How to perform a Bacterial CRISPR Cas9 Knockout Experiment

E. coli and other prokaryotes are often metabolically
engineered to produce valuable substances such as biofuels, pharmaceuticals, and biochemicals. For example, engineering E. coli to produce1,3-propanediol involved as many as 26 genetic modifications. This illustrates the need for a rapid and
cost-effective gene editing method for use in bacteria! One gene editing method that is gaining a
lot of traction is the CRISPR Cas9 system. Although the CRISPR system originated in bacteria,
it is more commonly used to edit eukaryotic genomes. This is because most bacteria do not possess
robust DNA repair systems, and therefore CRISPR-induced double-stranded DNA breaks are typically lethal
to bacteria. Some may see this as a barrier to bacterial
genome editing, but others have found it to be a benefit. How? CRISPR Cas9 can be used as form of negative
selection to dramatically improve the efficiency of another gene editing technique called recombineering. Gene editing experiments can be designed so
that when a bacterium is edited via recombineering it will lose a PAM sequence at the insert
site. This will prevent Cas9 from cleaving the DNA
of edited cells. In this way, CRISPR Cas9 is used to eliminate
unedited cells, creating a highly precise and robust bacterial gene editing system. In this video, we will demonstrate how to
perform a CRISPR-assisted knockout of the chloramphenicol resistance cassette in E.
coli. This case study will cover three phases:
Phase 1, the design and cloning of Cas9 and sgRNA for bacterial systems Phase 2, the preparation of electrocompetent cells and transformation Phase 3, screening and confirmation of gene knockout In phase 1, three components are prepared for CRISPR Gene Editing in E. coli. 1) A plasmid with the sgRNA 2) A plasmid carrying Cas9 and the Lambda
red genes and 3) the repair template First, sgRNAs are required to guide the Cas9 nuclease to the target locus. An sgRNA was designed against a chloramphenicol acetyltransferase cassette or CAT gene which was previously introduced into the E. coli genome. When intact, this gene gives resistance to
the antibiotic chloramphenicol. The sgRNA was cloned into the pTarget plasmid. Second, a construct carrying the Lambda red
genes is required. The Lambda red genes encode phage-derived
proteins that are crucial for enabling recombination of DNA fragments. The pCas9 plasmid carries inducible lambda
red genes, as well as constitutively expressed Cas9 and a Kanamycin resistance gene. Finally, to facilitate recombination, a repair
template is required. This template contains left and right homology
arms that are complementary to the target DNA. In this case study, the repair template carries
three stop codons for early termination of the CAT gene and a unique restriction site
for screening purposes. Most importantly, upon recombination, the
repair template eliminates the PAM site, thereby preventing Cas9 from targeting and cleaving
edited cells. In phase two, all the components are transformed into E. coli. pCas9 is transformed first, and grown at 30’C on kanamycin plates to
select for Cas9-expressing bacterial cells. The lambda red genes are induced to begin
expression and the cells are made electrocompetent. Electroporation is used to co-transform pTarget,
which carries the sgRNA, and the repair template into the cells. Once all the components are present, the lambda
red proteins enable the recombination of the repair template DNA with the bacterial genome. The repair template is inserted, causing the
CAT gene and PAM sequence to be disrupted. Without an intact PAM sequence, successfully
edited DNA will not be targeted and cleaved by Cas9. However, unedited cells will be targeted by Cas9 and will die when their DNA is cleaved. With this combination of recombineering and
CRISPR, the number of background unedited colonies can be greatly reduced in comparison
to recombineering alone. In phase three, cells are screened for CAT
gene knockouts. Transformants are replica picked onto kanamycin
and chloramphenicol agar plates. Cells that are able to grow on kanamycin plates
but not on chloramphenicol plates indicate E. coli cells that had the CAT gene successfully
knocked out. In this case, 36/45 transformants are successfully
edited. Successfully edited transformants are also
verified by restriction enzyme digest using the unique SpeI site introduced by the repair
template. The target locus is PCR amplified then digested
using SpeI and NcoI to reveal a unique digest profile. Three bands are observed to indicate a positive clone and two bands to indicate a negative clone. Finally, PCR products from positive clones
are subjected to Sanger sequencing to confirm correct repair template insertion and knockout
of the CAT gene. In this example, the sequence alignment shows
that colonies 1, 2, and 3 have the correct knockout insertion sequence containing three
stop codons and the SpeI restriction site. Using CRISPR Cas9 coupled with lambda red
recombineering technology, scientists have been able to achieve editing efficiencies
from 65% to close to 100%, making this method an excellent strategy for efficient genome
editing in E. coli. At abm, we offer CRISPR Knockout and Knock-in
Services in E. coli based on this game-changing technology. Our service includes: 1) Your E. coli strain of choice 2) sgRNA vector design and construction 3) Gene knockout or knock-in, at the locus of your choice 4) Clonal screening and selection 5) Validation of edits via diagnostic PCR
and Sanger Sequencing Simply send us your project and we can deliver
your sequence verified clones in just 8 weeks! For more information about this service or
any other CRISPR gene editing product or service, visit our website at Thanks for watching and don’t forget to
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  1. Hey everyone! If you're interested to learn more about CRISPR, we'd love for you to check out our new 4-week CRISPR Crash Course. It's completely free and you'll be ready to perform a successful CRISPR knockout experiment by the end of it. Sign up at:

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