Biology

Gene Editing for New Therapeutics

Abstract

The development of gene editing technologies based on programmable nucleases has recently made significant progress, making it possible to induce precise changes in the genomes of organisms. These tools can help elucidate the genetics behind various diseases and have immense promise as new therapeutics for correcting genetic mutations and permanently cure diseases that are resistant to traditional therapies. In this paper,  different nuclease-based gene editing technologies were reviewed, current progress toward the development of these technologies was presented, and delivery challenges that need to be addressed as well as future trends were discussed. Each nuclease has its own advantages and disadvantages, and the choice of which gene editing tool depends on the actual situation. With the progress in genetic research, in vivo delivery using novel vector and non-vector approaches need to be further developed to facilitate the delivery of the editing reagents and increase efficiency.

Introduction

Disease is everywhere. The delivery of proteins (monoclonal antibodies, enzymes, growth hormones, etc) as biotherapeutics is still popular. However, protein delivery has several limitations.[1] First, it is difficult to deliver proteins to across cell membranes and the blood-brain barrier due to the large size of the proteins; second, protein therapy is transient in nature and needs to be repeatedly administered to provide therapeutic benefit. These limitations drive scientists to focus on alternative ways, leading to the rise of gene editing. Scientists have discovered that over 3,000 mutations in the 25,000 interpreted genes of the human genome are connected to ill-causing phenotypes, and efforts are being made to directly manipulate DNA with the goal of permanently curing gene-related diseases in the near future. This process of genetic engineering in which DNA is modified, deleted, inserted, or replaced in the genome of an organism is known as gene editing.[2] 

Genome editing was developed in the late 20th century and is still a relatively new field. Despite that, it has been a heavily studied topic since its arrival. The first nuclease technology, called as zinc-finger nucleases (ZFN), was introduced in a 1991 and was the predominant genome targeting technology for over a decade.[3] In 2009, Transcription Activator-Like Effector Nucleases (TALENs) came into existence when the genome targeting ability of TAL effectors was discovered.[3] TALEN has a better way of targeting many sites and a simple method of building TAL effector arrays.[4] In 2012, Clustered Regulatory Interspaced Short Palindromic Repeats (CRISPR) was discovered as a new genome-editing tool. It formulates a complex containing the Cas-9 nuclease and small guide RNAs and gets rid of the need to build a completely customised endonuclease for each target, something TALEN and ZFN cannot accomplish. Because of this discovery, the entry barrier to genome editing has been lowered significantly, allowing for more users and more innovation.[3]

In this review, the different nuclease-based gene editing technologies will be described along with its advantages and disadvantages. Also, challenges will be discussed for each tool and the latest developments will be presented to realise the clinical benefits. 

Different Types of Gene Editing

There are currently two types of gene editing. One is known as “targeted gene replacement” and the end goal is to change localised sequences, usually ones that will result in null mutations. Simply, it means replacing an existing sequence with a different one premade in a laboratory. The other method involves extensive alterations in a subtler way, which could result in a deep change to the natural genome of species.[5] Both types of gene editing require the use of a type of enzyme called nucleases, which can cut the genome at specific places. There are four major nucleases so far in the industry: meganucleases, ZFNs, TALENs, and CRISPR/Cas-9. 

Meganuclease

Meganucleases are endodeoxyribonucleases with an ample recognition site of double-stranded DNA sequences.[6] This usually consists of 12 to 40 base pairs, and this site generally occurs only once in any given genome. Meganucleases (Figure 1) are therefore considered to be the most specific naturally occurring restriction enzymes.[7] 

Figure 1: Schematic diagram of four genome editing tools. (A) meganuclease has a homodimer structure. (B) Zinc finger nuclease (ZFN) is composed of two monomers, and pentagons represent a zinc finger DNA-binding domain.  (C). Transcription activator-like effector nuclease (TALEN) also comprises two monomers, and light green rectangles represent the DNA bind domain. (D) CRISPR/Cas9 system comprises a Cas9 protein (depicted in light gold) with two nuclease domains (HuvC and HNH), and a single guide RNA (sgRNA). The sgRNA guides the Cas9 protein to the complementary sequences of the DNA target. Arrowheads in red indicate cleave sites to each system. Diagram reprinted from Reference 8.

