I have a method for sterilising the ex plants. 10 minute rinse under running water. Then dipped in 70% ethanol for 1 minute, then 25% bleach + few drops of liquid detergent for 10 mins.
Followed by rinsing in boiled and cooled water.
I am looking at a biocide for the storage itself. Plant Preservative Mixture is one, but not available.
I guess adding a few drops of an antibiotic injection like amoxyillin to the medium might help.
I have Formalin tabs which you can use for sterilising the external holding container.
This Is another good read I found for surface sterilization, almost covering all the points suggested by Madan JI:
Surface-sterilizing Plant Material
1. Preparation of Stock Plants
Prior good care of stock plants may lessen the amount of contamination that is present on
explants. Plants grown in the field are typically more “dirty” than those grown in a greenhouse or
growth chamber, particularly in humid areas like Florida. Overhead watering increases
contamination of initial explants. Likewise, splashing soil on the plant during watering will
increase initial contamination. Treatment of stock plants with fungicides and/or bacteriocides is
sometimes helpful. It is sometimes possible to harvest shoots and force buds from them in clean
conditions. The forced shoots may then be free of contaminants when surface-sterilized in a
normal manner. Seeds may be sterilized and germinated in vitro to provide clean material.
Covering growing shoots for several days or weeks prior to harvesting tissue for culture may
supply cleaner material. Explants or material from which material will be cut can be washed in
soapy water and then placed under running water for 1 to 2 hours.
2. Sodium Hypochlorite
Sodium hypochlorite, usually purchased as laundry bleach, is the most frequent choice for
surface sterilization. It is readily available and can be diluted to proper concentrations.
Commercial laundry bleach is 5.25% sodium hypochlorite. It is usually diluted to 10% - 20% of
the original concentration, resulting in a final concentration of 0.5 - 1.0% sodium hypchlorite.
Plant material is usually immersed in this solution for 10 - 20 minutes. A balance between
concentration and time must be determined empirically for each type of explant, because of
3. Ethanol (or Isopropyl Alcohol)
Ethanol is a powerful sterilizing agent but also extremely phytotoxic. Therefore, plant material is
typically exposed to it for only seconds or minutes. The more tender the tissue, the more it will
be damaged by alcohol. Tissues such as dormant buds, seeds, or unopened flower buds can be
treated for longer periods of time since the tissue that will be explanted or that will develop is
actually within the structure that is being surface-sterilized. Generally 70% ethanol is used prior
to treatment with other compounds.
4. Calcium Hypochlorite
Calcium hypochlorite is used more in Europe than in the U.S. It is obtained as a powder and
must be dissolved in water. The concentration that is generally used is 3.25 %. The solution must
be filtered prior to use since not all of the compound goes into solution. Calcium hypochlorite
may be less injurious to plant tissues than sodium hypochlorite.
5. Mercuric Chloride
Mercuric chloride is used only as a last resort in the U.S. It is extremely toxic to both plants and humans and must be disposed of with care. Since mercury is so phytotoxic, it is critical that
many rinses be used to remove all traces of the mineral from the plant material.
6. Hydrogen Peroxide
The concentration of hydrogen peroxide used for surface sterilization of plant material is 30%,
ten times stronger than that obtained in a pharmacy. Some researchers have found that hydrogen
peroxide is useful for surface-sterilizing material while in the field.
7. Enhancing Effectiveness of Sterilization Procedure
• Surfactant (e.g.Tween 20) is frequently added to the sodium hypochlorite.
• A mild vacuum may be used during the procedure.
• The solutions that the explants are in are often shaken or continuously stirred.
After plant material is sterilized with one of the above compounds, it must be rinsed thoroughly
with sterile water. Typically three to four separate rinses are done.
Using Sterile water is a big precautionary measure that must be ensured while doing the process.
For those who have no idea how to culture, a very basic read on how to do culturing without a laboratory and high tech euipments:
Sterile Culture Techniques
Successful control of contamination depends largely upon the operator’s techniques in aseptic culture. You should
always be aware of potential sources of contamination such as dust, hair, hands, and clothes. Obviously, your hands
should be washed, sleeves rolled up, long hair tied back, etc. Washing your hands with 95% ethyl alcohol is an easy
precautionary measure that can be taken. Talking or sneezing while culture material is exposed also can lead to
contamination. When transferring plant parts from one container to another, do not touch the inside edges of either
vessel. By observing where contamination arises in a culture vessel, you may be able to determine the source of
In plant tissue culture, small pieces of plant tissue are placed on or in a medium rich in nutrients and sugar. If bacteria
or fungi come in contact with the plant tissue or the medium, the culture becomes contaminated. The contaminants
(bacteria and fungi) will use nutrients from the medium and the plant, which quickly destroys the plant tissue. Our aim
is to surface sterilize the plant tissue and put it on a sterile growth medium without any bacteria or fungi getting on the
plant or medium. This is not easy because bacteria and fungal spores are in the air, on us, in us, and under us!
