To combat issues related to immunogenicity and short-term expression, other groups have employed adeno-associated virus (AAVs). AAVs are able to transduce both dividing and nondividing cells. In dividing cells, AAVs are able to integrate transgenes into the host's genome at a specific site on chromosome 19 [73]. Within non-dividing cells, the AAV genetic cargo remains largely episomal, as the chromatin is less accessible. AAVs are less immunogenic than adenoviral vectors and are reported to cause relatively long-term transgene expression in nondividing cells. The primary disadvantage of using AAVs is that their packaging capacity is less than 5 kb, limiting the use of larger expression cassettes with greater complexity for regulated expression. However, the preproinsulin gene is quite small, so even when the gene is accom‐ panied with multiple regulatory elements in a complex expression cassette, the maximum size limitation is unlikely to become an issue for insulin gene therapy applications. Indeed, AAVs have been used to successfully drive insulin expression within non-β cells. Park *et al*. used AAV to deliver insulin under control of the CMV promoter into STZ-induced diabetic rats and found improved glucose tolerance (at 2 g/kg) comparable to that of non-diabetic control rats [74]. Additionally, they observed a less pronounced immune response using AAV when compared to the same treatment using adenoviral vectors.

Retroviral vectors are another widely used gene delivery vehicle, owing to their ability to integrate their genetic cargo into a host's genome and attain sustained gene expression. However, retroviral vectors are greatly limited by their inability to integrate their cargo into the chromosomes of non-dividing cells, a problem that severely hinders their utility in insulin gene therapy applications. In cases where retroviral vectors have been used to deliver insulin, hepatocyte proliferation must first be stimulated [75]. This, of course, greatly limits the translation of retroviral vectors for treatment of T1DM. Retroviral vectors have also been shown to have a preference to integrate their genetic cargo in close proximity to the transcrip‐ tional regulatory sequences of proto-oncogenes, as observed in 1999 following the treatment of nine severe-combined immunodeficiency patients [76]. Insertional mutagenesis led to the development of leukemia in four of the nine patients, ultimately halting the field of clinical gene therapy temporarily. All in all, retroviral vectors do not possess favorable features for insulin gene therapy applications.

Lentiviral vectors are a type of retrovirus that provide two key advantages over other retro‐ viruses: (1) they are able to integrate their genetic cargo into the genome of both dividing and non-dividing cells and (2) have less preference to integrate near regulatory sequences of protooncogenes, reducing the risk of insertional mutagenesis [77, 78]. Their ability to transduce nondividing cells is critical for gene therapy strategies, as most cells of the body are either nondividing or slowly dividing. An additional advantage of lentiviral vectors is that they do not elicit a strong immune response. Unfortunately, lentiviral vectors possess two major pitfalls limiting their widespread translation to human clinical trials: (1) Lentiviral vectors are difficult to produce in high titer [79] and (2) the efficiency of lentiviral transduction *in vivo* is signifi‐ cantly lower than other vectors, especially adenoviral vectors. Given the relative instability of insulin, the need to transduce a larger number of cells than other gene therapy applications, and the fact that—unlike other diseases—a partial correction of hyperglycemia is not sufficient to adequately treat T1DM, these pitfalls pose some limitations to the use of lentiviral vectors for insulin gene therapy applications. Nonetheless, lentiviral vectors offer long-term transgene expression with reduced immunogenicity and an improved biosafety profile and are thus a viable candidate as a therapeutic tool for treatment of T1DM.

To combat issues related to lentivirus infectivity, researchers have modified lentiviral vectors to improve their *in vivo* efficacy. For example, Naldini *et al*. has previously shown that inclusion of viral protein R—a viral protein present in native HIV-1 particles—within synthetic lentiviral particles is critical for hepatocyte transduction [80]. Conversely, Schaffer *et al*. has generated vesicular stromatitis virus envelope proteins that show improved serum resistance [81], which could improve lentiviral efficacy *in vivo*. A combination of modifications will hopefully yield a more potent lentiviral particle *in vivo*. Interestingly, Ren and colleagues used lentiviral vectors to deliver insulin to the livers of STZ-induced diabetic rats [82] and NOD mice [83] and observed long-term correction of diabetic hyperglycemia with no evidence of impaired liver function, intrahepatic inflammation, or recurring autoimmunity against the newly formed insulin-producing cells. While their work gives validity to the use of lentiviral vectors for insulin gene therapy applications, it should be noted that insulin expression was driven by the CMV promoter and displayed no responsiveness to circulating glucose levels.

#### **3. Concluding remarks**

Overall, insulin gene therapy provides a promising alternative to current treatments for T1DM. Although this treatment option will inevitably lack the full sophistication of native β cells, it would certainly improve upon current treatment options. Insulin gene therapy opens the possibility of having a one-time treatment option that can provide long-term correction of diabetic hyperglycemia through physiologically relevant mechanisms, like glucose-dependent alterations in insulin transcription and translation. Further, the simplicity of the treatment should yield reproducible results with excellent success rates and additionally help newlyformed insulin-producing surrogate β cells evade recurring autoimmunity. However, several hurdles must still be overcome.

In order for the treatment to be a viable option, long-term insulin expression must be driven to avoid repeated injection. However, the viral vectors currently used to drive long-term transgene expression, like lentivirus, are generally difficult to produce in high titer and limited by their transduction efficiency. As a result, most successful long-term efforts in the field have employed the CMV promoter, which can only drive strong constitutive expression of insulin. In order to take those research efforts to the next level, weaker tissue-specific promoters must be used that drive low basal levels of insulin expression during fasting periods and substan‐ tially upregulated insulin expression upon increases in glucose availability. To date, this has yet to be achieved. We are currently exploring several viral vectors for their capacity to deliver our insulin expression cassette—which has elements to drive liver-specific, glucose-responsive insulin expression —at a therapeutic level. In so doing, we hope to produce an affordable, long-term treatment option for patients with T1DM.

#### **Author details**

Andrew M Handorf, Hans W Sollinger\* and Tausif Alam

\*Address all correspondence to<EMAIL_ADDRESS>
University of Wisconsin-Madison, Madison, WI, USA

#### **References**

in insulin-dependent diabetes mellitus. The New England Journal of Medicine. 1993;329(14):977–86. PubMed PMID: 8366922.

of insulin mRNA stability. The Journal of Biological Chemistry. 1985;260(25):13590–4. PubMed PMID: 3902821.

## **Application of Genome Editing Technology to MicroRNA Research in Mammalians**

Lei Yu, Jennifer Batara and Biao Lu

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/

#### **Abstract**

Targeted nucleases have recently emerged as a powerful genome editing tool. The ability of introducing targeted, desired changes into mammalian genome makes them an invalu‐ able tool to unravel functions of miRNAs in biology and disease. In combination with ho‐ mologous donor vector, targeted nucleases can achieve high efficiency and precision, enabling bi-allelic ablation of miRNA in cultured somatic cells. Here we review the struc‐ ture and function of miRNA as well as the unique implementation of genome editing technology in modifying miRNA sequences in mammalians. This chapter discusses the four mainstay genome editing technologies: meganuclease, zinc finger nuclease (ZFN), transcription activator-like effector nuclease (TALEN) and clustered regularly inter‐ spaced short palindromic repeat-associated nuclease Cas9 (CRISPR-Cas9), focusing on TALEN.

**Keywords:** Genome editing, miroRNA, TALEN, ZFN, CRISPR-Cas9, Meganuclease, Non-homologous end joining, Homologous recombination

#### **1. Introduction**

Recent achievements in genome editing have resulted in progress beyond any imagination decades ago. This new technology provides tools for fast and precise alterations into genome with unprecedented efficiency and specificity. For instance, a group of targeted nucleases have been successfully used to modify genomic sequences in a wide variety of cells and organisms, including mammalians.[1-4] This modification is realized by the combination of conventional gene knockout technology with genome editing, empowering the researchers to make any desired changes in a given gene or its regulatory elements to establish casual linkage between genetic variations and biological phenotypes. This new methodology completes a framework for studying gene function and regulation *in vitro* and *in vivo*.

© 2016 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

#### **1.1. Conventional gene knockout technology**

Mammalian genomes contain billions of DNA bases and are difficulty to manipulate.[5-7] Conventional genome editing is inefficient and must undergo a number of complicated and time-consuming procedures to obtain double knockouts. Because this technology had been built upon homologous recombination (HR), the targeted gene modifications occur at an extremely low frequency (1 in 106 ~ 109 cells).[8] This is the reason why this conventional approach is mainly used for producing knockout mice, for which a large amount of high quality embryonic stem cells is available. Although knockout mice have provided great insights into the fundamentals of developmental biology and physiology; the pathological roles of genes have yet to be addressed in relevant disease models or human samples.

#### **1.2. Novel genome editing technology: Targeted nucleases**

To overcome limitations of conventional gene knockout technology, a number of nucleasebased methods have been developed in the past decade.[9-12] These novel technologies exploit the ability of endonucleases to induce double-stranded DNA breaks (DSBs) and stimulate subsequent damage repair mechanisms in mammalian cells.[11, 13-17] Remarkably, this nuclease-induced DSBs not only make sequence changes at the break points but also enhance HR rate to an astonish frequency of 1-40%.[7, 18] This approach provides a simple and effective way to streamline the genome editing with the potential to generate double knockout models in both cell lines and animals. Because of this technical breakthrough, nuclease-based methods quickly gain popularity for gene editing in variety of cell types and species. To date, four major classes of targeted nucleases are prominent: meganucleases derived from microbial mobile elements, zinc finger nuclease (ZFN) based on eukaryotic transcription factor DNA binding motif, transcription activator-like effector nuclease (TALEN) derived from a plan-invasive bacterial protein and most recently the bacterial type II clustered, regularly interspaced, short palindromic repeat-associated nuclease Cas (CRISPRE-Cas).[19] These technologies are collectively termed as targeted nucleases because they all have more or less a programmable DNA binding domain for specific genome targeting. The first three nucleases all recognize specific DNA sequences through protein-DNA interaction, whereas the last CRISPRE-Cas is targeted by a short guide RNA that recognizes the unwind DNA via Watson-Crick base pairing. The purpose of the nuclease is to make localized DSBs. These DSBs will invoke powerful non-homologous end-joining as well as homologous-direct recombination repair pathway for the versatile and precise modification of complicated mammalian genome.[14, 18, 20-32]

