rna-guided editing of bacterial genomes using crispr-cas … · rna-guided editing of bacterial...
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Jiang et al. 2013, Nature Biotechnology
RNA-guided editing of bacterial genomes using CRISPR-Cas systems
Experiment 1.
Null hypothesis: When the genome contains the appropriate target DNA, the CRISPR-Cas
system will cleave the target. The resulting double strand break (genome damage, followed by
DNA degradation) will result in cell lethality.
Rationale: Let us construct two strains, one with a susceptible genome, the other with a non-
susceptible genome, and test whether the hypothesis is valid or not valid.
This experiment utilizes three Streptococcus pneumoniae strains. For simplicity let us
call them strains 1, 2 and 3. Note that Streptococcus takes up exogenously supplied DNA
efficiently, and provided there is homology, incorporates it into its chromosome by homologous
recombination at relatively high frequency.
Strain 1 contains a Type II CRISPR-Cas system that targets the DNA of a bacteriophage that
attacks Streptococcus.
Strain 2 contains the target phage DNA inserted at the SrtA locus of its chromosome.
Strain 3 contains an altered sequence of the target DNA (containing a mutation in the PAM
sequence) inserted at the SrtA locus.
Genomic DNA isolated from strain 1 was used to transform strains 2 and 3.
Simple expectation: The CRISPR-Cas9 system will kill strain 2 by cleaving the phage target
DNA present in its chromosome. Therefore, no transformants will be obtained.
Strain 3 will give transformants. Its genome should be resistant to cleavage by CRISPR-
Cas9 as the modified target contains a mutation in the PAM motif.
Experimental result: Unexpectedly, strain 2 also gave transformants at a reasonable
frequency, although about 10-fold less efficient than strain 3.
Analysis of the transformants showed that most of them arose by replacement of the
target DNA by the wild type SrtA gene present in strain 1.
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[If the same experiment was done with a CRISPR-Cas9 with a mutant version of Cas9 that
cannot cleave DNA, both strain 2 and strain 3 should have yielded the same frequency of
transformants. This control is not shown in the paper. We shall assume that this is the case]
Conclusion: The CRISPR-Cas9 system, together with the target DNA, appears to promote the
editing of the genome via replacement of the endogenous locus by a homologous DNA
sequence introduced from an exogenous source.
The general outline of the experiment and the results obtained from them are presented
in the figure below.
[Question: Based on the experimental scheme shown in this figure (panel a), you would expect
a key difference between the Kanamycin resistant transformants formed in strain 2 and most of
the similar transformants formed in strain 3. Based on our discussions on recombination, can
you point out this difference? Can you explain how this comes about?]
The relevant features of the donor strain(s), CrR6 and CrR6M are shown above. The
recipients are schematically drawn below.
The results from the experiments are in panel b on the following page.
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In the figure showing the results, the donor strains of genomic DNA are functionally the
same with respect to CRISPR. The dark and light grey bars in the graphs can be treated as
duplicates (two very similar experiments). The unshaded bar is a control transformation done
with DNA containing the gene for streptomycin resistance. The transformation by this gene is
not affected by CRISPR.
Experiment 2
Purpose: To further test expectations from the editing hypothesis.
Rationale: If we increase the frequency of the editing cassette in the transforming DNA, will we
increase the frequency of replacement of the susceptible target locus in the genome by the
edited form?
We refine the experiment 1 as follows.
(A) We transform strain 2 with the genomic DNA from strain 1 (as in experiment 1). No extra
DNA is included.
(B) In the above transformation, we also include a certain molar amount of the wild type SrtA
locus (in the form of PCR amplified DNA).
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(C) In the transformation, instead of the SrtA DNA, we include the same molar amount of
modified target DNA (resistant to CRISPR-Cas9 cleavage).
[Question: What would be a good control to have been included as a fourth assay (D)? This
relates to the location of the target site (at the edited locus) and the identity of the editing locus]
The experimental results consistent with the predictions are shown in the figure below.
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Experiment 3
Purpose: How to optimize the target requirements for editing a genome of choice. When one
introduces specific mutations in a gene by the CRISPR-Cas9 technology, the engineered or
edited gene should not be subject to cleavage by CRISPR-Cas9. One can ensure this by
choosing the PAM or proto-spacer sequences in the editing cassette to be resistant to CRISPR-
Cas9 cleavage.
