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InvisibleJean-Luc Picard
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Questions on isolating monokaryotic mycelium and "spore safe" surfactants
    #9825775 - 02/19/09 11:00 AM (15 years, 1 month ago)

So i want to work with monokaryotic mycelium of P. cubensis...and i was wondering if anybody had a method for either isolating single spores prior to germination, or a method for finding and isolating monokaryotic mycelium post-germination.

Also, what are some surfactants that i can use as an "anti-clumping" agent in my spore solutions, that will not harm or drastically reduce germination rate of the spores?

Another few questions I have is how long with monokaryotic mycelium grow without mating? What is the average germination rate for P. cubensis spores, and out of those germinations, what is the rate of mating among the monokaryotes floating around?

Thanks for all of your help in advance :cool::thumbup:


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Invisiblespacel0rd
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9825917 - 02/19/09 11:31 AM (15 years, 1 month ago)

I think getting single spores is hardly possible. You could monitor petris after inoculation an look for germination.  A common method for getting monokaryotic mycelium is described by Stamets/Chilton in TMC. Spore solution is diluted to different extents and several dishes inoculated with it. There is a good chance that you also get a dish/dishes with the 'proper' amount of spores where you just have like maybe 3 germination points. There it is a lot less likely that monokaryons mate fast compared to normal inoculated dishes. You can check for clamp connection/the lack thereof and transfer the hopefully monokaryotic mycelium.

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InvisibleJean-Luc Picard
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: spacel0rd]
    #9825984 - 02/19/09 11:44 AM (15 years, 1 month ago)

OK thanks, that is almost what i was thinking about doing...also does anyone know where i could get any sort of chitinase enzyme, like a biochemical supply or something?

also still no word on the surfactant, which would probably be necessary for diluting spore solution since spores tend to clump together.

thanks in advance...


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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9826153 - 02/19/09 12:12 PM (15 years, 1 month ago)

There is no need for a surfactant, in your serial dilution use a very weak agar like 0.2% and give each bottle a good shake before transferring to the next dilution.  At 10 to the minus 5 you will definitely have single spores. 

An alternative is a a high powered stereo microscope on a laminar bench. You make a spore slurry spread it on a petri wait for a day and look for germinating spores.  Then using a really ultra  fine needle transfer one of the spores to a clean petri and voila.  Of course you think that you do not shake but wait till you magnify your movements with a stereo microscope. :evil:

a monokaryon should grow until the petri dish is full without mating assuming that you only have a single spore colony on the dish. 

You will be able to determine germination rate from your serial dilutions.

good luck !


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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: solumvita]
    #9826317 - 02/19/09 12:40 PM (15 years, 1 month ago)

Simply put a drop of sterile water on a print and dip your flame sterilized inoculating loop into it.  Swipe the spores in a repetitive zigzag pattern back and forth from the top of the dish to the bottom without ever lifting the loop off the agar. By the time your loop reaches the bottom of the dish,  you'll be depositing single spores.  Watch the dish almost hourly with a microscope for germination, and grab the hyphae right as they emerge before they have a chance to mate.  It's not that hard.

If your spores are very fresh, you can expect about one out of a hundred or perhaps a bit less to germinate.
RR


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InvisibleJean-Luc Picard
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: RogerRabbit]
    #9827690 - 02/19/09 04:48 PM (15 years, 1 month ago)

so the spores wont clump together in a water solution? I am trying to gather information because i would like to attempt a protoplast fusion experiment sometime this fall, as well as a couple of P. cubensis strain hybrids...Im still in the research to see if its possible phase...so please provide any input you think is relevant...also are there any readily available needles smaller than 33 gauge...i mean i think that would work just fine since the nominal I.O. is right around 8 times the length of a cubensis spore...to allow for hyphae growing out from the spores


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OfflineRogerRabbitM
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9828541 - 02/19/09 06:41 PM (15 years, 1 month ago)

You certainly don't need protoplast fusion to cross cube strains, since most will readily cross without it or anything else. Just put them together on petri dishes. Most useful would probably be to cross isolated good performing fruiting strains, thus you'd be crossing dikaryotic mycelium rather than monokaryons.

If you want a surfactant, use a small amount of sex lube or the spot preventing stuff they sell for automatic dishwashers. You don't need it though if you use the technique I gave you above.
RR


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InvisibleJean-Luc Picard
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: RogerRabbit]
    #9829065 - 02/19/09 07:53 PM (15 years, 1 month ago)

well on top of the strain crossing in cubensis species I would also like to do inter-genera crossing (terminology is not good?) :rolleyes:

again I'm just messing around because i love the science behind this...and if I'm successful then I would obviously post the entire experiment with pictures on the shroomery to add to the knowledge base.

