm rearing because at present I do not know what level (citronella oil active ingredient percentage) is lethal to the larvae , nor if there is a sub-lethal dose which would still be enough to kill the fungal species. If you are able to isolate several test subject larvae in controlled experimentation then you might find a usable dose.

    With fungal outbreaks it is often the transition from after yeast-like state to actual hyphae (root-like) growth that non-synthetic anti-fungals shut down. Then the fungal hyphae don't begin (&/or can not sustain) to modify their immediate surroundings into conditions favorable for increased fungal colonization. Interfering with the ability of budding hyphae to go forward , rather than contact killing of spores (which is harder to accomplish), is a way to knock-back fungal spread & lead to progressive die-back of existing hy

  • OAgain victorsb, - There is another thread currently about fungal contamination (w/ crickets). I've been looking for non-insecticidal agents & want to suggest you investigate flavonoids. Have come across several reports of this wih the general theme being that plant leaf flavenoids in their cuticle wax serve several purposes. One function is to be part of plant defense against fungal growth taking hold. Of course I am not sure all types of plant leaf flavenoids (or a simple tea made to extract leaf flavenoids) is always benign to larvae/insects.

  • Hi gringojay, thanks a lot for your suggestions. I will have a look at flavonoids, I hope I can find any suitable for lepidoptera.

  • RgetHi vivtorsb, - I think you may be able to "spot" treat your fungal incidents since your larvae are presumably not reared in a loose shifting substrate (like bedding for cricket egg laying or bran for mealworm larvae). Hydrogen peroxide or even stronger peroxyacetic acid (described in thread 'very strange colored locusts" 17th comment down Mar. 2015). We use peroxyacetic acid on early spots of contamination that might show up on the agar substrate in petri dishes of plant micropropagation. Hydrogen peroxide is easier to use at 3% strength if want to paint brush it on a spot. When we want to knock out mold throughout a room (if suddenly needed for acclimating micropropagate over production) & not spray fungicide then use 35% hydrogen peroxide concentrate (observing handling precautions) diluted in water so that it (35% hydrogen peroxide) is 1.5% solution (1/4 cup per gallon of water). This premixed solution is poured into a humidifier for dispersal in the room &/or directed toward surface(s) want to target.

  • Tincture of Wax Moth is made commercially in Russia for sale. You can find 2 sellers on eBay (USA site at least) priced around U$ 25/100 ml., or one also has Wax Moth capsules. One seller translates the Russian listing of uses ranging from cardiovascular, bronchopulmonary, genitourinary, inflammation, atherosclerosis, metabolism, prophylaxis, mind, endurance, etc.

  • I don't know if anyone still follows this discussion, but before I start a new discussion, I will try it here. I'm working in a lab at the university and I also have the problem, that the production of silk is disturbing some experiments. I followed the discussion, but I still have some questions. @Entojesse: Sounds like you just started to try to cool the larvaes, do you have now more experience? Didn't the larvaes produce silk until they puppet? Are they puppeting after the cold-treatment? How many days after hatching you put them into -18C? @gringojay: Thanks for all the interesting information, so I can start to understand, how the production of silk is working. But what you wrote is more theoretical. Do you have any idea how to inhibit the ecdysterone in practice? Coul I give some PI3K into the food? (It's very expensive and I don't know, what concentration I will need). Or can I control the production of ecdysteron with the amount of sugar in the food. This point I didn't understand completely and I also don't understand, what you exactly mean with "last instar"? Are there different stages in the development of the larvaes and how can I distingish them? Or can you tell me something about how many days after hatching I have to increase the sugar-amount in the food? (I rear most of my waxmoths at 37C in glasses, but some of them I rear at 30C to have a permanent breed.) Would be nice, if some of you could answer some of my questions and I guess, there will come a lot of questions more

  • Hi aluevera - Wax worms ecdysteroid hormone (ecdysone, 20-E, 20-hydroxy-ecdysone) levels peak 24 hours before it sheds the old cuticle & lives as a 6th instar. In contrast their juvenile hormone levels peak at the time of shedding that old cuticle & they live as a 6th instar.

    A strange coincidence occurs in relation to shedding the 6th instar cuticle. Both ecdysone hormone and juvenile hormone peak 24 hours before the shed that cuticle and they live as a 7th instar with their new cuticle. However I suspect it would be complicated to use 20-E for that & expensive determining dosage.

    This sets up a peculiar paradigm. If artificially pump up the level of ecdysone (20-E) at the starting time of 7th instar (normally wax worms' last instar) a further instar will come about. Instead of sequencing to pupation another cuticle will form constraing the end sector of the silk gland that moves silk proteins long during active silk extrusion.

    Likewise, at that same key instar transitional phase artificially pumping up (administering with food) the juvenile hormone level then will also instigate an extra instar. One tactic to ramp up juvenile hormone synthesis by late instars is to expose them to a lot of frass, which has a compound insect larvae are programmed to use as colony size response activator (lots of frass means lots eating & thus many around; so individuals should hold back in case they are needed for back-up should something happen to their elders). A chemical mimetic of juvenile hormone is fenoxycarb.

    Ecdysteroids come in different configurations, so there are variables regarding their respective receptors. The appropriate receptor (iso-form) has to be generated (spliced from gene) inside an insect cell & actually move toward that cell membrane for exposure to a compatible ecdysone configuration.

    20-E is capable of eliciting the generation of more than on format (iso-form) of ecdysone receptors. In the silk gland during larval instar shifts there are time phases of which ecdysone receptor kicks into action (transcribes).

