Home Plant Tissue Culture - Australian Trigger Plants

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Introduction: Home Plant Tissue Culture - Australian Trigger Plants

This Instructable will show you how to set up a home tissue culture lab, and will enable you to grow large numbers of many species of plants. I have focused here on growing a species of trigger plant, Stylidium corymbosum. If you are outside of Australia, just substitute African violet, the process works fine with that.

The work described in this Instructable formed part of the activities of a high school Tissue Culture Club, which I ran in 2018. I chose Stylidium (Trigger plants) because they are so cool! They can detect when an insect lands on their flower. When this happens they bash the insect with a fused stamen and pistil, that transfers pollen to the poor unfortunate. There are many Stylidium species, and to avoid undesirable or less fit hybrids, some are head specialists (hit the insects head), some are bum specialists (hit the insects rear end) and some hit only the left or right flank. Western Australia is a magic place for botanists - it has been geologically and climatically stable for tens of millions of years and is a global biodiversity hot spot for plants. You should come and have a look around in spring, at least once in your life!

This work can easily be done in a kitchen. There are a few technical tools that you need, but these can be purchased cheaply through ebay. A list of links showing you how to find a suitable cheap milligram balance for $15, and a pipettor for $25, as well as other resources is given at the end of this article.

I wish to acknowledge that my work here builds upon the skills of others. I publicly want to acknowledge the plant lore and insight of the Western Australian Wildflower Societies Hazel Dempster, (http://www.ns.wsowa.org.au/ ), who suggested the trivially easy Stylidium corymbosum as a species suitable to grow, and gave insight into its growth, non-tissue culture propagation, and who provided me with stock. I also acknowledge the support of other members of that association, (https://www.facebook.com/groups/129636970391772/), for their generous advice and comments.

I also wish to acknowledge Perth DIYBio group, (https://www.facebook.com/groups/diybioperth/) , for encouragement, fellowship and support, as well as my employer, Willetton Senior High School who supported my after-school Tissue Culture Club.

Step 1: Preparing Equipment

Tissue culture requires:

  • sterile workplace
  • suitable sterile media
  • suitable incubator/light chamber (see https://www.instructables.com/id/DIY-Plant-Tissue-Culture-Incubator/)

Firstly, we will look at making a sterile chamber and keeping it sterile.

Then we will look at how to make the media needed.

Then we will go through the steps needed to grow many plants from one cutting via tissue culture.

I will not show the details of how to make a suitable incubator/light chamber - that is covered in my previous Instructable https://www.instructables.com/id/DIY-Plant-Tissue-Culture-Incubator/

Step 2: Philosophy and Overview of Sterile Work

You are the biggest source of disease for your plants.

To mitigate this

  • Open bottles just before you need them, even in the sterile chamber.
  • Close bottles and containers as soon as you have finished with them
  • Don't allow your bare skin to be above anything you want sterile - your skin cells constantly fall off and carry germs with them
  • Check cultures for contamination
  • Prepare media a week in advance if you can and check it for microbial growth before using
  • Generously spray everything with 70% alcohol, repeatedly (don't contaminate your media though!)
  • Even with sterile sprayed gloves be careful not to touch the sterile inside surfaces of bottles and lids etc - pick them up from outside
  • Make sure that you are working in a room with still air - drafts are fatal and must be avoided.

Finally, cultivate your sense of paranoia. Germs are everywhere and they want to eat your plants!

Step 3: Making a Sterile Work Environment (aka Poor Man's Laminar Flow Chamber)

Materials for this step:

  • 1 x plastic box with lid ($10 from Reject Shop)
  • 1 x 3mm drill bit and drill
  • 1 x jig saw (box cutter can be substituted but work slowly to avoid shattering plastic)
  • 1 x light (look for a stable cheap LED torch)

Procedure

The goal of this part is to make an enclosed air space that does not allow movements of air and the microbial spores that come with that to come in contact with your media. Other people have used aquaria tipped on their side, but this design is better at preventing air movement.

Purchase a plastic box, of approximately 20l volume. The one you see above cost me $10 at The Reject Shop.

Drill a hole (3mm is fine, it is just to be able to start a cut) in a side panel. The one above is 11cm in from the side, and 3 cm up from the bottom. The dimensions are not critical, but you need to be able to fit both your hands in at the same time. The hole you see above is 18cm x 19cm. You can cut the saw with a very sharp knife, but these plastic boxes tend to shatter so work slowly and gently. Alternatively, a hand-held electric jigsaw is good for this task.

To use the chamber, mist it with a 70% ethanol solution as shown in the next step. Spray so that every bit of surface is covered (both the walls and the black lid which you use as a floor, and leave for 5 minutes. A battery powered light placed on the top will make it easy to see and work.

