Flasking Cool Growing Orchid Seed by Bob Hamilton

Recently, there have been questions about sterilizing media for growing orchids in vitro -- flasking. Some discussion of microwave ovens came up. My answer is rather oblique to the question of micro waving and definitely not terse!.

 I have been doing my own "bottles" for about 6 years. Most of what I have learned has been through empirical methods. Using the algorithm, ready, fire, aim, I've learned to reliably flask Odonts and Masdevallias. Basically, I had to learn to flask cool growing orchids because there were no cool commercial laboratories available to me. I had disastrous results with the labs I tried. I also like to doing things for myself.

 First off, let me say no one will put more care and effort into your flasks than you will. In addition, techniques such a ploidy conversion or selecting plants for replate based on vigor are not available from a commercial lab. You will be amazed at how many of your crosses germinate when you are in control. When I sent pods to outside labs, I got low germination reports. When I switched to doing my own flask work the percentage went well over 90%.

 I try and sow all my orchid seed "dry pod". This means the pod is mature on the plant, the seed was ready to dehisce. For reliable sowing of dry seed it is important not to get the seed pod wet after it begins to split or pathogenic fungus can grow into the embryos making sterilization impossible. There are times when a seed pods must be taken "green". In the oncidinae, some intergeneric crosses never mature. I understand fertility in Phals can be much higher when pods are harvested immature -- green.

 Should the maternal parent in green pod culture be virused, (sadly, I heard Gary Gallup of Gallup & Stribling say that almost 50% of Phal meristems bought from wholesale growers are virused; hence, Gallup now propagates only from seed) cutting the seed case can spread virus to the uninfected embryos. To prevent this, I take efforts to sanitize the surface of the seed pod and use a red hot scalpel when I cut it into two halves.

 Dry seed is sterilized with 20% by volume, Clorox bleach in tap water. Dry seed is placed in a 16 mm X 150 mm glass test tube. A drop of wetting agent and the Clorox/water mixture is added. About 1 inch is left empty at the top of the test tube and a cotton ball is placed in the top. This is shaken for 5 minutes and then the cotton is pushed to the bottom. The cotton compresses the seed at the bottom while the Clorox/water is poured off. Sterile water is added and the cotton plug is pulled back up toward the top. This action is repeated several times rinses away the Clorox/water. The final time the cotton is pushed down to the bottom it is not pushed all the way. A little water is left so that after the cotton is removed the seed can be poured into a mother bottle.

 (Be sure and bring water reservoirs to a boil before placing in an autoclave. A standard 15 minute autoclave cycle will not heat a large volume of water sufficiently to sterilize it. In addition, replace Clorox bottle monthly after opening. This material is volatile and subject to reduction in strength through dissipation of the chlorine)

 I sow my seed in 1/2 pint wide mouth canning jars. These have a low profile, about 3" so I can stack the several high in my pressure cooker. I sow 2 -3 mother bottled of each cross to reduce the chance of loosing a cross through contamination. If the contamination rate from sowing is 5% per bottle, 2 bottles mothers reduces the chance of loosing a cross from 5% to .25% and 3 mothers even less. For mother media, I use Sigma Vacin & Went media with 20 grams of sugar, 2-3 grams of charcoal and 7 grams of Sigma type E agar, pH adjusted is adjusted with nitric acid to 5.4. (I understand from my Paph friends (people who grow Paphs) that a higher pH, 5.8-6.2 plus in the dark is more effective for Paphs. Cost is around $1.50 per liter. I use about 3/4" of media per mother bottle.

 I sterilize in a pressure cooker. I have two large ones to save time. After flasks are filled with media I wipe the inside top walls near the lids with a towel. I believe removing any splashed agar during filling from these areas lessons the chance of later contamination. Flasks are placed in the pressure cooker and the lid secured. With the pressure cookers vent open, bring it to a boil and allow the media to come up to temperature (5-10 minutes). It is very important to replace all the air in the pressure cooker with steam. Hot air is not effective for sterilization at the normal time/temperature specified. I sterilize with 15 psi of steam for 20 minutes. I have also seen 15 psi used for 15 minutes (15 psi of steam is the equilibrium pressure for water at 121C).

 To prevent the lids of the canning jars from forming a vacuum when cooling, I give each lid a slight twist before capping so they do not lay flat. I don't tighten down the screw bands, I only give them about 1/3 of a turn so they are in place. After autoclaving, I place the pressure cooker under my laminar flow bench to cool. I open the pressure cooker vent when the pressure is about 0-1 psi. After unloading and cooling I give each lid a final twist tight. By waiting for these jars to cool before tightening I prevent a vacuum from forming.

 In answer to questions about using a microwave oven to sterilize I offer a this guess why it doesn't work well. Microwaves heat water by rotating the molecules 180 degrees rapidly. Water molecules are dipoles (opposite charge on each end of the molecule) and this causes them to align to the electromagnetic field. The field reverses 2.3 X 10 8 times per second causing the water to heat by friction. Materials such as plastic and glass are not directly heated by microwaves.

 I suspect microwaves will effectively kill micro-organisms if they are uniformly in the field; however, the distribution of microwaves in a typical oven is poor and there are substantial area where nodes occur with effectively no energy. We have all had to rotate our food to get it to evenly heat. Perhaps someone with more experience can elaborate. Bottom line, microwave ovens don't do a good job at sterilizing flasks. I do use mine to boil water and media prior to autoclaving).

