Dionaea muscipula Micropropagation Manual by Gary Gipson

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Dionaea muscipula
Micropropagation Manual

by GARY GIPSON

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TABLE OF CONTENTS

FORWARD
I. PREPARING THE MEDIA
Media Preparation Technique, Sterilization and Storage.
Stage 1: Media Recipe for Seed, Leaf and Flower Stalk Initiation
Stage 2: Media Recipe for Multiplication
Stage 3: Media Recipe for Growing
Gelling agents: Agar and Carageenans
Adjusting PH and Care of Your Electronic Meter
II. EXPLANT SELECTION
Seed
leaf
flower stalk
III. CLEAN ROOM
Equipment Sterilization
Clean room Techniques
IV. STERILIZATION OF EXPLANT
Common accepted method using Sodium Hypo-chlorate
Alternative method
Seeds
Leafs
Flower Stalks
V. CARE OF CULTURES; IN VITRO TISSUE REQUIREMENTS
STAGE 1: Initiation
Seeds
Leaf
Flower Stalk
STAGE 2: Multiplication
Seeds
Leaf
Flower Stalk
STAGE 3: Growth
STAGE 4: Ex-vitro Planting and Hardening
VI CONTAMINATION
Types and Treatment

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Forward
This Manual will never be finished. Micro-propagation; even with the focused study of a specific Species of Plant, is an ongoing process of experimentation with absolute perfection never achieved. All One can hope for is a compatible or acceptable environment in which to propagate a particular plant species in a temporary environment at accelerated growth in which makes it beneficial to do so in the first place.

Therefore, the purpose of this manual is to be a ‘go to’ source of methods that have been proven or has been evident to be at least successful to a certain degree for the Plant species Dionaea Muscipula.

This manual is geared for Wholesale and Retail environment of Dionaea Muscipula, therefore the methods described are for maximizing productivity for the least amount of time and therefore cost.

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Media Preparation Technique
I have only used distilled, RO, or De-ionized Water, as I wanted uniformity. Purity of the water also affects clarity of final product. Clarity is important for early detection of infections BEFORE One multiplys it out over many vessels. For better dissolving properties, add all ingredients (Basal Salts, Vitamins, PGR‘s, etc.) into the distilled water before adding the Sugar.

Add the Gelcarin (Carageenan GP-812 or Hi Clarity) Stir until all lumps are gone in the media. It is important that the media is cool to room temperature for the Gelcarin to dissolve quickly. If the media is even slightly warm, it considerably adds to the time it takes to get rid of the Gelcarin lumps when stirring.

Test the PH with ALL the ingredients added, including the Gelcarin.
High clarity Carageenan requires a pre-autoclave PH of 4.4 to acheive a post autoclave PH of 5.5/6.
Test with an electric Meter only and adjust PH down with White Vinegar, Citric Acid (better as it is an antioxidant), or HCL. Raise PH with Sodium Bicarbonate. A little goes a long way!

Dispense into vessels, stirring the media every now and then. I currently use a magnetic stir plate. Constant stirring prevents gelling agent settling which can cause inconsistant gel firmness from vessel to vessel.

Autoclave at 15psi for 15 minutes for 25 to 30ml. Media vessel content. Add a couple of minutes for 40ml or higher media containing vessels.

Store in baggies or tubs until use; store in refrigerator if sensitive PGR’S are used to extend their life. Storing in Laminar hood is best, but needs to be used as soon as possible as the water slowly evaporates, hardening the media.

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MEDIA RECIPE for Dionaea Muscipula
Stage 1: Seed, leaf, and flower stalk initiation

1,000 ml distilled water
1/3 suggested label strength ms basal salts (without vitamins)
1 ml ms vitamins 1000x stock
.50 ml kinetin (1mg/1ml) or 1.0ml bap/ .1ml naa combo.
1 to 1.5 ml ppm (optional)
30 grams sugar
7.2 grams gelcarin gp812 carageenan or high clarity carageenan (hi clarity carageenan is crystal clear).
Adjust ph to 4.6 for gp812 ( 4.4 for high clarity carageenan) with citric acid, or hcl.
(electric meter only) this will make the final ph after autoclaving to be around 5.6 as prior tests have shown.
Dispense.
Autoclave for 18 minutes at 15 psi. Cool in laminar flowhood.

