Prunus Mume Propagation by Hardwood Cuttings During Early Winter - The Peter Adams Method

[Canada Bonsai]

Really hoping I didnt waste my time with these

No such thing as time wasted with propagation, only time invested in either production or learning

how long did the callous take?

I did a test. I don't yet understand significance of the test or whether it should guide future practice, but here is what I did:

Using 1 parent plant, I created two batches of cuttings composed of material from different areas, and with different thickness:

50% were taken on November 19, 2020 and put into the fridge.
50% were taken on December 19, 2020 and put into the fridge.

Both batches went into the greenhouse on January 19, 2021. At this time, callous was not detectable with the naked eye (as opposed to other cultivars I am using whose callous is remarkable!).

Both batches started opening their buds on January 27, 2021. I attached a picture of what they look like today. The back for 4 rows are the Nov 19 batch (up to the ID tag). The front 4 rows are the Dec 19 batch (after the ID tag). All of them are very resistant to tugging, which means they have either formed substantial callous, or even roots, post-planting. The success rate, at this stage, is 100%, but this can change rapidly over the next 18 months from what I understand. To respond to your concern @JoeR , i still can't tell whether I should have been more concerned that these 2 batches did not have visible callous prior to planting. Right now, it seems to have been unimportant but that opinion may change if these cuttings start dying on me. The importance of callous prior to planing might also change from cultivar to cultivar. I really don't know.

To be clear, I am merely sharing my observations. I still do not have enough experience or data to start drawing conclusions.

 

[River's Edge]

For those that have had success, how long did the callous take? Someone mentioned a month earlier in the thread. These have been in the fridge for a month and a week or so. Little to no signs of callous on them yet, they are all standard pink mume. If they are a cultivar I dont know it. Really hoping I didnt waste my time with these. Not a great picture but:

I would spray with a rinse of hydrogen peroxide and water, shake dry and place back in the fridge in a plastic bag with either a damp paper towel or sphagnum moss. Check in two weeks, then two weeks later! The brown callus should be readily visible on the circumference of the cuttings bottom edge.
If they have been in the fridge for one month at this point then the cuttings were taken two months later than I normally take them. I have not experimented with the later timing. I am in zone 8b and usually take them November 1st. What I can tell you for sure is that timing with mume cuttings is usually variable. The timing for callus or roots can be very dependant on the size of the cutting, length, number of nodes on the cutting, diameter of the cutting, age of the cutting. Not to mention the cultivar and its overall condition. Smaller thinner cuttings with fewer nodes tend to be later or less successful. I have only planted cuttings with visible callus and I remove the flower buds. If there is any evidence of leaf then they should be planted at that point for any chance of success! I have not experimented with planting cuttings that do not show callus.
So definitely worth continuing. Darkness is also a factor so if your fridge has a glass door make sure they are wrapped in dark plastic or dark container.
Wishing you later success!

 

[JoeR]

I would spray with a rinse of hydrogen peroxide and water, shake dry and place back in the fridge in a plastic bag with either a damp paper towel or sphagnum moss. Check in two weeks, then two weeks later! The brown callus should be readily visible on the circumference of the cuttings bottom edge.
If they have been in the fridge for one month at this point then the cuttings were taken two months later than I normally take them. I have not experimented with the later timing. I am in zone 8b and usually take them November 1st. What I can tell you for sure is that timing with mume cuttings is usually variable. The timing for callus or roots can be very dependant on the size of the cutting, length, number of nodes on the cutting, diameter of the cutting, age of the cutting. Not to mention the cultivar and its overall condition. Smaller thinner cuttings with fewer nodes tend to be later or less successful. I have only planted cuttings with visible callus and I remove the flower buds. If there is any evidence of leaf then they should be planted at that point for any chance of success! I have not experimented with planting cuttings that do not show callus.
So definitely worth continuing. Darkness is also a factor so if your fridge has a glass door make sure they are wrapped in dark plastic or dark container.
Wishing you later success!

Awesome thanks for the input, I was taking them out today to spray peroxide anyway, and they've been in the fridge in a bag with sphagnum or sand. One cutting did have a small amount of mold, so I cut the moisture back some after the peroxide.