Is there an easy way to get a good chitinase enzyme, I've heard that you can cultivate Trichoderma harzianum in a LC with chitin samples added to the LC, and this mold excretes certain chitinase enzymes...and i know that the most common contaminate in button mushroom farms is Trichoderma aggressivum also goes by some name like Trichoderma harzianum type 4 or something similar...so it should be relatively easy to cultivate...Is there a way perhaps to use a type of pressure-assisted filter to filter out the trichoderma cells and spores out of the LC medium, thus allowing one to harvest the chitinase enzymes from the trich so it can be used without contaminating the culture medium in my other experiments...I was thinking one of the syringe filters that filters contaminates out of liquids for the syringe.

could this work, and if not any suggestions for another method?

thanks in advance


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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9833632 - 02/20/09 01:36 PM (15 years, 30 days ago)

Wow, a lot of good questions here.  I was once on a protoplast fusion kick for a long while.  I eventually decided it would be easier to manipulate them genetically.

As for getting SBIs (single basidiospore isolates) I've used two methods that work well.  One method is to dilute your spore solution to a fairly dilute state and plate it out.  Then using a scope just find single isolated spores.  Then I use a micropipette tip under the scope to grab a tiny plug of agar around the spore, then I squirt it out onto another dish using a glass pipette bulb or the like.

That method is tedious.  A better way is to simply dilute your spores so that you get 0-3 spores per a micropipette volume (or per drop).  Then you drop the solution in several places on your dish.  Then check under the scope and find the ones that got 1 spore in their drop site and circle them with a marker.  Then when they germinate just take a wedge of agar from the single spore sites and transfer to another dish.

Tips:  Use a surfactant like "jetdry", tween 20, etc..  You can also pour your dishes right to the top so that you can use higher magnification to look right through the top of the dish.

The mating rate should be 1 in 4 for a given monokaryon from the same print.  This could be higher when mixing different prints or different strains.


Look up some of my past posts on protoplast fusion.  As far as the chitinase, it's widely available but expensive.  It's worth making it yourself if you're going to do any number of experiments.  Just use spent substrate as a part of your Trichoderma growth medium as chitin, not surprisingly, induces chitinase production.  Then extract with a proper solvent, or just use the liquid medium possibly, and precipitate using ammonium sulfate.  There is a paper on this exact procedure and they claim being able to produce decent chitinase of good concentration.  Once you have your chitinase resuspend it in a proper buffer or solution and filter sterilize.  Alternately you could probably wash the precipitate with some type of sterilizing solution and then resuspend it.  If you find the paper please post it or a link because I would like to find it again.

For protoplast fusion you are going to need to create auxotrophs of each species (lookup filtration enrichment).  Then you need to suspend them in an osmotically stabilized solution.  Then digest with chitinase.  Then fuse protoplasts with PEG.  Then regenerate them on minimal regeneration medium.

There was also a patent we discussed here about simply grinding myc up in some soil with some PEG.  They claimed to create fusants using this method but I can't find any literature on the technique and it sounds questionable.

For further reading I suggest "Fungal protoplast: A biotechnological tool".  They have summarized a lot of info on optimal concentrations, times, and media for a variety of species.  You might also check out "Genetics and Breeding of Edible Mushrooms" there is a lot of protoplast fusion information there also.

Good luck, and keep us posted.


-FF

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: fastfred]
    #9835722 - 02/20/09 09:42 PM (15 years, 30 days ago)

wow thanks fastfred :thumbup:

I was looking around and found a method of cultivating trichoderma for optimal chitinase production.

Apparently a nutrient solution containing:
  1.5% colloidal chitin
  .42% peptone for nitrogen source, although increasing slightly increased yield

inoculated with 36 hour-old trich mycelium, and incubated at 30oC for 96 hours produced a fair amount of chitinase


I also ran across an article for a good isotonic chitin dissolving solution:

1. Mix .163 mg lyophilized chitosanase (Sigma C-9830) in 16.3 mL of .1M phosphate buffer at pH 5

2. Mix .3 mg powdered chitinase (Sigma C-7809) in 15 mL of .1M Phosphate buffer at pH 6

3. Mix 25 parts of solution 1 with 25 parts of solution 2 into 50 parts of a 3M solution of sorbitol containing 1% of Tween 20 (polysorbate 20)

This produces a pH 5.5 solution containing:
  .25% chitosanase
  .5% chitinase
  1.5M sorbitol
  .5% Tween 20

Optimal Chitin digestion occurs at 30oC for 4-34 hours...depending on spore type.


