As I understand it, professional orchid growers propogate orchids in sterile labs, and even then, it may not work, even if a particular plant is usually easy to propogate. Big plug for Meyers Conservatory, who's been among the best growers to deal with - for me, anyway. From his site: Meyers Conservatory's mission is to provide laboratory-grown orchid species at the lowest possible price in an effort to aid in orchid species conservation. Since we started in 1999, we've never operated as a for-profit business, instead charging just enough to cover the costs of the flasks we produce and to pay our employees. https://lab.troymeyers.com/flasking/donate-funds.php They accept donations. From the flasking page: We usually charge the same amount for each flask, regardless of its rarity. Rare orchids or particularly nice specimens usually command a higher price, but unless there are difficulties growing a species we don't charge more; we will charge more if propagation requires extra time or materials, but not just because it's desirable. By not charging a lot for something rare, we can rest easy knowing that we're not helping to perpetuate the high prices that make collection of wild endangered orchids profitable. __________ I check the site every couple of weeks. Troy doesn't have a large selection, but he's had orchids that I couldn't find anywhere else in the U.S. WC8 Here is a link that might be useful: Troy Meyers' flasking page

I found this on the web via the orchid forum, I know that it is a bit involved but orchid seed by its nature (no endosperm) is like that! Home Orchid Flasking This material may be copied for personal use, or for use by societies or research groups, providing that it is accompanied by a notice acknowledging authorship. Cedric Maunder This guide is based on myown experiences here in the UK, and the methods and materials described may not suit other people. Over a period of about two years I have had good results with a variety of orchid species and hybrids, and have suffered very little contamination of flasks. These notes are provided not because I consider myself an expert, far from it but because this comparatively cheap and simple set-up has worked well for me and might be helpful to anyone contemplating having a go at flasking. I am very grateful for help and advice received from other people - notably George Black of Brize Norton, England and Robert M. Hamilton of Berkeley, California, USA A very important factor is to be very aware that bacteria, moulds and other potential "nasties" are everywhere - in huge numbers. Awareness of this fact, together with recognition of the existence of gravity and air currents, will help to ensure that the said nasties do not gain access to your flasks. I should say at this stage that I do not possess a laminar-flow cabinet. The initial purchases needed for starting out in flasking are sufficiently expensive without the fairly large outlay that such a cabinet requires. Having seen the results obtained by George Black - a noted orchid hybridiser and flasker - using only a very simple glove box, I decided to try that method first. After a couple of years of this I see no reason to change to anything more complex and expensive unless I become sufficiently fool-hardy to consider commercial ventures! GLOVE BOX: This is a very simple device. Various people use cardboard boxes, glass carboys etc.! I thought that wood or plastic (acrylic) would probably be the best choices for me. (A simple wooden box can have a piece of glass for a lid and does not necessarily need a frame or hinges for the glass top. George Black has raised many, many seedlings from such a set-up.) I finally settled for using acrylic sheet; most pieces of the box can be made from off-cuts which are very cheap, though I used thicker sheets for the base and lid - these being at full price. The only drawback of this material is the adhesive it requires. This should be used out of doors or in a very well ventilated place. The material is transparent if one does not spill too much of the adhesive on the visually critical parts - lid and front panel. The one I made has the following dimensions (not very critical, I wanted it to fit on the end of the kitchen table when in use and on the top of the freezer for storage) Width - 24 inches; 810mm. Depth - 18 inches; 457mm. Height at rear - 12 inches; 305mm. --- 14 inches; 356mm. Might be better. Height at front - 8 inches; 203mm. --- 10 inches; 263mm. Might be better. The lid is made of two pieces, a rear fixed part and a hinged part at the front. I also used a fair number of small off-cuts to strengthen the corners and rear part of the lid. Diameter of arm holes :- 4.25 inches; 108 mm. (I used a saucer as a pattern!) I used pieces of shower curtain material cut to shape and size to make sleeves. A hem is made at the inside end with elastic inserted and tightened to give a comfortable fit for the wrist, and cuts made at the other end to make radial flaps with which the sleeve can be glued to the outside rim of the box. If one wears rubber gloves when using the box, the sleeves and gloves keep one's skin fairly