    It is when both the ecdysone receptor "A" iso-form & "B" iso-form get active that pupation starts. Otherwise the production of ecdysone receptor "A" iso-form inside silk gland cells of the end that moves silk proteins along during silk extrusion is activated 1st & only later the ecdysine receptor "B" iso-form is brought into action.

    Continues ... Tablet having problems now.

  • Continuation:

    The silk gland has 3 sectors. One end is the portion having cells where different molecular weight proteins (fibroins) of silk are made (mostly containing amino acids serine & cysteine). The cells then put out these proteins into that portions interior chamber (lumen)

    From that sector the fibroins (proteins) go to a middle portion of the silk gland where sticky substances (sericins) are made. These compounds (sericins) can be 25% of the spun silk & cover the fibers (fibroins account for 75% of silk).

    Now, if the accumulation of protein fibers (fibroins) inside the middle part of the silk gland is not kept moving along to the other end of the silk gland there is a back log that generates (presumably via mechano-sensitive receptors) signals to stop making as much silk proteins (fibroin). This would interfere with the ability to spin a large cocoon or, in theory, completely finish a cocoon.

    The 3rd part of the silk gland (called the anterior part) has structured locations inside the cells called vesicles. It is these vesicles in that end of the silk gland which must be activated in order for the silk passed along to be secreted out. These vesicles are activated by Calcium ion (Ca++) alterations being induced by primitive neurological processes.

    Which means to me that (potentially ?) inhibitors of neurotransmitter stimulated Ca++ release can stop the silk gland vesicles from performing their function. The compound Ryanodine is a Ca++ release inhibitor that has long been an ingredient in formulating insecticides, so obviously this kind of inhibition is complicated.

    What may (?) do the job is extracts from roots of Valerian herb (Valeriana edulis) since this plant inhibits Ca++ release in neurological context. Of course it is important to know that Valerian roots have been experimentally tested with some success as a mosquito insecticide.

    Both the dose of Valerian and the mosquito variety influenced how much of the root extract actually killed 50% of these bugs. In other words this strategy should be considered experimental and probably a valerian root extract could be trialed on only a test larva ready to spin silk starting at less than dose 0.14 mg valerian concentrate extract per sq. cm. Conceivably you can find a non-toxic dose.

    A guideline for how much active ingredients are in Valerian root depends on which solvent used to extract the active compounds. One research team found that in 100 grams of dried Valerian root using methanol they got 0.75% of active ingredients in weight (0.75% of 100 grams) after filtering out solids and evaporating off the methanol. When they used petroleum ether for 100 grams dried Valerian root they only got 0.17% active ingredient after filtering out solids & evaporating away that solvent.

  • edited November 2017

    Again aluevera, - As mentioned elsewhere in Forum frass has farnesol which in quantity prolongs larval duration instead of normal pupation. I think accumulating frass from your wax worm & introducing extra quantities of that to them is the most logical tactic to try for starters.

    Farnesol activates gene copying (transcription) factors & blockers of gene copying (repressor), so that signal pathways (transduction) are influenced leading to genetic outcomes (expression). The result is a stage of specific protein making (synthesis) being stalled (inhibited).

    Where there are protein changes, including which made & how folded for function, "stress" compensation genes come into action in a reversable manner. Which means the larvae can go back to their usual life cycle genetic programming. Under "stress" adaption paradigm growth stops, in part due to how farnesol alters key protein production.

    Farnesol acts on what is called a "guanine nucleotide exchange factor", which is associated (factor) with messenger RNA translocation. Farnesol stops this kind of factor from normal coordination with a ribosome, so that some kind(s) of guanine nucleotide exchange factor(s) can not form up as complexes with a certain ribosome(s) in the manner normally required for the next step.

    This (lack of active complex) short circuits the start of specific protein making (synthesis) & that means, in this context, the larval metamorphosis to pupation is blocked. The reorganization of genetic copying means that instead of going for pupation there is different gene translation actively controlling the cells; which I assume are in this context cells involved in juvenile hormone levels.

    Farnesol induces the enzymes (protein kin-ase C) that modulate other proteins' action to move from a cell membrane into the cell interior (cytosol). It (farnesol) holds back another compound inside the cytosol from naturally moving that kind of enzyme (protein kin-ase C) outward from interior cell cytosol to the cell membrane. This exerts modification on cell signalling.

    I would be hesitant to use laboratory grade farnesol (not water soluble) in this tactic since farnesol is used in insecticides against aphids. Whereas an insect is able to tolerate it's own frass content of farnesol. I assume this is due to how the farnesol is complexed with other compounds & as such would not use another insect's frass without small lot experimentation.

  • @gringojay: Thanx so much for all these intersting informations. Guess, you're a scientist? So don't wonder if I will have a lot of questions in the future. ;) So I think to try it with different concentrations of Farnesol might be a good and practial posibility for my lab-conditions. I also found another interesting substance I can try: Azadirachtin. It's a compound in "Neemtree-oil", also used as insecticide. I'll let you know, what will be the results. (Excuse me, if my english is not so perfect)

  • Hi aluevera, - Neem tree seed compound Azadirachtin is an insecticide. There is no reason I can think of why you would find it suitable to use on any insect being bred for production.

  • Hi aluevera, - I am having on-line problems. Will only post this for now.

    However, saw one article discussing "new wax comb" (not aged bee wax comb or any pollen) "... prolonged immature stage duration ...." However, "new wax" diet was also associated with a greater rate of mortality.

    The following report's first part also may interest you regarding WaxMoths, if not seen it yet. Injection technique, color photos & identifying male vs. female adults is covered.

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