Spore Print to Agar - The easiest and most efficient way to use and clean-up spore prints.

[@samrees275]

Love the maniacal laughter! ❤😂

[@loveskully15]

Your chuckles are freaking amazing and legit made my day lol, thanks for sharing. I'm doing this today!

[@SporeLuckUK]

Your laugh is contagious

[@thaduke8633]

I just did my 1st spore to agar may 3rd and now have growth today I'm so happy

[@themushroombloke]

I was actually going to ask you if there was a better way than an ennoculation loop and now I don't have to because here it is. Makes so much sense and preserves your precious prints. Fantastic! Keep em coming brother!

[@doloinc]

What's the issue with an inoculation loop? It's vastly more simple than this technique

[@trevorbraden5448]

​@doloinc because when you try to drop spores onto a plate in front of a flow hood they usually blow away..... it's really difficult to get them to land on the plate

[@doloinc]

@@trevorbraden5448 I've never seen any issues spores blowing away. Ever. It sounds like inoculation technique issues

[@trevorbraden5448]

@doloinc I lose alot of spores in front of my hood..... some still land on the agar..... but a lot get waisted..... blowing past the plate...... spores don't weigh anything so doesn't take much for them to blow around

[@themushroombloke]

@doloinc if you don't need a loop that's one less piece of equipment to buy and one less piece of equipment to sterilise.

[@trevorbraden5448]

Man this video was right on time I've been struggling with placing spore prints on plates for awhile now.... they always blow away in front of a flow hood......This is genius....

[@toddwmac]

Thanks Ed...great stuff as always. Also, it is so good hearing you say things like "i have no idea what it is...we'll see" when you are working with contaminated plates and such. Thanks!

[@cubertbensin]

Your videos are my favorite way to start the morning!

[@WhiteBeardHashtag1205]

good stuff indeed

[@starburstmemories]

OMG I never saw or thought of this technique to transfer spores. Looks so much better than scraping with my blade... with the added bonus perhaps of some sort of trap door analog that might get rid of potential contaminants? I'm also wondering, how does a much smaller surface of germination affect colony competition.. as compared to spreading out spores on a plate.. maybe it allows for dominant colonies to express their advantage earlier? Avoiding your T0 plate ending with lots of colonies, strong and weak, competing for resources? Another kick ass video from Ed, I keep thinking I saw them all😆

[@edwardgrand]

Yes. Definitely a trapdoor aspect to it. I'm still up in the air about the second point. So many variables to consider. Lots to think about there. Thanks for watching. I will make more forever :)

[@starburstmemories]

@@edwardgrand Hey Ed, not sure if you reply to replies to your replies.😅 Understandable if you don't! Guess we'll find out. I used your technique with great success, nice plump cultures emerged from under there. Then again it was with a Cambodian spore print, which are eager to succeed... I'd like to try with shier growers, like timbit (a melmac selection apparently, slow grower) and other difficult children. But I've only got swabs for those. Have you ever thought of doing this with a swab, somehow? Would contact with the agar ruin the whole swab in your opinion?

[@edwardgrand]

@@starburstmemories I would rip pieces off the swab and sandwich them between the surface and a cut out square of agar from another place on the plate. I have used that technique when I only had one valuable swab left and didn't want to use it all in one go. I would not suggest streaking that last swab as it will get agar and moisture on it. That will probably cause it to contaminate later from the chunks of agar it picked up from the plate. Or you could just do the grab and drag with a few tufts of cotton from the swab. I then recombine the germinated colonies on a new plate. Even if they are monos, you can reconstitute a dikaryon for fruiting. You can also keep the monokaryons separate for later experiments.