free of Domestos. (See below ) STERILIZING THE INTERIOR OF THE GLOVE BOX: This is done using Domestos or similar bleach diluted 1:4 with water. A 500ml spray bottle contains this solution so that It can be sprayed across all parts of the box to make a mist which will remove all organisms. (If the spraying is done correctly, all interior surfaces will have been coated with the bleach and organisms in the air will be carried down in the spray into a layer of bleach on the floor of the box. The sleeves and gloves must also be well sprayed, preferably at both the start and end of the sterilizing. My experience is that by the time this progress is completed, the box is essentially sterile. Wait a further five minutes or so to be absolutely sure. All materials, tools etc. needed for the flasking session must be put into the box before the spraying begins. Tools should be laid in a shallow dish full of the bleach solution before closing the lid. All flasks or bottles that are going to be worked with before starting, together with a flask of sterilised water, capped with foil. All must be thoroughly sprayed during the sterilising procedure. If some essential item is forgotten it will be necessary to do the whole procedure over again, SO have a check list and consult it before sterilising! BASIC REQUIREMENTS FOR FLASKING: Access to Sigma/Aldrich or some other reliable supplier of Chemicals and apparatus. [In the UK this firm is probably the largest and most comprehensive one engaged in this field, and I have found them to very efficient and helpful. Free catalogues - very well produced.] These firms can provide. almost everything needed, and thousands of other things, too! UK Addresses and Phone Numbers:- Telephone:- Orders :- 0800 373731 Customer Service :- 0800447788 The company appears to be active in most other parts of the world, but I'm sure there are alternatives in most places. Items Required:- 1. Chemicals: (Approx UK prices shown (1998) - subject to change) Medium:- Phytamax Orchid Maintenance Medium with charcoal and banana, P1056. 1 Litre size = £1.60 - Phytamax P0931 is without charcoal. Agar:- A 1296, 500g - £47.20; 100g. -£14.30 Calcium hypochlorite - Fluka 21182, 250g @ £5.10; lkg = £14.30 (This is 65% to 70% strength, 3.5g per 100ml for five minutes is probably sufficient for seed sterilisation). Suncaps (flask closures with filter vent) C 6795- foil with 3.00 mm.. filter; 500 £ 43.30 Domestic bleach - Domestos or similar, hand sprayer to use for spraying Domestos 1 :4 solution in glove box. 2. Hardware: Can all be obtained from Sigma but most can be obtained from other sources. Measuring jug, 500 ml., pyrex or similar. - kitchen departments of department stores. Glass funnel for filtering. Filter paper - Whatman 185mm. Grade 1 or 4 (faster) A few 5OOml glass flasks, with rubber bungs, are useful for making and filtering the hypochlorite solution, water, and so on. Large bag or elastic bands for temporary securing of Suncaps. Micropore tape for final securing of Suncaps. Shallow oblong dishes for holding flasking tools (see below) for sterilising (inside glove box) Supply of suitable bottles - jam jars, honey jars, baby food bottles (Sigma) etc. Labels for bottles. Small self adhesive ones made for labelling preserves/jams etc. Small stoppered bottle to hold seed during sterilisation. Pressure cooker high enough to hold 500mL flasks. Scales to read down to 0.05 gm. (0.1gm. divisions). (For making up the majority of solutions needed for flasking, colchicine use CSMColchicine.htm chromosome counting, etc. this should be adequate. To make up one's own medium really requires 0.01 gm. divisions. ) A microwave cooker can be used instead of a pressure cooker but, on balance, the pressure cooker is quicker and more convenient. For good results from sterilising in a microwave oven it is necessary to include about one litre of water in bottles with each batch of flasks. This acts as a thermal ballast so that enough time can be given to ensure sterility without boiling over of flasks or loss of their seals. Reference:- Microwave Sterilization of Plant Tissue Culture Media; BrentTisserat, Danny Jones, and Paul D. Galleta. Hortscience 27 (4): 358 - 361 1992.) MAKING UP THE MEDIUM: Sigma medium is clearly labelled with the weight required to make one litre. I usually make batches of 500mL. This is sufficient for ten to a dozen bottles. I usually prepare a few bottles with a deeper layer of medium for use when re-plating, others more shallow for seed sowing. Do not be tempted to make the layer of medium very thin, however, as it will soon become too dry and stiff! CABINET FOR FLASKS: Fluorescent tubes 4 foot - daylight, one or two ( at the most) for each shelf. Doesn't seem to be any marked advantage using UV lights at this stage. Plastic trays to place flasks on -- 9 inches from lights gives good results. Some cool growing genera are prone to produce protocorms with little or no chlorophyll if light to heat ratio is too low. Bringing flasks to lower (cooler) level results in some or all of these 'greening up' Time clock to give 12 or 14 hour days. This timing seems to be optimum