Experimental rationale: The experimental rationale is based on previous observations that the
PAM sequence (proto-spacer adjacent motif) proximal to the 3’-end of the proto-spacer appear
to have the following consensus: 5’NGG3’.
To test the nature of the consensus PAM more critically, we can assemble a randomized
PAM: ‘5’NNN3’. To be more conservative, we will assemble a randomized five-nucleotide
sequence: 5’NNNNN3’. We can then assemble an editing cassette library by assembling them
using PCR based methods in which DNA synthesis is initiated by the randomized primer set.
The library construction is also designed to link the editing sequences to the gene encoding
chloramphenicol resistance (Cam-R). Theoretically, there will be 45 = 1024 combinations of a
pentamer sequence. Assuming that the different sequences are uniformly represented in the
library of editing DNAs, we can transform the recipient strain (engineered to encode the
CRISPR-Cas9 system) with this library, and obtain a large number of Cam-R transformants, say
approximately 105 or more. If all the editing sequences are equally effective in transforming the
recipient strain (that is, their resistance to CRISPR-Cas9 is more or less the same), one would
expect each editing sequence to be present in approximately 100 transformants in the library.
To account for potential differences in the representation of individual sequences in the
library, and other experimental variations as well, the same library was transformed into an
isogenic strain that lacked the CRISPR-Cas system (or contained the CRISPR cassette lacking
the spacer sequence). In this strain, all sequences, regardless of their PAM sequences, are
expected to produce Cam-R transformants.
The latest technology to assess the representation of sequences is called deep
sequencing or next generation sequencing. Here the DNA from the entire pool of transformants
is isolated, and the sequence output from the pool in the region of interest is collected. This
region can be enriched by PCR amplification, and it is the amplified DNA that is analyzed by
sequencing.
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The data sets are expressed in terms of ‘sequence reads’, where the number or reads
convey the frequency of a particular sequence of interest present in the DNA pool. In other
words, high reads indicate frequent presence of a sequence and low reads indicate the rare
presence of a sequence. The data can be presented as the ratio of the reads between the
experimental strain and the control strain.
We can then arrange a three letter (triplet) Table, as shown below, for the first three position of
the PAM sequence: 5’NNN—3’.
For each first position, say A, there are four second positions (A, G, C or T) and four
third positions (A, G, C or T), that is a total of 16 triplets that start with A. There will be the same
number of triplets that start with G, C and T, or a total of 64 triplets.
The ratio between the reads for a functional PAM position between the experimental
strain and the control strain will be quite low, as this sequence will be eliminated by the action of
CRISPR-Cas9.
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If all three positions were critical, there should be only one triplet that is starkly
underrepresented in the Table. If two positons were critical, there would be four such triplets. If
only one position was critical, this number would be 16.
The most underrepresented set (shown in red) are clustered in one cell, with second and
third positions as G, G. The effect of the first position is not changed for A, G, C or T.
The consensus for the Pam sequence is 5’NGG3’. The next two positions
are not critical (5’NGGNN3’).
Experiment 4
Purpose: How strict is the requirement of the ‘seed sequences’ for CRISPR-Cas9 action?
‘Seed sequences’ are the first 8-10 positions of the spacer immediately 5’ to the PAM
sequence. It was observed that base pairing at some or all of the spacer positions with the
crRNA may be important for destruction of invader DNA.
Rationale: The rationale is the same as in experiment 3 (above).
Randomize the seed positions. [In this experiment, all the 20 pairing positions of the
spacer were chosen for randomization.] Build transforming DNA cassettes with the randomized
sequences linked to the Cam-R gene. Transform the two isogenic strains (only one of which has
the targeting activity) with this DNA pool. Collect large number of transformants, and subject the
isolated DNA (amplified for the region of interest) to deep sequencing. The relative proportion of
the reads for each spacer position in the experimental versus control strains can be plotted as
shown below.
Note:
1. In this experiment the PAM sequence is kept the same: 5’TGG3’.
2. Although it may not be obvious from the description of the methodology, the randomization of
the 20 positions was likely done in several sets (either 20 separate experiments or fewer
experiments targeting four or five nucleotides at a time). Randomizing 20 positions all at once
would comprise, 420 possibilities. That number is close to a trillion (1012). You will have to screen
1012 transformants to get a single representation of each sequence.
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At the first two positions, there is a strong preference for the base complementary to that
in the crRNA. However, as we move further away, the degeneracy increases. After position 11
or 12, all four bases seem to work well.