Meganucleases can be used to replace, eliminate or modify sequences in a highly-targeted way since they can change their recognition sequence through protein engineering. Meganucleases are used to modify all genome types, whether bacterial, plant or animal and they open up opportunities for innovation in humans.[6]

The meganuclease makes it easy for in vivo delivery due to its small size. The high specificity  of meganucleases allows a high degree of precision and a much lower cell toxicity. However, its application has a few limitations. First, targeting novel sequence is difficult due to the limited number of the meganuclease available naturally, thus the creation of new meganucleases for novel sequences will require substantial protein engineering. Also, the immunogenicity is unknown, as meganucleases may be derived from many organisms, including eukaryotes.

ZFNs

ZFNs are fusions of the nonspecific DNA cleavage domain from the restriction endonuclease with zinc-finger proteins. ZFNs can target specific DNA orders and this allows the ZFN to address and accurately change unique sequences inside higher organisms.[5]

As the second most famously encoded protein domain, a single zinc-finger is made up of around 30 amino acids in a conserved ββα figure. Some amino acids on the surface of the α-helix usually select three base pairs within the DNA smooth groove.[9] Figure 2 shows the structure of  α-helix and β sheet in protein secondary structure. 

Figure 2: Main secondary structure of protein. α-helix consists of a peptide chain coiled into a right-handed spiral conformation and stabilised by hydrogen bonds between the N-H and the C=O groups in the backbone. Beta sheets consist of beta strands (also β-strands) connected laterally by at least two or three backbone hydrogen bonds, forming a generally twisted, pleated sheet. See http://www.nslc.wustl.edu/courses/bio2960/labs/02Protein_Structure/PS2011.htm.

Zinc-finger proteins have become an important framework for the design of custom DNA-binding proteins, as the development of unnatural arrays with more than three domains progressed. After the finding of a highly-conserved linker sequence that allowed synthetic zinc-finger proteins, which recognise DNA sequences 9 to 18 bps in length, the production dramatically sped up. These 18 bps have the amazing capacity to attain specificity up to billions of base pairs of DNA.[10] This design has proven to be the best way of making these proteins. Thus, “modular assembly” was made with the use of an already established selection of ZFNs prepared by rational designs or combinations. This is very helpful since almost all of the 64 nucleotide trifectas have been paired with a ZFN, which was the only approach available for a long period of time.[11]

Engineered zinc-fingers are currently available to the public, and CompoZr of Sangamo Biosciences is in zinc-finger construction with Sigma-Aldrich, allowing investigators to bypass zinc-finger construction and validation altogether.[5] So far, an exciting utilisation of ZFNs has been in curative HIV research. The CCR5-ZFN trial is promising for the field, and more trials are being planned by Sangamo to investigate the dosing of the modified T-cells, with and without cyclophosphamide pre-treatment.[10]

ZFNs make ex vivo and in vivo delivery easy, and the probability of resulting immunogenicity is very low. However, the ease of multiplexing is difficult, it may require quite some efforts on protein engineering, and it is difficult to target non-G-rich sequences.[12]

TALENs

TALENS are hugely versatile and can be combined with numerous effector domains to affect genomic structure and function, including nucleases, transcriptional activators and repressors, recombinases, transposases, DNA and histone methyltransferases, and histone acetyltransferases.[5] They are transcription activator-like effector nucleases which are fusions of the Fokl cleavage domain and DNA-binding domains.

TALENs/TALEs are naturally occurring proteins from bacteria with genus Xanthomonas and contain DNA-binding domains made up of a series of 33–35 amino acid repeat domains that each recognise a single base pair. TALE specificity is determined by two hypervariable amino acids (Figure 1) that are known as repeat-variable di-residues (RVDs).[5]

Like ZFNs, TALE chains are tied for recognising contiguous DNA sequences. There is no re-engineering of the chains between repeats so that TALEs can potentially address single sites in the genome. Several large-scale, systematic studies have shown that TALE repeats can be combined to recognize virtually any user-defined sequence. One of the targeting limitations for TALE arrays is that TALE binding sites should start with a T base. Several methods have also been developed to allow rapid assembly of custom TALE arrays. These strategies include “Golden Gate” molecular cloning, high-throughput solid-phase assembly, and ligation-independent cloning techniques.[5]

In addition, the ease in which TALE repeats can be linked is demonstrated through the recent construction of a storage area comprised of TALENs targeting 18,740 human protein-coding genes. Equally as important, custom-designed TALE arrays are also readily available through Cellectis Bioresearch, Transposagen Biopharmaceuticals, and Life Technologies.[5]

TALENs enable an easy ex vivo delivery, but it is relatively challenging in terms of multiplexing and in vivo delivery. TALEs are sometimes preferred over ZFNs since they are more flexible than triplet-confined zinc-finger proteins. However, the cloning of repeat TALE arrays is a higher technical challenge due to their extensive identical repeat sequences. 