#### **2. miRNA**

Large fraction of the genome is transcribed without protein coding potential. [33, 34] These non-coding transcripts can be broadly categorized into short and long non-coding RNAs, in which the arbitrary size delineation is at ~ 200 bases in length. Short non-coding RNAs are less 200 bases including microRNA, tRNAs and snoRNAs. An abundant class of small regulatory RNAs are termed microRNAs (miRNAs) that have been identified to play major roles in gene regulation.[35-38] Like coding mRNAs, miRNAs are transcribed by polymerase II and processed into ~ 22 nucleotide non-coding transcripts, which repress translation by binding to target sites within the 3' untranslated region of mRNA.[39-41] Recent genomic studies have discovered over 1000 miRNAs in human and mouse genome.[34] It is estimated that up to 80% of human genes are regulated by miRNAs.[36, 42] Some mRNAs are clustered in polycistronic transcripts and allow coordinated regulation, while others are expressed in a tissue-specific and developmental stage-specific manner.[35, 40] The roles of miRNAs have been extensively studied at molecular and cellular levels in a number of species including mammals.[43, 44] Furthermore, their roles in physiology and development in animals have been established by conditional knockins and knockouts.[45-47] In fact, miRNAs are expressed across genome, and many of them show spatial and temporal expression. Similar to transcription factors, miRNAs may function as master regulators of embryonic pluripotency, differentiation, and tissue/organ formation.[37] Recently, increasing evidence suggests that miRNAs are implicated in numer‐ ous disease states as well. For instance, miR-21 was found to be overexpressed in virtually all types of human cancers and thus has emerged as an important therapeutic target in cancer treatment.[45, 48, 49] Additionally, miR-21 and other miRNAs have been shown to play crucial regulatory roles in basic cell functions such as cell growth, proliferation and apoptosis.[48, 49] Their pathological roles in multiple human diseases such as autoimmune, cardiovascular and neurological disorders and obesity are emerging.[50, 51]

#### **2.1. Genome organization and biogenesis**

MicroRNA genes have a great diversity in mammalian genomes. They are located in either introns of annotated protein-coding genes or outside the context of an annotated genes.[34] Gemone analysis studies reveal that up to 42% of human miRNAs are in clusters of two or more with pairwise chromosome distance of at most 3000 nucleotides. This pattern of cluster‐ ing allows similar levels of expression and coordinated regulation. Examples of the two most famous clusters are miR-17~92 and miR-302~367. While the deletion of miR-17~92 cluster causes skeletal and growth defects, overexpression of miR-302-367 cluster enhances somatic reprogramming to pluripotent status. These clustered miRNAs appear to function together; therefore alteration of a specific member may not result in expected changes in physiology.

Although the exact mechanism of the regulation of each miRNA remains to be determined, the biogenesis of miRNA is becoming more apparent. From their gene loci, miRANs are initially transcribed by RNA polymerase II as long primary transcripts, which are processed into approximately 70-nucleotide precursors by the RNAse III enzyme Drosha in the nucleus. [52] These highly structured precursors are termed pre-miRNAs and subsequently transported from the nucleus to the cytoplasm by an Exportin-5 protein shuttler.[40] In cytoplasm, these pre-miRNAs are further cleaved by another RNase III enzyme, Dicer, resulting in imperfect miRNA:miRNA duplexes of about 18 ~ 24 bp in length.[53, 54] Although either strand of the duplex may potentially mature as a functional miRNA, only one strand is usually chosen and subsequently incorporated into the RNA-induced silence complex (RISC) where the miRNA and its mRNA target interact.[55, 56]

#### **2.2. Molecular tools for miRNA study**

Despite their fundamental importance, the exact role of miRNAs in the context of human development and disease processes remain largely unknown. This is partly due to lack of effective methods for completely abolishing their expression in human cells and diseaserelevant models. Although miRNA knockdown by short-interfering RNAs (siRNAs) provides a rapid and inexpensive tool to study most protein-coding genes, it cannot be used to reduce mature miRNAs in a sensible way at the cellular level. Other alternatives include small molecule inhibitors, antisense oligo-nucleotides, anti-miR vectors and miRNA sponges.[57, 58] However, the major limitations of these methods are transient nature of their effects and a high risk of off-target effects as well as toxicity. It is no surprise to see reports on discrepancies between the effects of miRNA inhibitors and genetic knockouts. With the completion of the genome sequencing, genome-wide gene targeting knockout of miRNA have been taken.[59] As a result, resources for the conditional ablation of miRNAs in mouse have been attempted. [60] In one study, generation of 162 miRNA targeting vectors and 46 germline-transmitted miRNA knockout mice was reported.[60] However, this homologous donor based knockout technology is expected to have tremendous hurdles to apply to other mammalian species such as rat, pig and monkeys. These larger animals are more close to humans; however, they all lack large scale culture of high quality embryonic cells to make use of this conventional technology. Nevertheless, the work in mice will provide an important basis for elucidating the physiological roles of certain miRNAs in at least one animal species.

#### **3. Targeted nucleases**

Programmable DNA-binding proteins have emerged as an exciting platform for engineering targeted nucleases for precise genome editing.[61] The key component of these engineered nucleases is the DNA recognition domain that is capable of directing the nucleases to the specific genome loci therefore generating DNA double-strand breaks (DSBs) near or at the target sites.[62] In mammalian cells, these DSBs are repaired by one of two mechanisms, nonhomologous end-joining (NHEJ) or homologous recombination (HR).[18] Repair by NHEJ is error-prone and often results in small insertions or deletion mutations, termed indels, at the break point. At the same time, DSBs can also greatly stimulate high-fidelity HDR repair mechanism in fast dividing cells. In mammalian cells, NHEJ is dominant over HR, but the frequency of latter can be substantially increased when large amount of exogenous homolo‐ gous donors are co-delivered into cells.[25]

With these strategies, various methods, including meganucleases, ZFNs, TALENs and CRISPRE-Cas have been reported for genome editing in a wide variety of mammalian species. [63-70] For these methods, a DNA endonuclease enzyme for generating DSBs is brought in place by a guide molecule. In the first three scenarios, the guider is a protein, whereas the last (CRESPRE-Cas) is a short stretch of RNA. Figure 1 illustrates the action of targeted nucleases and the binding mode of guide molecules. In general, the protein guider is more specific than the RNA guider, where degeneration is governed by hybridization mechanism. The pros and cons as well as their features of four main technologies are discussed below.

**Figure 1.** Action of the Targeted Nucleases and Their DNA Binding Mode. Genomic DNA is shown horizontally in black and double stranded, with the site of DNA cleavage indicated by arrowheads. For meganuclease, the holoen‐ zyme binds and cleaves the target DNA. For ZFN and TALENs, they function as a pair; with one zinc finger DNA binding domain (ZF) binding to the upper strand while the other ZF binding to the lower strand. Once this fused FokI enzyme (purple oval) is oriented to form a homodimer, it is activated to cut DNA. For CRISPR-Cas, the Cas9 holoen‐ zyme (orange oval) is directed to the target site by the guide RNA and cleave the DNA at the position close to the PAM motif (grey arrow).

#### **3.1. Meganuclease**

Meganucleases are endonucleases with a large DNA recognition site of 12 ~ 45 bp in length.[71] As a result, this site may occur only once in most mammalian genome.[6] Although Meganu‐ clease possesses high degree precision and low toxicity, its target range is limited. Moreover, the intertwined DNA-binding domain and nuclease domain restrict its capacity to reprogram for other targets, and the probability of finding a meganuclease for cutting a desired locus is extremely slim.[72] For example, the phiC31 integrase mediates recombination between a donor DNA of two 34 bp sequences, termed as attachment sites (att) and the other in mam‐ malian genome. In the introduction of phage integrase, a phiC31 integrase can insert a plasmid donor DNA of any size and requires no additional co-factors. Other advantages of using phiC31 integrase include non-viral delivery, sustainable transgene expression, and functions in species like bacteria, yeast, plants, frogs, chickens, mice, rats, pigs, cows, and humans.[2, 73] However, potential sites of phiC31-based genome editing are limited. In human genome, of the 106 mapped integration sites of phiC31, ~ 39% are within coding genes and ~ 61% are in the intergenic regions.[74]

#### **3.2. Zinc finger nuclease**

First described in 1996, ZFN is a chimeric protein that is composed of two distinctive domains, a programmable DNA binding domain and an endonuclease, FokI.[75] Because of its pro‐ grammability, ZFN have been successfully employed to modify almost all genome types including bacteria, plants, and animals.[1, 76, 77] In fact, ZFNs have been used for the correc‐ tion of a number of hereditary diseases such as hemophilia B, sickle cell anaemia, a-1 anti‐ trypsin deficiency and gene therapies for viral infections.[77-80] ZFN-based HIV gene therapy is already under clinical trials, because of its specificity and safety profile.[81, 82] However, the use of ZFNs has been partly hindered by the complex and time-consuming strategies to generate highly specific zinc-finger arrays with sufficient affinity and specificity. It is worthy to note that ZFNs also suffer some constrains in targeting range with about one potential target site every 500 bp.[83] It is conceivable that one may find it difficult to design ZFNs to precisely targeting a smaller gene in genome, such as miRNAs.

#### **3.3. Transcription activator-like effector nuclease**

Similar to ZFNs, TALENs are also fusion proteins that comprise of a programmable DNA binding domain with an endonuclease, FokI. The DNA-binding domain of TALEN consists of ~34 amino acid repeats, followed by a single half repeat of 20 amino acids.[84] Interestingly, the tandem repeats are nearly identical, except for two amino acid codons at positions 12 and 13, referred to as "repeat variable di-residue" (RVD). [85]Each of the four most common RVDs specifies the binding to one of the four nucleotide bases (Table 1). The natural RVD for G is NN with asparagine at positions 12 and 13. NN binds G with high affinity, but also recognizes and binds A with relative low affinity.[86] Although artificial NH or NK provides good specificity for G, the binding affinity to G is relative low as compared to NN.[86] It is worth to note that TALE proteins bind DNA sequences with an invariable base T in the first position of the target. The corresponding module is not the repeat but the cryptic sequences flanking the repeats. Because the first binding base is invariable, the DAN-binding domain can theatrically be programmed to bind any sequences starting with a "T". Taking advantage of the simplicity of the coding principle, the DNA-binding domain can be easily designed to allow binding of almost any sequences within genomes. Owing to the repetitive nature of the DNA binding domain, the assembly of the custom TALENs by direct synthesis or traditional cloning is expensive and technically challenging.[87] Realizing the potential of TALEN technology, a number of approaches for TALEN assembly have been devised to allow low ~ medium throughput, or high-throughput with automation. Fortunately, a number of Biotech compa‐ nies provide either assembly kit and/or service due to its technical difficulty (Table 2). Like ZFN, this genome editing technology has been shown to function in a wide variety of cells and organisms, including bacteria, yeast, plant, insect, zebrafish and mammal.[21, 88-92] Further‐ more, unlike meganuclease or ZFN that limit the choice of targets, TALEN can virtually bind any loci in the genome with new design that removes the 5' first "T" base constrain.