I had to take the cuttings later than I wanted, but I was waiting until the leaves started to fall off. When I took them, I had to clean the fading leaves off all of the branches. My trees all held on to their leaves much longer this year than normal it seemed like.

 

[JoeR]

No such thing as time wasted with propagation, only time invested in either production or learning



I did a test. I don't yet understand significance of the test or whether it should guide future practice, but here is what I did:

Using 1 parent plant, I created two batches of cuttings composed of material from different areas, and with different thickness:

50% were taken on November 19, 2020 and put into the fridge.
50% were taken on December 19, 2020 and put into the fridge.

Both batches went into the greenhouse on January 19, 2021. At this time, callous was not detectable with the naked eye (as opposed to other cultivars I am using whose callous is remarkable!).

Both batches started opening their buds on January 27, 2021. I attached a picture of what they look like today. The back for 4 rows are the Nov 19 batch (up to the ID tag). The front 4 rows are the Dec 19 batch (after the ID tag). All of them are very resistant to tugging, which means they have either formed substantial callous, or even roots, post-planting. The success rate, at this stage, is 100%, but this can change rapidly over the next 18 months from what I understand. To respond to your concern @JoeR , i still can't tell whether I should have been more concerned that these 2 batches did not have visible callous prior to planting. Right now, it seems to have been unimportant but that opinion may change if these cuttings start dying on me. The importance of callous prior to planing might also change from cultivar to cultivar. I really don't know.

To be clear, I am merely sharing my observations. I still do not have enough experience or data to start drawing conclusions.

Very cool thanks for sharing the experiment. What light are you using? Looks like you used akadama and lava, wonder how that compares to perlite or other standard rooting soils.

 

[River's Edge]

Awesome thanks for the input, I was taking them out today to spray peroxide anyway, and they've been in the fridge in a bag with sphagnum or sand. One cutting did have a small amount of mold, so I cut the moisture back some after the peroxide.

I had to take the cuttings later than I wanted, but I was waiting until the leaves started to fall off. When I took them, I had to clean the fading leaves off all of the branches. My trees all held on to their leaves much longer this year than normal it seemed like.

I was not sure if the white specks were mold or something else! When I use a small saw to rough cut the stems it leaves small specks on the cuttings like that! I prefer the small saw as it does not crush as much tissue as side cutters! Easier to clean the cut with a grafting knife before applying rooting hormone.

 

[JoeR]

I was not sure if the white specks were mold or something else! When I use a small saw to rough cut the stems it leaves small specks on the cuttings like that! I prefer the small saw as it does not crush as much tissue as side cutters! Easier to clean the cut with a grafting knife before applying rooting hormone.

Nope I'm not sure what the specks are, one cutting just had a small tuft of mold at the end. A saw definitely seems like the better tool for some of the larger cuttings, these things have strong wood. Did you apply the hormone before storing in the fridge or after?

 

[River's Edge]

Nope I'm not sure what the specks are, one cutting just had a small tuft of mold at the end. A saw definitely seems like the better tool for some of the larger cuttings, these things have strong wood. Did you apply the hormone before storing in the fridge or after?

I have done both over the years. The process I practise now is using the rooting hormone after the callus is formed.

 

[Canada Bonsai]

I have been trying to correct chlorotic leaves for almost 2 weeks. (Status update: some of the cuttings have roots, others only have callous. The volume of leaves being produced seems to have no relation to whether the cutting has roots or not. At the risk of damaging a few cuttings, i'm discovering this by removing substrate from the surface with tweezers until i see the bottom of the cutting)

I initially tried dissolved biogold that i watered into the substrate and used as a foliar spray. I thought that I initially saw improvement, but things seemed to get worse over 3-4 days.

I then tried Pro-Mix's 20-08-08, again by watering as well as foliar spray (https://www.promixgardening.com/en/...ulti-purpose-soluble-garden-fertilizer-20-8-8). This has not worked either.

I have doubts about whether it might be the PH of my water since the uptake must be minimal at this stage, and i can't think that PH in my humidity should be an issue? Could this perhaps be caused by deficiencies in the quality of the resources that my parent plant was producing last summer/fall, which the cuttings taken from it are not relying on?

I was wondering if you might have encountered a similar issue @River's Edge ?

@0soyoung any 0solutions in mind?

