FlytrapCare Carnivorous Plant Forums

I made two more liters of media tonight and I used gelcarin in 10 of the flasks. It definitely gels firmer than the phytotech agar (a111) for me. I'll need to use less of it next time as I think the media is a bit on the firm side.

I do really like how clear it sets up though. Hopefully the plants grow as well on it.

I'm thinking 5.5 to 6 Grams per Liter, at the most, is what I'm gonna do.

It's good that We all do different and We can compare notes.

Thanks for Your input, Nep and Matt!

Matt wrote:Thanks for posting this here Gary. I've been following your and Gregorio's posts on the HTCG list serve. It definitely sounds promising. If you can find an alternative to bleach that works as well or even better, that would be great!

As for phenolic bleeding, at least in the case when I use sodium hypochlorite (aka bleach), the phenolic bleeding and dying of the tissue generally only happens the first few days. If you're already seeing that there is no bleeding and the dying of the tissue has stopped, that's a good sign.

Definitely keep us posted Gary and thanks for sharing the results of your experiments with us.

DAY 4: No Mold, No creeping of dying tissue whatsoever; the brown crisp margins are the same, or lack of in the case of the 2 minute dip.

The average 'chalk outline' of black inking Phenolic bleeding You would expect is present where the tissue touches the medium. Repositioned on media. Looking good so far.

Thing is, I did this very sloppily, just for kicks; not even under the hood, but there is ppm in the media and I did alcohol the caps before putting back on.

Wherever I saw a tiny bubble on the leaf (1 or 2)when I immersed in the PAA solution, there is a tiny brown spot there and I've been told that that is an Oxygen bubble where the solution really 'got after it' on whatever the contaminate on that particular spot. In one case, it was actually a tiny piece of moss, (that's how sloppy I did this; no initial rinse). If this solution 'targets' the microbes, and doesn't react with healthy tissue, that is VERY good!

I had a confused Common rescued from a Grocery Store at Halloween that grew a nice, flower stalk, hacked that one and cut a common leaf and used the leftover 4 jars from a good batch and did a plating tonight, following the protocols laid forth by the Author (Gregorio) to the letter. I did a 2 minute swish of PAA on the leaf, cut in half, plated 2 pieces of that, and I did a 1.5 minute on the stalk (it was very clean; only 1 bubble) chopped it and plated the top part with the bud, and a piece of stalk.

We'll see; plating 4 isn't a good test, but a test nonetheless; I only have so many to chop up right now!

Here's another thing; the explants were plated within 5 minutes of cutting. The exposure of the explants in the open under the hood was only seconds. Here is the procedure:

2 drops of Antibacterial soap in 100ml water. Drop explants in; shake good for about a couple of minutes.
Pour out explants and solution in a strainer over a cup, so the explants are suspended over the discarded solution.
NO RINSE
Pull tweezers out of jar that also contains the PAA (no Alcohol), pluck the leaves, etc. and drop in PAA solution.
Start timer.
Swirl; shake, whatever for the given time (to be determined, but 2 - 2.5 looks good so far), then dump through the strainer again over another cup.
Grab the scalpel out of the PAA solution it was soaking in, cut ends, cut leaf, etc as desired. Grab the soaking tweezers, open cap and plate.
Done.

Gary

Sounds simple enough Gary. Thanks for keeping us up to date on that. If the sterilization with PAA works, you should get a strike with the flower stalk. I get more strikes with flower stalks than failures.

I might have missed it earlier in the thread, but could you also list the steps to create the PAA Gary? I remember reading it a while ago when Gregorio first starting posting about it. And since then I've read that the longer you let it set, the more concentrated it becomes (because the reaction is slow or something to that effect). But it would be great to have everything contained in this thread.

Gregorio's instructions were this:

In graduated glass container, add 100 ml Vinegar and microwave until boil. (This has possibly been deemed unecessary, but I think He still recommends it).
Add 400ml Hydrogen Peroxide to bring up to 500ml.
Store in Hydrogen Peroxide bottle in fridge. Mark date.

Use fresh or used, He didn't seem to care, I think it takes 7 to 10 days to 'mature' fully, but He tests with some strips or something and says it is the same strength no matter how old it is. (I guess that is how You measure strength) I used it once that night, and it did great (so far, day 6), but then I did it 48 hours later and it burned all tissue, but I added some harsh steps.

The last test run I did, the solution was 24 hours old and I followed his protocol to the letter and had no ill affects on the leaves at 2 minutes, stalk at 1.5 minutes, whatsoever.

PH = 2.8 at room temperature

ps: I've been using a glass jar with this same solution to sterilize the forceps and scalpel in the experiments; I just don't like the alcohol residue (that I try to be careful to not have) getting on the explant while cutting and handling.