\*Note: NH and NK favor specificity rather than activity. They bind to G more specifically but with less affinity as compared to NN.

**Table 1.** TALEN DNA binding repeat and its simple code scheme

**Table 2.** Selected companies with TALEN tool kit and service

#### **3.4. CISPRE-Cas9**

Unlike ZFNs or TALENs, CRISPR-Cas needs a short RNA for target site recognition, which is mediated by Watson-Crick DNA base pairing. To form a triplex with genomic DNA, CRISPR-Cas9 also requires a NGG protospacer adjacent motif (PAM) immediately downstream of the hybrid region.

The bacterial CRISPRE-Cas9 system can be reconstituted in mammalian cells using three components: a programmable, specificity-determining CRISPR RNA (crRNA), an auxiliary trans-activating (tracrRNA), and a CRISPRE-associated endonuclease Cas9 (Cas9).[93, 94] In current applications, crRNA and tracrRAN duplexes are fused to generate a chimeric sgRAN that mimics the natural crRNA-tracrRNA hybrid. The single sgRNA has been shown to interact with the holoenzyme Cas9 to generate efficient cleavage.[95] The adaptation of the CRISPR-Cas9 by the research community have been phenomenal, and the system has been used to modify genes in most model species, including drosophila, silkworm, plant, C elegan, zebrafish, Xenopus, rat, mouse, pig, and human.[96-106] The success is mainly due to some significant advantages. First, a single protein Cas9 is required and remains the same. This means no time-consuming protein engineering is required, in contrast to that of meganuclease, ZFN or TALEN. Second, genome targeting depends on an oligonucleotides (20 ~ 30 in length), which are very easy and cheap to produce. Third, among the established programmable DNAbinding domains, CRISPR-Cas9 is most easily to facilitate genome-scale perturbations owing to its one-binding to one target. Last, this is an open system, where most of them are established and can be purchased from the non-profit distributor Addgene in Cambridge Massachusetts. However, the sequence of individual guide RNA makes difference in terms of efficacy and specificity, not all guide RNA provides the same high levels of genome editing activities.[107] Bad guide RAN can elicit off-target effects, causing cyto-toxicity to cells.[108] This can be very problematic for therapeutic applications.

The technologies are still quickly evolving, and tools to better assess both efficiency and specificity will be essential to improve the system. Choosing certain method shall be carefully balanced to serve the need of research. For example, ZFNs, TALENs and CRISPR-Cas9 are reasonable options to target most coding genes. ZFNs and TALENs shall be preferred when specificity and low-toxicity are required. For genome-wide screen or in dealing with multiple targets, CRISCR-Cas9 is more suitable because of its robustness and one molecule for one target. For small gene manipulations, TALENs may be of choice since they have broad rang access in the genome and edit precisely the target with high specificity with much less toxicity and off-target effects as compared to CRISCR-Cas9.

#### **4. Design strategy and experimental approaches**

#### **4.1. Design strategies for miRNA targeting**

One of the challenges in knocking out a miRNA is that the mature and fully functional miRNA is only ~ 22 nucleotides in length. Therefore, sequence alterations outside this 22-nucleotide region may have little or no effects on function of miRNA. To design useful targeted nucleases, sites have to be carefully chosen so that targeted nucleases are directed to the critical region of miRNA gene loci. To this end, the relative small but highly structured pre-miRNA (~70 bp) appears appropriate since it contains sequences vital for miRNA biogenesis and function.[35, 43] Within pre-miRNA, the seed regions of 5p and 3p as well as the dicer processing sites are of choice (Figure 2). Indeed, a number of reports have used this strategy for miRNA knockouts; studies have shown that small indels targeting these sites are effective to abolish miRNA expression in mammalian cells.[45, 46, 59]

Here, we use miR-21 as an example to illustrate the valid target sites that were successfully used in knockout studies. We choose miR-21 for a number of reasons. First, the human miRNA-21 was one of the first mammalian genes identified. MicroRNA-21 is located on plus strand of chromosome within a protein coding gene TMEM49. It is independently transcribed as a ~3433 nuclotides long primary transcript, where the pre-miR-21 (72 nucleotides) has a typical stem-loop (hairpin) structure similar to other miRNAs.[109] Second, the mature miRNA sequence is typically processed from the 5'-arm of the miR-21 precursor.[110] Last, miR-21 is strongly conserved and its role in physiology and pathology has been extensively explored and firmly established.[45, 48, 49, 109] Together, miR-21 provides general design strategies for miRNA gene editing.

As illustrated in Figure 2, the region corresponding to 1-8 nucleotides of the mature miRNA is the most preferred site of cleavage. This is because a small indel in this seed region is expected to abolish miRNA activity.[111] The second choice would be the adjacent Dicer processing site of the seed region. In principle, the chosen of process sites shall be preferred to the one that close to the seed region, indels involves both process site and seed sequences would most likely results in complete knockout. According to above consideration, the 5' miRNA seed region and the adjacent Dicer processing site are usually chosen for miR-21 knockout. Similarly, the 3'-arm of the miRNA precursor shall be preferred when the mature miRNA is of 3p miRNA.

**Figure 2.** MicroRNA knockout strategy and the preferred cleavage sites. The upper panel shows the stem-loop struc‐ ture of miR-21, with mature miR-21 shown in red and seed region underlined. The middle panel shows the TALEN pair and their binding sequences, separated by a 15-bp spacer. The cleavage site normally falls in the middle of the spacer, where the FokI dimerizes and makes double-strand breaks (DSBs) at the seed and/or Dicer process site. These DSBs can lead to two potential consequences as illustrated in the low panel. The predominant route 1 leads to indels via non-homologous end joining mechanism, where the alternative route 2 may lead to a precise genome editing via homologous recombination. Repair by homologous recombination can be used to bring in any desired changes at the targeted site. For knocking out miRNA-21, replacement of pre-miRNA with selection markers can facilitate the enrich‐ ment and selection of edited cells.

Here we introduce the potential sites as a general strategy for miRNA knockout utilizing the dominant NHEJ mechanisms. In fact ZFNs, TALENs and CRISPR-Cas have been used to functionally knockout a number of miRNAs in mammalian cells.[45, 59, 112-116] Among the three types of programmable nucleases, TALENs are suitable for small loci with narrow targetable regions, whereas ZFNs and CRISPR-Cas are limited by availability of binding modules and the requirement of PAM motif respectively. It is important to keep in mind; the binding sites of targeted nucleases may be different from the cleavage sites in terms of design. For ZFNs and TALENs, the cleavage site is situated in the middle of their binding sites of ZFNs or TALENs. For CRSPR-Cas, the cleavage site is within the guider RNA and close to the PAM motif (Figure 2).

#### **4.2. TALEN design and assembly**

Free online tools such as TALEN Targeter and SAPTA are available to design DNA binding domain of TALEN that is specific and has a low risk of of-target effects.[117, 118] Similar to ZFNs, TALENS s requires the design of protein pairs, which bind two optimal anchoring positions on opposite strands, usually spaced 15 ~ 25 bp apart to allow for FokI dimerization and cutting.[119] Length of DNA binding sequences may vary, typically ranging from 14 ~ 20 bp. In humans, 20 bp may offer high specificity, considering the genome size. It is worthy to note that longer domains (18 ~ 20) may decrease cell toxicity by reducing the risk of off-target effects.[120]

A number of approaches have been developed for rapid assembly of custom TALENs. With these advances, TALEN pairs can be generated easily and economically in a matter of days. [121] Most of the methods rely on the ability of type IIS restriction enzymes to assemble premade repeats into fully functional TALEN scaffold (Figure 3). A TALEN scaffold is comprised of a number of domains from the N-terminus to the C-terminus, including a nuclear localization signal, part of the N-terminal sequences for the first "T" recognition, flowed by the last half-repeat and part of C-terminal sequences fused with a nuclease FokI. Because the last binding nucleotide can be any one of the four, the TALEN scaffolds normally have four different flavours. To make the TALEN fully functional, an eukaryotic gene promoter is normally placed at 5' of TALEN coding region and a poly A signal sequences at the 3' end of TALEN.

Most assembly protocol is based on the Golden Gate method, which relies on the ability of type IIS restriction enzymes to cut outside their recognition site. Type IIS recognition sites arranged in inverse orientation at the 5' and 3' end of DNA fragment will be removed upon cleavage, slowing simultaneous restriction and ligation. The continuous re-digestion of unwanted ligation products increases the formation of the desired construct.[122, 123] As type IIS fusion sites can be designed to have different sequences, Golden Gate cloning enables directional and seamless assembly of multiple DNA fragments. Based on this principle, onestep or two-step assembly protocol or kits are developed and commercially available to allow do-it-yourself assembly of TALEN in any molecular biology laboratories.[121]

**Figure 3.** Golden Gate assembly of TALENs and molecular structure. Two-step assembly strategy can increase the cor‐ rect joining of monomers into multimers and subsequently into the TALEN scaffold. A TALEN, from its N- to C-termi‐ nus, is composed of a nuclear localization signal peptide (NLS), modified N- and C-termini flanking the assembled RVDs, and fused nuclease FokI at C-terminus. CMV (cytomegaloviral) promoter situates at the 5' of the TALEN and drives its expression in mammalian cells. The polyadenylation signal (Poly A) is to add a poly(A) tail to the TALEN mRNA.

#### **4.3. HR donor design and construction**

While it is possible to disrupt genes in mammalian genome with TALENs alone, the frequency of gene editing is typically 2 ~ 40%, averaging ~16% for mono-allelic disruptions.[87] Cells carrying bi-allelic disruption are rare and require time-consuming signal cell-derivation and subsequent screening.[5] One strategy is to combine TALENs targeting to the miRNA seed region with a homologous recombination of donor vector carrying a selectable marker.[46, 59, 114] This approach enables convenient positive selection, and the combination of NHEJ with stem-loop deletion results in efficient bi-allelic miRNA gene ablation, which can be as high as >90%.[46]Additionally, by using HR donors, endogenous loci can be potentially modified with custom sequences such as IRES-florescent proteins to allow functional assessment of endoge‐ nous miRNA expression and regulation.[124]

#### **4.4. Generation of knockout cell clones**

All targeted nucleases can be used in mammalians to create miRNA knockouts.[45, 46, 59, 113-116] Success of miRNA gene editing depends largely on the ability to deliver all the reagents efficiently to the cells. For transgenics, direct injection of DNA vector or sometime *in vitro* transcribed nuclease mRNAs into embryos is effective.[88, 125] For cultured cells, the options include plasmid DNA transfection, viral deliver, and transfection with synthetic mRNA or proteins.[126-130] Normally, donor DNAs can also supplied via plasmid format and co-delivered with nucleases using the same methodology. To deal with difficult cell types, viral transduction appears to be effective. Although lentiviral system appears to work well for both ZFNs and CRISPR-Cas, it is not suitable for TALEN delivery due to its incompatiblility with the tandem repeats of TALEN.

