Welcome to the first ever Milton CRISPR exploration! Over the course of the DYO, we worked to knockout the lacZ gene located in plasmids (placZ plasmids) of E. coli bacteria. The LacZ gene encodes the enzyme β-galactosidase, known to dissociate lactose into the simpler monosaccharides glucose and galactose. β-galactosidase can also hydrolyze the substrate X-gal in a reaction that produces a blue by-product (1). Since the blue by-product forms only when β-galactosidase breaks down X-gal, blue-pigmented E. coli colonies indicate the presence of functional β-galactosidase enzymes and therefore an operative lacZ gene in the bacteria. Conversely, white E. coli bacteria did not have a functional lacZ gene. Such a clear visualization of gene expression allows great opportunities for genome investigation, since bacterial pigmentation can act as one marker for successful gene knockout.

To initiate gene knockout, we transformed plasmids containing the Cas9 gene as well as a 20bp strand of guide RNA (gRNA) for the lacZ gene (pKO plasmids) into a sample of E. coli grown on agar with X-gal. Once this plasmid permeates the bacterial membrane, the E. coli can use its machinery to synthesize the necessary components for the CRISPR-Cas9 complex introduced to the cell through the plasmids. With the direction of the gRNA for the lacZ, the Cas9 nucleases can target and cut the full lacZ genes already present in the placZ plasmids of the E. coli. As discussed in our explanation video, a double stranded break will initiate the process of the non homologous end joining. (NHEJ), a very error-prone repair system that will permanently shut down the lacZ gene. Without a functional lacZ gene, E. coli cannot manufacture β-galactosidase and break down the X-gal present on the agar plates. Thus, we hypothesized that colonies with successful knockout of the lacZ gene would be white after transformation and incubation, as they cannot produce the blue by-product of X-gal catalysis. We also transformed a control plasmid containing the same Cas9 machinery but a random gRNA (pCtrl plasmids) into another bacterial sample. Without gRNA that matched the lacZ gene, Cas9 nucleases would not cut and consequently silence the lacZ gene. As a result, these control colonies would still produce functional β-galactosidase that would catalyze the breakdown of X-gal and stain the bacteria blue with the reaction by-product.

Both the pKO and pCtrl plasmids also contained a chloramphenicol resistance (CmR) gene that was needed to signal the plasmids’ complete transformation into E. coli cells. We incorporated chloramphenicol, a bacterial antibiotic that kills E. coli, into the agar medium on which E. coli was grown after transformation. Thus, the bacteria that grew after incubation were known to contain their treatment plasmids which help the CmR gene, key to their survival. The original placZ plasmids also consisted of another resistance gene for ampicillin (ampR). To ensure these plasmids were present in the E. coli before initiating transformation, the bacteria was rehydrated using ampicillin. Aditionally, after one futile attempt at transformation, we added ampicillin to our agar and completed the transformation again with the same bacteria to ensure our results derived from a procedural hiccup rather than an issue with the placZ plasmid.

While colony incubation could signify successful gene silencing, we also confirmed lacZ knockout thorough PCR genotyping in the last step of our experimentation. We conducted a multiplexed PCR using pre-constructed primers that aimed to amplify 500bp of the Cas9 gene as well as about 750bp of the lacZ gene. We ran this PCR on extracted DNA from both the white and blue E. coli colonies along with control lacZ and Cas9 DNA. Then we aliquoted samples of the PCR products into wells of a gel and let the electrophoresis run. In all gel lanes, we expected to see evidence of the Cas9 gene, as all transformed plasmids contained that sequence. But we predicted to only see a lacZ gene present in the PCR product from the blue or control colony; no lacZ gene in the white colony PCR samples represented a fruitful gene knockout of lacZ!

In both the pKO and pCtrl plates, transformation was achieved in only four bacterial cells (Figure 4). This finding is indicated by the number of E. coli colonies, or circles of bacterial growth, on each plate. Each small colony reproduced from one cell that obtained a plasmid through transformation and successfully transcribed the CmR gene in order to subsist on chloramphenicol media; then that cell asexually reproduced through binary fission, bearing more offspring with the same contents of the transformed plasmid. While the success rate of this transformation, like all others, was low, only one cell needs to successfully uptake a plasmid, since it can then create a whole colony. As predicted, E. coli containing the pCtrl plasmid were colored blue (Figure 4A). Again this pigmentation suggests that the bacteria still possesses a functional lacZ gene and therefore operative β-galactosidase enzymes that generate a blue by-product during their reaction with X-gal. Also as hypothesized, the pKO plate exhibited growth of only white E. coli, implying that these bacteria no longer carry functional lacZ genes (Figure 4B).

This conclusion was further solidified through PCR and gel electrophoresis. In lanes 2 and 3, respectively, the Cas9 and lacZ genes, amplified in PCR, demonstrate the relative sizes of the target PCR product (Figure 5). Aligning bands indicate the presence of that specific gene in the bacterial genome. The lane 4 bands display the presence of both the Cas9 and lacZ genes in the pCtrl E. coli colony. Since CRISPR was never initiated in the pCtrl bacteria due to the lack of correct gRNA, these results make sense: CRISPR never located and silenced the lacZ gene. In lane 5, the single band of DNA fragments the size of the Cas9 gene uncovers that the lacZ gene no longer existed in the bacteria with pKO plasmids. In other words, the pKO plasmid induced knockout of the lacZ gene, which could not be located and amplified by primers in PCR because NHEJ rendered the gene dysfunctional.

The wide range of genetic diversity in CRISPR-Cas systems (2 classes, 6 types, 19 subtypes), used by bacteria for viral defense, gestures to their complex evolutionary history (2). New systems derive from beneficial mutations that occur during asexual reproduction of bacteria. A closer comparison of the spacer content (viral DNA incorporated in the bacterial genome) in bacterial colonies revealed that different bacterial populations of the same species had high genetic variation in spacer DNA. In other words, the bacterial environment selects for spacer content that maximizes the bacterial immune response (2). Bacteria from the same habitat may experience similar viral threats and thus receive advantages from the same spacer content, which is used to target specific phages.

Furthermore, outside of bacteria, human’s employment of the CRISPR-Cas system for gene knockout and editing has inherently posed new threats on evolution and natural selection. When humans begin to direct evolution through genetic modification, they assume the power to define which traits are “good” and “bad” (3). As a result, nature holds less of an effect on the discrimination of the most niche traits. Additionally, due to human intervention in the genome, the potential of random mutations to drive evolution may deplete as scientists determine favorable traits. In agriculture, for example, scientists are using CRISPR to bolster plant resilience. In increasing the frequency of traits that humans see agreeable for their own consumption, scientists can also limit the genetic diversity within species by eliminating the occurrence of processes like balancing selection, a phenomenon where disadvantageous traits are naturally allowed to persist if they demonstrate linkage disequilibrium with advantageous traits (3). As gene editing has advanced, so too have the conversations about its ethical concerns as well as its potential to disrupt the intrinsic process of evolution. Do the opportunities of gene modification trump the ramifications of CRISPR technology on evolution?