This article also described how a 30% PEG concentration is optimal for protoplast fusion, as is a pH of 9. Also the addition of Ca2+ ions may help in small amounts. But apparently the PEG increases the chance of fusion over 1000% what it would be without it.


My questions now are:

1. for the auxotrophs, is the purpose of this to slow down cell metabolism so you can fuse without worrying about the cell dividing?

2. If this is the case, would you be able to germinate single spores in a nutrient-deficient solution and allow the hyphae to use up stored energy passed down by the parents, then allow the mixture of chitinase and Tween20 to seperate individual monokaryotic protoplasts? Or would this method kill the hyphae or not work because the cells are stuck together.

3. How would one go about making a minimal regeneration medium?

4. How long will monokaryotic mycelium grow vegetatively on agar, or is it indefinite but slow?

Sorry if these are redundant questions I'm just trying to get a feel for what I'm getting myself into :cool::eek:


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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9841570 - 02/21/09 09:55 PM (15 years, 29 days ago)

This could be an interesting project.

To put in my 2 cents, the purpose of the auxotrophs is to have parents that each lack the ability to grow without supplementation of some (different) essential nutrient (mineral, amino acid, etc)

Then, when you have regenerants that may or may not be fusants, you grow them on a minimal medium to allow you to select the fused protoplasts, with the idea that each would complement the other so they could grow without the supplements required when they grow alone.

does that make sense? I am kind of just waking up from putting the kids to bed.

anyway, there is a lot that goes into it, but I think its a worthwhile project. I am very interested in the chitinase production you are talking about.

keep us posted

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: caricapapaya]
    #9849291 - 02/23/09 10:18 AM (15 years, 27 days ago)

Yep, that's a good explanation.  Basically you need a method to select fusants from non-fusants and self-fusants.  If you don't you'll end up with mostly non-fusants and self-fusants and you'll have to try screening a large number of regenerants based on morphology.  You also need a way to keep the fusants from segregating.  If you don't they'll simply sector out and revert to the parental types.  You'll still loose chromosomes, but you'll at least ensure that there is at least some DNA from each parent.  The longer you keep them together the greater chance that you'll develop a stable fusant or get some recombination.

A minimal regeneration medium would be mushroom minimal media (MMM) and an osmotic stabilizer.  I just made a post on MMM recently, if you can't find it I can repost it here.  Looking in my book, it seems that most experimenters plated on regeneration media and then transferred to minimal media after 1-3 days.  Regeneration media was CZYAM (Czapek's yeast extract agar) + stabilizer (sucrose+mannitol) or PDA + 0.6M sucrose + 0.6M mannitol.  Sorbitol is used instead of mannitol in some cases.


Your pH is off, it should be around 5.8 for production of protoplasts.  I would not adjust it up to 9 for fusing.

As far as growth characteristics of monokaryotic myc... IME I haven't had any problems propagating it.  Just use standard procedures (refrigerate master plates/slants) and you'll be fine.

A couple other points/ideas...  Snail enzyme is reported to be used with success.  Also it is reported that combinations of enzymes are more effective.  I know snails are easy to grow in a fish tank usually.  If you could get the right kind you could have a chitinase + snail enzyme combination that might work pretty well.  I'm not sure if you could just scrape up the snail trails off of the glass or if you have to crush/grind them up and extract the enzyme.

I highly recommend getting this book:
http://www.amazon.com/Fungal-Protoplast-Biotechnological-D-Lalithakumari/dp/1578080932/

They reviewed literally hundreds of journal articles to compile this information.  It contains a lot of concentrations, pHs, etc. for a wide variety of species.  There are tables and graphs showing optimal data and so on.  It's a lot of handy information if you're doing this sort of thing.  Definitely worth inter library loaning it at least.

It also covers a lot of alternate methods such as mechanical (french press) preparation of protoplasts, electrofusion, etc..  I don't think those ways are better or easier, but it is good to know about them.

Feel free to ask any questions here.  If you need something specific I can look it up for you if it's in the book.