Gary

goldslinger wrote:2 drops of Antibacterial soap in 100ml water. Drop explants in; shake good for about a couple of minutes.
Pour out explants and solution in a strainer over a cup, so the explants are suspended over the discarded solution.
NO RINSE
Pull tweezers out of jar that also contains the PAA (no Alcohol), pluck the leaves, etc. and drop in PAA solution.
Start timer.
Swirl; shake, whatever for the given time (to be determined, but 2 - 2.5 looks good so far), then dump through the strainer again over another cup.
Grab the scalpel out of the PAA solution it was soaking in, cut ends, cut leaf, etc as desired. Grab the soaking tweezers, open cap and plate.
Done.

Couldn't the strainer be a source of contamination, or is a sterile strainer used for the post-PAA step?

As soon as You flush out the PAA and their plants through it, it's sterile; but I did spray alcohol on it before doing everything under the hood.

Gregorio, the Author of this experimental protocol, does it all in the open which is what You can do with high enough amounts of PPM in the jars; at least that is what the producer suggests. He also uses PPM because he microwave sterilizes and it's more touch and go. He also squirts 1ml of the prepared PAA solution onto the media and dumps out the excess just before plating and He thinks this helps alot. It flash sterilizes it and quickly turns into vinegar and pure oxygen, so no harm done, I suspect.

Day 7 and mine are doing great, except for the one I did for 3 minutes; it's burnt pretty bad, but was that way from the first. I think 2.5 minutes tops for VFT leaves. 2 minutes seems to be pretty good, too, but remember, this is all VERY preliminary. It is gonna take alot of people quite some time and a heck of alot of wasted medium to see if this alternative is viable en mass.

This isn't the best time for Me to jack around with it, as I think I was honing in on the traditional proven method; just kind of jumped aboard, but so far, I'm pretty surprised of how well it's working so far. NO migration of dead rot, whatsoever on Day 7 and usually I start to see that by now to a minor degree.

We'll just have to see. Besides, I think I don't have it on good media as I used BAP which Matt and perhaps others have determined that not only is it not necessary, but is actually detrimental to the explant; at least in stage 1.

For what We do, I think it best to do it under the hood and keep the PPM levels low to 0. Like Nep or someone said somewhere, if You follow proper sterilization techniques, PPM isn't needed.

I think I read that Matt doesn't like to get the PPM level over .75ml per liter or something like that, but He can tell You for sure.

goldslinger wrote:I used BAP which Matt and perhaps others have determined that not only is it not necessary, but is actually detrimental to the explant; at least in stage 1.

Yep, that's been my experience. The only successful explants I've had are ones that I've put on media without any PGRs.

goldslinger wrote:For what We do, I think it best to do it under the hood and keep the PPM levels low to 0. Like Nep or someone said somewhere, if You follow proper sterilization techniques, PPM isn't needed.

There are many "hard core" TCers that despise PPM, and I think that Jens is one of those people. Personally, I almost have to use it otherwise I get too much contamination. Perhaps my technique is bad, but I don't have a laminar flow hood like Jens does and without PPM, I get contamination in about 5% of the jars. That's just too much for me, so I use PPM religiously now and will continue to do so until I build my LFH, hopefully sometime this spring.

goldslinger wrote:I think I read that Matt doesn't like to get the PPM level over .75ml per liter or something like that, but He can tell You for sure.

I have tried PPM at 0.5ml/L, but that wasn't strong enough and contamination was nearly as high as it was with none. I almost always use it at 1ml/L and that works well. I've used it at 2ml/L when I know that the tissue has some sort of "mild" contamination (it's contaminated but the tissue continues to grow anyway) and Dionaea seem to have no problem with that level of PPM either.

Matt wrote:I have tried PPM at 0.5ml/L, but that wasn't strong enough and contamination was nearly as high as it was with none. I almost always use it at 1ml/L and that works well. I've used it at 2ml/L when I know that the tissue has some sort of "mild" contamination (it's contaminated but the tissue continues to grow anyway) and Dionaea seem to have no problem with that level of PPM either.

That's good to know for sure; I'll put that in My notes and in My pipe and smoke it.

I made a Gelcarin half batch yesterday and used .5 mL in it; NO BAP, and the rest is correct.
I did the Gelcarin at the rate of 5.5g per Liter, and it is more gelatinous, but not real jiggly; I'm thinking it might be good to go. I'm going to replate the flower stalk, and the leaves that have been sitting on a BAP containing hard Gelcarin media right now; it's been 4 days and I probably need to move them anyway for the Phenolic factor; might as well be on the fresh, proper made recipe with no BAP, etc.

Thanks!

Gary

Sounds good Gary! Please let us know how it goes. I'm excited for you to get your first strike. You've definitely worked for it!