CRISPR/Cas

CRISPR/Cas is a method that can pass on acquired immunity to archaea and bacteria via RNA-guided DNA cleavage. Short segments of foreign DNA called “spacers” are integrated and transcribed into CRISPR RNA (crRNA). These crRNAs anneal to trans-activating sequence-specific cleavages and silence pathogenic DNA by Cas proteins. It can, therefore, be targeted to cleave basically any DNA chain by redesigning the crRNA.[5] CRISPR has shown promise in human cells, by co-delivery of plasmids expressing the Cas9 endonuclease and the necessary crRNA components. The Cas9 has also been converted into nickases, which allow a further level of control over the mechanism of DNA repair (Figure 1).[13] 

Cancer research scientists are currently using the technique to explore the biology of cancerous brain tumors with the aim of producing a specialised treatment. By picking apart cancer cells, researchers can decipher which genes are most important to the disease’s survival. In 2016, Chinese scientists began testing CRISPR edited immune cells in lung cancer sufferers[13], and China may end up being the first country to cure cancer using CRISPR with this technology

There has been some debate on ethics regarding embryo editing. Carrying out gene editing within human embryos could help to improve chances of pregnancy during in vitro fertilisation (IVF) treatments. In addition, scientists also hope to use CRISPR to reduce miscarriages. Since current genome editing technology does not have sufficient efficiency and specificity to be reliably safe, the potential risk is that mutations may be generated at non-target sites, resulting in permanent effects that may not always be benign or predictable. Therefore, CRISPR’s application to human cells is hotly debated, but in some countries like Sweden, it remains legal.[13] When the recent headline about the Chinese CRISPR twins came out, the global community was shocked.[14] With a goal of making the girls immune to the virus HIV by editing the genomes of the embryos using CRISPR/Cas9, He JianKui did not intend for controversy. Unfortunately, while gene editing can lead to life-changing results, it can also be used for detrimental causes. Scientists have gradually reached an understanding that implanting such an embryo is a boundary that should not be easily crossed until the risks are reduced or eliminated.

Another application has been stem cell editing, where genetic manipulation of patient-derived stem cells creates models for diseases. These include cystic fibrosis, Parkinson’s, and cardiomyopathy, etc. Now, it is possible to correct disease-causing mutations in donated patient stem cells to create isogenic cells for any target cell for research. Generating these isogenic lines makes it possible, for the first time, to unambiguously show the contribution of gene mutations to a disease phenotype.[15]

CRISPR-cas is considered to be faster, cheaper and more reliable technique than other gene editing tools. Because the target specificity of Cas9 depends on the guide RNA:DNA complementarity and not modifications to the protein itself (like the other three gene editing tools), engineering Cas9 to target new DNA sequences is as straightforward as it is easy to obtain different guide RNA sequences using standard cloning procedures and oligo synthesis. Also, CRISPR makes it easy for ex vivo delivery and multiplexing. However, the tool only has moderate ease of in vivo delivery because the Cas9 is relatively large and may have limited packaging capacity in the viral vectors. Also, it has a target constraint since the targeted sequence must precede a protospacer adjacent motif (PAM) which is mostly NGG (where N can be any of Adenine, Guanine, Cytosine or Thymine).[12] Even though DNA complementarity is highly specific based on base pair rule, there is still off-target cleavage occurrence with mismatches within the distal part of the PAM.  

The Need for Delivery Vehicles

Figure 3. Comparison of Ex Vivo and In Vivo gene editing therapy. In ex vivo therapy (top panel), cells are removed from a patient, corrected by gene editing and then transplanted back. It is critical that target cells must be capable of survival outside the body and post-transplantation. In in vivo therapy, cells are edited in situ (bottom panels), meaning the direct administration of a vector to the affected area. For in vivo systemic therapy, delivery system such as AAV vectors could be used for efficient editing in a wide range of tissue types. For in vivo targeted therapy, viral vectors could be injected locally to the affected tissue, such as the eye, brain, or muscle. Adapted from Reference 12.

