Last Sunday, I wrote about The case of the vanishing twist. Unless the plates are all contaminated, I’ll get clues tomorrow when I compare mutants grown at 20 and 25ºC. In the same post in passing, I mentioned another mystery: Charlie Anderson and collaborators stain arabidopsis roots nicely with fast scarlet but me and mine do not. I noticed from their paper that their staining solution was growth medium buffered to pH 5.6 and I wondered whether pH holds the solution to the mystery.
Figure 1.In lieu of pictures of plants, here is a shot of the inside of Wightwick Manor where Laura and I visited today. This was owned by wealthy makers of paint and ink who went all in for William Morris, Pre-raphaelites, and associated Arts and Crafts artists. They were papering their rooms with Morris papers (and textiles, and carvings, etc.) in the hey day of production. And the family donated the house and grounds in the 1930’s to National Trust intact (instead of as more usually after selling off all the good stuff for death duties). The house is crammed with paintings, textiles, pottery (lots by De Morgan. Absolutely amazing.
This week, I made up a buffer called 2-(N-morpholino)-ethanesulfonic acid. Actually, no one calls it that, instead the buffer is called MES. Yes, I know, “What a MESs!” ha, ha. I have to tell you that this MESs is cleaned up by a related buffer called, wait for it, MOPS.
With that out of the way, I adjusted the pH of the 25 mM MES to an even 6. I added some fast scarlet powder for a 0.01% solution, as used in the paper. As another treatment, I also took a 0.01% fast scarlet solution in water and added some detergent. For the same reasons that a detergent is good at helping dirt slide off of dishes, a detergent helps remove crud in the roots that could be blocking fast scarlet from reaching its target (cellulose). To my surprise, the detergent changed the lovely scarlet color of the dye solution to orange. A lovely orange to be sure but the chemistry behind that transformation eludes me completely.
Not so surprisingly, adding the detergent did perk up the staining but this is because the insides of the cell took up a lot of stain. I could see fluorescing nuclei. Not good for looking at cell walls. I might try a lower concentration.
Happily, staining in the pH 6 buffer was distinctly brighter than the staining I got in water, particularly in the all important elongation zone. Apparently I have uncovered the reason (well, a reason) for the good staining in the Anderson et al. paper. Buoyed by this success, I made a pH 7 buffer (PIPES) and a pH 8 buffer (HEPES). Yes, the person who named these buffers was droll. (He was also Good, David Good). Staining in these was all better than water and more or less about the same as pH 6. Staining for 45 min vs 5 hr seemed to make no difference. More or less equivalent staining from pH 6 through 8 is reasonable insofar as many processes have a broad pH optimum. Maybe pH 8 was a tad weaker but there was root-to-root variability, and only three roots per treatment, so not headed to the bank.
Alas, I have no pictures of the stained roots. There is no camera on the microscope I am using. For that reason, I am trusting my eyeballs and looking for large effects. I think that will suffice. I am not trying to understand what is going on (although that would be welcome); instead, I just want bright roots. Are the roots bright enough? Maybe. I am going to try increasing the concentration of stain by a factor of three. And, I am going to try to add a little salt. Another difference between staining in growth medium and water is the former has about 15 to 20 mM worth of ions (nitrate, phosphate, potassium, etc.). This adds up to a modest value of ionic strength, which I can emulate with potassium chloride. I think it is worth doing a little more checking. For now.