Stage 2: Multiplication
Make 1/3 MS with Full Vitamin and full Sugar (30g/Liter).
1-1 liquid strength = 1ml/L BAP, .1ml/Liter NAA
It is said another excellent Multiplication media is to use Phytotech Labs Dionaea/ Drosera multiplication media. I am currently experimenting with it.
Kinetin is a proven Cytokinin for Dionaea for multiplication, however BAP in My early tests gives better results in individual corms and doesn't have a tendancy to form a tumerous like solid callous.
After experimenting with varying concentrations of Kinetin in multiplication stocks, it is evident that Kinetin is much more powerful than I gave it credit for. I also underestimated the natural inclination of Dionaea to multiply on it’s own even without any Cytokinins. I have gone full circle from .5ml/Liter to 2.5ml/L and finally back to .5ml/Liter.
The reason for finally ‘settling’ on the .5ml/Liter on multiplication stocks is that in Dionaea propagation, it is not necessary to have large amounts of Kinetin to get the desired multiplication rate necessary even for large production. My ex-plants on higher amounts of Kinetin continued to multiply for several platings afterward on the grow rack, and I was turning one Ga-7 Grow Box re-plate to 4 additional vessels with virtually no maturation or significant rooting (vital to growth) evident. This multiplied out to hundreds of vessels with virtually nothing to de-flask. If I continued on this trend, I would eventually have tens of thousands of Plants coming out of vitro at One time, not to mention losses from contamination inevitable from the multiple platings and labor and materials involved.
I am currently experimenting with 1mL BAP and .1mL NAA.

Stage 3: Growing and/or Acclimation for Vivo
Makes 1/3 MS w/ full Vitamin
1,000 ML DISTILLED WATER
1/3 MS BASAL SALTS
1.00 ML MS VITAMINS 1000X STOCK
1 ML PPM (optional)
20 GRAMS SUGAR ( 30 Grams if just Growing to increase size)
7.3 GRAMS CARAGEENAN
Alternatively, use PHYTOTECH LABS VFT/DROSERA Pre-transplant media.

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Agar
The staple and proven Gelling media for use on Dionaea ex-plants Use 6-7 Grams per Liter.
Tends to lower PH some depending on purity. Micro-propagation Grade Agar possess’ better clarity and doesn’t affect the PH as much.
Agar is better for shipping in vitro plants.

Gelcarin
Gelcarin, specifically Carageenan GP-812 is used with great success by many Culturists. The clarity (that can vary from batch to batch) is as good or slightly better than Micro-propagational grade Agar, and can have a slight golden hue. Clarity is important for catching phenols, contamination, etc.
Another advantage is that it is partially water soluble, so clean up is easy. It’s recommended usage is between 6 and 8 Grams per Liter in strength. I’m finding that 7-7.5 Grams/Liter is a good happy medium concentration for baby food jars. 7.5 -8 Grams is more suitable for larger volumes of media in larger vessels.
Gelcarin GP812 does not readily clog the ph meter, at least the glass bulb type, as it is water soluble (partially), and the electronic meter could be used to test ph with all ingredients added including the Gelcarin before boiling. I was told by Gary of Phytotech labs that the industry standard is to test and adjust the PH of the media with ALL the ingredients added including the Agar or Gelcarin, but before boiling.
Phytotech introduced a new ‘High Clarity’ Carageenan and has proven to be ultra clear and mixes much easier.
Both Carageenans (GP812 and Hi-clarity) raises the PH of the autoclaved media much Higher. This needs to be compensated for. I have found adjusting ½ MS media to 4.6 for GP812 and 4.4 for the new Hi-Clarity Carageenan before autoclaving will yield the recommended PH of 5.5 to 5.6 after autoclaving.

PH of the Media
PH is important to the health of explants, but to what degree is debatable and depends on what is being cultivated. The proper PH for Dionaea media seems best at 5.6 more or less. I talked to Gary at Phytotech Labs and His take on PH and its importance echoed what a lot of experienced TC’rs that I’ve talked to have said. It can be important, but there tends to be an overemphasis on the importance of getting the PH just right. Don’t over think it because a culture won’t crash if it isn’t spot on, at least with most plants, anyway. The PH drops over time, anyway, as the explant extracts nutrients from the media, yet the explants are thriving?