To achieve a therapeutic effect, programmable nucleases need to be delivered directly to the nucleus of the target cells to enable gene editing. As a large biomolecule, nuclease is too large to readily diffuse across the membrane into the cell or the nucleus. Therefore, efficient methods of delivery will be critical to the success of therapeutic genome editing applications. Figure 3 lists the current nucleases delivery approach including both ex vivo and in vivo method. 

The ex vivo method involves taking target cell populations out of the body, modifying with programmable nucleases, and then transplanting back into the original host.[12] This way, the cells can be manipulated with several delivery methods including electroporation, cationic lipids, cell-penetrating peptides, carbon nanowires, and viral vectors. This is useful, since off-target modifications are a problem, as holding back the amount of nuclease could lower such mutations.[16]

In vivo methods are also used to deliver nucleases to diseased cells in their native tissues. The most common technique employs viral vectors to deliver these nucleases. Scientists initially remove the damaging genes from the virus, inject it with the desired genes, and then inject the virus into the body to insert its good genome into the target population.[17] Of course, this has its risks. The new genes may be inserted into the wrong part of the DNA, creating new problems. The virus could also overexpress, creating more proteins than originally planned.[7] Adeno-Associated Virus (AAV) is the most commonly used viral vector as it has high delivery efficacy for several types of tissue including the eye, liver, muscle, etc. However, the packaging capacity of AAV vectors is relatively small, which makes it challenging for the delivery of large nucleases.[18] To reduce the potential risks associated with the use of viral vectors, non-viral delivery methods, e.g. the use of lipid nanoparticles (LNP) or polymers as the major delivery vehicle, are also being investigated. Current strategies available for both physical and chemical delivery of CRISPR-Cas gene editing components were recently discussed and reviewed[14], and utilisation of a new delivery vehicle that can achieve higher efficiency, higher specificity, and lower toxicity will improve the therapeutic outcomes of nuclease delivery.

Conclusion and Extensions

Much progress has been made in the improvement of gene editing tools since their discovery, and many potential applications have been initiated, including the creation of animal disease models, treatment of infectious diseases, treatment of cancers, and modification of stem cells. Of these major nucleases used to cut and edit the genome, each nuclease has its own advantages and disadvantages, and the choice of which gene editing tool depends on the actual situation. First, none of the gene editing tools have perfect specificity and each need to be considered for the specific sequence. Secondly, the tolerability level of off-target cleavage and mutagenesis can vary, depending on the application. In some model organisms, there are ways to validate the effects of newly introduced sequences. Off-target mutations may be tolerable even in some medical applications if they do not lead to a novel clinical condition.

Looking forward, research in gene editing  is going to continue and advance. Genome editing will most likely continue to be a widely-used tool in research, commercial, and medical applications. One question that arises is whether CRISPR-Cas is the last tool in programmable nucleases, or perhaps something even better will be discovered soon.

Another trend down the road is the expansion of medical treatment. For example, trials with ZFNs used to target the CCR5 co-receptor gene in T cells of HIV-positive patients were encouraging and has spurred more clinical tests in the future. TALENs have also been used to enhance the efficacy of therapeutic CAR-T cells, and at least two trials using CRISPR-Cas9 for this purpose have started. These examples rely on the ex vivo delivery method, i.e., first edit  cells in the laboratory and then transfer back to the patient. With the progress in genetic research, in vivo delivery using novel vector and non-vector approaches need to be further developed to facilitate the delivery of the editing reagents and increase efficiency. 

Gene editing is a field that is already thriving and expecting more in the future. With the ZFN, TALEN, CRISPR, meganuclease, new innovations are being constantly produced. Humankind can only wait to see what the potential of these technologies will be.

Acknowledgements

I would like to thank Professor Qiaobing Xu at Tufts University ([email protected]) for his excellent mentorship and guidance in the preparation of this paper.

References

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About the Author

Daisy Wang, USA

Daisy Wang is currently a 12th grade student at Boston Latin School. She enjoys studying scientific topics and doing science projects. She is the finalist of Intel International Science & Engineering Fair (2019), the second place winner of Massachusetts Science & Engineering Fair (2019), and the first place winner in Boston Science & Engineering Fair (2018 & 2019). She promotes interest in science to many younger students in the school and helps them with their science projects. She successfully meets her academic goals, scoring high in her regular and AP classes. She also participates in community service in the local hospital and senior centers. She looks forward to studying and researching biology-related topics in college.

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