[@edwardgrand]

I just posted this to FB. Thanks for the inspiration :) How to save that last swab (without streaking) and reconstitute a dikaryon from stubborn (or dirty) spore swabs. What happens when you’re on that last swab and you don’t want to risk getting it wet with agar/gellan by streaking? If that last swab is just too valuable and you’re afraid of blowing it all on one streak plate, ‘Grab and Drag’ to the rescue! This also sometimes works well for spores that are old, dirty or difficult to germinate. The G & D not only separates the spores from each other (and possible contams), but they get wedged in the crevices and have more contact with the moist agar, increasing the chance for germination. You can just grab and drag with a few tufts of cotton from the swab, thereby saving the rest for further attempts (e.g. on different media) if necessary. After they germinate, I recombine the germinated colonies on a new plate. These widely dispersed spores might be monos or small colonies of dikaryotic mycelium. Either way, you can pile them up and reconstitute a new dikaryon for fruiting. You can also keep the monokaryons separate for later experiments. 😊 So the basic idea is as follows: 1. Ripe a few tufts of cotton off the swab with sterilized, cooled tweezers. 2. Drag those fibers through the agar/gellan in a serpentine pattern or several times in parallel lines (using the same tufts of cotton fibers). Keep the tweezers tight and the tuft in one orientation. As you pull the fibers through the media, spores will be dislodged and deposit in the crevice created as you move the tweezers through the media. 3. Label and wrap the plate. 4. Allow the spores to germinate. 3 days for fresh spores, up to 2 months for old spores. If they are being stubborn, try adding a few mL of sterile water to the plate to re-hydrate the spores. 5. When you see colonies start to form, pick small (3-5 mm square) pieces from each and transfer them to a pile on a new plate. 6. Chop Chop!!! If the colonies represent a bunch of monokaryons, you need to chop them up in the pile on the fresh plate so that they can undergo fusion/somatogamy and form a new dikaryon. If everything goes well, they will shoot out a portion/sector of rhizomorphic dikaryotic growth which can then be subcultured to a new plate, or just allowed to continue growing if it looks clean. Pros: -Allows you to more efficiently use that precious last swab. -Helps separate contaminant colonies from mushroom colonies. You will get good at recognizing contams when they germinate and avoiding them. -Simultaneously allows you to isolate monokaryons, as well as get a fresh dikaryon for normal fruiting (and collecting fresh spores). -Once you’ve used your portion of the swab, you can pass it on to your buddies or save it for future use. -BTW, if you only have spore prints, you can just simply swab the print with sterile swabs and use the procedure above. This allows you to use a smaller quantity of spores and for them to be dispersed on the swab, making it easier to isolate single spores during the G & D (which separates them further). Another option: You can rip pieces off the swab and sandwich them between the agar surface and a cut out square of agar from another portion of the plate. I have used this technique when I only had one valuable swab left and didn't want to use it all in one go. I would not suggest streaking that last swab as it will get agar/gellan gum and moisture on it. That will probably cause it to contaminate during storage from the chunks of agar it picked up from the plate during the streaking. Remember to have fun. That last swab has often caused me panic in the past. With this procedure, you can usually put those worries to rest! -Ed

[@starburstmemories]

@@edwardgrand Thanks for your detailed answers man! While transferring multiple colonies or reconstituting dikaryons works great for me with some spores, with harder growers (tidbit, mr peanut) I've had mitigated results. It feels like some of the weaker cultures take nutrients/space away from the good ones and generally slow things down or even stall my plates. This was from transferring separated cultures to one plate but not mixing them together. So my next step is definitely to try your "trap door germination" with these swabs, or your "chop chop" approach of culture transfers 🤣 Trap door germination from prints has shown very good results for me, and whether using this or chopping things up together, I feel like proximity plays an important role, maybe not for germination but for plasmogamy and early culture growth (mono or di). My train of thought is that perhaps it forces cultures to compete earlier on and leads to the strongest ones making it out of the pack more efficiently. Totally hypothetical, but that's my hunch. (realise I'm kinda repeating myself here from first post, sry about that) I'd love to hear if you or others have similar experiences. Thanks again for everything dr Ed!

[@PaganPope]

just brilliant with the agar. learn something new every time tnx ED

[@DangerRussDayZ6533]

I've been using a cool little dental pick with a ball on the end instead of a point. Used for some kind of cavity filling. I just dab the spores with the ball end and used the flat end to spread them. It's actually probably using too few spores. I'm going to try this method from now on. Thanks!

[@MonsterMushhead]

I’m so glad you created this video. I’m going to be do some agar projects and this will be the tek I use ty Ed!