for Oncidiinae genera. SEED SOWING: This can be ripe/dry seed or 'green pod' - (unripe capsule! ) DRY SEED: Keep seed as clean and dry as possible before use. Some people say to dry the seed in a dessicator - I haven't found this to be a good thing. Sow as soon after harvest as possible - but will keep for several weeks if dry. Presumably gradually loses viability. I use a 50 ml. bottle or flask with screw cap or tight fitting bung. Make a 'solution' of Calcium Hypochlorite (60-70 % available chlorine) using 7.0 or 3.5 gm. of Hypochlorite in 100ml water . STRENGTH OF CALCIUM HYPOCHLORITE TO USE: (Traditionally 7 gm. of calcium Hypochlorite in 100 ml. of water has been used. Recent articles by Fred Bergman suggest that this is too strong. I've recently tried batches of the same seed at 7gm., 3.5 gm., and 1.75 gm., in 100 ml. All have germinated - better germination in weaker bleach, one batch at 1.75/100ml. showed contamination. All batches at 3.5gm/100ml. are O.K. I'll use 3.5 gm. routinely now, continuing to try out some at 1.75 gm/100 ml.) Filter into a flask. Place seed in the 50ml bottle/tube and add about 15ml Hypochlorite and ONE drop of kitchen detergent (Squeezie). Shake well and after 5 minutes filter the contents (Cover the funnel top with clean sheet of filter paper.) Rinse twice with STERILE water- keeping the funnel and the open bottle of water covered with filter paper during transfer to the funnel. The covered funnel is then transferred to the glove box. Use a spatula (sterilised in Domestos 1 part,. water 4 parts) to transfer seed to flasks. After everything has been sterilised by spraying as described above, carefully remove the foil top from one of the bottles of medium, placing the top, top surface up, on the floor of the box. The bleach solution will prevent contamination of the inside of the foil top and it can be replaced on the bottle after sowing. The foil will have turned down edges and corners from being secured to the bottle during and since sterilising. It will sit on these turned down corners so that the inside of this lid will remain clean and dry (in most cases! ) . GREEN POD: Capsules sometimes abort before they are ripe. These can often be salvaged. Also, some seeds will not germinate after being sterilised but will germinate if sown from green pod. Soak and brush the capsule in 1 :4 Domestos solution. Give good 10 minutes in the bleach. Make sure that a suitable scalpel blade or small knife is in the glove box before starting! It is best to put the capsule, spatula and blade together into a dish or bleach inside the box, together with flasks, bottles etc. needed for sowing, before spraying as above. Open the capsule inside sterile glove box (after sterilising as described above), and scrape seeds with the end of the spatula into flasks. Check your flasks/bottles every week or so. Germination often occurs quite quickly so that tiny green protocorms can be seen on top of the medium, Some seed takes a long time to germinate, so do not be too hasty to throw out seed that does this, I have heard of seed germinating a year or more after being sown! Any contamination will also usually be apparent in the first few weeks. It is wise to sow at least two bottles of each lot of seed so that throwing out the contents of a contaminated bottle does not negate the whole sowing. If a very precious batch of seed does become contaminated.. you could try treating this with 1:4 Domestos. IN THE STERILE BOX open the flask, apply the bleach to the patches of contaminant and an area around them. Reseal the flask and hope. If caught early and the contamination is slight, this does work, but not always. REPLATING: Replating is usually beneficial Even with charcoal in the medium waste products will accumulate. Also the medium dries out due to air exchange through the filter and uptake of water by the seedlings. This is an opportunity for picking out the most vigorous seedlings and giving them more room I replate whenever seedlings seem to have slowed their progress, grown too large for the bottle, or if the medium seems to have become too stiff. I also tend to replate bought-in flasks unless they are growing very well. I use the same type of medium for sowing and replating. If you are sowing or replating Paphiopedilums, Phragmipediums or terrestrial orchids you will need to modify these methods, very likely needing a different medium. I have no personal experience of these types of orchid. You might be able to obtain advice from Kew regarding terrestrial genera. DE-FLASKING: Doing one's own flasking allows one to de-flask at optimum times and also allows replating of any smaller backward seedlings. Thus the yield is usually much higher. The process is essentially simple. Have a large bowl of clean tepid water to hand. Remove the foil cap from the bottle or flask and use a wire hook to gently ease the seedlings out into the bowl. If you use flasks rather than bottles, this can be difficult as the seedlings jam in the narrow neck. For this reason I much prefer bottles. It helps to fill the flask with tepid water and allow this to melt the agar. It is usually then