Dear Chlamy folks:
Recently, we obtained a gene, gsp1, for a putative transcription factor in
Chlamy. We have no mutants of the gene and would like to develop a
PCR-based method for obtaining knockouts. We have thought about 2 related
approaches, one using insertional mutagenesis (I) to delete a large portion
of the gene and one using EMS or UV to create smaller deletions within the
gene (II). This latter approach is used in C. elegans
(http://www.nki.nl/nkidep/h8/protocols/del.html). We would appreciate
responses to the ideas below. We feel relatively naive about this, so go
easy on us if we make really obvious mistakes in our thinking.
I. Insertional mutagenesis combined with PCR to obtain large deletions.
We would decide on the best gene to use for insertion; let's say we used
the ble gene. We would cut the gene out of the plasmid and use only the
gene for insertion. (When insertional mutagenesis is done in Chlamy we
understand that 10-20kb of DNA often is deleted.) We would choose an
antisense primer from the 5' end and a sense primer from the 3' end. We
would choose several primers from our gene, which is about 7 kb. Let's say
we choose 5 sense primers and 5 antisense primers evenly spaced along the
entire length of the gene. The insertion we would look for would knock out
about 1/2 of the gsp1 gene. If the insertion were at the 5' end, when we
did PCR on the DNA from the cell clone with this insertion using an
antisense primer from the 3' side of the middle of the gene, we would get a
novel PCR product of about 0.2-1.0 kb (depending on how far from the end of
the ble gene our ble primer was, etc.). The problem we are having is trying
to determine how feasible this approach would be. That is, how many clones
would we have to pick to find an insertion that knocked out ~1/2 the gene.
Or, in other words how large an insertional library of Chlamy clones
would we have to screen? ( I'm sure that this is a relatively simple
problem for some of you out there in Chlamy cyberspace.) One way I can
think about it is that (assuming the Chlamy genome is about 70,000 kb)
knocking out 699,996.5 kb of DNA not in our gene will not give us a signal.
Or, knocking out the proper 3.5/70,000 kb = 1/20,000 of the genome would
give us a signal. Does this mean that on average, we would have to screen
20,000 clones to get one positive? Please let us know if there is a
fundamental flaw in the reasoning. It seems that we would have to screen
more, because the insertions will not be even, but stochastic, and thus
some insertions will be at the same sites, and some sites will not be
knocked out. I don't know how to make this calculation. I also don't know
if it makes a difference if one uses a large (eg. Nit1) or a small (eg.
ble) gene for the insertions or if the amount of DNA that gets deleted is
For this to be a usable approach requires that each portion of the genome
have the same probability of receiving an inserted piece of DNA. I don't
know if that is true, either.
We would pick clones into microtiter plates. If we had to screen only
~10,000 clones (100 microtiter plates), we would probably pool the 100
plates into 10 plates (ie., 10 clones per well) for screening. Further, we
would pool and screen rows and columns, to find the intersection that gave
us a positive. We would probably do 2 separate PCR reactions on each pool;
one PCR reaction would contain both sense and antisense primers from the
ble gene and the set of 5 sense primers from gsp1. The other PCR reaction
would have both ble primers and the set of 5 antisense primers from gsp1.
Once we had a positive, we would go back to the originals and screen that
well from each of the 10 plates used to make the pool that gave us a
What factors should be considered in selecting a gene to use for insertion?
I believe that multiple copies of the inserted DNA sometimes are found
adjacent to each other. They would give novel PCR products if we did PCR
with the ble primers alone, though, and I think we could detect those.
At least one highly respected Chlamy researcher has told me several times
that s/he doesn't think that this is a feasible approach. (It may have
been said less tactfully than that.)
II. EMS or UV combined with PCR to obtain smaller deletions.
In this method, cells would be mutagenized with EMS or UV under conditions
that cause deletions. According to the worm people between 10-50% of
mutants have deletions using the described conditions. After mutagenesis,
the cells would be allowed to undergo several divisions (how many?) and
then divided into pools for screening. In this case a pair of primers
from gsp1 would be used that were spaced about 2 kb apart (larger, smaller
spacing?) and we would screen for PCR products that were smaller than
We look forward to your criticisms and ideas about these approaches.
William J. Snell, Ph.D.
Professor Cell Biology and Neuroscience
University of Texas Southwestern Medical Center
5323 Harry Hines Blvd.
Dallas, Texas, 75235-9039
email-William.Snell at email.swmed.edu