Conclusions based on experiment 3 and 4:
For genome editing experiments, to protect the editing DNA cassette, it is easier (and also
sufficient) to mutate the PAM sequence. Additionally, one may also prevent base pairing at the
first two or three positions of the seed sequence.
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Experiment 5
Purpose: Engineering a strain into which spacers can be easily introduced to facilitate the
editing of individual genes.
Two strains are utilized for this purpose. One of these contains the cassette for the
expression of tracrRNA, a single repeat sequence, and the Cas9 gene linked to a drug
resistance marker inserted at a genomic locus. By PCR amplification of two DNA segments
using suitable primer pairs, one can construct two amplified DNA products, each containing the
desired spacer flanked by a copy of the repeat sequence at either end. By a process called
‘Gibson assembly’, one can generate a single cassette containing the spacer and the antibiotic
marker. The second strain is similar to the first in the integrated cassette for tracrRNA
expression, Cas9 and the single spacer. However the gene for a second antibiotic marker is
linked to this cassette.
The second strain serves as the experimental strain for editing. The amplified spacer
containing DNA along with the editing DNA (directed to a desired gene) can be introduced into
this strain. One recombination event will introduce the spacer sequence along with the new
antibiotic marker, which can be selected for. The second will replace the edited gene where the
targeted parental gene was located on the genome.
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Experiment 6
Purpose: To test how practical is targeted editing of a genome, and how efficient it is.
Rationale: Mutate the endogenous LacZ gene (bga according to Streptococcus naming
scheme) of Streptococcus. LacZ can be easily assayed, so the introduction of the mutation can
be readily confirmed.
The editing cassettes are of three types
A. In one the mutation R481A inactivates an active site residue.
B. In the second case, two mutation N563A and E564A, inactivate the enzyme.
C. In the third case, a large portion is deleted, creating a mini-lacZ gene that is inactive.
A
B
The top line in this figure shows the targeted endogenous LacZ gene. The coding strand
of LacZ contains the complement of the PAM sequence (3’ACC5’; complement of 5’TGG3’). In
the editing gene A, the seed sequence is altered by the R481A mutation, so that it is protected
against destruction. Furthermore the mutation introduces the recognition site for the restriction
enzyme BtgZ1, so the edited gene can be monitored by enzyme digestion of the DNA. In the
editing gene B, the PAM sequence is mutated, so it is safe from CRISPR-Cas9. The mutations
N563A, N564A introduces the restriction site for TseI.
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Transformation was performed with the spacer cassette along with an editing cassette.
In the control transformation, the wild type lacZ was used as the editor. Transformants were
selected by Kan-R selection. The results summarized below demonstrates that the modified
editors (not subject to degradation) gave ~10 fold-more transformants than the wild type editor
(subject to degradation).
The DNA from the transformants and control DNA (unedited) were amplified for the Lac
Z gene region, treated with BtgZ1 (in the case of editor A) and TseI (in the case of editor B),
and analyzed by electrophoresis on gels. The size difference between the editor and the
parental gene can also be readily detected by gel electrophoresis. The results of this analysis
(shown below) satisfy the expectations for the edited LacZ in all transformants except 1.
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Finally, the enzyme levels in the transformants were assayed. As expected the
transformants showed no more LacZ activity than background levels. Thus editing works, and
quite efficiently!!
Note: For the time being, ignore the double deletion results (bgaA, srtA) in the figures labeled
B and D (we will come to the double deletion in the next experiment).
Experiment 7
Purpose: To test if one can perform sequential editing of genes.
Rationale: As we saw in the previous experiment, the introduction of the spacer cassette in the
transformant replaces a preexisting drug marker by the newly introduced one. By switching back
and forth between the two markers, one can perform a series of sequential editing experiments.
In the first experiment, the spacer targeted the srtA gene with a srt gene as the editor.
The expectation is that the transformants will contain a deletion of the srtA gene. The srtA gene
(sortase A) is required to anchor LacZ to the cell wall. In its absence, the enzyme escapes from
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the cell surface into the supernatant. Thus srtA deletion can be identified by a considerable
decrease in the cellular LacZ activity.
In the subsequent experiment, the target was the lacZ gene, using a lacZ gene as the
editor. The co-transformation was performed in the transformant obtained in the previous
experiment, and already verified as srtA. These transformants, analyzed by the size of the lac
Z, verified the intended deletion. Furthermore, the transformants showed no LacZ activity
consistent with the double deletion. (Now, see whether the rightmost data in graphs ‘b’ and ‘d’
above make sense.)