Thank you in advance!

 

[0soyoung]

I favor the view that photosynthesis is necessary for rooting, although I am aware of many exceptions (analogous to some seeds will germinate without stratification, but a higher percentage do when they are). So I, like you, think the chlorosis is worrying.

Chlorosis is most often caused by a lack of iron, but magnesium is at the active center of the PSII chlorophyll. So you could try foliar sprays of iron sulfate (dissolving a dietary iron supplement pill in water will do) and/or Epsom salt (magnesium sulfate). I would aim for solutions of less than 1% conc., though the exact conc. isn't critical). Leaves should green up in a matter of days if either is effective.

 

[River's Edge]

I have been trying to correct chlorotic leaves for almost 2 weeks. (Status update: some of the cuttings have roots, others only have callous. The volume of leaves being produced seems to have no relation to whether the cutting has roots or not. At the risk of damaging a few cuttings, i'm discovering this by removing substrate from the surface with tweezers until i see the bottom of the cutting)

I initially tried dissolved biogold that i watered into the substrate and used as a foliar spray. I thought that I initially saw improvement, but things seemed to get worse over 3-4 days.

I then tried Pro-Mix's 20-08-08, again by watering as well as foliar spray (https://www.promixgardening.com/en/...ulti-purpose-soluble-garden-fertilizer-20-8-8). This has not worked either.

I have doubts about whether it might be the PH of my water since the uptake must be minimal at this stage, and i can't think that PH in my humidity should be an issue? Could this perhaps be caused by deficiencies in the quality of the resources that my parent plant was producing last summer/fall, which the cuttings taken from it are not relying on?

I was wondering if you might have encountered a similar issue @River's Edge ?

@0soyoung any 0solutions in mind?

Thank you in advance!

In my experience the two most common issues are overwatering and high PH in the soil or water, affecting uptake!
Wet compacted substrate can be a common cause, so I would monitor the watering in this situation. When working with cuttings in controlled situations there is a tendency to mist frequently, water often and not let things even begin to dry out!
Toxicity could be present if the concentration of foliar spray is too high or applications too frequent. I have practiced rinsing leaves between foliar applications to reduce salt build up, that may be of assistance in your situation.
Excess light can also affect the chloroplasts, strength of light, duration of exposure!

 

[Canada Bonsai]

@0soyoung @River's Edge thank you both so much!

i made adjustements and followed both of your ideas and advice, and things are already looking a lot greener

thank you guys!!

 

[JoeR]

Thank you to everyone who dropped knowledge in this thread, looks like a success so far for me! Still in the fridge for a couple weeks while I build the cutting chamber. They constantly had fungal growth but seems to not have affected them, just cosmetic.

 

[JoeR]

A few observations for callus formation.

Length of cuttings did not seem to matter, and neither did girth (directly). Unsurprisingly though, success was much better with shoots taken from the apex. Many of these were really thick cuttings however. Branches that had flower buds also seemed to do well, can't say better or worse definitively but possibly better. I did not cut them off.

Cuttings stored in the fridge did better than ones in a pot of sand outside. This is possibly due to inconsistent moisture, drying out, inconsistent temperature, etc. For the cuttings in the fridge, it was critical the ends of the cuttings made contact with the damp media. In one bundle, maybe half or so of my cutting ends weren't actually touching the media and subsequently none made callous.


So, if I were to do it again, id:
-take only apical shoots, floral or vegetative
-store in the fridge with damp sphagnum, over paper towels or sand
-ensure cut ends are buried in the sphagnum


First picture is well calloused cuttings (top) and unsuccessful or low callus cover (bottom). Second picture is the bundle where half missed the media. Pretty happy with it!

 

[River's Edge]

A few observations for callus formation.

Length of cuttings did not seem to matter, and neither did girth (directly). Unsurprisingly though, success was much better with shoots taken from the apex. Many of these were really thick cuttings however. Branches that had flower buds also seemed to do well, can't say better or worse definitively but possibly better. I did not cut them off.

Cuttings stored in the fridge did better than ones in a pot of sand outside. This is possibly due to inconsistent moisture, drying out, inconsistent temperature, etc. For the cuttings in the fridge, it was critical the ends of the cuttings made contact with the damp media. In one bundle, maybe half or so of my cutting ends weren't actually touching the media and subsequently none made callous.