When you see sunlight shining in a window you can, from certain angles, see dust particles in the air. There are
hundreds of bacteria attached to each dust particle. A horizontal laminar flow unit is designed to remove the particles
from the air. Room air is pulled into the top of the unit and pushed through a HEPA (High Energy Particle Air) filter
with a uniform velocity of 90 ft/min across the work surface. The air is filtered by the HEPA filter so nothing larger
than 0.3 um (micrometer) can pass through. This renders the air sterile. The flow of air from the unit discourages any
fungal spores of bacteria from entering. All items going inside the unit should be sterile or sprayed with ethanol or
isopropyl alcohol. They will remain sterile unless you contaminate them.
A transfer cabinet provides an enclosed environment that is not sterile but can be sterilized. A cardboard box lined
with aluminum foil or plastic, or a well-cleaned aquarium, provides a shield to reduce contamination. A box that is 20-
24 inches wide, 20-24 inches high, and 12-16 inches deep provides a good work area. Working inside any of these
does not guarantee success.
The following precautions are necessary for all work areas.
1. The room should be swept and if possible, mopped.
2. Each work surface should be washed with a 10% Clorox, or lysol or other disinfectant solution.
3. Doors and windows should be closed.
4. Air conditioners and fans should be turned off.
5. If possible, each student must have a work space that can be properly treated against contamination. For
example, the box or aquarium mentioned earlier, or a piece of poster paper lying on the table to indicate the
student’s sterile workspace.
6. Have spray bottles filled with 70% ethanol or isopropanol (never methanol) placed so each student has access
to one bottle. Spray everything going into the sterile area. 7. Have a central supply area so all necessary items can be picked up and taken to the workspace as needed.
Items can be returned to the central supply area after being used.
8. Sterile instruments will be needed for each person. One way to accomplish this is to have a ½-pint jar of 70
% ethanol for scalpels and short forceps. When tissue has to be positioned in a vessel, long 10-inch forceps
are needed. The long forceps need to be placed deep enough in alcohol so that any part of the forceps that
might come into contact with the vessel is sterilized. A 100-ml graduated cylinder can be used to hold the
alcohol and long forceps. A ½-pint jar of sterile water is needed for dipping the instruments to remove the
residual alcohol that might dry out plant tissues.
9. A sterile work surface is needed on which to place the sterile tissue to trim it. The easiest thing to use is a
sterile petri dish. If you have glass ones, you can autoclave and reuse them. Presterilized plastic dishes are
used and discarded. Spray the bag of dishes with 70 % alcohol before you open it and place the desired
number of unopened dishes at each station. Each dish has two sterile surfaces-the inside top and inside
10. Long hair should be tied back or covered. Hands should be washed, not scrubbed (scrubbing dries hands and
creates flakes of skin that have bacteria) and sprayed with 70 % ethyl or isopropyl alcohol or coated with
isopropyl alcohol gel. Gloves and masks provide extra protection. Do not talk while performing sterile
operations. Do not lean over your work. Keep your back against the backrest of your chair. Try to work
with your arms straight: this position may feel awkward, but it will reduce contamination. Do not pass
nonsterile items over sterile areas or items. Reach around rather than over. Make your movements smooth
and graceful so that you do not disturb the air more than is necessary. Work quickly though, the longer it
takes to manipulate the tissues the greater the chance of contamination. Have only the necessary items in
sterile work area. Remove items that are no longer needed as quickly as possible. Act out each step before
beginning so that you understand what you are about to do.
Store cultures in a well-lit area (not in direct sunlight), and do not allow the temperature to exceed 80 degree F where the
cultures are stored. If you are placing cultures under lights, use only fluorescent light. The preferred schedule is 16
hours of light and 8 hours of dark. Check the temperature prior to placing the cultures under the lights because
temperature will build even under fluorescent lights.
Check cultures every 3-5 days for contamination. Slimy areas mean bacterial contamination while fuzzy areas are due
to fungal contamination. Do not open containers that are contaminated. The contaminants could be disease causing or
pathogenic. The only safe way to dispose of these is to autoclave (or pressure-cook) them for 15 minutes at 15 psi.
Contaminated plastic dishes can be placed inside a large can or autoclavable bag to be sterilized before discarding.
I have a single runner of Hydrocotyle vulgaris, given to me by Asif (Zipper boy). We can try with it as it seems to be emersed, not sure though. The leaves were waxy and exposed on water surface.
Will check with VASA and get back to you about DMS, Agar and some glass containers.
I will also pick up scalpel, Gloves and forceps from them if necessary(for cutting and replanting). Let me know about it.
Joined: Jun 29, 2003 Posts: 7087 Location: Bengaluru, India
Posted: Thu Jul 26, 2012 12:04 am Post subject: Re: Aquatic Plants Tissue Culture
I'll be seeing my chemist tomorrow about the DMS and agarose and some containers, I don't think the
fellow has containers, and scalpels, check with Vasa too and update, we'll need both the guys it looks like,
or will have to head elsewhere for a scalpel.
Posted: Thu Jul 26, 2012 11:16 am Post subject: Re: Aquatic Plants Tissue Culture
Madan Ji, last year I have already worked on Tissue Culture of Aquatic plants. But in fourth week, due to fungus, I have lost all the work. I have all the essentials required for tissue culture of plants. This year I will start again the process in the end of September.
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