Experiment 8
Purpose: If sequential editing of genes is possible, how about simultaneous editing of more
than one gene?
Rationale: Introduce the spacers for the intended targets in the targeting cassette along with
their corresponding editing cassettes in a single transformation experiment.
The experiment 7 is modified to deliver the spacers for LacZ and srtA along with the
deleted editor genes (as diagrammed below). The question is whether the Kan-R transformants
obtained contain the double deletion (deletions of the LacZ and srtA genes).
???
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The results shown below bear out the presence of the double deletion in the
predominant majority of the transformants. In rare cases, only one of the deletions (either in
LacZ or in srtA) is obtained.
Experiment 9
Purpose: What factors are responsible for the CRISPR-Cas mediated gene editing? What is
the level of background editing? What are the limitations of the method?
Rationale: The target of editing here is a gene for the resistance against erythromycin (ermAM)
containing an ‘in-frame’ stop codon placed within the srtA locus. We can call this gene erm-
AM(stop). The strain is therefore erythromycin sensitive (Erm-S). This experimental strain is
called JEN53.
[Question: Can you design an experimental scheme for constructing this strain using the
CRISPR-Cas editing system. Pattern your steps after those that we discussed in earlier
experiments.]
Our assay for editing is the restoration of the wild type ermAM gene. Since the ermAM(stop)
contains an artificially introduced PAM (see figure below), we can use the adjacent sequence as
the spacer for targeting this locus. The premature stop codon is ‘TAA’.
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The standard method we have followed for editing can be illustrated by the figure below. The
targeting cassette with the spacer will give Kan-R transformants after integration into the
chromosome. If the co-transforming editor gene performs its expected function, the Kan-r
transformants will be Erm-R as well.
Note: I have to warn you that the description of this set of experiments somewhat convoluted,
and the interpretations rather difficult to follow.
A. First, they want to assess the background level of recombination, that is, recombination not
assisted by the targeting system.
The transformation is performed with the editor DNA together with either (a) the CRISPR
construct with the targeting spacer (ermAM(Stop) or (b) the CRISPR construct lacking the
spacer (O). Note that the Kan-R gene is linked to the CRISPR construct.
They look for Erm-R colonies directly without selecting for Kan-R. The frequency of Erm-
R colonies is ~10-2 of the total colony forming units. [Total colony forming units = the number of
cells growing in the medium without erythromycin. 8.5 x 10-3 or 9.4 x 10-3 is close enough to 10-
2] The result is presented in the histogram plot below.
It is not mentioned what fraction of the Erm-R cells are also Kan-R, and what fraction is Kan-S.
Another missing fact: If they did the transformation with just the editing cassette, in the
absence of the CRISPR construct, what is the frequency of Erm-R colonies? Is it 10-2 or is it
different?
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B. They transform cells with the editor DNA in the presence of the control CRISPR (0), lacking
the targeting spacer. They select for Kan-R colonies first, then they estimate the frequency of
edited colonies (Erm-R) within this population, that is, Erm-R cfu (colony forming units)/Kan-R
cfu). This frequency is ~10-1 (see graph above). [7.9 x 10-2 can be taken as = ~1 x 10-1.]
Let us assume that the frequency of Kan-R recombinants (transformants) is 10-2 of the
total colony forming units. Then one tenth of these, 10-3 of total cfus, are also Erm-R. Based on
10-2 frequency of single Kan-R or Erm-R transformants, the theoretical frequency of the double
transformants (Kam-R and Erm-R) is 10-4 of total cfus. This value is a factor 10 lower than what
is experimentally observed. To account for this discrepancy, the authors suggest that there is a
subpopulation of the recipient cells that are more competent than the rest of the population for
DNA uptake or recombination or both.
C. Next they transform the recipient cells with the CRISPR cassette: ermAM(Stop) along with
the editing cassette. Now the frequency of Kan-R resistant transformants that are also Erm-R
(indicating editing) is 99%. That is, almost every transformant that received the CRISPR
cassette edited the ermAM(Stop) gene to wild type (see graph above).
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The most likely explanation for the result above is that an unedited ermAM(Stop) gene
would be targeted by the CRISPR system, causing chromosome damage and cell death.