It is worth mentioning that more important than the accuracy of the PH to a specific target range, is the CONSISTANCY of the PH from batch to batch for the sake of not stressing the cultures that have acclimated to a particular PH in the media.

The only real opportunity to adjust the PH in the media is in the preparation of the media. The standard accepted protocol for adjusting the PH in media in the TC industry is such that One has ALL the ingredients in the media, including the Gelling agent such as Agar or Gelcarin, but have not boiled the media. Adjust the PH with an electronic meter at room temperature and You are done! Now You can boil, dispense, and sterilize.

Gelcarin and Agar are polysaccharides and won’t clog an electronic meter. When finished, rinse probe under warm tap water, and store moist in it’s solution, if applicable. Gelatin will clog a meter as will any other protein based compound.

PH meters will clog over time even in the best of environments. The Supplier may tell You to replace the probe/meter, but all it really needs in most cases is just a cleaning. Meters should be cleaned every 6 months or so, especially if You are noticing ‘drift,’ meaning it is taking several minutes instead of 2 minutes to get a solid reading on the meter. You can buy cleaning solution for soaking, but I was also told that You can simply clean the probe in a jewelers ultrasonic cleaner (You can buy a cheap one at Wal-Mart) in plain water for a minute or two and let the cavitation explode off the particles in the porous glass probe bulb. If You have a powerful industry ultrasonic, care must be given not to break the glass bulb.


PH STRIPS for Testing Prepared Media
The practice of checking PH with PH strips are useless in the testing of the prepared media as the PH strips were not designed for the high content of the various Salts that We have in the media. If You have a plain glass of tap water that is a PH of say, 5.5 and prepared media that is 5.5, drop a PH strip in both and they will be two completely different colors. It is impossible to get an accurate reading in Our application. PH strips also change color under different light. Check the strip under fluorescent lighting and under an incandescent and it is readily apparent. The color code reference chart won’t change color much, but the wet strip will change color dramatically, from muddy brown to bright apple green.

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CLEAN ROOM

Equipment Sterilization


Autoclaved vessels, instruments, paper towels, etc. are best cooled and even stored in the Laminar flow hood or the like with a front cover intact. A running flow hood tends to dry out the media quicker, however.

Whenever possible, One should deep sterilize the utensils in an autoclave. Periodically, even the closed laminar flow hood should be wiped down with 70% alcohol and a sterile cloth or paper towel. Start from the top and work down. Do this with the fan running.

Clean Room Techniques

Prior to starting working under the laminar flow hood, ensure All air conditioning vents and any other source of moving air in the room other than the laminar flow hood should be shut or shut off. All doors, windows, etc. should be shut as well. You need 10 minutes or so for all air movement to still and contaminants to drop. Turn on the UV lamp if so equipped and leave that on for 15 minutes while You wait for the air to settle. Leave the room or avoid exposure to the UV lamp.

Next turn on the laminar flow hood fan and let it run about 10 to 15 minutes prior to working. Turn on the bacteriocinator or glass bead sterilizer at this time.

While in the hood area, avoid opening and closing doors, cabinet doors, etc. rapidly. Keep Your own movement slow and deliberate as well. The 100 to 110 fpm airflow out of the unit can be easily overwhelmed by forces greater introducing contaminants into the unit.

Wear short sleeved shirts or lab coat. Wash hands and forearms up to the elbows. Alcohol hands and arms and put latex or similar gloves on. Hairnet should also be worn. If You suspect You will have to communicate while under the hood, then wear a mask as well. Put on a clean, short sleeved lab coat, rubber gloves, and hair net. Spray gloved hands and forearms with 70% alcohol. Anything introduced into the hood should be sprayed with alcohol or bleach solution.

Vessels with plantlets taken off of the shelves for re-plating should be sprayed with bleach solution or at least alcohol. I currently use 70% alcohol and spray vessel liberally.

Bleach solution (1 part 6% Clorox to 10 parts water, and 2 drops Tween 20 per 100ml) should be made fresh each time or used within 24 hours or so. I currently use heat for instrument sterilization. It is best.
 
Explant Selection

Do not touch the tissue to harvest with Your bare hands. Wear gloves or use tweezers. Have a glass of water handy to keep the tissue hydrated, and always keep the cutting wet. Cut off as much of the tissue as possible to ensure You have enough to sacrifice when the final cutting is done for vitro.


