You've convinced me about leaf guard cells It is really not possible to know for certain if a plant is dip, trip, or tet by guard cell size, breeding habit or external plant characteristics, registration info, offspring, and so on.

Where chromosome counts are made, there is still no absolute assurance that the cultivar's ploidy is stable, that it is named correctly or identified accurately. So much for database and hybridizer certainty ... I think I will name my first registered daylilies "Best Guess For Now" and "Embracing Uncertainty" :} Still, I do have, supposedly, the dip and tet versions of the cultivar "Swedish Girl," so you've also convinced me that it would be great to see if my two fans differ in any way.

Do I understand correctly that the need for chemical pre-treament is just for 1) Colchicine to make chromosomes easier to see, and 2) HCI to soften the cells so that they can be squashed more easily into a single layer without bursting? If a household alternate (say, vinegar) were possible to substitute for the HCI, softening the cells over days instead of minutes, and we took out the Colchicine step altogether during pre-treatment ... would the chromosomes still be too difficult to see, even at higher magnification?

The current source of my continuing curiosity, where Colchicine is not mentioned: http://people.bridgewater.edu/...
and a "yahoo" answer about HCI substitutes that piques my curiosity: "vinegar is a weak acid and is used in high school chem labs all the time because it is a lot safer than using stronger acids"

chalyse said:Do I understand correctly that the need for chemical pre-treament is just for 1) Colchicine to make chromosomes easier to see, and 2) HCI to soften the cells so that they can be squashed more easily into a single layer without bursting? If a household alternate (say, vinegar) were possible to substitute for the HCI, softening the cells over days instead of minutes, and we took out the Colchicine step altogether during pre-treatment ... would the chromosomes still be too difficult to see, even at higher magnification?

The current source of my continuing curiosity, where Colchicine is not mentioned: http://people.bridgewater.edu/...
and a "yahoo" answer about HCI substitutes that piques my curiosity: "vinegar is a weak acid and is used in high school chem labs all the time because it is a lot safer than using stronger acids"

For part of the cell cycle the chromosomes are not easily visible as they are completely elongated. Then during cell division the chromosomes condense and replicate. Cell division is only part of the cell cycle so in a group of hundreds of cells only a few will be in the right parts of the cell cycle so that their chromosomes are visible and countable. The colchicine stops the cell cycle at a stage that allows the chromosomes to be visible because they are condensed and causes cells to accumulate at that stage so there are more cells at the best stage for counting.

If colchicine (or equivalent treatment) is not used then there will be fewer cells at the right stage for chromosome counting (perhaps some root tips will not have any cells at the correct stage). One could work around that by looking at more root tips from the same plant. I'm not sure whether weak acids are able to break down the cell walls and other materials that prevent the cell from being squashed well enough so that chromosomes are not all on top of each other. It is possible to replace the hydrochloric acid with cell wall digesting enzymes. Those are available to the home-owner for making wine at home. One would have to experiment through trial and error to determine how long daylily root tips would need to be enzyme treated. It is possible to buy vinegar that is stronger than the regular household vinegar and one could experiment by trial and error with using it at temperatures similar to that for HCl.

"Where chromosome counts are made, there is still no absolute assurance that the cultivar's ploidy is stable..."

If the chromosomes of a cultivar have been counted properly then the count can be considered stable for that plant and its divisions, if it is diploid, triploid or tetraploid; it would be one in a million or lower for that number to change naturally.

heaven!! (... ... look away while I beam a little ... and dance a little...)

This is just the kind of process-building that fills me with joy. So, 500x or higher I will go, at the most accessible price, with some great alternative over-the-counter acids to try out, and some experimenting with how to best get cells to soften and spread under slides so that flatter single layers might display chromosomes more clearly. Even if it takes me a year to find a single cell containing chromosomes at meta- or anaphase to observe, it will be a spellbinding moment.

You guys are awesome. Thanks is hardly enough to say when a new endeavor appears and keys are lent to open the door to a fascinating new vista. Already, I have learned more than I ever did in high school biology ... not that it wasn't right there all along ... but because it had no real-world meaning to me then.

Now, it does.

Anyone who is interested in making diploid x triploid, triploid x diploid, diploid x tetraploid or tetraploid x diploid, etc., crosses should keep in mind that although natural pollinations are not very common in daylilies because those crosses will fail most of the time, many of the viable pods they produce will have been caused by natural pollinations.

As an example of the amount of contamination possible, of 37 plants produced from diploid-tetraploid crosses only 31 were triploids and the remaining six were diploids and tetraploids (probably from natural self or cross pollinations). Li ZW, Pinkham L, Campbell NF, Espinosa AC, Conev R. Development of triploid daylily (Hemerocallis) germplasm by embryo rescue. Euphytica 2009;169:313-318

When making cross-ploidy pollinations one should reduce the possibility of natural pollinations, for example by opening buds the day before, removing the stamens or anthers, bagging the flowers after hand-pollination, removing petals, covering the pistil, etc.

I've always wondered about waiting to remove stamens the day the flower is open and pollen is already viable ... since waiting for bloom opening means pollen gets sprinkled around while trying to get them out even if snips are used. I didn't realize that opening the bud a day prior and removing stamens or anthers would not irreparably damage the pistil's ability to successfully form pods!

Same deal for removing petals, but now I see that makes sense (they will fall off anyway). The hard part I have is with bagging ... humor me with another outlandish idea, for which I truly want feedback?