 I store "mothers bottles" on a shelf, ready for use. Because they are unvented, they do not dry out or contaminate with time. When I do open them, they are not under vacuum which keeps them from "sucking" in a volume of air at high velocity upon prying off the lid, lessoning the chance of contamination. I also autoclave erlenmeyer flasks the same way, i.e. the stoppers canted and laid on the top, not sealed. In this position there is no danger of them "popping off" during sterilization. I push the stoppers in place when I unload the pressure cooker and let them cool in the flow bench.

 After sowing mother bottles I place the solid lid back on and screw down the band. I don't vent these containers until after contamination free germination has occurred, 4-6 weeks typically.

 When it is time to vent these I remove the band and metal lid and replace it with a "Suncap". Suncaps are sold by Sigma and are clear, mylar 5" x 5" squares with a 4 mm teflon sub micron filter in the center. There is a technique for doing this. Suncaps are sterilized, about 30 at a time between the pages of 5" x 7" telephone notepads These are wrapped in foil. After autoclaving, I store these under the laminar flow hood and just tear off a page and use the Suncap - voila, they stay sterile this way until use. It is hard to sterilize more than 30 Suncaps per notepad as the paper is a poor thermal conductor and if it is too thick, 15-20 minutes in a pressure cooker won't drive the heat into the center of this mass. I also use fresh steel bands on the suncap. I have two sets of bands. Those that I use for sterilizing and those I use for "Suncapping". The ones that go through the pressure cooker get rusty and the high friction from the rust tends to warp and twist the Suncap. A steel band that isn't rusty works much better. Jars and the bands are recycled.

 I add these insights about filters. Suncaps are made of a stretched teflon membrane similar to Gore-Tex. The pore size is sub micron, hence bacteria and pathogenic fungus cannot enter. This prevents contamination. The Suncap filter membrane is very thin and has a very high pore density thus suncaps tend to allow a lot of evaporation. It is almost like having a 4 mm hole in the top of a flask; hence, one has to watch flasks for excessive dehydration. I have begun blocking off the open area with a small round paper label to minimize this problem. An alternative is micro porous tape used for bandgaging wounds.

 I use cotton in rubber stoppers on erlenmeyer's and here the pore size is actually rather large; however, the path length is very long and evaporation is much lower than with Suncaps. One can refer to the suncap as a membrene filter while cotton is a "tortuous path" filter. Both work well. Diffusion, temperature fluctuations and changes in barometric pressure assure 1 or 2 percent of the flask volume is exchanged daily. I might add a caveat, if you use rubber stoppers and cotton, unwashed cotton (available from an upholstery supply) reduces the chance of contamination. Unwashed cotton contains oils that repel water. Wet cotton can grow pathogens through it. You may also want to put a drop of CuSO4 in water (saturated) or some picric acid in water (a few percent) on the cotton. This acts as a biocide and prevents pathogens from growing through the cotton. (Yes, I know picric acid is used in making hand grenades; however, I have never had a rubber stopper explode and picric acid a great biocide.)

 When germinated embryos develop into small balls 2-3 mm in diameter and begin the show the emergence of a tip (called the leaf primordial) it is time to spread them onto replate media. I use several replate medias. Gallup & Stribling distributes Hill's Replate Media with Banana:

Gallup & Stibling

645 Stoddard Ln.

Santa Barbara, CA 93108

1-805-969-5991

 It is popular around the world because it is great media. It is also fairly expensive. A good alternative is a media worked out by Terry Root of the Orchid Zone, my variation follows:

Sigma Phytomax Maintenance Media, M6668 (contains sugar and charcoal)

7 grams of type E agar

1 jar of banana baby food (113 grams or 4 oz.)

pH adjusted to 5.4 (higher for Paphs?)

For salt sensitive plants such as Masdevallia:

Vacin and Went

7 grams of type E agar

1 jar of banana baby food (113 grams or 4 oz.)

pH adjusted to 5.4

 I move to 1 pint narrow mouth mason jars for spreads and final replates. I recommend the protocorms be spread sparsely for the 1st spread. It is likely if this is done correctly final replates can be done from the 1st. spread. A spread will take several months before it is ready for replating. This final replate will take a few additional months before plants are ready to come out of the bottle. Protocorms can be kept in the mothers for a fairly long period of time so one can time the replate process so flasks are ready to plant out in the spring (avoid taking flasks out in the winter).

 The procedure for spreads and replates is fairly simple. I use a very small replate hook, a 1/8" stainless rod, about 12" long, with a small bent hook on one end. I fashioned this with jewelers saw and some files and heat. I do not use tweezers; however, some flaskers do (they cause cramps in my hands). The "hook" is easy to sterilize in a flame and cools quickly. I use Suncaps on spreads and replates.

 Growing takes place on Metro wire shelves under fluorescent lights. The shelves are 2'x 4' with two single lamps fixtures per shelf. The ballast are electronic to prevent local heating and the cheapest, cool-white bulbs (cheap ones) are used. Light levels are ~ 300 ft. candles. The lights are run for 8 hours per day so as not to disturb my neighbors or arouse the nepherious instincts of kids in the neighborhood. I try and keep the flask room under 75F during the day and over 60F at night. Excessive day temperatures causes problems with proliferation in the flask. (I grow cool orchids so I do not have experience with warmer growers.) I do know that the flask rooms I have visited at commercial growers are always comfortable.

Hope the above information proves useful.

 Bob Hamilton - Dec 94