[@AzimuthAviation]

Another excellent and educational video. Coming from a physical science/engineering background I've found you're a valuable resource as I am picking up experience in mycology. I'm backing out of a liquid culture that has at least yielded opportunity to make spore prints and doing no pour agar later today. Now, if I only had some color shift plasti-dip pearls to go along with the light malt extract ;)

[@josephfuller2716]

This helps greatly I have a spore )

[@nikoashe711]

WoW this technique is just awesome ❤ Thank you for sharing that !!!

[@WhiteBeardHashtag1205]

another great video brother, keep it up, youre good for the community

[@The_Broke_Mycologist]

Just received some prints , heading to sab now ! Thanks for video !!

[@RawTubeTV]

Another great tip that I will use next time. Thanks, Ed!

[@RawTubeTV]

Hey Ed! what are you working on today? Hope you are doing GREAT!!! 🍄❤

[@Dirt_Worm]

I like this, I also like the inoculation loops for this. I used your rip swab tek last night.

[@easttnfungi2144]

Nice one bro I appreciate the time an work yall put n I love learning new things.

[@jethrohetero4990]

Thanks Ed. You are my hero

[@sonnybrasco9735]

Nice video fam

[@JohnSmoke-ih1ow]

Thanks Ed

[@Mycophilia]

Awesome, I will definitely try this out when I get some wild prints!

[@briancunningham3283]

Tweezers eye surgeon uses would be awesome for this. Super fine point have 3 pairs now have a new use for them besides digging out splinters lol

[@wizardofgreen]

Very cool man I like the way you make it simple . So when you grab just the little cotton hair for xfer, are you ever thinking that just that tiny little piece, vs a chunk, will be a great narrowing of the gene pool? Or would the fact that it's maybe still a T1 or T2 mitigate that?

[@edwardgrand]

The idea is to narrow down the gene pool so you can fruit something consistent. It really depends on what it looks like on the new plate. Could be ready for spawn, but some people like variety. The main idea is to make sure it's clean. A smaller piece will more likely eliminate competition from other mono- and dikaryons and possible contaminants that may be lurking.

[@hillbillyherb]

Dang! Wicked technique!

[@growingjourney2755]

Nice I gotta try this!!!

[@oldboots75]

Nice one Ed. Cheers

[@dennispayne3865]

Very nice! Thank you sir!

Oh wow that looks so much simpler

[@gangsterbuilder]

I cut the a little aluminum and drop it in a dropper bottle but I like this better

[@basement_culture]

My partner was just asking me about this the other day! Word.

[@stayanddrown]

Great video! Are you doing this all in front of a hood? I wasnt sure if it's a good idea to do spore and early culture stuff in front of a hood because im worried about contams getting blown all over my work place. And since this stage seems highly likely for contams, just wanted to get others opinions.

[@vincentwu2848]

Whoa! I've never seen this technique anywhere else. I've seen people recommend swiping spore swabs in a Z shape across the plate. What's the reason for the Z-shape swipe, as opposed to your technique of touching the spores to three places on the plate? Also, is your laminar flow hood delivering sterile air from the top? Do you have any thoughts on which type (vertical or horizontal) are better suited for mycology? Thanks!

[@edwardgrand]

The idea of 'streaking', as it's called in microbiology, is to get single spore (or bacterium) separated from possible contaminates and other spores. It was also the impetus behind me doing the 'grab and drag' technique. With the grab and drag, the idea is to get a single spore released from the swab cotton tuft, which then germinates, and can be carefully isolated and hopefully will develop into a monokaryotic colony that can be used for mating. In bacteria, streaking can be used as a rough estimate of CFU (colony forming units), but serial dilution is a much more precise way to do that. In fungi breeding, all we need is one monokaryon to proceed, so we don't need to be too concerned with how we get it as long as it's a monokaryon.

[@edwardgrand]

Re: laminar flow hoods It doesn't matter if it is vertical or horizontal. I have worked with both and it is simply personal preference and individual work flow. If you are doing most mycology stuff (e.g. grain, bags, etc.) I would suggest a horizontal flow FFU. I frequently use a vertical type, but that is because of some logistical issues, not really my preference. TBH, a still air box (SAB) is fine, but when you get to a certain volume of work, it becomes awkward and impractical. I've actually done a fair amount of work using the 'cone of sterility' techniques using a Bunsen burner and alcohol wiped desk. I wouldn't suggest it for long-term, but you can get away with it sometimes (isolated away from contams is one time).