Following the delivery, small-scale sequence changes are introduced at the break by NHEJ. [18] These indels are typically assayed by polymerase chain reaction (PCR) amplification of the region, followed by DNA sequencing or by a gel electrophoresis assay using T7E1 endonuclease, or alternatively by high-resolution melting analysis.[131] In addition to detecting changes at genomic level, a reverse transcripts-PCR on the expression can also be performed to confirm the reduced/ablated expression of miRNA at transcript level. PCRbased genotyping can be used to make distinctions between HR and NHEJ events when donor DNA is used.[131]

#### **5. Applications of targeted nucleases for miRNA research**

Targeted nucleases are powerful genome editing tools for uncovering gene functions. Though relatively new, they have been successfully employed in a broad variety of systems and produced exciting results for miRNA research.

#### **5.1. Knockout of the miR-200 family**

To understand the biological and pathological significance of miRNAs, a talen-based knockout library for 274 highly conserved human miRNAs has been established.[59] To demonstrate the genome editing activities of the TALEN library, 66 TALEN pairs against 33 miRNA loci are selected. All TALEN pairs tested induce mutations as assessed by a mismatch-senstivie T7EI assay with a frequency above 0.5%.[132] To gain some insight of functional role miRNAs, the authors conduct detail analysis on members of the miR-200 family.

There are at least two members in the highly conserved miR-200 family, miR-141 and miR-200c. [59] Interestingly, miR-141 and miR-200c have largely indistinguishable activity and differ only in the seed region by one nucleotide. This imposes a great difficulty to use either over‐ expression or complementary inhibitor-based knockdown to investigate the potential func‐ tional divergence without complications. Using TALEN technology, however, the authors can target the seed region of the 5p strand for miR-141 but choose the Drosha processing site in the 3p strand for miR-200c, hence avoid cross-targeting.[52, 110] With this design, both single and double knockout clones were obtained and their corresponding expression of mature miRNAs was confirmed by RT-PCR.[59] Using these cell models, the authors found that miR-141 represses the expression of mRNAs that have miR-141 motif at the 3'-untranslated region. Similarly, miR-200c represses expression of mRNAs that have miR-200c motif. These data indicate the two closely related miRNAs do not cross-react notably and may control largely nonoverlapping group of genes. Together, TALEN-based method may provide unprecedented tools for miRNA research with great precision and specificity.

#### **5.2. miR-21 ablation and cancer research**

MicroRNA-21 gene knockout in the cultured human cells was achieved independently by two research groups.[45, 131, 133] The first group used a combination of TALEN pair with a HR donor. TALENs were designed to position the miR-21 seed region in the central portion of the spacer, directing the cleavage to the functionally essential miRNA motif. The HR donor construct was created corresponding to the cleavage location of the TALEN pair and carried 509-bp (5' arm) and 600-bp (3' arm) regions of homology to the miR-21 genome sequences. TALEN pair and HR donor were delivered together using transfection proto‐ col. In the case of HR events, the donor replaces the entire miR-21 precursor with two selectable marker genes (red florescent protein and puromycin resistant gene). Clonal population of cells in which an HR event occurred can be easily selected by puromycin treatment. Because NJEJ is the predominant repair mechanism induced by DSBs, selecting for HR events would most likely produce clones that harbour bi-allelic modifications, with the second allele carrying an indel in the seed region. In fact, this approach demonstrated bi-allelic miR-21 gene disruption at very high frequency of 87% in cultured HEK293 cells. [131] Analysis of three independent clones showed a total loss of miR-21expression. Phenotypical examination revealed an increase in miR-21 target gene expression, reduced cell proliferation, and alteration of global miRNA expression profiles, which is in agree‐ ment with the role of miR-21 in cancer biology.[45, 48]

#### **5.3. miRNA knockout for transgenics**

To explore the gene specific-function *in vivo*, generation of gene-targeting animal is powerful strategy. To this end, targeted nucleases have demonstrated that all nucleases are applicable for generating knockout animal and have several advantages over the conventional embryonic stem cells or nuclear transfer technologies. Recently, ZFNs, TALEN and CRISPRE-Cas were able to generate knockout animals in several model species in addition to the established spices of mice.[19] One of the strong merit is that they are free of constrain from using embryonic stem cells. The genome of the fertilized zygotes can be directly modified by injecting the targeted nuclease. Additionally, these new tools are robust enough to allow a simultaneous knockout of multiple genes, thus greatly accelerating the experimental speed and answering complicated biological questions.

Targeted knockout of miRNAs in mice by TALEN has also been reported.[113] In this study, microinjection of synthesized mRNA of TALEN was carried out in one cell stage of embryo. Embryos were allowed to develop to two-cell stage and subsequently transferred into pseudopregnant female mice. With optimized protocol, these mice were able to produce 29.6% mono-allelic offspring. Further cross-mating of the heterozygous founder mice with the wild type strain was able to produced heterozygote offspring, suggesting transmittable of the mutated miRNA.

#### **6. Prospects and challenges**

Targeted nuclease technology has become one the most powerful and versatile platforms for engineering biology. The new technology is enabling systematic interrogation of genome, including miRNAs in mammals with high precision and efficacy. It is superb over other technologies by creating null mutations that lead to complete suppression of gene expression. Furthermore, bi-allelic ablation of miRNAs in cultured somatic cells opens new avenue to study the pathological role of miRNAs in relevant disease samples. Knowledge gained from these studies is more likely lead to discovery of novel drug targets, accelerating new thera‐ peutics towards clinic application. In deed, a number of animal studies as well as clinical trials using targeted nucleases already provide encouraging results. In addition to repairing mutations underlying inherited diseases, targeted nucleases can be used to create productive mutations in tissues to combat viral infections or complex diseases such as familial hypercho‐ lesterolemia and hypertension.

Targeted nucleases can be tweaked to carry out other functions, such as modifying DNA associated histones, activating or inhibiting gene transcriptions and monitoring chromatin dynamics in living cells. It becomes increasing apparent that targeted nuclease is a versatile and common platform for elucidating gene function and epigenetic regulation.

However, many challenges still lie ahead. The most prominent issue may be the off-target effect. In this aspect, ZFNs and TALENs appear less problematic owing to the intimate interaction with their binding sites and the requirement of correct binding of two molecules. In contrast, CRISPRE-Cas tends to have higher off-target effects, because it binds DNA via base pairing. One or more mispairings may be tolerated, which may cause detrimental unintended effects.[134] Attempts to solve this issue include improvement of guide RNA scaffold, use of computational algorithms to predict off-targets, and development of high throughput method to assess unintended cuts. The challenge, though, is difficult to detect the off-target cleavage. Hence, one shall interpret the data with caution, and is aware of the potential off-target effects and its related toxicity. Secondly, although nuclease molecule may have high level of activity within cells, one may find that some tissue types are still difficult to deliver into. In this case, multiple delivery methods shall be tried, suitable one shall be optimized. Finally, for further development toward clinical application, it will be essential to thoroughly the safety and toxicity profile using a variety of mammalian models.

#### **7. Conclusion**

Advance in genome engineering technology based on the targeted nucleases are enabling the systematic interrogation of mammalian gene function. Using this system, miRNA gene sequences within mammalian genome can be easily edited with high efficacy and precision. Targeted miRNA editing will empower researcher to reveal the complex regulatory circuits governed by miRNAs and to realize, in the long-term, their full diagnostic and therapeutic potential.

#### **Author details**

Lei Yu1 , Jennifer Batara2 and Biao Lu2\*

\*Address all correspondence to<EMAIL_ADDRESS>
1 Institute for Advanced Interdisciplinary Research in Science and Technology, East China Normal University, Shanghai, China

2 Department of Bioengineering, Santa Clara University, Santa Clara, California, USA

#### **References**

## **Emerging Gene Correction Strategies for Muscular Dystrophies: Scientific Progress and Regulatory Impact**

Houria Bachtarzi and Tim Farries

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/62282

#### **Abstract**

Muscular dystrophies comprise a heterogeneous cluster of inherited muscle degenera‐ tive disorders with the common feature of progressive muscle weakness. These represent good candidates for treatment with gene-based therapies. Progress in gene transfer technologies has raised hopes for successful therapeutic restoration of mutated genes such as dystrophin in Duchenne muscular dystrophy. Delivery to enough muscle cells, however, remains a challenge for a successful gene replacement therapy. Other approaches based on exon skipping to correct mutant dystrophin's pre-mRNA splicing patterns have been tried, and partial restoration of dystrophin expression was reported in late-stage clinical trials, but full therapeutic efficacy is yet to be confirmed. The emergence of gene editing and its recent success in AIDS have opened a new therapeutic era for muscular dystrophies. This chapter will cover new gene correction strategies for muscular dystrophies and their regulatory challenges before they can become routine treatment modalities in the clinic.

**Keywords:** gene therapy, muscular dystrophies, regulatory impact, inherited muscle degenerative disorders, progressive muscle weakness

#### **1. Introduction**

Muscular dystrophy refers to a range of conditions of progressive muscle weakening generally due to genetic defects in proteins that are critical for muscle functioning. The most common form of the disease, and one of the more common of any seriously debilitating genetic disorders, is Duchenne muscular dystrophy (DMD), which is caused by the mutation of the

© 2016 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

dystrophin gene on the X chromosome and found in approximately 1 in 3600 of male births. The dystrophin gene product is a large protein with repeated elements and is involved in connecting the muscle fibres to the extracellular matrix [1]. Other proteins, mutations of which may cause forms of muscular dystrophy, include poly(A) binding protein nuclear 1 (PABPN1), myotonic dystrophy (DM) protein kinase (DMPK), or the product of the Emery-Dreifuss muscular dystrophy gene.

This chapter focuses on opportunities to develop treatments for these conditions with therapies that target the genetic defect that is the cause of the disease. The promise shown by new technologies for gene therapeutics makes this a particularly propitious time to consider the impact that such scientific progress may have on this clinically important set of conditions.

Notable among the new technological advances of relevance are the development of methods of specifically changing the sequence of the human genome, either by introducing a new gene sequence, which may repair the function of a defective gene, or by correcting the defect in the endogenous gene. Although the means to deliver genes to human cells still rely on the use of viral vectors, the methodology for the effective manufacture of these complex biologicals has developed. The approvals by the European Union (EU) of Glybera® and Imlygic® show that viral products can be manufactured to the quality standards acceptable for commercial products, with acceptable profiles of safety and efficacy. Another transformative technology is the derivation from stem cell precursors of different differentiated cell types with the potential to repair damaged or otherwise defective tissue. Putting these technologies together results in a possibility to create and manufacture to GMP standard, well-characterised patientspecific (or at least patient-compatible), genetically modified cells, a combination that may be anticipated in the near future to enable the viable therapeutic treatment of many previously intractable genetic diseases.

