Here are a couple of things I think will help you. Please take the time to read this page as it will help you understand & successfully use the rest of the Tissue Culture pages.

First of all you must get the baby food jars clean. I clean mine by rinsing in hot water after dumping the food. Stuff a rag in and swirl it around to clean the walls good. I then run them through an empty dish washer without soap. One grain of food left in these jars will crash the culture. I learned that the hard way. Take your time on the jars, make sure they are clean.

For any tissue culture sterilization you need to have 10% bleach. Make it by using 1/4 cup bleach + 2 1/4 cups water then add 3 drops liquid dish soap. These instructions should be included with your kit but I wanted to put this here for folks just wanting to sterilize seeds and do not have a kit.

Use new sterilized water and new 10% bleach for each sitting and make them the same day you are going to use them. I use baby food jars with autoclave lids for my sterile water now. I make up several jars and use each one one time.

For parts that are hard to sterilize sometimes we soak them in a 10% peroxide solution. Make this by adding 1/4 cup peroxide to 2 1/4 cups RO/bottled water. No need to boil to sterilize.

For parts that are hard to sterilize sometimes we soak them in a PPM solution after rinsing. Its best to use MS as noted below. After the PPM soak place directly into culture jars, do not rinse again.

Make a 2% PPM solution by adding *1 mL PPM to 50 mL water or MS*, sterilize.
For really tough ones you can use 20%-50% PPM and a 12-24 hr soak but make sure to use the MS as noted at the bottom of the page. I make it in baby food jars with auto-claive lids. You can usually get a few to several uses out of it depending on how contaminated the parts are you place in it. Store for later use in the fridge. Sterilize before each use. If it becomes cloudy, discard. You can strain gunk out of it as long as it remains clear. Always allow it to cool to room temperature before using.

PPM % Calculator <-- Pops a small window to enter PPM mL and Water/Media mL to give PPM percent

If you use a spray bottle and mist the room you are working in right before you get started your contamination rate should drop considerably. If possible leave the top of your chamber and the table you are working on slightly misted. It will help keep micro-particles down while your moving around.

Make sure there is no air movement. Turn off the air/heat vents in the room you are working in. If possible close doors so you are closed off from the rest of the house. Turn off any fans that may be in the room, including ceiling fans. You don't want ANY air movement from any source. Don't let anyone in the room but yourself. Avoid opening and closing doors to the room while you are working. Don't talk while you are working. If you have to cough or sneeze get out of the room.

Use 90% alcohol to sterilize tools and your arms/hands. I spray alcohol from my elbows to my hands but I have tough skin. If your skin is irritated try the 70% and if that still bothers you sterilize latex gloves and be extremely careful not to contaminate the culture. I have found 90% alcohol at Walgreens and Walmart, but they just started carrying it. Check around and with some luck you will find it.

Should contamination occur you can pull the seeds, leaves or roots. Use a soft bristle tooth brush under running water to remove all visible mold. Rinse well in Ro or bottled water. Soak in *50% PPM* for an hour and transfer to a clean culture. DO NOT RINSE, take straight from PPM to culture using sterile forceps. It is said roughly 50% will survive. My experience has been rather limited but I have saved both seeds and leaves this way. Out of my three times it succeeded twice, the failure was with D. tracyi leaves. I assume for roots you must decrease the soak time to a few minutes. You need to catch and treat the contamination within three days and the sooner the better.

A chemical found in some pool shock also works well for sterilizing plant tissue, NaDCC. Please look in the tissue culture sections in the forum for details.

Seeds - If you are working in caps like I do you must make sure the caps and the pipette are sterile. You must also take great lengths not to contaminate the sterile water. Thoroughly clean seed lids then soak in alcohol for a few hours before using. If you are using a seed strainer soak it in alcohol before using. Never shoot liquid back into any of the quart jars with a pipette for any reason. I use a paper-towel lined bowl to empty the pipettes.

I place the seeds in a lid then use a pipette to cover them with alcohol, bleach and then the rinse. When the time is up after each soak I use the pipette to suck the liquid back out of the lids. I fill the pipettes I am going to use with alcohol an hour or so before I start, and I stand them in alcohol with the forceps. Use 90% alcohol for this too.

I have not had any problems sucking up seeds even when working with tiny Drosera seeds. Use common sense and it should go fine, don't touch the small seed lids or you can contaminate them.

I set my seed lids on a tupperware lid big enough I can label what is in each seed lid. This works excellent for soaking several different seeds in Gibberellic Acid overnight.

To sterilize seeds they need to set in 10% bleach for 2 - 15 minutes followed by a rinse lasting the same. You should rinse them in sterile water (nuke to boil and allow to cool). Generally, the bigger the seed the longer the soak. It is really the harder/rougher the shell the longer the soak but it seems to go hand in hand and bigger is easier to judge. If outside contamination is going to occur it will most likely happen in the rinse. Just take your time, think and take precautions and all will go well. Always make the 10% bleach fresh, don't store for later use. That goes for the sterile water too, make it the same day you are going to use it.

Hard/rough seeds such as Sarracenia I dip in 70% alcohol then rinse in sterile water before soaking in Gibberellic acid. This is just a fast dip, a few seconds, and a fast rinse.

By far the easiest way I have found so far to sterilize carnivorous plant seeds accross the board is:
Soak in 2%-4% PPM w/basal salts(MS) for 4 hours. Do not rinse, transfer directly to media. Lids like above work well for this. Hard seeds should be soaked in water for an hour or so before GA3 and/or sterilization. This works on a huge range of seed and is my preferred method of seed sterilization.

Leaves - Some leaves are very easy to clean while other are not. The vast majority of carnivorous plant leaves are in the same difficulty range, fairly difficult. Some leaves may need to be soaked in a 10% Hydrogen Peroxide solution for a few minutes before soaking in bleach. Make the solution by adding 1/4 cup hydrogen peroxide to 2 1/4 cups water.

Dip in 70% alcohol and swirl it around a bit. Make sure to let go of it for a second so you don't miss the little spot your holding with the forceps.

Next set it in the 10% bleach for the listed time. It needs to be submerged which can be accomplished a couple of ways. One way is to lean the forceps on it to hold it down, another is to cap the jar and gently swirl it, I do the latter. Then rinse for listed time.

Roots - Rinse in water good then Sterilize by soaking in 10% bleach for 5 minutes and rinsing well in sterile water. The roots I have done have not been very tolerant to sterilization, but at the same time are hard to sterilize. The best method I have found is to soak them in a 1 part PPM/50 parts water mix (sterilized) for 15 minutes after sterilization. Transfer to medium without rinsing.