Seed

Seeds have a very high success rate in Tissue Culturing, and a lot of beginners start with them. Although a Clone of the Mother plant isn’t possible, it still gives the gratification of numerous plants for trade or sell, and is a good primer to hone Ones skills with all stages of TC, so that when a precious strike does happen on a leaf of a prized plant, You will be better prepared for success.

As with most plant species, seeds utilized as soon as possible after produced by the parent plants, have a better germination rate.

It is said that Dionaea seed require cold stratification, but I think it is generally accepted that stratification isn’t required if One possesses Fresh seeds, stored properly, so that they have retained a higher internal moisture content and the outer ‘shells’ are still acceptably permeable.

A batch of seeds that are older than a few months, or weren’t properly stored in a sealed vessel just above freezing, may require a natural stratification, or as an alternative, a 24 to 48 hour soak in a Giberellic acid solution that will duplicate the natural processes that may be required of the aged seed.

Seeds should be shiny, black, and plump. I don’t think the test of seeing which ones float and which ones sink is a reliable test of viability because of the surface tension of the water on these tiny, light seeds, cannot always be broken.



Leaves

Compared to flower stalks, the leaves are the hardest to be successful on, but are generally used as they are available most Year round.

The best time to harvest leaves for TC is in Spring (or when the Mother plant ‘thinks’ it’s Spring) as this is when the Plant is at it’s fastest growth rate. Inversely, leaves shouldn’t be harvested in the Fall or In Dormancy, as the plants internal chemistry has it programmed to stop growing, so shall it be with the results of trying to culture it in Vitro. So would it also stand to reason to not harvest leaves off of a plant that possess’ a flower stalk, as leaf production has halted at that moment, as well, not to mention that the plant is more stressed. Utilize the prized flower stalk, instead. If You are getting the ex-plants from the leaves of artificially lit plants, then Your collection may be at various stages of growth, and some may even be confused enough to be flowering.

The age of the leaf to be harvested doesn’t have to be the youngest, and has been suggested that middle aged leaves seem to give the highest success rate, similar to the percentages seen in leaf pulling propagation. A newly opened trap and leaf may be the best to harvest; it is hardier, still growing, and large enough to work with.

Harvest the entire leaf above the soil line. Unlike leaf pullings, most strikes in vitro on leaves have been on the upper half of the leaf and petiole area, so don’t think You have to preserve the lowest part of the leaf, or have the whitish rhizome to have success.

After sterilization, cut only the bottom part margin of the sterilized leaf, leaving the top and perhaps, even the trap. The ‘crease’ in the immature trap lobe tends to be a hiding place for contaminants and I have had success cutting the trap off.


Flower Stalk

The flower stalk on the Dionaea is treasured as the most viable tissue for Cloning in Vitro, with success percentages reported above 90%, far better than leaves with a current success rate of about 10%. Flower stalks are also more robust and can withstand a more rigorous sterilization process, which gives a better margin for error. The fact that the flower stalk grows erect and probably never has had contact with the soil can only benefit, as well.

Although Many have successfully propagated a flower stalk at all stages of growth, even post seed, (providing the stalk is still green) Most agree that the best time to harvest a Dionaea flower stalk is in pre-flower stage, just before the first flower opens (see illustration 1) with the most viable part and greatest chances of callous success being the junction of where the main stalk separates into the individual flower stalks. It has been suggested by an experienced Dionaea propagator, that the absolute best part of the flower stalk that He’s had 100% success with is the little ‘spur’ or node that sometimes grows midway up the main flower stalk. The main part of the stalk can also be cut and put into vitro. These cut ‘logs’ tend to have a lower success rate, but is a good back up plan if the main flowering segment succumbs to contamination (as can happen with all of the hiding places for contaminants in this part).


Utilize the upper ½ or 2/3 of the entire stalk, cutting it into 2 sections, if need be, to better fit the sterilization jars. This excess will give You something to hold with tweezers or gloved hand and can be cut much shorter later after sterilization.

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Illustration 1

There are several reasons why they turn black or just sit there without doing nothing. The black tips are dead tissues due to sterilization and from phenols. TC’rs only mention their success but not their fall. Remember that tissue don't usually strike. You are lucky if you get one strike from one culture. TC’rs usually take 10 to 20 ex-plants and only a small quantity would strike. Young parts of the plant doesn't necessarily means that it would strike, sometimes the older the better. It's like traditional leaf cutting, young ones wouldn't strike the most. Look for a middle aged ex-plant. Example is a newly opened trap on VFTs, they are good for ex-plants. I would suggest using the growing tips like the flower stalk that is just branching out as the inter-nodes and actively developing cells.