If I were to follow the practice of prior day removal of anthers, then pollinate (same day? or wait till petals open fully?), would putting a thin strip of masking tape over the end of the pollinated pistil (or around its tip, with the top end crimped off) not damage things too far, and allow some pollen underneath to remain viable? I use 1/8 width masking tape to record the cross and wrap it just just below the flower on the short stem, and it would be easy enough to pull off a bit to "cap" the pistil after pollen is set on it.

Is leaving the petals to open for the day of display, after anthers are routinely removed before maturation, then an option if some form of pistil-only protection is applied?

And (tries to bite tongue but there are just sooo many questions! lol) ... if I remove the anthers the day before will they mature if kept overnight so that they can be used for hand pollinating other cultivars?

Yes, and you can refrigerate and/or freeze pollen. Those little centrifuge tubes work wonderfully well!

Thanks, @Betja! Just to clarify, when I see anthers early enough, there is no visible pollen on them at all (they sometimes look just dark and smooth) like this:



But, I'm hoping if those are removed (to prevent unplanned pollination), and kept at 60-80 degrees for a period of time (hours? days?) the anthers will open up to reveal viable pollen, like below?



I will try it this summer, and use refrigeration to store the pollen, since my freezer attempts have not worked yet. Any idea of how long refrigerated pollen is viable (long enough to use them over the span of a summer, or just for a few days)?

Tina, Bill Waldrop has an excellent description of pollen collection and storage in his blog, and I think you should scroll down to Friday, August 2, 2013 and check it out. His blog is really wonderful and he seems to cover so many different aspects of daylily hybridizing, conversions, and just planting and growing in general.

http://billsdaylilycorner.blog...

I do it a little differently, since my climate is usually warmer and much drier. I remove the anthers from the stamens with tweezers while still attached to the bloom just as the anthers are opening/turning the golden color, and I make sure to remove any portion of the stamens that might still be attached, and I place them in a centrifuge tube and air dry them for a few hours if I'm going to freeze the pollen, or I leave it in the fridge for 2-3 days first if I'm going to use it right away. When the pollen is dry and fluffy, ready to go as your second picture shows, I close the top of the tube and gently shake it to separate the pollen and then VERY GENTLY apply it to the pistil of the flower I'm going to pollinate with a little paint brush. I'm sure there are others here who do it differently, but this works really well for me.

Yes, the pollen is actually produced and developmentally mature quite some time before the flower opens (just not 'dry'). You can open the flower before it does so itself and remove the stamens. Kept at room temperature and not in extremely high humidity the pollen will become fluffy and useable.

Some do use 'caps' of various sorts for the end of the pistil (usually to protect from rain). The most important factor would be to not damage the end (the style). Some form tiny tubes using aluminum foil wrapped around a nail, etc to make the pistil cover. Geneticists typically use small paper bags to 'bag' the flower.

If the pistil (or specifically the stigma) is protected from insect visits (since the day before the flower opens on its own) then the petals do not need to be removed. Removing the petals is a precaution that reduces insect visits to the flower. Some cultivars have pistils that push through the tip of the bud up to several days before the flower opens. Those cultivars need to have the end of the pistil protected from insect visits ideally from before the pistil becomes visible.

Checking daylily cultivar ploidies.

There are businesses, laboratories, that will examine plant ploidy for a fee (using flow cytometry to measure the amount of DNA in the cells). It is not extremely expensive (at least in some labs). I found one estimate from 2003 that indicated $11 per plant. I requested a current estimate from a lab. If the lab is outside the US then there would be the added cost of having the leaf samples inspected by the state/federal agencies for a phytosanitary certificate and the cost of couriering the package.

For one lab the current cost for determining the ploidy of plant samples is a basic flat charge of approximately (exchange rates fluctuate) $70US plus $4-$6 for each plant in the test. Testing 25 different cultivars/seedlings, etc. at the same time would then cost $70 + $100-$150 plus phytosanitary inspection and shipping fees. I would expect that there are laboratories in India and other countries with lower charges. There are probably commercial labs in the US that will analyze plant ploidy.

Cool! Yes, I often find my favorite double daylilies have their pistil/stigma visible a day or two before blooming ... so I'll get to work on some caps to try out for them as well (dang visiting insects). Having the blooming petals left intact to enjoy is an awesome bonus.

I see from Betty's link that having an absorbent surface is mentioned as being important for helping pollen fluff and dry - I'd always just dropped the anthers into a small plastic pill box, but could add a corner of paper towel if needed, and try both fridge and freezer, to see what happens?

Just in case anyone has more input to add, here's an over-the-counter root-tip fixer alternative I've found to try out:

Immerse cut root tips overnight in a fixative solution of 50% Horticultural Vinegar (20% acetic acid) and 50% Methyl Alcohol (Hardware Store Shellac Thinner) with about 1% Kosher Salt.
Remove rut root tips and put on a slide; blot away as much of the fixative as possible.
Add a drop of Methylene Blue (Fish medicine) to stain the chromosomes, then blot off as much of the stain as possible.
Then add a drop or two of glycerin or water to make a temporary mounting media. Put a cover glass over the root tip and press down on it gently but firmly using just your thumb or a soft, clean pencil-end eraser. Plastic cover-slips are best for this procedure as they will not break under the pressure.