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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).