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Just thought I'd let you know: I have had a 100% sterile success rate with the peroxide agar kitchen tek, both with the cooked method ( no autoclaving ) and the autoclaved method. Methods and comparison below Dispensing and media transfer was done under absolutely not sterile conditions, in order to see if the tek could be replicated anywhere ( those of you who know my kitchen can stop laughing ). I was a tad sloppy with my transfer tek as well to see what would happen Reporting it here because sometimes the signal/noise ratio can be a little loud elsewhere Batch 1- autoclaved + 6ml/L 3% H2O2 Chinese sauce containers were autoclaved in a bag for 20 min 100ml liquid MEA autoclaved for 20 min, consisting of the following 20g/L light malt extract 0.1g/L garden lime 0.1g/L potassium phosphate dibasic 15g/L gelcarin ( 20g/L agar works just as well ) Media was not subject to a pH test Post autoclave the media was allowed to cool and just above setting point 6ml/L 3% H2O2 was added. Media was swirled heavily so that the inner surfaces of the bottle were fully coated Dispensed immediately into sauce containers *on the kitchen bench* in the open air Lids were placed lightly over the containers and 1hr later the plates were completely sealed after inoculation with various species Batch 2- cooked 1hr + 6ml/L 3% H2O2 Chinese sauce containers were autoclaved in a bag for 20 min 100ml liquid MEA cooked for 1hr by placing media container in an open saucepan. Media container lids were left loose. Water came up the the media level- bottles weren't more than 3/4 immersed. Cooked at a slow boil for 1hr 20g/L light malt extract 0.1g/L garden lime 0.1g/L potassium phosphate dibasic 15g/L gelcarin ( 20g/L agar works just as well ) Media was not subject to a pH test Post autoclave the media was allowed to cool and just above setting point 6ml/L 3% H2O2 was added. Media was swirled heavily so that the inner surfaces of the bottle were fully coated Dispensed immediately into sauce containers *on the kitchen bench* in the open air Lids were placed lightly over the containers and 1hr later the plates were completely sealed after inoculation with various species Sterile ( non-peroxide MEA ) library cultures were opened and haphazardly used ( left open for much of the inoculation process ) to inoculate the plates above using a scalpel blade which was only cleaned and flamed between species The following species were placed in the centre of the agar of each container Reishi ( Ganoderma lucida ) Blue Oyster ( Pleurotus spp ) Lion's mane ( Hericium erinaceus ) Elm Oyster ( Hypsizgus ulmanarius ) Plates were sealed with Austraseal and incubated in the dark at 22C 2 plates from the cooked group and 2 plates from the autoclaved group were left uninoculated as controls to check for contamination during dispensing By my judgement it was all a bit haphazard and I didn't believe it would work. Even a contam rate of 10% per batch would have been acceptable At +1 week there is no contamination, anywhere, and growth is good for the Reishi and Elm Oyster. Still waiting on the Blue Oyster and Lion's Mane, but plenty of time yet- those parent cultures could have been a little old- I have storage/ library cultures and can reinoculate from them easily at +3 weeks If you are thinking about the peroxide tek for agar- give it a go. I've only made it sound complex cos I wanted to write it all up so you know I took all the steps. I now pronounce this part of the tek piss easy Edited very fast, because I am an idiot and forgot to put the decimal point in
Darklight posted a topic in MycologyI'm isolating some local woodloving species onto agar and Trichoderma contam is a constant hassle with one of them. Trouble is the little buggers don't show up as discernable until they sporulate. And the mycelia I am working with is slow to start. What are your best options for working against Tricoderma spp on agar? I couldn't find anything specific on it. My best options so far are Serial dilution- a huge pain in the arse, because Trich is sporulating once it's visible, so the spores go into solution, and there are gazillions of them. Plus a good serial dilution run takes heeeeeeeaps of containers. Over a month. Minimum a hundred. Subculturing ahead- taking only the tiniest fragment of the leading edge, culturing it on, repeat Cardboard slopes haven't worked, as I'm uncertain of this species' ability to grow on cardboard. Still waiting at +10 days to see signs of mycelia. And I've seen Trich grow on cardboard heaps of times. The mycelia would have to grow much faster than the Trich to outrun it and I don't think that's going to happen Is there an agar type media selective against Trichoderma I could try?
(1) For the love of all things good in this world, do not store more grain than you have to; or at least store it properly. For most of my commercial cultures I use organic wheat grain. Some time ago I bought 30kg sack, thinking I would cut down on costs by buying in bulk. The first sign of trouble was a few pantry moths flying around the lab. No problems I thought, I'll leave the sack out in the sun for the day to kill off the weevils and store it in another room. Well, it turns out the sack was infested, and with the humid warm conditions of late summer, I soon had 100's of moths in all my rooms, in all my medias, in all my grains, in all my supplements, in all my everything! (2) Put your agar plates in the fridge as soon as they have set. Leaving agar plates out on the bench for extended periods of time gives mites (YES MITES) a chance to crawl in, contaminate your plates with asper. sp and/ or lay eggs that will ruin your culture later. Don't try to save your plates; freeze them to kill the mites and throw away. Again, this time of year is particularly bad with high humidity. (3) If you can avoid it, don't have any timber in your clean space- given enough time and moisture you can have a mold city growing under benches, which you wont know about, absolutely loading the air with spores. Keep it stainless or plastic. Learn from my mistakes fellow mycologists. With love, Mimz