-FF

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: fastfred]
    #9859650 - 02/24/09 10:16 PM (15 years, 26 days ago)

what method should be used to select auxotrophs?...I'm sorry but I cant quite grasp how one would select a parent that cant live without something without killing it. Sorry if this is a dumb question :eek:

If you can point me in the way of an article or link for auxotroph selection that would be awesome...otherwise i may buy that book you posted a link to fastfred.

again, much thanks to everyone involved...you guys have been a huge help...and for anyone else...please dont be afraid to throw your 2 cents into this thread...

Thanks in advance, :cool::thumbup:
agmotes165


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InvisibleJean-Luc Picard
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9861609 - 02/25/09 09:48 AM (15 years, 25 days ago)

i found an article on filter enrichment...lemme see if i understand this:

so basically you take a number of spores and attempt to germinate them in a minimal medium, shaking stirring vigorously the whole time...and every 4-12 hours you filter the mix through a filter medium that is porous enough to allow ungerminated spores to pass through...but hyphae are filtered out. After several runs you have all spores that cant germinate on the minimal medium alone.

Then I'm guessing you would split these up into seperate containers and add an amino acid or essential mineral that is common in other articles on filter enrichment...and watch for germinated spores. Then you can isolate auxotrophic monokaryotes and grow them out for the protoplast isolation.

I'm also guessing that you need to do this for both species that you are trying to fuse.

The question I have is:

When an auxotroph from species A that requires amino acid B fuses with another auxotroph from species A that requires B, that self-fusant would still required amino acid B in order to survive right?

When auxotroph A requiring amino acid B fuses with another auxotroph from species C that requires amino acid D, then do these traits cancel each other out, and could you could grow them out on a minimal regeneration media.

Also...once you have a solution containing fusants...they should germinated and grow just like dikaryotic mycelium, so could you take a drop of fusants and place it on an agar plate (assuming the cell walls have regenerated) and let it grow out...would it sector like normal mycelium...I'm looking for a way to test several hundred fusants after the fusion...and isolation of each sector on several hundred agar plates seems to be the best solution if it will work.

Please feel free to correct me if im wrong and make suggestions :tongue:

Thanks in advance,
agmotes165


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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9863976 - 02/25/09 03:58 PM (15 years, 25 days ago)

Quote:

When an auxotroph from species A that requires amino acid B fuses with another auxotroph from species A that requires B, that self-fusant would still required amino acid B in order to survive right?




Yep.  There is a mutation in some gene that is key in a synthetic pathway for a certain amino acid so the organism must take it up from the media because it can't make it's own.  There is also revertants (another mutation that reverses the original one) and suppression mutations (one that affects another gene that re-enables the pathway), but those are beyond the scope of what you need to worry about.

Quote:

When auxotroph A requiring amino acid B fuses with another auxotroph from species C that requires amino acid D, then do these traits cancel each other out, and could you could grow them out on a minimal regeneration media.




Yep, that's the idea.  It's called "nutritional complementation".

Quote:

Also...once you have a solution containing fusants...they should germinated and grow just like dikaryotic mycelium, so could you take a drop of fusants and place it on an agar plate (assuming the cell walls have regenerated) and let it grow out...would it sector like normal mycelium...I'm looking for a way to test several hundred fusants after the fusion...and isolation of each sector on several hundred agar plates seems to be the best solution if it will work.




Well they are already germinated.  They likely won't grow like the dikaryotic mycelium they will likely show morphology different from both parental types.  They will likely sector out because they will be losing extra chromosomes as they grow.  Keeping them on minimal media insures that you will have at least some genetic material from each parent because if they loose their nutritional complementation they will stop growing or at least slow down and be overtaken.

A usual reason for losing chromosomes is that they will not segregate properly because they're segregation is out of sync with the cell cycle or certain spindle components, etc. don't function properly with each other.  So then you get a new cell with some of the chromosomes, but some left behind in the parent cell.  Chromosomes can also be lost during nuclear division and get chewed up in the cytoplasm.

But sometimes you will get recombination between chromosomes or insertions, inversions, duplications, etc..  In those cases you will have the genetic material mixed up between each species chromosomes and you'll get a brand new phenotype.

There have been examples of new inter-species hybrids created which car stable, will fruit, and display characteristics of both parental types.  This HAS been done before, so don't think that it's impossible to achieve.  But it will probably take a fair bit of work and supplies.


As for suggestions...  Once you get your protoplast generation, fusion, and regeneration down your issue is going to be screening them.  The way I've seen this is mostly in dishes.  They look for stable fusants then try to fruit them.  I don't see why you couldn't pool them in minimal liquid media, grow them out a bit, then try to fruit them by inoculating this into standard substrate.  You'd have less control over them but you should be able to screen a lot more of them at once.