They measured the fraction of Kan-R cfus produced after transforming the recipient
strain with functional CRISPR: ermAM(Stop) plus editing DNA or the dummy CRISPR:0 plus
editing DNA. The latter transformation gave 5 to 6 times more Kan-R cfus than the former (see
graph below). In other words, the CRISPR causes lethality in cells containing the target DNA,
unless the target gets edited and escapes CRISPR attack.
Editing template:
However, the above result also shows that not all unedited cells are killed, presumably
because there is some inactivating mutation in the CRISPR cassette or an escaper mutation in
the targeted gene (in the proto-spacer). Otherwise, 100% of the Kan-R trasformants obtained in
the transformation with CRISPR-ermAM(Stop) should have been Erm-R (which was not the
experimental result). To estimate the frequency at which escaper cells arise, we can transform
the recipient strain with CRISPR-ermAM(Stop) or CRISPR-(0) and count the frequency of Kan-
R transformants per unit amount of DNA. Here, no editor DNA is included in the transformation.
The ratio of the numbers gives the frequency of escaper cells. In the result shown below, the
frequency of escapers is ~3 x 10-3.
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Editing template: (None)
The explanation for the combined results is the following. The CRISPR cassette and the
editor cassette recombine with similar efficiencies with the chromosome. In the absence of the
CRISPR, the edited transformants and the transformants containing the unedited gene will both
survive on Kan-plates. What CRISPR does is to target the unedited gene and kill off the cells
containing them. In other words, one obtains apparently high-frequency editing within the Kan-R
population by CRISPR-mediated selection.
E. Does the CRISPR-induced double strand DNA break trigger recombination by the break
repair mechanism contributing to the total extent of CRISPR-mediated editing observed? The
co-transformation assays were performed as usual: (a) editor sequence plus CRISPR:
ermAM(Stop) and (b) editor plus CRISPR(0). The fraction of Kan-R, Erm-R cfus/total cfus were
plotted (see below). This fraction was roughly 2 to 3 time higher for (a) than for (b). The authors
interpret this result to mean that the CRISPR has a modest effect on recombination mediated
repair.
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Experiment 10
Purpose: Can we develop CRISPR-base techniques for editing genes of bacteria in general?
Rationale: Streptococcus has a strong homologous recombination system. Most bacteria, E.
coli or Salmonella, for example, are not as robust in recombination. Can one adapt the CRISPR
system to edit the genomes of the latter type of bacteria by placing the necessary components
in plasmids which can be efficiently introduced by transformation?
Since efficient integration of DNA segments into E. coli chromosomes by homologous
recombination requires special techniques, the initial trials are done by using the CRISPR-Cas
systems housed in plasmids. In a low copy plasmid harboring chloramphenicol resistance
(Cam-R), the tracrRNA, Cas9 and leader-direct repeats are housed. One can bring in a second
plasmid (compatible with the first plasmid) harboring kanamycin resistance which contains the
spacer sequence for targeting the chromosomal gene to be edited. The editor sequence is
introduced along with the spacer containing plasmid in the form of an oligonucleotide. In the
schematic diagram below, the design of the dual plasmid system is outlined.
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Let us now perform an editing experiment in which we wish to introduce an A-to-C
change in the rpsL gene (coding for a ribosomal protein). This mutation, which results in a
Lysine-to-threonine substitution, confers resistance to the drug streptomycin. Thus, we can
identify editing by selection on Srm-plates.
The recipient E. coli strain, containing the tracR-Cas9 plasmid, is transformed with the
spacer containing plasmid (pCRISPR-rpsL) and the oligonucleotide containing the A-to-C
mutation. Srm-R colonies were obtained, only when the oligonucleotide was included in the
transformation. In a control experiment, the spacer sequence was omitted from the transforming
plasmid (pCRISPR-0).
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The results obtained are plotted below. In the editing experiment, the fraction of Srm-R
cfus relative to total cfus is two orders of magnitude lower than the fraction of Kan-R cfus
relative to total cfus. If all the unedited cells (Srm-S) were killed by the CRISPR-Cas9, the
fraction of Srm-R cfus should have been the same as Kan-R cfus. That is, there are a lot of
Kan-R cells that escape CRISPR-Cas9 attack even though the rpsL gene is unedited. Thus, if
your edited gene cannot be selected, identifying an edited colony simply by screening the Kan-
R transformants would be an arduous task. One will have to screen a few hundred
transformants before hitting upon the right one.
When the control plasmid (pCRISPR-0) is used in the second transformation, there is a relative
increase in the Kan-R cfus by four orders of magnitude. Clearly, the CRISPR-Cas9 is killing off
a large fraction of cells with the wild type rpsL gene. However, the killing is not good enough for
routine editing of E. coli genes.