Step 4: Preparing Sterilising Alcohol

Materials needed for this step:

  • 1 refillable aluminium hair spray ($2 from Reject Shop)
  • Methylated spirits (1l, $4 from Coles).
  • water.

Preparing 70% ethanol.

70% is better at killing germs than 100%. By using 70% you have a much more efficient killing solution, as it is better at disrupting bacterial and spore membranes, and you save money. I mark out on an old methylated spirits bottle the height that corresponds to 700ml and the height that corresponds to 1l, so that I can make this solution really quickly.

  1. Measure out 700ml of methylated spirits
  2. Add 300 mL of tap water to it
  3. Mix well, then fill your sprayer with it (the pink cylinder in the photo above).

Generously spray everything you want sterile with this, and then leave for 5 minutes. After this time it will be sterile.

Step 5: Preparing Stock Solutions for Media

Procedure

Preparing Stock Hormone solutions (supplier details are on the Preparing the Media section)


Stock solutions

Stock 6-Benzaminopurine (this is a kinetin hormone): Obtain 6-benzylaminopurine (catalog number B800 from Austratec). Prepare 1mg/ml BAP (dissolve 0.2g in 200ml using a 2 or 3 decimal place balance). Split up into smaller lots of 1ml and freeze until use. Use an insulin syringe (which you can buy from a pharmacy) or a pipettor to measure out when adding to medium.

Stock naphthaleneacetic acid (this is an auxin hormone): Obtain Naphthaleneacetic acid, potassium salt (catalog number N610 from Austratec). Prepare 1mg/ml NAA. Weigh out 0.2g into 200ml using a 2 or 3 decimal place balance.Split up into smaller lots of 1ml and freeze until use. Use an insulin syringe
(which you can buy from a pharmacy) or a pipettor (see photo above) to measure out when adding to medium.

Step 6: Preparing the Media

Materials required

I recommend Austratec as a supplier of media components, hormones and agar. https://www.austratec.com.au/

They are sympathetic to the citizen scientist/amateur/school, and supplied good quality, free technical support to me when asked. If you are in Australia, then these guys are your best option at this time. If you are outside of Australia, Phytotech Laboratories (https://phytotechlab.com/ ) in the USA stocks all of the reagents listed here (Austratec has the Australian agency for Phytotech)

  • 6-benzylaminopurine (catalog number B800 Austratec)
  • Naphthaleneacetic acid, potassium salt (catalog number N610 from Austratec).

  • Murashige & Skoog basal medium with vitamins (catalog number M519 Austratec)
  • Tissue culture-grade agar (catalog numberA296 Austratec )
  • Plant Preservative Medium (PPM) (catalog number P820 Austratec) This is the "secret sauce" that will bring you success as it suppresses small amounts of contamination. While it can be omitted from the mix, your losses of plants will be much higher and you may find the process of learning good sterile technique very disheartening)
  • Sucrose - use ordinary white sugar from the supermarket.
  • Distilled or rain water
  • milligram balance (eg this $17.80 one from ebay https://www.ebay.com/itm/0-001g-20g-Digital-LCD-B... )
  • popstick (as spatula)
  • Insulin syringe from local pharmacy (for measuring 10/ 100 ul) or pipettor ( http://www.the-odin.com/pipette/ )
  • teaspoon
  • microwave oven
  • Pressure cooker (should heat to 121 degrees celsius at 2 atmospheres of pressure - they almost all do)
  • pickling jar (200-300ml, with seal)
  • culture tubes 25mm by 100mm or similar (must be autoclavable - most plastics are not). Note that other containers can be used instead eg really small pickling/preserving jars. eg https://www.austratec.com.au/product/culture-tube...
  • 250ml Schott clone bottles ( eg

    https://www.ebay.com/itm/250ml-Glass-Reagent-bottl... ). Pickling jars with seals also will do if you are short of money.

optional pH meter: You can get a $USD 7.80 pH meter from ebay here (https://www.ebay.com/itm/Portable-Digital-PH-Meter... )

To calibrate it, place a heaped teaspoon of cream of tartar (supermarket grade is fine) in a small cup of water and stir for a minute. As long as there is still solid powder in your cup, the pH of the solution will be 3.56. This is a cheap way of purchasing standards, and works just as well. Remember to rinse your pH meter before and after calibrating it, so that it doesn't become clogged with calcium tartrate deposits.

Recipes

0.05M HCl for pH adjusting (optional)

  1. Take 1 drop of fuming pool acid.
  2. Add to 100ml of water
  3. Take 1 drop of the resulting mixture and add it to 200ml of water.
  4. This is approximately 0.05M HCl, and is suitable for adjusting pH.