So, if I were to do it again, id:
-take only apical shoots, floral or vegetative
-store in the fridge with damp sphagnum, over paper towels or sand
-ensure cut ends are buried in the sphagnum


First picture is well calloused cuttings (top) and unsuccessful or low callus cover (bottom). Second picture is the bundle where half missed the media. Pretty happy with it!

You might wish to wait and see the end result. Apical shoots tend to be too thin for adequate reserves to root properly a high percentage of the time. Callus formation is the beginning stage, leaf formation is another thing, root formation does not always follow! For more detailed comments you can check my article in the November issue of the ABS journal.
If you take only apical shoots the end results will be poor as opposed to taking thicker cuttings from one year old growth.

 

[JoeR]

You might wish to wait and see the end result. Apical shoots tend to be too thin for adequate reserves to root properly a high percentage of the time. Callus formation is the beginning stage, leaf formation is another thing, root formation does not always follow! For more detailed comments you can check my article in the November issue of the ABS journal.
If you take only apical shoots the end results will be poor as opposed to taking thicker cuttings from one year old growth.

Absolutely, I was just making notes for the callous stage. The apical branches were all one year old but also the thickest, these were taken from ground grown plants though. I'll check out the article!

 

[River's Edge]

Absolutely, I was just making notes for the callous stage. The apical branches were all one year old but also the thickest, these were taken from ground grown plants though. I'll check out the article!

The ground grown will often produce thicker shoots. I have found best results with cuttings of approximately 3/8 " as a guesstimate for size and five to six inches in length.

 

[River's Edge]

Hi all
I thought you might be interested in the following observation.
One of the difficulties with hard to propagate species is root development and differentiation within the period of time the cutting develops from reserves within the cutting itself.
This past winter I conducted a side experiment with two variations in substrate to see if there was a difference in root development.
For some i used seived medium particle pure pumice, for an alternate group i used seived medium particle pure akadama.
Both substrates produced ample roots but of very differrent characteristics.
The pumice produced thicker less differentiated root structure which resulted in less foliage production over the same period of time.
The Akadama produced thinner roots with substantially more differentiation that resulted in more additional foliage development within the same period of time.
I suspect the roots produced in pumice will suffer more damage in repotting compared to the roots produced in Akadama affecting survival rate and recovery time.
Both groups housed in the same propagation unit received the same care with respect to light, temperature, fertilizer and watering levels. The pumice required more frequent watering as it dried out faster than the akadama. I found this interesting because the two substrates have similar water retention percentage ratings. Interestingly this does not translate to a similar period of time the moisture is retained.
I would postulate that this is due to variation in pore size or possibly the apparent slight swelling characteristic of akadama. Other factors could be PH of substrate.
I have included two pictures, same size cuttings of same species and cultivar. Although I am including only two pictures the observations above were consistent for the substrates discussed.
It will be interesting to compare growth as the cuttings progress over the next few months. Based on previous experience I would predict the cuttings with advanced root differentiation will have better short term results and the others may eventually catch up.
The first picture show the roots from pumice, they are thicker with fewer feeder roots, and more easily damaged at this stage. The second picture shows the roots structure from akadama. Much more differentiated with greater foliage development with the same period of time. Stronger roots that are more pliable and easier to transplant without damage. This was very evident as I took the time to spread the radial roots for better nebari development in the next stages.

 

[エドガー]

Very interesting. Akadama is after-all clay (albeit fire baked), like the clay in native ground-soil... and clay has super high CEC levels and stay moist for a long time compared to pumice or lava rock.

From what I've read and seen... after akadama breaks down and compacts after a couple years, it looks super similar to native ground soil. (at least the brown clay we have here in CA)

 

[Pitoon]

I wonder if akadama that has reached the end state (broken down) could be collected and fired to make a pot?

 

[River's Edge]

Akadama is after-all clay

A bit misleading in the sense that Akadama has volcanic origins and contains clay minerals but lacks other clay like properties which are present in organic process formed clay. I believe it is lacking in the characteristics that would make it suitable for pottery, but I will leave that to the experts.
Great in depth description in the resource section for those interested in specifics.

Inorganic Soil Reference Sheet 1.2