This chapter looks at the current status and future prospects of how the latest gene-based technologies are being applied to the alleviation, or even cure, of inherited muscular dystro‐ phies. Furthermore, we consider the challenge of translating these treatment modalities into medicines that can be approved for commercial use by the regulatory authorities, notably within the EU. To do this, developers must go beyond the scientific mechanisms of efficacy to establish how the products will be manufactured to consistent quality standards and to demonstrate that they are clinically safe.

#### **2. Muscular dystrophies**

Muscular dystrophies comprise a heterogeneous cluster of inherited muscle degenerative disorders, each caused by a distinct gene mutation. More than 30 genes have been identified, each causing a different type of muscle pathology with different patterns of muscle weakness and disease progression (**Table 1**). This includes but is not limited to Duchenne, Becker, congenital, myotonic, Emery-Dreifuss, facioscapulohumeral, oculopharyngeal, and limbgirdle muscular dystrophies. Each of these varies in terms of pattern of inheritance, age of disease onset, biochemical markers (such as creatinine kinase's upper limits), types of muscles affected, and complications at other organ sites, including cardiac and pulmonary problems. These give rise to varied symptoms, including muscle weakness and wasting (a common feature of a number of muscular dystrophies), joint stiffness, and scoliosis in addition to respiratory complications (such as chest infections and shortness of breath). Other symptoms include ankle swelling often linked with cardiomyopathy, fainting, eyelid drooping, and dysphagia. Ophthalmological symptoms, such as myopia in facioscapulohumeral muscular dystrophy and severe congenital muscular dystrophy (CMD) variants, cataracts in DM, and eyelid drooping in oculopharyngeal muscular dystrophy (OPMD), have also been reported. Hearing loss and skin lesions are also common in facioscapulohumeral muscular dystrophy and Ullrich CMD, respectively. Overall, these cause profound impairments in physical activity and quality of life [2, 3].

Emerging Gene Correction Strategies for Muscular Dystrophies: Scientific Progress and Regulatory Impact http://dx.doi.org/10.5772/62282 191

loss reported.

MDC1A = congenital muscular dystrophy with merosin deficiency; MDC1C = congenital muscular dystrophy and abnormal glycosylation of dystroglycan; FCMD = Fukuyama congenital muscular dystrophy; WWS = Walker-Warburg syndrome; X-R = X-linked recessive; AD = autosomal dominant; AR = autosomal recessive.

**Table 1.** Summary of major muscular dystrophies.

With an estimated incidence of 1 in 3600 to 6000 boys, DMD is one of the most common and severe forms of muscular dystrophies with an early-onset and progressive muscle weakness leading to the loss of ambulation by the second decade. This X-linked recessive condition is caused by mutations in the dystrophin gene (*DMD*, locus Xp21.2), which is expressed in skeletal, smooth, and cardiac muscles, and hence the pathological involvement of different organs beyond skeletal muscle weakness, including respiratory and cardiac systems. The *DMD* gene comprises 79 exons and encodes a 14-kb mRNA transcript. This gives rise to a large protein product (427 kDa), a crucial cytoskeletal protein that mediates major structural and signalling functions within muscles. Four functional units make up the dystrophin pro‐ tein. These include the actin-binding domain at the N-terminus, the rod domain consisting of 24 spectrin-like domains with four interspersing hinges, the cysteine-rich domain that mediates binding to β-dystroglycan, and the C-terminal domain mediating binding to syn‐ trophin and dystrobrevin. These binding units confer a principal structural role for dystro‐ phin facilitating its assembly with other proteins to form the dystrophin-associated glycoprotein complex (DAGC) (**Figure 1**) [1, 4, 5].

**Figure 1. The role of dystrophin in muscle physiology and disease**. (A) The Dystrophin-associated Glycoprotein Complex (DAGC) in skeletal muscle; (B) schematic illustration of exons that make up the *DMD* gene; (C) consequences of a frameshift mutation in the *DMD* gene illustrated by exon 50 deletion; and restoration of dystrophin expression with exon skipping therapeutics.

Deletions in one or more of the 79 exons that make up the *DMD* gene are common in DMD and account for approximately two thirds of the reported mutations. Other documented disruptions within the *DMD* gene include duplications (~10%), point mutations (~10%), and smaller rearrangements (15%) [6]. These lead to loss of dystrophin, which in turn destabilises the DAGC, resulting in the weakening of muscle fibre strength, increased susceptibility to stretch-induced damage, and raised intracellular calcium influx. These physiological distur‐ bances account for the underlying histopathological features often observed in skeletal and cardiac muscles from affected patients, including muscle fibre necrosis, inflammation, and substitution with fibroadipose tissue [7, 8].

No curative treatment is currently available for most muscular dystrophies, including DMD. Current approaches involve relieving symptoms, delaying disease progress, and preventing complications [2]. Although these interventions proved beneficial in the short term, none of them can provide a long-term treatment and a permanent correction of the underlying pathological features. From a molecular point of view and based on advances in the identifi‐ cation of genes behind the observed phenotypes, most of these muscle pathologies represent good candidates for treatment with gene-based therapies.

#### **3. Gene therapy for DMD**

DMD has been the main focus and the proof-of-principle model for most gene therapy strategies targeting neuromuscular disorders, over the past years, with proof-of-concept validated in preclinical and clinical settings. The first clinical trial in the neuromuscular field, in fact, involved a gene replacement approach to deliver full-length dystrophin via a nonviral method of transfer [9]. The observed low expression levels, however, at local injections sites, highlighted the need for potent gene replacement transfer systems. Of these, adeno-associated viruses (AAVs) are seen as the best available option. The efficiency of functional dystrophin expression in patients with DMD, using AAV as a vector, was assessed in a phase I clinical trial [10]. The approach was shown to be safe with no concerning side effects, although overall efficiency was compromised by the development of dystrophin-specific T-cell-mediated immune responses. This finding was complemented by observations that preexisting T-cellmediated immune responses to AAV were present, which, together with the dystrophinspecific T-cell responses, could have contributed to the low transgene expression levels detected following intramuscular injection. These early conclusions highlighted the need to circumvent immune destruction of therapeutic transgenes, by delivering unaffected homologs of dystrophin. Of these, a microutrophin-expressing recombinant AAV2/6 was shown to restore the dystrophin-glycoprotein complex and revert pathology in dystrophin (-/-)/utrophin (-/-) double-knockout mice model [11]. Delivering truncated versions of target proteins or their homologs, however, does not reconstitute a full gene replacement approach for DMD. Microdystrophin and minidystrophin transgenes often lack some crucial rod and hinge domains of full-length dystrophin, including neuronal nitric oxide synthase, syntrophin, and dystrobrevin, hence compromising maximal dystrophin functionality and membrane rigidity. This led to the engineering of triple AAV constructs using trans-splicing and hybrid methods, capable of delivering full-length dystrophin following coinfection of the tri-vectors in affected muscles *in vivo* [12, 13]. In trans-splicing, each vector acts as an independent construct holding sequential exonic sequences of human dystrophin's coding sequence. Coinjection of the vectors cause the constructs to cojoin via their inverted terminal repeats and deliver a fulllength therapeutic transgene. These results circumvent the limited packaging capacity of AAVs and offer clinical hopes for boys with severe forms of DMD, for which the reversion of phenotype to a milder disease form [Becker muscular dystrophy (BMD)] using microdystro‐ phin or minidystrophin-expressing AVV is not sufficient.

Although these gene replacement therapies are still in the early clinical phase of research and development, exon skipping-based therapeutics for DMD is progressing faster. Antisense oligonucleotides (AONs) have long been an effective alternative to dystrophin gene replace‐ ment therapy. These work by binding to the dystrophin transcript at sites that interfere with normal RNA processing, so that exons containing the mutations are bypassed, giving rise to in-frame transcripts capable of producing shorter yet functional protein products (**Figure 1C**). This was based on observations that BMD-like patients have in-frame transcripts with shorter dystrophin yet still maintain ambulation [14]. Hence, manipulating the splicing pattern of mutant dystrophin in DMD patients with an AON-based approach has the potential to alter disease phenotype from a clinically severe form of DMD to a milder BMD phenotype. For instance, the reading frame of dystrophin could be restored by blocking specific enhancers or splicing regulatory elements responsible for controlling the gene's exon recognitions. The approach was initially demonstrated *in vivo* in the *mdx* mouse model of DMD, bearing a singlepoint mutation in exon 23 that creates a stop codon with subsequent absence of dystrophin expression. Using a 2′-*O*-methyl (2′OMe) oligoribonucleotide complementary to the murine intron 22's 3′ splice site, it was possible to restore sarcolemmal expression of dystrophin in transfected myotubes *in vivo* [15]. The proof-of-concept has also been demonstrated in relevant human cell culture models derived from DMD patients bearing an exon 45 deletion, one of the most frequently deleted exons in DMD. The efficient restoration of dystrophin's coding frame was achieved by targeting splicing regulatory elements in exon 46 using a 2′OMe oligonu‐ cleotide [16]. Similarly, a successful correction was achieved through exon 51 skipping in a phase II clinical trial, where a successful restoration of both dystrophin and the DAGC was observed in patients with a deletion in exons 45 to 50 or exons 48 to 50 [8, 17], opening up great therapeutic hopes and paving the way towards late-stage clinical trials. A Biologics License Application (BLA) was submitted to the Food and Drug Administration (FDA) for approval of the antisense agent PRO051, 2′OMe (drisapersen) targeting exon 51 for DMD. However, this application was recently rejected, as a phase III study (NCT01803412) of long-term intake failed to meet its primary efficacy endpoint and also showed evidence of significant toxicity to a number of organs. Another similar antisense agent, eteplirsen, is currently under the FDA review for approval.

Although AON-based therapy for DMD has long been the prime attention of scientists and clinicians, this approach does not provide a long-term cure for DMD. Regular administrations of high doses are required to achieve a constant skipping and redirection of gene expression.

#### **4. Gene editing strategies**

With advances in human genome and increasing need to provide a simplified long-term curative approach for genetic muscle diseases, new sophisticated technologies based on gene editing have emerged. These aim to permanently correct disease phenotypes in affected individuals using site-directed endonucleases such as zinc finger nucleases (ZFNs), transcrip‐ tion activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR). These corrections occur through the generation of doublestranded DNA breaks that will eventually induce intrinsic cellular DNA repair mechanisms mostly via nonhomologous end joining (NHEJ) [18, 19].