Should the media look all wet up on top, and maybe a drop or 2 of liquid that I've been draining off should be left there?

Your media will eventually evaporate so you can leave the excess liquid in the jar.

They are warm white, but probably should be the 6500K lights, is that important?

You don't need bright light for your TC. I usually get more strike success leaving the ex-plant in a cool dark place for 3 months then introducing it to light after that. Even the strike'd ones shouldn't be exposed to bright light as they are very fragile.

Not a problem Gary. With regards to your vented phytocaps, it should be fine as the vent does not release that much of air exchange. I would not recommend the humidifier, it might introduce microscopic spores to your cultures.

About the leaves and cutting it, yes it is OK to cut some part of the rhizome to remove excess sterilizing liquid that might still be present even after a good rinse. I tried cutting parts of the leaves to promote more strike but instead got phenolic exudation in a matter of days. I wouldn't recommend it. I would suggest to have the traps intact, had great success with it. Just a reminder, it is not good to do TC on VFTs right now as their genetic structure is going to tell the tissue that it is not the right time to grow. The best way to do ex-plants is in Spring when plant knows that they need to grow. Seeds are OK for now and for other tropical plants that doesn't need dormancy. Patience is a virtue. I killed lots of tissue but learned not from any mistakes but from understanding the main biology of plants. Sometimes it's common sense. Once you get one strike going, you will face another problem...Where to place all of them! As you said, you just need one. Last night I re-plated a Shark's Teeth, took me 30 minutes and divided at least 50 plants after only 1 month in multiplication medium. VFTs multiply by 10 fold every week.

Seeds need to be in a warm bright location even in TC. Ex-plants on the other hand can be in a cool dark place to avoid phenolic exudation.
 
TISSUE STERILIZATION

Clorox Bleach Protocol



Below is a method used by a very successful Dionaea Horticulturist for Flower Stalks:


VFT FLOWER STALK METHOD:
I use any age flower stalks, some have set seed and are turning black at the tips and still have been ok. However the older they are there is an increased chance of contamination. If seed is not required the younger the better. As for the stem itself I do not use it anymore for us it is too time consuming when we are having great results from the flower head. If using the stem use it as young as possible while it is still very soft.
1. I wait until the flowers have fully extended from the stalk.
When collecting the flower and the main stalk I wear latex gloves and place the whole cut stem with the flower head into a mild solution of hydrogen peroxide of .1%. I am already starting to work as clean as possible. This keeps the stem hydrated as well.
2. Cut all flowers off as close to the base of the flower as possible leaving as much of the flower stalk as possible (this is the stem the flower is attached to not the main stem). Cut the flower head from the main stalk about 25mm below where the flowers join the main stem This gives something to grip while working with it. You will have a stick with a number of little branches on it. One other thing, sometimes on a flower stalk half way up the stem a spur appears this section is excellent to use as well. This is my favorite piece to work with, it is not too big and always strikes.
3. Standing by a sink with running water grab the base of the stem with forceps and dip everything in methylated spirits. Use enough metho to completely submerge it and vigorously swish it around for no more than 30 seconds try to get all the air bubbles out of the joints etc. This is not so much a sterilization process (which it is as well) but a de-waxing of the plant material. Quickly wash it under the water making sure to remove all the metho. Holding it with fingers can be used for this (wearing gloves). One more thing use a strainer in the sink there is nothing worse to see your only flower head disappear down the plug hole.
4. Place the flower head in a solution ( I use 100ml per flower head) of water with a couple of drops of wetting agent. We use the liquid wetting agent used for soil and pots

















Seed Sterilization



Matt Miller’s Protocol

My protocol for Dionaea seed is:
1) 3 to 5 minutes in H2O2
2) 10 to 12 minutes in 10% bleach with 1 drop of Tween 20 per 100ml
3) Three 3 minute rinses in sterile H2O
4) Plate


Geoff’s method involves a 10 to 15 minute soak in 10% Bleach solution with 1 drop of Surfactant (such as Tween 20) per 100ml solution. Agitate occasionally. The seeds are then rinsed in tap water, then soaked in 3% H2O2 solution for 15 to 20 minutes. A sterile Water rinse finishes this method.