possible to use the wire hook to tease out one or two seedlings initially, followed gradually by more until the flask is empty. I have no doubt that de-flasking from narrow necked containers causes more damage to the seedlings, If you have very precious seedlings in such a flask, it may be worthwhile carefully breaking the flask before attempting removal. AFTER-CARE OF THE SEEDLINGS: We plant seedlings out into community pots rather than individual containers. We plant them in perlite as per the instructions in these pages. After planting the seedling, ensure the compost is moist, cover the container with a bell-jar type plastic lid, Or a plastic bag. Ensure that the seedlings do not become too dry. During the dark English winter, at least, we keep community pots under lights for some (It is useful to have some spare shelf space in the flask cabinet for this purpose. We find that this suits most seedlings very well.) PHYSAN OR SIMILAR FUNGICIDES DURING DE-FLASKING: Unless there are signs of contamination this does not seem to be needed . FLASKING TOOLS: I use two types of tool, both home made. A fine Spatula. A fine fork. Both are made from eighth or quarter inch brass rod (wire,) from a toy/model shop. Very cheap and very easy to work. For the spatula, a 12 inch length of quarter inch rod has one inch of one end flattened by hammering with a light hammer - very easy - don't hit too hard! The flattened area is then gently bent to a suitable curve with pliers. For the fork, use either quarter inch or eighth inch rod - depending on size fork required Use a fret-saw with fine blade to make a cut centrally in one end of the rod for a depth of half an inch, Pliers and a fine metal file are used to make points, separate the ends and produce a suitable curve. It is useful to have two sizes or fork, one for small seedlings, the other for the large ones. Stainless steel is mentioned by most authors, but I have had no problems with brass. It stands up to bleach solutions well, and does not appear to have any bad effect on seedlings. It is cheaper and easier to obtain as well as being much easier to work. ©Cedric Maunder.

Question 1 - I was really blown away by how deep and big the taproot is growing on these trees. It's like most of the energy is going to create this big root. It's thicker than the trunks in every case, and the roots are easily twice to sometimes 3x as long as the seedling itself. So on on of them, I cut back the taproot a little bit... I took about a half an inch. I had read Al mentioning that slowly cutting back a taproot over time can help spur the tree to grow more roots higher up. I hope I didn't take it out of context. The first day the lowest leaf of that seedling, which has started to go brown on the rim, became pale and collapsed on one side of the leaf. Then yesterday it went dry. Today there has been no further progress of the leaf's collapse and no further signs of stress. The seedling seems strong... Was I wrong to try and trim it back just a little bit? Or did I do alright? Is this seedling out of the woods for now do you think? Young plants are nearly all dynamic mass, and as such, have a very strong "will to live". If your plant is currently doing well, there is no reason it will take a turn unless something cultural forces a change in course. Bonsai practitioners have learned that lifting seedlings as soon as the first set of true leaves emerge and removing the seed radicle immediately below the root to shoot transition zone and treating the top as a cutting eliminated taproot issues entirely in most plants. In stubborn plants with a hard tendency toward vertical roots, you'll need to regularly remove all first order roots (attached to main stem) growing downward from the root to shoot transition zone as well as all second order roots growing up or down off the first order roots. Question 2 - related to question 1, regarding the other seedling I root pruned a little. This one I took a little more off of because of root rot. I know I had to take it off, and I also took off a bit of taproot for this one too. The seedling itself seems fine. The uppermost leaf - which is fairly new and thin yet - went limp the first day, but has become stronger and greener these last two days. I know I had to take off the bad roots, but I was worried I went too far. I'm hoping I'm seeing indications that I didn't? Only way to tell is by putting a waiting period behind you during which the plant retains its turgidity throughout the daylight hours. Stomata close at night and slow water loss, so unhealthy plants have opportunity to recover from low internal water pressure and wilting. No wilt during the day is a good indication to bet on full recovery. Question 3 - When I was done, I investigated the 5 'dead' seedlings. Two of them looked like some of Al's bare-root pictures. The main part of the taproot was healthy, seemed alive, and had small alive roots still growing from it. but the end of the root was truncated in mush, much of which fell off into the soil. I washed them as clean as I could in a mist spray and cut off the mushy end to clean root. Then I potted those in 