Another thing you might try is mating your fusants back to either or both of the parental types.  This might overcome fruiting problems while still keeping some of the DNA from each of the parents.

As for the filtration enrichment...  This is the way to go.  The other method is to make replica plates from complete media with mutants on it onto minimal media, finding which ones don't grow, and going back to your complete media plates to pick off those colonies.  Believe me, that is quite tedious and you won't be able to pull it off unless you have a pretty good mutation system which produces a good deal of mutants vs. non-mutants. 

There is also a "rescue" method which operates in this manner...  You mutate a population then add an agent (such as an antibiotic, fungicide, or poison) which only kills actively growing/dividing cells.  The idea is that the auxotrophs will go dormant since they run out of amino acids and their respiration will drop low, which makes them less vulnerable to the agent than the faster growing prototrophs.  Then you remove the agent and media, perhaps wash them off a bit (depending on their sensitivity to the agent and concentrations used), then add complete media and/or plate them out, then test them for auxotrophy.

Shit, here is actually a book I read describing the rescue method...
http://books.google.com/books?id=XO4EGzpp1M0C&pg=PA121&lpg=PA121&dq=auxotroph+characterization+method&source=bl&ots=cQkHYUkbpl&sig=Et8gK4i4yRKCZBQkk_vsmh7H4Ko&hl=en&ei=orClSdrQAoTSnQepoImVBQ&sa=X&oi=book_result&resnum=6&ct=result
Could have saved typing that out.

Anyhow, the rescue method would be fairly easy if more was known about the species involved.  But, you'd probably have to figure it our yourself and find the proper agent and concentrations.  If you do any of this work please be sure to post your findings.

If you want to test the nature of your auxotrophs I suggest getting a kit or reviewing the methods.  Basically there are 20 amino acids so by making your medias with certain combinations of the aminos you can screen them quickly.  You basically create overlapping combinations so that if you see it growing in square or wells 3 and 5 but not 1 or 4 you can id the deficiency without screening them on 20 different types of media.  I forget the name for this method, but I'm sure you can find the details if you look hard enough.

Back to filtration enrichment...  There are many different ways to do it.  You can try doing it on spores or on myc.  To do it on myc you grow your mutated spores in complete media, blend them up until they are 1-3 cells long, let them recover for a few hours, filter them out, wash them, transfer them to minimal media, grow them out for a day or so, then filter them again saving the filtrate.  This should filter out all the ones that grew in the minimal media while giving you the ones that didn't.  Then you plate them out on complete media and replica plate those to minimal media, going back to your master plate to pick the ones that didn't grow on the minimal media.

The spore method may seem easier, but I think you'll run into problems with protrophic spores germinating at different times.  Nevertheless filtration enrichment is just an enrichment, you'll still have to screen them regardless of how you do it.  The idea is simply to greatly increase your ratio of auxotrophs to prototrophs so that you can find them without screening 10,000 colonies.

As far as doing the filtering... you might have to try a few different methods.  You might be able to just use a very coarse/fast filter or I have seen people using glass wool or cotton/polyfill plugs in funnels or even filtering through sand.  You could also try using different kinds of fabric (maybe like t-shirt material).

As for creating auxotrophs for both parental species, you might not have to.  If you can find something else to use for the selection.  Perhaps one strain is (more) resistant to a fungicide.  In that case you could simply get an auxotroph for that strain.  Then you'd have strain A that is fungicide resistant AND an auxotroph.  So species A wouldn't be able to grow in minimal media and species B couldn't grow in the fungicide media.  But their fusants should be able to grow in minimal media with fungicide.

There are other ways also, if you know enough about your species.  For example if one is sensitive to fungicide A and resistant to B, and your other species is the reverse you should be able to make media with A + B that only a fusant could grow on.  But you'd have to be careful that it is a resistance gene rather than a sensitivity gene, otherwise you might just get no growth at all.


Good luck with everything and keep us posted.  In your quest it might also be useful to start working with genetic markers such as albino, redspore, and PE phenotypes.  That way you can check your crosses and look for parental phenotypes in your crosses/fusants.