[@experimental-mycologia]

Wow!

[@tmhalula]

Where are you located? Interested in meeting to share techniques?

[@edwardgrand]

I'm far away from the Western world. Except this computer terminal. LOL

[@theresahalula2904]

@@edwardgrand I live in the san francisco bay area. I am enjoying your videos about breeding.

[@kevinfromdevon]

Ed I love your videos BUT.... Only joking. I have a question. I made myself a methylated spirits burner with a hemp wick. It doesn't burn very hot. I went back to using my map gas torch. However, I notice that your burner isn't very ferocious either. Does the item that Iam trying to sterilise need to be glowing red hot? Other mycology KZbinrs often say to get them red hot. Thanks for your time 🖤

[@edwardgrand]

You can listen for the scorching sound when you touch it to agar. If it makes a sound, it is more than hot enough. I also let them rest in 70% alcohol between uses. You can use a cigarette lighter in a pinch. Between transfers, you're really only trying to kill the last thing it touched, which hopefully is clean mycelium, so you don't need to get it crazy hot.

[@kevinfromdevon]

@@edwardgrand thanks for the answer Ed. I really appreciate it. Last time I used the torch I ended up with black deposits (I don't think it was soot) off of the scalpel. I'm guessing I over did it that time, or these scalpels are a sub standard material

[@kevinfromdevon]

@@edwardgrand I will keep a small jar of iso on hand for that purpose👌

[@edwardgrand]

@@kevinfromdevon Soot is sterile. The fungus doesn't mind. Some people put carbon in their media. :)

[@kevinfromdevon]

@@edwardgrand I'm pretty sure that the black deposits were burnt myc or agar. Doesn't appear to have affected anything and I've since transferred away from those dishes again. Thanks for the reply Ed

[@kevinfromdevon]

I tried this technique on 4 different prints and just ended up with bacteria growing on my agar. I'm guessing the prints are not clean enough to start with.

[@edwardgrand]

Yes. I hate to say it, but spore printing requires good technique. I have seen things that send shivers up my spine...

[@kevinfromdevon]

@@edwardgrand I think I will keep using the inoculation loops until I am working with my own prints. Later today I will be trying out your tweezer method on some swabs that just came in the mail

[@edwardgrand]

@Kevinfromdevon if the prints are dirty, it might take some proper streaking technique to isolate clean spores. It sucks. Adds weeks to the effort.

[@kevinfromdevon]

@@edwardgrand I received them from experienced cultivators, so Iam hopeful that I can get something from them. I can only cross my fingers at this point and see what I can do. Thanks for the replies ✌️

[@aldoevortega5224]

Do you have a preferred agar recipe for spore germination?

[@edwardgrand]

I have been using this for everything: SRYA recipe 3% Sticky Rice powder/flour 2% Agar 0.1% Baker's or Nutritional Yeast Food Coloring (if desired) FOR 500 ml Total volume 15 g sticky rice powder 10 g Agar 0.5 g Baker's Yeast Food Coloring or 1g charcoal if desired. 500 ml tap water (not distilled or DI) - Should make between 20-25 standard size plates. But 2 % MEA works fine, too. There is no magic recipe. If you have trouble germinating on MEA, it is likely the spore's problem, not the media.

[@nrolled2888]

Wouldn’t that contaminate the spore if the agar was bad? Id scrape spores onto agar.

[@edwardgrand]

I usually pour my plates a week before using them. No reason they would be contaminated. The idea is to separate the spores from possible contams on the print by 'streaking' them across the plate.

[@trancenaari]

Not Really ! even a little bit agar left of the print will be enough to ruin that print. Tricky, Not Recommendd !

[@edwardgrand]

I make sure the agar spot is dry before packing it back up. Seems fine.

[@ZenAndPsychedelicHealingCenter]

It wouldn't ruin the print at all. A tiny bit of agar, in dry conditions, won't be enough for the mycelium to really start growing.

[@Steveshappylittletrees]

Excellent content. Follows the KISS method to the T (K eep I t S imple S tupid)👍