#### *Zinc finger nucleases (ZFNs)*

ZFNs are currently the highly advanced gene editing system. Initially, these were rationally designed taking advantage of key biological properties of zinc finger transcription factors such as their DNA sequence recognition function. Each zinc finger component is composed of ~30 amino acids and is capable of recognising 3 bp of DNA. The overall structure is arranged in tandem repeats of zinc finger motifs, hence allowing the recognition of longer sequences of DNA. The nuclease activity of ZFNs is conferred through fusion to *Fok*I endonuclease's catalytic domain. This design ensures that enzymatic and subsequent DNA cleavage activities are only targeted to sites recognised by the binding domains of ZFNs. The latter consist of Cys2His2 zinc finger structures, in which a single zinc atom is surrounded by 30 amino acids and is capable of recognising 3 bp of DNA. More often, however, three to six zinc finger units are assembled, which allows the recognition of 9- to 18-bp DNA sequences. This is usually regarded as acceptable for a single locus targeting in a human genome. From a structureactivity relationship, however, full functionality is not achieved unless *Fok*I is presented in a dimerised form so as two DNA cleavage domains are dimerised around the target DNA sequence [19, 20]. In an attempt to maximise the biosafety profile of ZFNs, several methods have been employed to increase specificity. These include limiting the spacer length between the recognition sites of chimeric ZFN subunits [20]. In fact, ZFNs with shorter interdomain linkers connecting the Cys2-His2 zinc finger and the nuclease domains were shown to have a restricted activity with a 6-amino acid linker exerting the most selective activity at a target DNA site with a 6-bp spacer [21]. Another approach has been to generate obligate heterodimer nuclease domains with decreased off-target effects [22–24]. This is associated with a relatively weaker interaction between the heterodimer cleavage domains, which would necessitate stronger interactions between each monomer and the target site to achieve a site-specific cleavage while minimising cleavage at weakly bound nontarget sites. This low-affinity highavidity approach has been proposed as a plausible mechanistic approach behind the sitespecific cleavage activity of the newly engineered obligate heterodimer *Fok*I domains. An alternative to the obligate heterodimer method has been the expression of autonomous ZFN pairs, the combined expression of which was as effective as obligate heterodimer ZFN domains at inducing targeted chromosomal deletion in mammalian cells, with reduced toxic effects that are often thought to be linked with unwanted individual ZFN subunits' cross-reaction [25]. The enhancement of *Fok*I's enzymatic activity has also been reported, whereby an *in vivo* evolution-based method was employed to further increase the cleavage activity of *Fok*I. The incorporation of the enhanced domain in heterodimer ZFN structures resulted in a potent product with improved overall cleavage profile [26].

The successful correction of different genetic mutations associated with various diseases, including sickle cell anaemia [27], haemophilia [28], α1-antitrypsin deficiency [29], X-linked severe combined immunodeficiency [30], and, more recently, HIV [31, 32], has led to its application in DMD, in which precise gene editing can be achieved by deleting targeted exons from the dystrophin gene. For instance, using extended Modular Assembly (eMA)/Context-Dependent Assembly (CoDA) methods, it was possible to generate several exon 51 targeted ZFNs. Two of which demonstrated a remarkable activity with mild off-target mutagenic effects in myoblast cells from DMD patients, harbouring a deletion of exons 48 to 50. When implanted into the hind limb of immunodeficient mice, the corrected myoblasts were capable of main‐ taining dystrophin expression *in vivo*, with a correct sarcolemmal localisation [33]. This proofof-concept study has shown that the ZFN-based approach could potentially be adopted under a cell-based therapy approach for DMD, hence holding a promising faster translation into the clinic, considering that the approach is already in clinical trials for *ex vivo* cell modifications in HIV [32].

#### *Transcription activator-like effector nucleases (TALENs)*

TALENs were originally isolated from *Xanthomonas* bacteria, plant pathogens capable of employing up to 40 effector proteins to circumvent eukaryotic cell defences during a host infection [34]. These are composed of repeated motifs of 33 to 35 amino acid residues, identical with the exception of the 12th and 13th residues, which are often known as the repeat-variable di-residues. The latter play a key role in TALEN's DNA-binding specificity, whereby a different pair of amino acids would exhibit a specific binding to a corresponding nucleotide in the target sequence. For instance, the asparagine (Asn; N)-isoleucine (Ile; I) NI, the histidine (His; H)-aspartate (Asp; D) HD, the Asn-Asn NN or the Asn-lysine (Lys; K) NK, and the Asnglycine (Gly; G) NG pairs preferentially bind to adenine, cytosine, guanine, and thymine, respectively. These constitute TALENs' DNA-specific binding domains, which, like ZFNs, are conjugated to nonspecific *Fok*I cleavage domains, hence directing them to the target site for gene editing and subsequent correction of the final protein product. A recognition sequence of 14 to 20 bp, together with an appropriately spaced *Fok*I subunits separated by 12 to 19 bp, will ensure good genomic target recognition with maximal cleavage activity owing to efficient *Fok*I dimerisation [35, 36].

Although not yet being as clinically advanced as ZFNs, the lower cost and ease of production of TALENs have generated an increasing interest in the technology. Recent work in DMD has shown the feasibility of the approach using optimised exon 51 targeted TALEN-encoding plasmids transfected into myoblast cells isolated from two different patients with deletions in exons 48 to 50. A satisfactory gene correction was observed with up to 12.7% and 6.8% of alleles confirmed to have indels in the two treated patient myoblast cell lines, respectively. This correction was further correlated with a good restoration of dystrophin expression at the expected predicted size of ~412 kDa compared to its expression in the wild-type isolated myoblasts [37]. Whole exome sequencing of the successfully corrected TALEN-treated cells revealed no insertions or deletions, except at the exon 51 target locus. Further analysis using the TALE-NT 2.0 Paired Target Site Prediction web server confirmed the nature of the observed single-nucleotide variants occurring. These were in fact related to expected genomic mutations that normally occur during cell clonal expansion, as none of these showed any similarity to the employed TALEN target site with spacers of 1 to 30 bases [37].

Besides the dystrophin gene as a main target in DMD, other genes have received comparable interest as targets for TALEN-based gene editing. Of these, myostatin (MSTN) has been the centre of attention as a member of the transforming growth factor-β family. This is mostly linked to its recognised role in muscle physiology, acting as a negative regulator of skeletal muscle mass. In fact, studies have shown that mutations in the *MSTN* gene cause an increase in skeletal muscle fibre numbers and sizes, which in turn lead to muscle hypertrophy without any alarming consequences [38–40]. These observations were recently translated into thera‐ peutic interventions, whereby inhibiting the MSTN signalling pathway using pharmacological agents has shown real benefits in DMD but also in other muscle wasting pathologies such as sarcopenia and cancer cachexia [41–45]. A phase I study is currently under way to assess the safety of an anti-MSTN monoclonal antibody in advanced cancer patients with cachexia (NCT01524224). The same approach using a different pharmacological agent, BMS-986089 (Bristol-Myers Squibb), will soon be tested in a first clinical trial involving patients with DMD (NCT02515669). From a molecular therapeutic point view, *MSTN* gene editing provides a longterm control to switch off MSTN signalling in the wasted affected muscles. The approach was successfully demonstrated recently using a pair of TALEN-expressing plasmids targeting the human exon 2 locus, a highly conserved region within the coding sequence of the *MSTN* gene. Consistent with previous ZFN-related toxicity studies, the reported MSTN-TALEN targeted system was engineered using obligate heterodimers of the *Fok*I domain to minimise off-target effects often seen with homodimer variants. Initial experiments in the HEK293 cell line revealed that the mutation induced by the TALEN approach was persistent with indels still detected 1 month after transfection using the T7E1 assay, a commonly used enzyme mismatch cleavage method for detecting mutations. A similar finding was reported in four different cell lines from different species including human, bovine, and murine cells. Further investigation in primary myoblast cultures derived from a dysferlin-deficient mouse model as well as from patients with dysferlinopathy confirmed an efficiency of 10.3% to 24.6% of gene editing after treatment [46]. Although most ZFN- and TALEN-based gene editing systems for muscle diseases to date have been engineered based on NHEJ mechanisms, proof-of-concept data are now emerging on TALEN-mediated homology-directed DNA repair (HDR). Targeted integration of dysferlin, a mutation of which is associated with limb-girdle muscular dystrophy (LGMD) type 2B and Miyoshi myopathy, has been shown following cotransfection of HEK293 with MSTN targeted TALEN-expressing plasmid and a donor plasmid expressing dysferlin tagged to an enhanced cyan fluorescent protein under a cytomegalovirus (CMV) promoter [46]. Although this approach is still at its early research and development stage, the results obtained *in vitro* remain promising and could offer the basis for future TALEN-corrected myoblast transplantation therapy for a number of severe muscular dystrophies.

#### *Clustered regularly interspaced short palindromic repeats (CRISPR)*

Recent achievement in the field of gene editing has seen the emergence of CRISPR as a novel correction tool in biomedicine. CRISPRs were first described in prokaryotes as part of an RNAguided adaptive immune system to protect against foreign intrusions such as viruses and plasmids. In response to these, bacteria and archaea are programmed to incorporate short sequences of foreign DNA at one end of a repeat element, which is often referred to as the CRISPR. The integrated inserts between the repeat elements of prokaryotic CRISPR-associated (Cas) loci hence confer a permanent future mechanism of defence against the past invaders [47]. Three types of CRISPR/Cas systems (I–III) have been described, with differing structural, functional, and mechanistic characteristics. Each system, however, shares a similar sequence arrangement of short repeated units of 30 to 40 nucleotides, separated by a unique "target" specific nonrepeated sequence (a spacer) of equal length [48, 49]. A leader sequence incorpo‐ rating a promoter is often present, which initiates a unidirectional transcription of CRISPR sequences. Whereas types I and III systems are present in both bacteria and archaea, type II systems have been reported in bacteria only [50]. Type II systems that use a unique Cas protein referred to as Cas9 are of particular interest in medical application. The RNA-guided, nucleasemediated genome editing of a type II CRISPR system is mediated by Cas9, a nuclease that is directed to the target DNA site by a single-guide RNA recognising a specific locus next to the protospacer adjacent motif. This subsequently creates a double-stranded break, which could be repaired by NHEJ or by a homology-directed mechanism of repair provided that an exogenous plasmid donor is codelivered [51].

