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I understand that yeast integrating plasmids (YIps) must be linearized in order to promote homology-directed recombination. However I don't quite understand where.

In the end, the external regions of the linear constructs must be homologous to the genomic target; but quite often in a naked YIp backbone the only region that has homology to the host genome is the resistance marker... and cutting it would defeat the whole purpose of having a selection marker, would it not?

A basic example (from the Bartel Lab, 2009): enter image description here - The region 500-2000 is homologous to some weird, unrelated species. No homology with yeast genome. - An insert would go to the multiple cloning site (XbaI, NotI etc) - AmpR and ori are just there for the bacterial step

So all that's left is URA3, which one could cut with, say, NcoI... but then the yeast will get a negative phenotype (loss of uracile metabolism, by disruption of the biosynthetic gene), which is annoying to select for - and could be the result of a spontaneous KO mutation (unlikely but not impossible).

In the precise case of URA3, one could select for the loss of the gene with 5-Fluoroorotic acid, but other markers like LEU2 and HIS3 don't have such a straightforward loss-of-function selection strategy, as far as I know (?)

So, is there a way to select for YIp integration based on gain-of-function phenotypes in this example? or would one have to incorporate one in the transgene, like a fluorescent protein?

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  • $\begingroup$ I think the idea is to linearize the plasmid in the URA3 region and using this homology for homologous recombination thus disrupting the endogenous URA3 gene of the recombinant yeast. Negative selection would be made using 5-FOA as you suggest. Other sites on the plasmid would be used for cloning the gene to be inserted. $\endgroup$ – user22542 Feb 19 at 15:24

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