CRISPR (clustered regularly interspaced short palindromic repeats) technology can make precise, permanent changes to DNA in animals and plants. Even though just seven years have passed since the first in vitro demonstration of targeted DNA editing by CRISPR, early-phase clinical trials are already underway for several different genetic disorders in humans.
In therapeutic applications, CRISPR may help overcome some of the most persistent challenges in global health and development, including previously untreatable diseases. In diagnostic applications, it may help identify a host of infections, diseases, disorders, and other pathologies. Yet other kinds of CRISPR applications, including those in agriculture and the control of disease vectors, show promise.
CRISPR gene editing can make permanent modifications to DNA at a single, specific location. Use of CRISPR for gene editing is versatile and more precise, more cost effective, and faster than previous gene editing technologies (including gene targeting and zinc finger nucleases). It comprises a nuclease (for example, Cas9), which cleaves dsDNA; an RNA guide, which guides the nuclease complex to a specific DNA sequence; and (in case of homology-directed repair, aka HDR) an additional template DNA, which specifies for the cell’s machinery what edits to make at the target DNA site.
Cas9 is the most commonly used nuclease in CRISPR gene editing. With an appropriate guide RNA, it can target specific stretches of genetic code and cut DNA at precise locations, which may make it possible to correct the pathological DNA mutations underlying diseases like sickle-cell disease (SCD), Huntington’s disease, and cystic fibrosis. However, unintended DNA cuts and edits can sometimes occur at genomic sites that have DNA sequences similar to those of targeted sites. Such modifications are called off-target effects (OTEs).
Although CRISPR-Cas9 gene editing offers unparalleled genome editing efficiency and can function in various cell types and species, challenges remain due to insufficient target site specificity and low on-target efficiency. Many researchers have tried modifying the guide RNA and generating mutant Cas9 proteins to improve target specificity, but these alterations often also reduce on-target editing performance.
Here at IDT, we have successfully engineered a new Cas9 nuclease by devising an unbiased bacterial screen to isolate a high-fidelity Cas9 that has greater targeting specificity than the wild-type (WT) Cas9 while retaining nuclease activity comparable to that of WT Cas9. In fact, our HiFi Cas9 is the most active and specific high-fidelity Cas9 enzyme available, and it is delivered as a ribonucleo-protein complex, which provides optimal targeting specificity.
Although Cas9 is the most commonly used CRISPR nuclease, others do exist, and applications are being developed for them as well. These other enzymes can work in tandem with Cas9 or be used to serve radically different functions. For example, a newly discovered Cas14 protein can identify single-stranded DNA,1 whereas Cas3 can precisely recognize specific genetic sequences and subsequently degrade the DNA, leading to deletions of up to 100 kilobases.2,3
The use of multiple Cas nucleases means that a larger range of sequences can be recognized, enabling new applications. For example, panels of CRISPR components can contain multiple nucleases and guide RNAs to provide diagnostic information on a broad range of diseases, conditions, and pathogens.
Introducing Cas12a
While Cas9 remains the best-characterized and most widely used nuclease for gene editing, Cas12a (previously named Cpf1) has recently emerged as an alternative.4 Cas12a has several unique features (Figure 1) that distinguish it from Cas9 and expand the range of CRISPR-based genome editing tools. Most notable is the fact that Cas12a targets AT-rich regions of the genome, in contrast with Cas9, which targets GC-rich sequences. However, Cas12a suffers from dramatically lower nuclease activity than Cas9.
At IDT, we have recently focused on developing the Alt-R CRISPR-Cas12a (Cpf1) Ultra system, which targets alternative sites that are not available to the CRISPR-Cas9 system, to open up CRISPR editing to additional sequences in the genome. The new variant, Cas12a Ultra, has enhanced editing activity, reaching or exceeding the performance of Cas9 (Figure 2).
Moreover, Cas12a Ultra retains activity across a wider temperature range than the WT Cas12a enzyme (Figure 3), making it useful for genome editing in a wide variety of organisms, including plants and mammals. Given the unique characteristics demonstrated, what are some potential uses for Cas12a in gene editing and how do these differ from Cas9?
Applications for Cas12a
In the case of agriculture, the potential applications are endless. Gene editing using Cas12a could be used to make crops more resilient, increase yields, and boost nutritional value. Mushrooms with longer shelf lives, potatoes that are low in acrylamide (a potential carcinogen), and soybeans that produce healthier oil are already being developed and tested in the field, to name just a few.5
These are particularly interesting scenarios where Cas12a may be the more effective tool than Cas9, given both Cas12a’s affinity for AT-rich areas of the genome and the abundance of AT-rich regions in plant genomes. The technology can also be applied to livestock, and there is work currently ongoing to help researchers understand how gene editing might be used to help farmers in Africa breed more productive chickens and cows.5
Additionally, the human malaria parasite, Plasmodium falciparum, carried by the Anopheles gambiae mosquito, has a genome mostly made up of AT sequences—about 82%—which may be another appropriate target for CRISPR-Cas12a over CRISPR-Cas9. Research is being conducted along a variety of avenues to analyze the potential of using gene editing to prevent malaria transmission. Although current efforts and tools to prevent transmission must continue, full eradication will require novel technological advances. And CRISPR-based gene editing is one of the most promising tools being investigated.
Potential paths to eradication could involve targeting only female mosquitoes with gene drives—a technique that makes heritable edits to their genes that then pass down to offspring at higher than Mendelian rates—to either render future generations of females sterile (via inducing an intersex phenotype) or skew them toward producing predominantly male offspring, which do not bite humans, and therefore do not transmit the disease.
The path forward
As CRISPR technology advances rapidly and generates excitement among scientists and the general public, it is important to balance perspectives on both the potential benefits and risks. It is paramount that we have not only the tools to edit more precisely and effectively, but also tools to check for and eventually address OTEs, particularly when considering gene editing in animals and, eventually, humans.
Like other potentially powerful technologies and scientific breakthroughs, gene editing raises legitimate questions and fears about possible risks and misuse. Alleviating these fears to benefit from the opportunities afforded will require open, transparent, and continuous debate, as well as education of stakeholders—from scientists and governments to civil society and local communities.
References
1. Shieber J. Mammoth Biosciences adds the final piece of the CRISPR diagnostics puzzle to its toolkit. TechCrunch. Accessed August 6, 2019.
2. Makarova KS et al. An updated evolutionary classification of CRISPR-Cas systems. Nat. Rev. Microbiol. 2015; 13(11): 722–736.
3. Dolan AE et al. Introducing a Spectrum of Long-Range Genomic Deletions in Human Embryonic Stem Cells Using Type I CRISPR-Cas. Molec. Cell 2019; 74(5): 936–950.
4. Swarts DC, Jinek M. Cas9 versus Cas12a/Cpf1: Structure-function comparisons and implications for genome editing. Wiley Interdiscip. Rev. RNA 2018; 9(5).
5. Gates B. Gene Editing for Good: How CRISPR Could Transform Global Development. Foreign Affairs. Accessed August 7, 2019.
Chris Vakulskas, PhD, is senior staff scientist at Integrated DNA Technologies (IDT).