Hackerberry’s method is as follows:

Here's my protocol for VFT seeds. Never had any contamination and 90% germination.

10 second dip in alcohol [70% isopropyl]
10 minutes in 10% bleach
Rinse 3 times in sterile water.
Plate.



I have noticed that A coffee filter packet folded over the seeds and a paper clip to keep it shut helps. A nylon stocking might work as well.


08/30/2011
My method for Dionaea seeds recently was placing seeds in a folded rectangular piece of coffee filter, fastening shut with paperclip or the like; stirring in a 10% solution of 6% household bleach for 10 minutes, transferring the packet of seeds under the hood to straight 3% hydrogen peroxide (to neutralize the chlorox and further sterilize the seeds), then finally 2 - 3 minute sterile rinses; then plate. The seeds were layed on top, not buried onto 1/3 MS, full vitamin, full sugar media with 7.5g/L High Clarity Carageenan. I will post the results. The seeds were sowed 45 Days ago, shipped to me in Mid August in a plastic bag, and I refrigerated them until the date above.
Dionaea seeds need more light than tissue cultures for best results.

UPDATE: 9/15/11 1 seed has germinated. No contamination on any of the 4 separate vessels each containing 1 seed.


Flower Stalk Sterilization


This is Matt Miller’s technique:
• Harvest ex-plant and rinse thoroughly under running water. Soak it in clean RO or distilled water with a bit of anti-bacterial soap (triclosan)
2) Quick alcohol dip. Skip this for small or fragile leaves. Can go up to 40 seconds with flower stalks.
3) 2 to 4 minutes in H2O2.
4) Rinse in sterile water.
5) 4 to 12 minutes in 10% bleach with 2 drops tween 20 per 100ml. 4 minutes for small and fragile leaves (pings, Drosera, thin Dionaea, etc.). 8 to 12 minutes for big flower stalks. All other tissue is somewhere between those times.
6) Two 3 minute rinses in sterile water then plate.




JASON KOCH’S METHOD
Flower Stalk
(I have good success with this method)



I have been successful on flower stalks with this method:

Rinse in running tap water for 8 minutes or so
40 Seconds in 91% Alcohol
7-8 minutes in Hydrogen Peroxide (drug store strength).
8 minutes in 10% Clorox solution; doesn’t use tween 20 (I do)
2 Rinses for 5 minutes each in sterile water
Cut stalk/s including flower buds off of the little ‘stalk lets’ and the main stalk into 1 inch sections
1 final sterile rinse in water for 5 minutes
Plate FLAT on the media (1/3 or ½ MS with .5ml/Liter Kinetin






TISSUE STERILIZATION
PERACETIC ACID


Peracetic Acid solution is a very powerful oxidizer and promises to be a much simpler and Dionaea friendly solution to sterilization as well as tissue sterilization, eliminating the Clorox, rinses, etc. associated with the traditional methodology.

Peracetic Acid is easily made by boiling 1 part White Vinegar of grocery store strength, then adding 4 parts Hydrogen Peroxide (H2O2) to the hot Vinegar. Peracetic acid gains strength over time, and I read maturity can take up to 1 month. Because of this variable, it is best to use it at it’s weakest, or the same Day the solution is prepared. This eliminates the soak timing variable to give consistent results on the ex-plants.





‘VINOXIDE’ Flower Stalk Sterilization
(I have had problems with latent contamination with this so far)

Dip in 70% alcohol for 10 to 20 seconds, then immediately rinse off under running water thoroughly.
Agitate in PAA for 5 to 6 minutes. Shake off excess a bit, then cut off the excess lower main stalk, leaving just enough to anchor the ex-plant into the media. Excise the little leaflets, too, then the flower buds, leaving as much little stalk as possible. Plate while still wet with PAA.



VINOXIDE Leaf Sterilization
(works very good!)

