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From bob@argon.berkeley.edu Mon Jul 12 21:28:29 1993
Subject: A brief description of how I flask...

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 microwaving and definitely 
not terse! "Netters" not interested in flasking may want to type "delete" 
at this point.

I have been doing my own "bottles" for about 6 years. Most of what I
have learned has been through imperical 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 disastorous 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 dihesce. 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
intergenric crosses never mature. I understand fertility in Phals can be
much higher when pods are harvested immature -- green.

Should the maternal parent in greenpod culture be virused, (sadly, I heard Gary
Gallup of Gallup & Stribling say that almost 50% of Phal merristems bought 
from wholesale growers are virused; hence, Gallup now propogates 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, Chlorox 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 Chlorox/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 pusshed to the bottom.
The cotton compresses the seed at the bottom while the Chlorox/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
Cholorx/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
 sufficienty to sterilize it. In addition, replace Chlorox 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 lossing 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 meida 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 equalibrium 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 
(oposite charge on each end of the molecule) and this causes them to align 
to the electromagnetic field. The fieled 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 microrganisms if they are uniformely 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 erlynmeyer 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 occurreed, 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 submicron 
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 submicron, 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 microporous tape used for bandgaging
wounds.

I use cotton in rubber stoppers on erlymeyers 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, un-washed 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 primodia) 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.

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