In muscular dystrophy, the CRISPR/Cas9-based technology could theoretically be employed to correct germ-line DNA prenatally in one-cell zygotes or as a postnatal treatment. The first approach was recently tested *in vivo* in *mdx* zygotes. By injecting a single-guide RNA-guided CRISPR/Cas9 together with an appropriate HDR template, Long et al. demonstrated for the first time the feasibility of the approach to correct a nonsense mutation in exon 23 of the *DMD* gene [52]. Although the CRISPR/Cas9-treated *mdx* zygotes produced mice with varying degrees of gene correction, those showing 41% of gene correction by HDR and 83% correction by NHEJ repair mechanisms demonstrated a complete disease-free phenotype with normali‐ sation of dystrophin expression at the histological level [52]. Despite showing some promising results in DMD, a successful germ-line DNA gene editing strategy would (if ever ethically permissible) require a good knowledge of the nature of mutation affecting the maternal disease carrier to allow a site-specific targeted correction. This prior knowledge, however, could be problematic in X-linked DMD in which it is estimated that one third of all mutations in the *DMD* gene arise spontaneously [53] and hence would not qualify for a Cas9 gene editing-based treatment. This would restrict the approach to the correction of well-characterised known point mutations, which only make 15% of DMD mutations. One alternative approach to consider would be a postnatal *DMD* gene correction. This was recently demonstrated using multiplexed single-guided RNAs to direct the Cas9 nuclease to mutations at exons 45 to 55 of the dystrophin gene [54], an approach that has the potential to correct more than 60% of mutations in DMD patients owing to the large hotspot mutation deletion achieved. Despite a good restoration of dystrophin expression both *in vitro* and *in vivo* following transplantation of the CRISPR/Cas9 treated DMD patient myoblasts into immunodeficient mice, the overall deletion efficiency, however, was less than that obtained following exon 51 deletion [54]. This suggests that the size of the targeted sequence is crucial in dictating Cas9-associated gene editing capacity to mediate an efficient repair by NHEJ or HDR, where a size-dependent decrease in nucleasemediated gene deletion was previously noted [55]. Although proof-of-concept has eloquently been shown in research settings, CRISPR/Cas9-based gene editing in DMD would only be envisioned in the clinic should appropriate gene delivery methods are employed. Gene transfer systems capable of directing the elements of the CRISPR/Cas9 cassette to the diseased muscles *in vivo* are needed to ensure a satisfactory *DMD* gene editing and a full therapeutic outcome. For instance, AAV has previously been shown to be a safe and an effective gene transfer system in some clinical trials for gene replacement therapy [56]. In this regard, the AAV8 and AAV9 serotypes would be useful delivery tools for CRISPR/Cas9-mediated gene editing for DMD, owing to their reported efficient gene transfer to skeletal muscles and the heart following systemic administration [57, 58], which therefore could potentially result in a robust gene correction at the key target tissues harbouring the mutation in DMD patients. Recent studies have demonstrated the efficiency of AAV as a good platform tool for CRISPR/ Cas9-mediated gene editing transfer to dystrophic muscles in DMD disease models *in vivo*. The approach was recently tested following administration by three different systemic routes, including intraperitoneal, intramuscular, and retro-orbital injections in postnatal mdx mice, and proved to be successful with an increasing dystrophin expression reported from weeks 3 to 12 after virus injection. This correlated with an overall improvement in skeletal muscle function [59]. Mutated exon 23 targeted deletion was also demonstrated to restore disease phenotype in both neonatal and adult DMD models *in vivo* by systemic and local delivery of a CRISPR/Cas9-expressing AAV [60]. A similar viral-mediated transfer of a CRISPR/Cas9 system coupled with paired guide RNAs flanking the mutated exon 23 in DMD proved that AAV is a good gene editing transfer system for local and systemic restoration of dystrophin expression in muscle cells but also myogenic stem cells in diseased muscle *in vivo* [61].

#### **5. Application in trinucleotide repeat expansion muscular dystrophies**

Proof-of-concept studies in DMD have opened new horizons in the application of gene editing in other muscular pathologies for which the correction of toxic genes is believed to address the underlying dynamic mutations often associated with triplet repeat expansion and some locus contractions disorders.

#### *Oculopharyngeal muscular dystrophy*

OPMD is an inherited autosomal dominant, slow-progressing, late-onset degenerative muscle disorder characterised by progressive eyelid drooping (ptosis), swallowing difficulties (dysphagia), and proximal limb weakness. Whereas the incidence in Europe is 1/100,000, the disease has been largely reported in the Bukhara Jew population in Israel (1/700) and the French Canadian population in Quebec (1/000) [62].

The underlying genetic defect behind OPMD is an abnormal expansion of a (GCG)n trinu‐ cleotide repeat in exon 1 of the PABPN1 gene, which leads to an expanded polyalanine tract at the N-terminal of the PABPN1 protein (12–17 repeats are often detected in mutant PABPN1 compared with only 10 repeats in the wild-type protein) [62].

From a molecular therapeutic point of view, OPMD is a good candidate for gene editing-based treatment. Previous studies have shown the importance of abrogating intranuclear inclusions (INIs) of mutant PABPN1, the main pathological hallmark of the disease, with neutralising agents capable of binding aggregated mutant proteins. Of these, intrabodies were shown to be effective in reducing mutant PABPN1-associated INIs and restoring a normal pattern of gene expression in a *Drosophila* OPMD model [63]. Although these show antiaggregate properties by tackling the disease phenotype at the protein level, they do not address the underlying molecular pathology behind it, hence necessitating a repeated administration approach to keep the disease under control. In this regard, gene editing could offer a long-term permanent therapeutic advantage. Allele-specific correction of expPABPN1 can be achieved by CRISPR/Cas9-mediated editing targeting the expanded GCG moiety. Unlike recently tested short hairpin RNA (shRNA)-based knockdown of PABPN1 [64], this gene level of correction could result in a permanent production of functional PABPN1 protein that is capable of correcting the OPMD phenotype at both histological (reduction of pathological aggregates) and molecular (abrogation of mutant PABPN1) levels. The need for an adjunct gene replace‐ ment therapy as is often the case in knockdown approaches [62] could no longer be necessary. However, thorough characterisation and mechanistic studies would need to be conducted both *in vitro* and *in vivo* to confirm CRISPR/Cas9 editing specificity towards the expanded mutant PABPN1 with no obvious off-target effects on the wild-type PABPN1 gene or indeed other genes within the treated tissues.

#### *Myotonic dystrophies*

DM is an autosomal dominant, slow-progressing inherited multisystem genetic disorder affecting the muscles (causing wasting), the eyes (leading to cataracts), and the heart (causing conduction defects). The disease is also characterised by metabolic disturbances (endocrine changes) and prolonged contraction of skeletal muscles (myotonia).

Two types of DM have been defined to date. These include DM type 1 (DM1) and DM type 2 (DM2), with DM1 being the most severe and most common form of DM. Overall incidence has been estimated at 3 to 15 per 100,000 in Europe with a higher prevalence in Iceland (1:10,000) and a reported incidence of as high as 1:500 in Quebec [65]. The disease is caused by a CTG trinucleotide expansion in the 3′-untranslated region of the DMPK gene. CTG repeats exceed‐ ing 37 are considered abnormal with larger repeats (below 400 repeats) leading to a more severe disease [66, 67]. Like most debilitating muscular pathologies, no curative treatment currently exists for DM. The use of pharmacological agents [68, 69] and antisense RNA-based therapies [70–72] have shown improvements in disease phenotype, but overall benefits on the long-term remain, in some cases, limited and full therapeutic efficacy is yet to be demonstrated in the clinic. Although still in its early infancy, gene editing for DM treatment is an appealing alternative to most pharmacological and gene therapy approaches reported to date. Correction of mutant DMPK at the gene level offers the possibility for a permanent modulation of pathological phenotypes and long-term disease control. A recent study has shown the feasibility of editing intron 9 in the DMPK gene in DM1 neural stem cells derived from human DM1 induced pluripotent stem cells (iPSCs) using a TALEN-mediated homologous recombi‐ nation-expressing cassette integrated upstream of the CTG repeats [73]. A significant reduction in nuclear RNA foci, together with restoration of normal microtubule-associated protein τ (MAPT) and muscleblind-like (MBNL) splicing patterns, were observed [73]. Although transition into clinical use may be long and difficult, this proof-of-concept *ex vivo* study offers a rationale for the genetic correction of DM1-dervied stem cells as a potential autologous cell therapy for DM1 patients in the future.

#### **6. Regulatory challenges and pathways into the clinic**

In October 29, 2015, the United Kingdom became the first country in the world to legally approve one type of a human germ-line gene modification based on mitochondrial replace‐ ment. This approach is thought to save at least 10 children each year from mitochondrial diseases by preventing maternal transfer of mutations in mitochondrial DNA to offspring (Department of Health, 2014). Human germ-line genetic engineering, however, is currently not permitted in the United Kingdom and in other European countries. Although a somatic gene editing-based approach would be less questionable from an ethical point of view, a number of safety concerns would need to be addressed from a regulatory perspective. In Emerging Gene Correction Strategies for Muscular Dystrophies: Scientific Progress and Regulatory Impact http://dx.doi.org/10.5772/62282 203

**Figure 2. Regulatory pathway for advanced therapy medicinal products**. MAA = Marketing Authorisation Applica‐ tion; BLA = Biologics License Application; EMA = European Medicines Agency; FDA = U.S. Food and Drug Adminis‐ tration; PIP = Paediatric Investigation Plan; ATMPs = Advanced Therapy Medicinal Products.

accordance with Article 2(1)(a) of Regulation (EC) No. 1394/2007 and as per the European Medicines Agency (EMA) classification, gene editing-based products, including cells modified *ex vivo*, will be regulated as advanced therapy medicinal products (ATMPs) for which a central authorisation procedure governed by the EMA would apply (**Figure 2**). This would eventually lead to a single marketing authorisation that is valid across the entire EU and the European Economic Area (EEA) countries. A number of guidelines on the requirements for product quality and preclinical and clinical studies have been issued by the EMA over the past years to facilitate the transition of promising experimental advanced therapies into the clinic.

#### *Safety considerations for gene editing*

The risk of inducing modifications at off-target genes, leading to unwanted side effects, is a major safety concern. The degree of off-target events and their clinical implications are crucial questions that need to be carefully addressed as part of a regulatory new investigational drug development plan. The primary concern is that modifications could be carcinogenic. It must also be remembered that (unless the therapy is based on a cell type that is selected and clonally expanded after modification) the off-target effects will be heterogeneous from cell to cell, so the analysis must encompass a suitably large (and potentially diverse) population of cells exposed to the modifying agent to evaluate if any cells may be adversely transformed even at low frequency. There may also be risks associated with changes that have cytotoxic effects, and if such effects are frequent in the same cells that undergo correct gene editing, the consequences could include loss of efficacy.