5:1:1 too. I'm hoping against hope that I might be able to save them. The top id dead back to the twiggy stump... Any thoughts if they might come back? Anything special I might do for them to give them the best shot - for instance a special fertilizer or watering strategy? Withhold fertilizer until the plant recovers. DAMP or MOIST are the operative words. If the fungal infection has gummed up the plumbing such that a vascular connection between roots and shoots isn't in the offing, it's a goner. If the infection isn't systemic, the possibility of retaining viability is still on the table. What kills cuttings is the inability to make that plumbing connection due to rot, or the cutting having an inadequate energy reserve. If the plant/cutting/seedling runs out of reserve energy while it still a net user of energy instead of a net producer of energy, it's over. 4: I have 5 I have not yet repotted. They seem to be okay... though 3 aren't doing much of anything, they're just existing and may have some first sign of browning on lowest leaves, but I might be looking too hard too... One is finishing it's fall flush of leaves, and one is start starting it's fall flush... I'm not sure if I should repot them so close to winter or if I should let them be until spring, since they seem okay right now. I especially am not sure I should mess with the one putting out new leaves. But on the other hand, what if I really am seeing some warning signs and some suddenly start taking a down turn in the next week or month? Wouldn't it be better to do it early, now, rather than later? The only reason I re-potted the ones I did was because I was sure they wouldn't make it through winter... I am not sure on these ones. If they were your plants, what would you do? A dearth of mobile nutrients, over-watering, root congestion, soil compaction, can all lead to loss of lower leaves. I'll ask you to spare me from having to do the research by telling me if these plants would normally enter a predictive dormancy during the fall to winter transition? If yes, then waiot until spring but guard against over-watering. If no, repot only if you suspect strongly that the plants will no longer be viable come late spring (June). 5: Fungus gnats. =( I manage to get rid of them for a while, but they always come back. Of course, I was always using compost mixes, particularly organic ones, and they're usually lousy with them no matter how much a brand claims they pasteurize/sterilize for pests. Do non-mineral soil media mixes have the same ability to attract and support these and other soil-borne pests? If yes, which ones to watch for? If you don't already have a good understanding of what constitutes the comparative degrees of container media (as in unusable, practically unusable, just plain poor, ok, better, and the variations of better from 'sort of better' to 'really a whole lot better'), making that a priority will almost certainly propel your advancement farther/faster than anything else you might apply yourself to in the near term. If you need the info that can tie soils up into a neat little conceptualization, I'll be really glad to help, but I think trying to do it in a thread that covers so many topics isn't the best format. Are you familiar with and understand the concept discussed here? Which brings my follow-up: If it can support pests that spent part or all of their life cycle in soil, then it should be able to support their predators too, right? I prefer to combat pests with their predators, like nematodes, predatory mites, aphid lions, etc. Would this be doable if needed? How about we work toward providing a home for roots that doesn't provide creature comforts? In the immediate, mosquito dunks and a restrained hand on the watering can are potential remedies. 6: For the future of these little babies of mine, what would a good strategy be for keeping them happy growing in a container? You'll find an outline here that will provide a plan for maintaining woody material in containers and in good health indefinitely. I know I need to keep them smaller than if they were in the landscape, and that means eventually removing the taproot. I also understand that taproot removal usually starts after a year. But these trees don't actually seem to have much besides taproot... smaller roots branch off of it, and a few have some smaller roots that come from the trunk matrix but not most from what I've seen. Would the best bet be to gradually prune the tap root back over a few years in an attempt to get the trees to put out more roots higher up? Or should I go for broke and just cut it way back and defoliate? Or is it possible that the poor media I have been growing in so far as not encouraged growth at the higher and mid root range and drove the roots downwards looking for air? I'm sorry I haven't taken any pictures to share to give you a better idea. If I end up re-potting any of the others, I will take some. I think I covered these questions above, but if not, ask again. Al

I'd love to try growing one from seed to test its hardiness in my zone. I have Texas betony, rue, and four nerve daisy seeds I could trade for it.