Thanks, Matt!

BTW, there might be something to this 7 Gram per Liter Gelcarin strength. Yea, it's harder, but it's different, for sure.

I really thought it was too hard, but I already had some made, so I put that seedling that Germinated on My first try back in October, (the One since then that has been subjected to all kind of Mad Scientist whims of Mine, and is a BAP,KINETIN junkie mass of whirly callous) that seems to have stalled and vitrified on too soft media I made on Agar last Month, and on the Gelcarin at full strength (7 Grams per Liter) it is taking off again something fierce in a short time.
I also put a couple others on the same batch of Gelcarin at full strength, and they too, are taking off, so I wonder if the transfer rate of this Gelcarin is alot better or something. I also got less water at the harder gel.

I think a very good experiment for Me will be with some Dionaea seeds and see what the germination rate is on varying strenghs of Gelcarin media It the transfer rate is really that good, it will get those little hard black seeds to germinating at full strength. Maybe 20 seeds on each from same seed packet and see.

I dumped some water out of this second Gelcarin batch made at the rate of 5.5Grams per Liter, already. It doesn't jiggle, but it can be pierced really easy, and makes a 'thud' when tapped, not sharp vibration.

It has different properties that We can't guage on what's right by the 'jiggle' method.

It is VERY CLEAR, with a slight golden hue, and I have been catching string mold (or some kind of mold) ridiculously early in some of My experiments, treating, and keeping the tissue alive.

Now My Poly test tubes aren't clear enough to help things, so I sprung for the boron silicate vials and 'Kim' caps, so now it should look like my cuttings will be in suspended animation now, or something.

Thanks for the info Gary.

I too have noticed that the properties of Gelcarin are different. I've only used it in about 20 jars now in a couple of batches (a few batches per jar), but I know what you mean about it seeming harder when tapping the jar, but then it's really easy to pierce. I haven't had any tissue on it long enough to know if it will perform better than agar or not. After hearing your results, I'm excited to see though.

Day 15 on the aahpaa treatment experiment; if You can call it that; I was just horsing around because of boredom that day.

I have two left that are viable out of 4. Not enough cultures to start with to do a real test, but so far, the explant that was soaked for 2.5 minutes still looks as fresh as the Day I put it in! I have good vibrations about that one.
The other one that was left in for 2 minutes, isn't looking so well, but still has alot of green on it, so We'll see. The 3 minute one is long gone, was badly burned.

The other Day, I plated 6 more leaves and am on 48 hours on them as I type this. All were soaked for 2.5 minutes and that seems to be holding true for soak time in the aahpaa treatment, as they seem just fine so far.

The batch I did on 12-12 don't look so good, and I think the A.B. soap in the jar of water is the reason for that. I really think You are just trapping the dirt and re-depositing it on the explants again if rinsing in the jar.

My next experiment will be to just run them under tap water for about 10 minutes, no soap or anything, just brushing off visible dirt, then into the aahpaa for 2.5 minutes, then plate. I'm not going to do the jar soak as described, though. I think it burns them, and I may have done this on the second try that crashed, but I didn't take notes on that one, either.

My curiosity is peaked on this, and has become my primary means, so far, as it is ridiculously simple and fast.
The first batch that wasn't overthinked and was sloppily done is doing the best!
There is always the chance that for once, I got a very clean explant to begin with and couldn't lose, so that's why I am still testing with a higher volume, and taking notes, this time.

Thanks for the update Gary. I'm glad there's someone in the CP world partaking in Gregorio's experiment. It could turn out to be nothing, but it's definitely sounding promising and even if Peracetic Acid isn't the next big thing in the TC world, it has definitely been a learning experience for you and everyone else partaking in the experience and fun for me to follow.

That's good to know for sure; I'll put that in My notes and in My pipe and smoke it.

I made a Gelcarin half batch yesterday and used .5 mL in it; NO BAP, and the rest is correct.
I did the Gelcarin at the rate of 5.5g per Liter, and it is more gelatinous, but not real jiggly; I'm thinking it might be good to go. I'm going to replate the flower stalk, and the leaves that have been sitting on a BAP containing hard Gelcarin media right now; it's been 4 days and I probably need to move them anyway for the Phenolic factor; might as well be on the fresh, proper made recipe with no BAP, etc.

I am going to re-cant this earlier statement and suggest what most already know, that this level is too soft, and exudes water too much.

I am going to use the suggested 7 Grams as the minimal level of Gelcarin in a whole batch. The few explant seedlings that I have on this concentration is doing just fine; I just have to get over the hardness factor and thinking it was too hard, but it is not Agar.
I am going to make a full batch and chop a lot of plants for the next few weeks, this time around. Hope they can handle it!