I fully recommend you OZONE. It's very easy, just bubble ozone in water mixed with vegetal oil (1% tea tree oil for example) for 30 minutes and spray your plants (abaxial and adaxial surfaces of each leaf). It must be a very fast process because ozone turns to O2 very fast but during the application time. Ozone will react with oil promoting a Criegee reaction generating peroxides attached to the tails of the oil molecules. Just try this and tell me if it was useful or not.

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Good luck!

This is an interesting question and I do not have an answer. The ozone answer sounds interesting. As an alternative, on a small scale, have you tried treating seeds with a hypochlorite solution, wash with autoclaved water, and then you grow plants from the seeds in sterile containers? The plants that grow might be clean and provide you with suitable explant material for tissue culture.

Dear Amina Ilyas as Peter M Bradley say, it's difficult to find a proper answer to your question, because of your limitations to use biocides in vitro. I would like to focus in the use of young small explants. It has been demonstrated (although you don't have mentioned the plant species with you're working with) that the younger/smaller the explants, the lower the content endophytic contaminants. The using of antibiotic under ex vitro conditions present a lot of problems, at least for me. Since the applications of fungicides have not functioned, why do you think that antibiotics could do?

I forgot to upload this work

Research Proposal SEEDLINGS AND OZONE. FOE OR FRIEND?

Bleach is a good option. You may need to try out a concentration and time series to find a level that reduces unwanted organisms but doesn't cause too much damage to your plants. A little tween or even dish detergent in with the bleach will help remove contaminants too. Just make sure the plant cuttings are rinsed really well after your decontamination treatment.

The growing of surface sterilized seeds in tissue culture (as mentioned above) is another good option.

You could also try bleach treatment of stem cuttings then get those established in sterile culture then use those for your tissue source rather than going straight from the greenhouse.

Is the goal of your work to have antibiotic free cultured plants? Otherwise, why the ban on using antibiotics?

as Amy suggested, try dilute hypochlorite - 100-1000 ppm at pH 6

Just be sure to use sterile water for any rinses, no point in doing all the hard work to try and get surface sterile plants just to add back contaminants from non-sterile water.

Careful handling and proper surface sterilization are only needed. Avoid the use of antibiotics at any stage of PTC.

Pretreatment of donor stock in the planthouse prior to excision and sterilisation is mission critical for optimal aseptic technique.

I encourage all clients to separate their donor stock well in advance, place in a well ventilated planthouse area with good spacing so material isn't crowded, and water at ground level only for at least 10 days. Misting should be avoided, as should high humidity.

Add appropriate liquid fertiliser to force new growth, which is usually less contaminated than older tissue

-12 days before excision, spray your plant with a compatible systemic fungicide, and respray with systemic fungicide at -3 days

If the parent material can handle it ( some tender leaved tropicals don't ) I apply the following protocol for sterilisation:

• Excise using scissors cleaned with 70% ETOH and place immediately into sterile water ( sometimes adding 100mg/L ascorbic acid can help, sometimes not- assess for suitability beforehand ). Take straight to the lab
• Sonicate plant material for 30s- 1 min in water + a drop of unscented detergent to dislodge larger contaminants- detergent helps with wax and other water insolubles that can prevent sterilising solution getting to tissue
• Swirl or shake in 70% ETOH + a drop of unscented detergent for 30s-1min
• Move explants into sodium dichloroIsocyanurate solution ( I use 1 tablet of Milton antibacterial treatment /100ml H2O + a drop of detergent but check product dose in your country and calculate equivalent ).
• No more than 20% of the final volume of solution in the container should be plant material- 10% is better. You don't want to overcrowd and exhaust your sterilising fluid
• Sterilise for 3-10 minutes depending on tissue type/ species. Make sure the entire inner surface of each container is wetted with the solution. Swirl for preference, rather than shake, to reduce tissue damage.
• In the flow hood, open your SDCN containers, remove tissue with sterile forceps and place on sterile filter paper or sterile paper towel, with no individual plantets touching, and air dry until ready to cut to shape and insert into media
• If you're *truly* desperate, add 2ml/L PPM to the media before autoclaving. Transfer plants to fresh PPM containing media at +2, +5, +7 days before reverting to normal subculture periods in PPM-free media. PPM is a biostat, and can exhaust in the face of excessive challenge, hence the frequent transfers

This is a longer process, the beauty of the SDCN/ baby bottle tablets is that they're gentler on plant material than bleach, less likely to cause chlorosis or secondary wounding, and don't require rinsing with water (a potential stage to re-introduce contaminants via operator error ) and the tablets are stored in discrete packaging which means they're less likely than bleach to exhaust while in storage

I've cleaned up some truly awful parent stock using these protocols. Obviously the more stages you can skip, the better. Do a few sterilising test runs without media and observe tissue damage/ chlorosis/ sterilising solution penetration etc before you throw precious germplasm into the void. And good luck

I never had any luck using ozone to sterilise explants, but combining it with an oil like tea tree is a novel addition and could work

Two more options to add to a protocol for sterilising tricky plants:

• For the final stage of sterilisation you can place your solution + drop of unscented detergent under mild vacuum to ensure closer contact with plant tissues. For hairy or waxy explants ( Davidsonia spp spring to mind here ) this can help
• Or give up on the whole large-tissue idea and go for the meristematic tissue only. I believe this was the only way that Kava plants were put into tissue culture for some time

Dear Amina Ilyas

Prevention of microorganism contamination in plant tissue culture is essential, it can be achieved by careful selection of explants...

Isolate the pots and grow them separately in greenhouse, do not mix with other plants.

Observe the health of the plants.

Avoid selecting explants from insect or mites infested plants, which have more chances of contamination.

Use Bavistin/ fungicide spray in the greenhouse before collecting explants.

Follow the standard sterilization procedures prior to inoculation (plenty of info available).

Avoid Antibiotics.

The use of antibiotics should be avoided even to clear latent microbes from the explant tissues. Again, the plants are growing in a clean environment the chances of such contaminants are rare. Use apical tips to initiate the cultures in such cases and the use of systematic fungicide or bactericide may help to avoid this type of contamination.