Experiment 11
Purpose: Increase the editing frequency, so that one may identify edited colonies even against
the background of escapers.
Rationale: The phage lambda ‘red’ recombination system is an efficient homologous
recombination system. It is comprised of the lambda Gam, Exo and Beta functions. E. coli
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strains genetically manipulated to express the ‘red’ system are called recombineering strains.
Such strains should be able to recombine the editor DNA in to the homologous chromosomal
locus at high frequency.
When a recombineering strain containing the Cam-R plasmid was transformed with the
control Kan-R (CRISPR-0) plasmid and the editing oligonucleotide, the fraction of Srm-R cfus
relative to the total cfus was ~5 x 10-5. This is the level of recombination of the editor DNA with
the chromosomal locus promoted by the ‘recombineering’ system (see graph below). When the
transformation was performed by replacing the Kan-R(CRISPR-0) plasmid with the Kan-
R(CRISPR-rpsL) plasmid, this fraction rose to ~2 x 10-4. The CRISPR stimulates recombination
of the editing DNA. This effect may result from the DNA break induced recombination, or the
killing of unedited cells or both.
If the results are expressed as the fractions of Srm-R
cfus per Kan-R cfus, the values for the control (CRIPR-0) and the experimental (CRISPR-rpsL)
are ~3 x 10-4 and ~6.5 x 10-1 (~65%), respectively (see, graph below). Now, two thirds of the
Kan-R transformants contain the edited gene. Instead of having to screen a couple of hundred
colonies, it is only necessary to screen a handful, say 8 or 10 colonies, to obtain the edited
gene.
What is the killing efficiency of the CRISPR system when the gene is unedited? The
cells containing the Cam-R plasmid were transformed with the Kan-R plasmid containing
CRISPR-rpsL or CRISPR-0 in the absence of the editing nucleotide. In the absence of killing,
one would expect roughly the same Kan-R cfus in both cases. As seen in the figure below, there
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is ~103 fold difference between the two, the absence of the functional CRISPR. The simple
explanation for this result is that CRISPR kills cells with the unedited rpsL gene, thus selecting
edited cells.
In the presence of the editing oligonucleotide, the number of cfus obtained with the CRISPR-
rpsL plasmid went up by a factor 4 to 5. Cells in which rpsL was edited by the oligonucleotide
escaped death at the hands of CRISPR-rpsL.
In theory, in the above experiment, when the editing oligonucleotide is omitted, one
should not have expected any Kan-R cells to survive in the presence of CRISPR-rpsL. Yet, we
did observe some escapers (1.2 x 102) in the graph above. Of course, the hundred odd colonies
are far fewer than the 4.8 x 105 colonies obtained in the absence of CRISPR. The fraction of
escapers is the ratio of the two numbers, 2.5 x 10-4. That is, two to three cells in every
10,000cells become resistant to CRISPR even without editing. [Think of the possibilities for
gaining resistance].
In the early part of this set of experiments (Experiment 11), we determined that the efficiency of
recombineering for replacing the rpsL gene by the mutant version (conferring Srm-R) is ~5 x 10-
5. We determined that the fraction of escapers (with unedited rpsL) is 2.5 x 10-4. There are
roughly five times more ‘escapers’ than ‘editees’ (I coined this term for a cell which underwent
editing at rpsL). Or among the Kan-R cfus, we should expect only one in every five (20%) to be
an ‘editee’, the other four being ‘escapers’. In reality, we saw 65% of the Kan-R cfus to be
edited. Therefore, the killing effect of CRISPR cannot completely account for the selection of
‘editees’. Perhaps the CRISPR mediated DNA break at rps-L may boost recombination
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efficiency over that due to recombineering alone. To test his possibility, the recipient strain was
cotransformed with the CRISPR-rpsL plasmid or the CRISPR-0 plasmid along with the editing
oligonucleotide. We can estimate the Kan-R, Srm-R transformants as a function of total cfus
(this way, we can get around the killing effect of the CRISPR). Indeed, there is a roughly six fold
increase in the frequency of Srm-R cfus with the CRISPR present (see graph on the right).
With the increase in recombineering aided by the CRISPR-assisted DNA break, nearly
every Kan-r colony is expected to have an edited rpsL gene, roughly in agreement with the
observe 65% ( six to seven out of 10).