For Dionaea leaves, the ex-plants are rinsed under cool, running tap water for several minutes. The ex-plants are then put into a cup of the Vin-Oxide solution that has been prepared 2 hours prior and is room temperature. The timer is set and the container capped and agitated continuously for 3 to 4.5 minutes depending on the size and hardness of the tissue. The solution containing the leaves are then dumped through a tea strainer suspended over an empty vessel, separating the ex-plants from the solution. The ex-plants are then laid on sterile paper towel, cut and plated onto the proper medium. Remove the traps, but leave on as much of the petiole as possible. The Petiole is the ‘hot spot’ of the ex-plant. Plate while still wet with the Vin-Oxide.




CARE OF CULTURES



STAGE 1
INITIATION STAGE


Day 1; Seeds: After Plating on 20 to 33% MS, place in well lit Vito Shelf.

Day 4: Look for phenolic (black, inky staining emitting from the cut ends of the ex-plant)on the freshly plated ex-plants, and under hood, transfer to a different part of the media that is fresh. Look for mold at this point. Bacteria ’may’ just start to be evident as a milkish liquid atop the media or outlining the ex-plant.

Approximately 4 weeks: Look for plumping, and/or curling of tissues or any other positive sign that the plant is in pre-callous. They may look goose pimply and may even have some reddish areas. It is a good idea to re-plate on same media formulation at this time, especially if they seem to be just living, as this really seems to give them a ‘boost’ and they may callous very quickly after doing this.

Approximately 4 Weeks; Seeds: Seeds should have been germinated by now; continue to grow on current media, unless something warrants re-plating, like drying media.

Approximately 8 weeks: Re-plate onto Plain 1/3 MS (no hormones) as there will be multiple callous and/or growing points at this point that it will be hard enough to do the first division when that times come, without the help of PGR’s. Plain ½ MS will allow the plants to mature to a large enough clump for division.

Approximately 8 Weeks; Seeds: Re-plate seedlings onto same initiation Media
Of plain 20% MS, full Sugar. Do this possibly one more time or when the plants are an inch or so big before going to the multiplication stage. The seedlings will be treated as leaves or Flower Stalks from now on.

An excerpt from Geoff; A professional VFT grower.

This is my method from seed:
1.Seeds germinated on 20%MS and transferred monthly for about 3 transfers to obtain a good working size plant.
2.Seedlings then transferred to a multiplication media, 50% MS + hormones, 50%MS is used from now on for all stages.
3. Multiplied plants are then transferred to a growth medium with 0.5mg/l Kinetin
4. Once clumps are big enough they can be divided into smaller workable clumps and can be graded for size etc. These clumps can be re-multiplied or placed on the final growth media for planting out.
5. Final growth media before planting out has no hormones, just 50%MS + 20g/l sugar.

Stage 2
Multiplication Stage

Divide clumps into smaller clumps and re-plate onto more Cytokinin containing ½ MS media for fast multiplication (like .5ml/L or more of Kinetin), or plain media as Dionaea will divide without PGR’s, bur not as vigorously. Phytotech’s prepared multiplication media is supposed to be a very good multiplication media. You May add .5ml/Liter Kinetin to the Phytotech media and see if growth and multiplication are improved even more.
Dionaea on Multiplication media may go well over the typical 4 to 5 week re-plating since no significant growth is experienced; this is depending on the size and/or number of ‘clumps’ that are utilizing this media. Any roots that may develop should be cut off upon re-plating to discourage rapid growth instead of rapid division of that particular plant. Same thing with flower stalks that sometimes sprout up.

Stage 3
Growing Stage

When division is no longer wanted, but maturity and size is. This is accomplished on 1/3 MS with Full Sugar, with no PGR’S added.

For production purposes, Dionaea are a species that do best with regular 4 to 5 week interval re-plating on growth media. Growth of the cultures significantly slows or stops after this period, depending on the ratio of plantlet to media ratio. Once growth stalls, it takes awhile for it to restart, slowing production. I have started using more media in the vessels to delay the frequency of replating on a given number of explants in the vessel.




Stage 4
Final Growth Stage

To prepare plants for VIVO by maturing and hardening, and to initiate the plants natural Photosynthesis process. This may take two or more platings depending on what size of plant You want to initiate in soil. It is recommended that Dionaea be the equivalent of a 3 year old plant when planting out as much larger, they seem too delicate. Be sure to plant out when plantlets are actively growing on non-expired media for best results out of vitro. This gives them the resources to adapt.