-FF

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: fastfred]
    #9864596 - 02/25/09 05:39 PM (15 years, 25 days ago)

Hey fastfred thanks so much for the info...i will definitely look through that article and probably buy that book you mentioned in a previous post...once i get everything together I will post a new thread in advanced mycology complete with pics...hopefully :grin:

and thanks to everyone else that contributed to this thread, you guys have all been a great help...I hope to give back to this board in the future with a successful protoplast fusion! I'm keeping my fingers crossed :tongue:

Wish me luck!!
agmotes165


--------------------
The universe is under no obligation to make sense to you - NDT

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Offlinecaricapapaya
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: Jean-Luc Picard]
    #9868829 - 02/26/09 11:06 AM (15 years, 24 days ago)

Quote:

Believe me, that is quite tedious and you won't be able to pull it off unless you have a pretty good mutation system which produces a good deal of mutants vs. non-mutants. 




One way that might work for getting mutations is to use short wave UV (ie germicidal lamps around 254 nm)

UV can be mutagenic and might be easier to get and control self-exposure than other mutagen like chemicals.

If you plate out a known number of spores or germlings and expose them to UV for varying time periods and then score each plate for survival of the colonies, an exposure time which kills a high percentage of the material plated out should (or could) produce a high number mutants among the survivors.

You would then screen these survivors for mutations you could use by using the methods outlined by fastfred.

The mutations caused by UV (or many other mutagens) are pretty random as far as what genes might be damaged, but it is one way the auxotrophs have been generated.

keep us updated.

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Invisiblefastfred
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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: caricapapaya]
    #9870675 - 02/26/09 04:05 PM (15 years, 24 days ago)

UV is kind of tough to work with.  Chemicals are easier IMHO, but more expensive.  You can get a decent UV bulb for >$10 at 1000bulbs.com.

I discovered that a standard longer fluorescent light bulb has a diameter that makes it a perfect sleeve to go over the UV bulb.  However I had little success with cutting it properly.  I was able to do it with a diamond wheel after 2-3 tries, but it broke when I tried to polish the ends a bit so they wouldn't be so sharp and likely to crack.  I ended up making a nice sleeve with a bulge and hose barb at one end out of pyrex.

The idea would be that you put a spore solution in it and the sleeve holds all the liquid against the UV bulb.  (About 1/16 inch gap between them.)  That way the UV will penetrate evenly into the solution.  You can also shake it a bit as it's going to get all sides of all spores evenly exposed.

I also have a UV strata-linker which will deliver a measured dose of UV, but I think there are too many variables to make knowing exact dosage very useful.  Each setup just needs to be determined individually by experimentation.

You're shooting for 1% (of normal) germination rate.  This will give you the highest percentage of mutants vs. non-mutants.  You can probably go even lower than that.  Just use a concentrated (dark) spore solution and go until 1-2 mL gives you 2-3 germinations per dish.


-FF

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: fastfred]
    #9876705 - 02/27/09 12:15 PM (15 years, 23 days ago)

ff,

what chemical mutagens have you worked with? I am wondering what would be used with fungi.

I have used colchicine (useful for doubling chromosomes in plants) I dont know if its been used in fungi or not.

Also, a compound which the discoverer called "morphocene" I was supposed to produce mutations by inhibiting mismatch repair during DNA synthesis. In our application (mutating plant material) it killed lots of stuff, and we didnt notice any mutations. although we were just screening for phenotypic variation.

That chemical could probably be used with fungi.

My concern with chemical mutagens is spilling it on myself, although I am probably overly paranoid about the dangers....

so, my question, again, after all the babbling, is what have you used (or what do you know to have been used)?

Thanks

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Re: Questions on isolating monokaryotic mycelium and "spore safe" surfactants [Re: caricapapaya]
    #9878055 - 02/27/09 03:18 PM (15 years, 23 days ago)

I've used EMS (ethyl methane sulfonate), ethidium bromide, and UV since these are commonly found around the lab.  I've also grown some gamma irradiated seeds out before, but didn't do the irradiating myself.  I'd also like to try using nitrous acid at some point.

I don't know if colchicine has been used with fungi.  I would guess that it has.  It would be interesting to see what could be done with it.  You might be able to get some interesting effects like self-fertile spores and the like.

> My concern with chemical mutagens is spilling it on myself,

Wear gloves and clean up well afterwards.  You DON'T want to be getting any quantity of these on yourself.  Cancer is not fun and you sure as hell don't want any mutated or deformed babies.

For most intents and purposes UV is probably the best bet.  It's pretty safe and cheaper than chemicals.  One thing to remember is to keep your irradiated spores/tissue in the dark after the UV treatment.  Photolyases use light to repair DNA damage caused by UV, so if you don't keep it in the dark you'll undo most of the mutations you've induced.


-FF

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