Callus forming medium


Place 50ml of water into a 200ml beaker

Add the following: 0.8g Agar (culture grade)

  • 0.22g Murashiga and Skoog salts (this is 1/2 strength for WA low nutrient soils- high phosphorus kills many of our plants)
  • 100 ul of BAP from stock solution
  • 10 ul NAA from stock solution
  • 3.0g Sucrose.
  • 100ul PPM solution.

Make up to 100ml using distilled water. Melt the agar by heating in microwave until solution starts to boil. Stir with a teaspoon and repeat heating until the solution is clear.

As soon as you have melted the agar, go on to making slopes.

Optional (as the Murashiga and Skoog salts from used have a pH of 5.7, but you might want to check anyway)
Use a pH meter and a pasteur pipette of weak, diluted acid 0.05M HCl, added dropwise, to adjust the pH to between 5.6 and 5.8. You will need at most a few small drops, so be slow and gentle)

Root forming medium

Place 50ml of distilled water into a 200ml beaker

Add the following:

  • 0.8g Agar (culture grade)
  • 0.22g Murashiga and Skoog salts (this is 1/2 strength for WA low nutrient soils)
  • 10 ul NAA from stock solution
  • 1.5g Sucrose.
  • 100ul PPM solution.

Make up to a final volume of 100ml using distilled water.

Melt the agar by heating in a microwave until solution starts to boil. Stir with a teaspoon and repeat heating until the solution is clear.

At this point, you can either transfer 3cm of the Rooting Media to pickling jars, or use it to make slopes (see next section).

Whatever you do, you must autoclave it soon to avoid microorganisms contaminating it. Twenty minutes after steam will do the job, followed by cool down to room temperature.

Rinsing water

(best to prepare a lot of 6 or 9 of these and leave ready to use. Sterile water is great for washing off mistakes as well as being important for rinsing off bleach in the plant preparation steps)

Place 100ml of tap water into a sealed glass jar. Autoclave it for 20 minutes, then allow it to cool overnight.

Step 7: Making Slopes

Procedure

Add either Callus or Rooting media to culture tubes (enough to 1/3 fill the container).
Place tubes in a pressure cooker, (filled with 4 cups of water) and heat until steam emerges and the pressure cooker starts to boil stably. Time for 20 minutes after this point, and then turn off. While still warm, remove the lids in the sterile chamber, and lean the agar tubes on a lid or something that will allow the tubes to set on a slope (see the above picture).

Step 8: Overview of the Process of Tissue Culture in Plants

This is an overview of how to culture your plants. Precise details are in the next section.

There are 5 steps to this.

  1. Harvest the plant tissue. Try to get growing tips, you need a plant organ called a meristem - a shoot or root tip. Hazel Dempster of the Western Australian Wildflower Society has a great idea for this - chop off the roots that poke out of the pot. Each root is a meristem, and you can get lots more than you can shoots. Personally, I go for the shoot tips, because I know they work.
  2. De-infest the plant material. Plants grow in ecosystems. They have their own microbiome. Organisms that they get on just fine as a whole plant will destroy them in tissue culture, so these must be removed.
  3. Once you have de-infested your plant, you need to grow it up into a callus. A callus is a confused mass of meristems, each of which is capable of becoming a new plant. You can think of this step as amplifying the number of plantlets you can grow.
  4. After you have grown a nice big callus, you will want to break it up into little plants. If it doesn't have roots - it aint a plant! So at this point you need to transfer the meristems from the broken up callus onto rooting medium. After 4 weeks, you should start to see roots.
  5. Once your new plantlets have good roots, they need to be removed from tissue culture and reintroduced to the world. You can expect most of your losses and deaths to occur at this stage.

Step 9: Collecting Plant Material for Callus Formation.

Materials needed:

  • Prepare 70% ethanol and sprayer as described before.
  • 3x autoclaved bottles of 100ml water. Prepared previously by placing 100ml of water in a pressure-resistant bottle in a pressure cooker. Heat until stable steam, then continue for 20 minutes.
  • 1 x jar of 200ml of 10% bleach solution (20ml bleach in 200ml water)
  • Sterilised forceps and scalpel
  • latex disposable gloves - Ansell are good, substitute similar if needed.
  • Detergent
  • Strainer (fine mesh to let water run through as you wash under the tap)
  • Culture tubes containing Callus Medium slopes prepared previously .

Procedure

Assemble and prepare the sterile chamber. Make sure it has been well sprayed with ethanol. Spray both sides of a plate, or use a sterile dish of the kind shown in the first photo in this section.

Your strategy here is to remove as much junk, bacteria and fungi as
you can from your plant material. You need tips with meristems (root tips should work too as long as they are properly clean), preferably young and actively growing.