Dionaea do not need the 'hardening' media of lowering sugar. They seem to adapt and root well enough without this intermediate step. If One wants to, then this Growing Media is accomplished on Plain ½ MS without or little PGR’s and lowering the Sugar to 20grams/Liter. Lowering this Carbon source forces the plant to start the photosynthesis process, via it’s leaves. Stronger lights should help in this stage. Eliminating the PGR’s will induce more roots. 1 to 2mg/Liter powdered charcoal can be used to help absorb residual PGR’s and phenolics at this point.


PLANTING OUT

Media should be rinsed off of the plants thoroughly so as not to induce fungus in the soil.
Soil should be the standard 5 parts Sphagnum Peat, 3 parts Silica Sand, and 2 part Perlite. The media should be thoroughly wet at this time, but with no standing water in the tray.
A Fungicide drench would be prudent at this point to curb losses to fungus that the plants are very susceptible to at this point. The safest Fungicide for Human exposure is the Sulfur type. The Sulfur type is most effective as a preventative measure. Cleary 3336wp or flow-able is systemic and has been proven to work, and CAPTAN as well. All listed above are proven safe for Venus Flytraps. Others may be as well, but I have not tested. DO NOT USE COPPER BASED FUNGICIDES, as it is lethal to most Carnivorous plants. It may be necessary to use more than 1 type in an outbreak. It is VERY IMPORTANT to act immediately upon signs of fungal attack as it can spread from plant to plant very quickly and can kill an entire tray/s.

The VFT’S stomata are 100% dilated in the cultures, so covering the tray with a clear hood with many ½ inch or so sized holes is a must for at least 10 Days out of vitro, then the Humidity allowed to fall slowly by means of larger openings or otherwise (I’m experimenting with taking the hood completely off overnight after 7 or 8 Days out of vitro) over the next Week or Two. This acclimation allows time for the Stomata to become functional as well as root development, both of which are necessary to keep the plant hydrated.




CONTAMINATION
Types and Treatment


There are two types of contamination in Cultures; Fungal and Bacterial. Fungal is pretty much all of the molds and are generally fuzzy, and vary in color. Typical is the White and Green types.

Bacterial is typically not fuzzy, but creamy and can be a variety of pastel colors as well.
Bacterial contamination is generally harder to deal with.

Molds typically are apparent on fresh cultures within 3 to 5 Days.
Bacteria’s can take from 3 Days to Weeks before they appear.

I am experimenting with a medicine containing the same basic formulas including sugar as the contaminated media, but without any PGR’S and Gelling agent. Then add 6ml/Liter of PPM into the solution. Autoclave this solution and layer this formula over the gel of the contaminated culture. This solution can be stored indefinitely in the refrigerator, I’m told. The infected culture remains on this 2 layer treatment, until re-plating. I Will update the results as they come in.


Gary:

If you notice that the media has some bacteria grow. You going to place a liquid media over the current solid media. The liquid media composition will be the same as the solid but without gel and with 5ml/L of PPM. The liquid media has to be sterilize the same way that you sterilize the solid.

If the bacteria spot is visible most probably you need to retreat like three times until you get a liquid media older than 48 hours that is not cloudy. If you notice cloudiness in the liquid media you discard the liquid and put more fresh liquid media. This is a treatment for contaminated ex-plants

Do not cover completely the ex-plants

So far, this treatment seems to be more of ‘buying time’ to plant out than a cure, especially if the bacterial infection is internal from the ex-plant I have been discarding small, clumped tissue and only saving plantlets that are in the advanced stages of growth, and the PPM solution seems to really help keep the infection at bay until de-flasking. The infection; especially mold needs to be caught on the earliest stages for this to be effective.
 
Great article - is there a link to the original article?. Who is Gary Gipson - not a member here I suppose?

Would love to see more photos of the process and a list of the equipment & chemicals. How much does it cost for a minimum order for such chemicals?

Thanks for the article upload.
 
Gary was the TC supplier for carnivorous California before they have hired the in-house lab technicians. He's switching to another hobby now, hence sharing the document with us. I have his permission to share here to help others getting started. You can find the original document on Facebook group, search carnivorous plant tissue culture.
 
just a side note on the carrageenan GP812 from photo tec , its not consistent and may need adjustments when you add to the media mix you use, when buying a new container, have used 3 large containers and all have been differnt amounts or powder to get the media to set to my perferred firmness , has been my experience so far , but good stuff to use
 
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