Hi, dear all,
Since I joined Dicty community, I encountered a lot of technique problems or confusions. Some of them may be not a real problem, and the question may be stupid, but they all bother me for a long time. I put them all here, hope you can help me clarify them.

1.      Blasticidin selection: for knock out, after transformation for around 5 days under blasticidin selection, I transfer single colonies under medium to 96-well plate. For 96 colonies, only a few of them (usually around 10) can really grow up in the wells again. It looks to me a lot of "fake" colonies survive through the drug selection. If the recombination rate happens to be low, this phenomenon will make the screening very difficult to me. 5ug/ml of blasticidin is used. (This concentration is enough to stop WT cells growing and kill them under shaking condition, though).
2.      G418 selection: for overexpression, after transformation for around 5 days under G418 selection, I spread single cell on KA plate, and then pick single colony to culture in 12-well plate. A lot of them can not expand in the well anymore. Again, I suspect some WT cells survive through the drug selection. 20ug/ml of G418 is used. Even for the real colonies growing up, sometimes only a few of them express target protein, others do not express but are G418 resistant. This will make me picking a stable overexpression cell line is as difficult as screening a knock-out strain!
3.      Colonies on KA plate: for picking single colony on KA plate, I found if I pick too much KA bacteria with dicty cells to the plate, then the well surface will be occupied by the bacteria (although there are antibiotics in the medium), and the dicty cells would not eat bacteria to proliferate. Instead, they looks "trapped" individually by the surrounding bacteria, and starve to die after a few days!  If I pick much less bacteria with dicty cells, then dicty cells will expand! Is it possible bacteria secreting something to anti dicty?
4.      Colonies size on KA plate: the colonies sizes are not uniform at all even for cells from the same strain. And the size is also dependent on the physiological stage of the cells. For example, old, unhealthy cells give smaller colonies, comparing with normal cells. So, like screening strains giving different size of colony, or presenting data showing KO cells giving smaller colony size, does not make a lot of sense to me. Am I wrong?
5.      Chemotaxis: to show a mutant strain has chemotaxis defect, we would quantify the chemotaxis speed and directionality. However, chemotaxis is also dependent on how cells are polarized. For example, pi3k- cells would not show a lot of chemotaxis defect if they are pulsed long enough. The thing is "pulsed enough" or "polarized well" is an arbitrary concept. If polarizing is not paralleling, how meaningful to calculate the absolute number of "chemotaxis index"? At least we should not expect to get the same number from different labs, right?
6.      Loading control of northern blot during development: to show a gene expression changes during development, usually we show an mRNA northern blot normalized to "total RNA". However, we all know that the cells size shrinks a lot during cells starvation and development, and the "total RNA" also decrease a lot in a cell. The real question in our mind actually is "how this gene changes in a cell during development", or "how this gene changes in a aggregate/slug/fruiting body during development". So I think normalization to total RNA is not accurate at all.  Should we normalize RNA-seq of dictyexpress to "cell number" instead of "total RNA"?

Thanks,       Xin-Hua Liao