  1. Trim a tip from a growing plant. 1 cm of material is enough, but be sure not to damage the growing tip.
  2. Dip your plant material into detergent (5 drops per glass of approximately 200ml). Stir and leave for 2 minutes.
  3. Rinse your plant material in a strainer under a running tap, until no foam bubbles are seen. Continue for 1 minute after this.
  4. Place your plant material into a jar of bleach solution (200ml containing 20ml of White King bleach and 180ml water). Leave for 10 minutes and shake occasionally. While you are waiting put your latex gloves on, spray them with the ethanol spray, and spray your dissecting tools (forceps and scalpel) with 70% ethanol.


    Your strategy in Steps 5-12 is to serially dilute out all of the bleach. Even a small amount of residue will kill your plants. So if 1/10th of a milliliter of fluid will stick to each plant, and each bottle of wash contains 100ml of water, you are diluting the bleach to 0.1/100 x 0.1/100 x 0.1/100 or to 1 billionth (1x 10^-9) of the concentration used to kill the bacteria and fungi. This should be safe, but remember to tap your plants between each wash, otherwise you will transfer too much bleach at each step.
  5. Pull out the plant material with an alcohol-sterilised pair of forceps (spray it), and transfer to the sterile chamber.
  6. Tap the plant material against the sterile dish to remove most of the bleach.
  7. Transfer the plant material to the first bottle of sterile water. Cap the bottle, and shake gently for 1 minute.
  8. Remove the plant material from the first water wash. Tap the material on a clean region of the sterile plate to remove the excess bleach.
  9. Transfer the plant material to the second bottle of sterile water. Cap the bottle, and shake gently for 1 minute.
  10. Remove the plant material from the second water wash. Tap the material on
    a clean region of the sterile plate to remove the excess water.
  11. Transfer the plant material to the third bottle or culture tube of sterile water. Cap the bottle, and shake gently for 1 minute.
  12. Remove the plant material from the third water wash. Tap the material on
    a clean region of the sterile plate to remove the excess water. You can pause at this step if you need to, as the final rinse water is no longer toxic to the plants.
  13. Open a culture slope containing Callus Medium. Push the stem end into the agar medium, cap and repeat until you have used up all the plant material.
  14. Place into the tissue culture incubator for 6 weeks, or until you have formed a good callus. Set the day/night cycle to 16h light/ 8h darkness.

Check your cultures every few days and discard any dead or sick ones by autoclaving. (This prevents build up of plant pathogens).

Step 10: Transferring to Rooting Medium

Once a suitably large callus (such as the one shown in the first photo in this section) has formed, it needs to be broken up and transferred to rooting medium.

Materials

  • sterile forceps
  • sterile scalpel
  • sterile dish
  • preserving jar with rooting medium
  • Sterile water.

Procedure

Fill the sterile dish with sterile water

Remove the callus from the agar, and slice it with the scalpel so that some of the inner material is attached to a meristem (here these are the ring or rosettes of leaves. Note that there are smaller ones between the larger ones, and that this callus contained approximately 25).

Shake each explant in the water to rinse off any media/liquid carry over, and tap it dry against the side of the sterile dish.

Place each explant into the preserving jar of rooting medium so that the cut end is down and the meristem end is up. Orientation is very important for plant development. Try not to get any leaves under the agar if possible.

When finished, seal the jar and place in the incubator for 4 weeks or until roots are formed. Alternatively, you might choose to give the plant 24h of darkness (store it in a cupboard) before transferring it to the incubator. The rooting medium contains all the nutrients necessary for plant growth, and the darkness may help start off the rooting process. I didn't bother with the dark, but some say that it helps.

After you have good root development, you may prepare your plants for planting out.

Step 11: Preparing to Plant Out

When the plantlets have developed good roots, it is time to plant them out.

Materials

  • Scalpel
  • forceps
  • Strainer (see photo)
  • Soup bowl
  • Clean Native potting mix
  • Propagating sand
  • square seedling pots

Procedure

  1. Remove the agar plug from the bottom of your rooting medium container. You can slice it with a non-sterile scalpel to help this.
  2. When you have isolated the individual plantlet, gently squeeze the agar block around its roots. The agar will break and you can most carefully pick it away. Do not break roots.
  3. After you have removed all the big bits, you will still find some small pieces stuck under the plant. Tease these out with forceps.
  4. It is critical to remove all agar. That medium is still full of goodies, and will feed up microorganisms that will destroy the delicate plantlets.
  5. Rinse the plantlets in a strainer held in a soup bowl by running tap water over it for 5 minutes. You must remove all traces of liquid from the plant, as any liquid carried over will contain nutrients that will feed disease causing pathogens.
  6. Plant into square pots, containing fresh potting mix / native potting mix mixed 50/50 with propagating sand. Cover the top of the pots with plastic wrap or other means to keep humidity high for the first week out of tissue culture.

Wait, and your plants will grow.

You have completed Tissue Culture!

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