In this regard, rigorous quality assurance tests would need to be conducted to identify any insertions or deletions that could result from a gene editing NHEJ-based treatment. These include gene sequencing, mismatch cleavage assays based on CelI or T7 endonuclease I enzymes, and the tracking of indels by decomposition (TIDE) method. These tests could be used as quality assurance tools to determine the number and location of indels or substitution events occurring after gene editing treatment, more often based on *in silico* predictions. However, as previously noted with ZFNs, some mutations could occur near cryptic off-target sites that are not predictable *in silico* [74]. Care should also be taken when validating the assays employed for product characterisation purposes. This follows from previous observations whereby initial analysis by whole genome sequencing (WGS) in gene-corrected human iPSCs (hiPSCs) revealed a large number of indels, of which some were confirmed to be false positive [75]. This highlights some limitations of WGS and calls for a multitesting approach employing different analytical methodologies.

Although current assays would be valuable in identifying the location of suspected off-target mutations that could arise from NHEJ or HDR-based gene editing treatments, they do not, however, provide precise information on the ability of these mutations to cause carcinogenicity in the long term. Hence, additional functional studies are required to assess the significance of these off-target events at the molecular and cellular levels. For instance, an unwanted insertion in the middle of an essential gene sequence could have dramatic consequences. One should have enough knowledge on the biological and physiological functions of the affected gene to draw an informed decision on the overall safety of the proposed approach. Similarly, an off-target insertion into an enhancer or a repressor region would disturb genetic homeo‐ stasis resulting in an unwanted down-regulation or up-regulation of genes that come under the affected promoter. This could subsequently interrupt normal cellular activity with potential alteration of phenotype at histological and physiological levels. Although not all mutations could affect cellular proliferation, an unwanted "indel" mutation occurring within genes known to regulate cell replication or involved in programmed cell death raise an alarming concern from a safety point of view. This is due to potential risks arising from a compromised cellular viability leading to severe toxicity in healthy tissues or an uncontrolled replication that could eventually lead to tumour formation. These potentially serious conse‐ quences warrant careful considerations during early product development stages; hence, basic biological studies to further characterise the sites at which these mutations have occurred should be conducted as ad hoc validation studies to rule out any unwanted insertional mutagenesis and/or tumourigenic consequences following a gene editing-based treatment.

Although clinical safety experience with biological therapies based on gene editing is lacking, current product development and regulatory strategies should also draw from past and available human clinical trial data on retroviral and lentiviral vectors' integration sites. A key safety parameter would be to satisfy the regulatory bodies that those detected off-target effects have been thoroughly characterised and occur at "low risk" sites unlikely to cause insertional oncogenesis following integration of foreign DNA. In fact, not all events at off-target genomic sites would result in serious side effects. For instance, those occurring at "extragenic" sites distant from essential gene regulatory sequences are less likely to cause harm than those affecting "intragenic" sites. Modifications to sites that do not fall within a gene transcription unit as well as those situated more than 50 kb away from the 5' end of any gene and more than 300 kb from genes linked with cancer or microRNA sites are generally considered safer. Genomic sites not within ultraconserved regions and outside long noncoding RNAs would also represent low-risk sites in the event of off-target DNA integration [19, 76].

Furthermore, validation studies should not only focus on the affected off-target sites but also neighbouring sites by measuring the effect of these mutations on neighbouring gene expres‐ sion. These studies should ideally be validated *in vivo*. The choice of appropriate animal models, however, remains a challenge and care should be taken when interpreting data [19, 76, 77].

#### *Validation and choice of functional studies for assessment of toxicity*

Although basic biological studies are valuable in assessing the extent of genome modification as a whole and increasing our understanding of different alterations taking place including those that are unlikely to result in clinical toxicity, they do not fill all the regulatory require‐ ments gaps. Functional toxicity studies would, therefore, need to be conducted in parallel early during the preclinical stage of product development. Two paramount questions would need to be carefully addressed to satisfy regulators' concerns on serious toxicity issues that are specific to genome editing-based medicines. These include cytotoxicity and genotoxicity. Tumourigenicity is the most important safety consequence to consider, but germ-line effects are also not expected to be permissible. Cytotoxicity may be caused by (i) the vector itself (including expression of viral vector antigens and potential persistence of the vector in cells), (ii) gene editing machinery (and any persistence thereof, especially DNase), (iii) off-target gene editing, or (iv) on target gene editing (possible if the editing is either not accurately restoring the wild-type or is generating new antigenicity and/or genetic instability).

Eloquent studies and approaches would have to be designed to address these issues. At preclinical stage, this could include viability assays based on GFP-positive (GFP+ ) cells, whereby the treated cells are cotransfected with the tested nuclease-expressing construct and a GFP-expressing plasmid. This would allow investigators to track and quantify any observed decline in the GFP+ cell population as a result of a nuclease-related toxicity. The approach would also be valuable in dose escalation studies when deciding on optimal dosage adminis‐ tration for subsequent first-in-human clinical studies. Monitoring of clonal changes has also been used as an informative way to assess genotoxicity *in vitro* in cells pretagged with unique short sequence identifiers to allow tracking of changes in starting clone dynamics over a period of time [78]. Similarly, other approaches have relied on fluorescently tagged cell cycle indica‐ tors in cell lines such as HeLa FUCCI cells as a complementary *in vitro* validation method to assess the genotoxicity effects of genome editing at the cell cycle level [79].

However, it is important to bear in mind that measuring cytotoxicity and genotoxicity using approaches based on reporter genes and tagging systems is restricted to preclinical assess‐ ments and cannot be employed at later stages of product development when assessing potential toxicity in human clinical trials. Hence, the need for clinical toxicity assays that are fit for purpose should not be neglected. Furthermore, correlation of these assays with clinical outcomes is yet to be demonstrated. For this reason, all functional toxicity studies conducted *in vitro* should use cellular models and gene transfer methods that are similar to those intended to be applied in clinical settings to fulfil some of the safety regulatory requirements.

#### *Vector considerations for in vivo and ex vivo gene editing*

Potential cytotoxic effects do arise not only from on-target and off-target effects but also from the employed gene transfer system itself. In this regard, the type of vector (viral or nonviral) and the gene (nuclease) transfer approach (direct *in vivo* delivery versus *ex vivo*) adopted will have to be implemented earlier during product development and considered as part of the constructed regulatory strategy for the nuclease-based medicine under consideration.

For severe neuromuscular disorders in which there is a widespread distribution of affected muscles, AAV vectors and in particular serotypes 8 and 9 are seen as ideal systems for gene editing-based nuclease transfer to diseased muscle tissues owing to their relatively high tropism for skeletal muscle cells and the heart [57, 58] as well as their documented safety profile in the clinic following recent approval of Glybera® in Europe [80]. From a product development point of view, this can be achieved through an HDR-based gene editing construct packaged within a recombinant AAV. Although technically this is achievable with ZFNs due to a relatively small monomer insert size of ~ 1.2 kb, this might not be the case for Cas9 nucleases. With an insert size of ~4.1 kb, this could be an issue when it comes to packaging a bicistronic construct harbouring a donor DNA template [81], considering the limited packaging capacity of rAAVs. Taking into account the EMA's guidelines on quality, nonclinical and clinical issues related to the development of recombinant adeno-associated viral vectors (EMEA/CHMP/ GTWP/587488/2007 Rev. 1) [82], caution should be taken during the manufacturing stage of the vector to avoid the possibility of producing AAV particles whose packaged DNA is greater than that of wild type virus. Technically, this could be overcome by splitting the Cas9 gene between two vectors. However, this markedly increases the developmental and regulatory complexity, with each product warranting a full characterisation and a thorough assessment of approaches employed during manufacturing, quality control, and preclinical evaluation (including choice of animal models for testing, vector persistence and tropism, reactivation of virus infection, and germ-line transmission) in addition to clinical studies. The latter would have to be based on a data-driven dose selection for each vector used and show a comprehen‐ sive picture of virus biodistribution and shedding, immunogenicity profile, and germ-line transmission considering the permanent gene modification achieved with gene editing-based nucleases and the risks associated if the virus expressing these genes accidentally infect germline cells. A long-term follow-up is therefore highly recommended and should not be neglected as part of a complete regulatory plan for a gene editing-expressing AAV vector.

Similarly, the use of genetically modified and nonmodified cells including myoblast cells and myogenic stem cells for the treatment of monogenic inherited neuromuscular diseases has been well validated in preclinical and clinical studies [83–86] and it is anticipated that these therapies would become normal treatment modalities in the clinic in the future.

Although gene edited-based cell therapies for HIV and leukaemia are seeing a rapid positive progress in clinical trials and are rapidly extending to other debilitating diseases, a smooth and safe progress of these life-changing advanced therapies in the market would require fulfilments of a number of criteria from a regulatory point of view. The ultimate goal is to ensure that these therapies would reach the severely affected individuals at minimal health and safety risks with regards to the treated patients themselves as well as third parties and the environment in large. According to the EMA's guidelines, a multifactorial risk assessment approach needs to be considered taking into account the origin of cells involved, the type of vector employed during the genetic modification procedure, the manufacturing process, the noncellular components used as part of the formulated product, and the intended specific therapeutic use of the final product.

Lentiviral vectors have long been the vector of choice for the *ex vivo* engineering of stem cells and muscle progenitor cells for the treatment of muscular dystrophies [87–89]. This adds an additional layer of regulatory requirements from quality, nonclinical and clinical perspectives. The manufacture of viral vectors to the GMP quality standards required for an approvable medicinal product is expensive and often inefficient, but developers are helped by the guidance issued in these areas by the EMA (for example, "Guideline on the quality, nonclinical and clinical aspects of gene therapy medicinal products (EMA/CAT/80183/2014)") [90], as well as the monograph of the European Pharmacopoeia (Ph. Eur. 5.14). A thorough product characterisation is required as part of this process that would include different but comple‐ mentary methods based on molecular, biological, and immunological assays with the aim of assuring the identity, purity, and potency of the produced genetically modified cells as a final product. Major considerations for viral vectors used *in vivo* are the biodistribution, potential for generation and/or shedding of infective virus, potential for germ-line modification, and the impact of interaction with the recipient's immune status. Clinical use of viral vectors and genetically modified cells should also comply with applicable regulations for genetically modified organisms.

Complexities arise from the genetic manipulation (gene correction) as well as the differentia‐ tion status and capacity of the modified cells, which could result in a mosaic cell population. Furthermore, intrinsic variations between cells as a result of donor differences give rise to massive batch variations in the final product. From a regulatory perspective and in line with the EMA's guidelines, nonclinical and clinical studies need to be conducted with cell medicinal products that are well characterised. The manufacturing process needs to be robust and quality control focused capable of maintaining consistency and reproducibility of the final cell-based product. For this, all starting materials need to be well defined and carefully documented. For treatment of muscle diseases, it is often reasonable to administer cell-based products that are in a differentiated state, which may pose less tumourigenic risks. However, one cannot exclude the existence of a subpopulation of cells in an undifferentiated proliferative state. For iPSCs, for example, it is empirical to conduct additional testing of cell transformation and tumour formation during the early manufacturing stage of the product as a precaution measure. This would often need to be combined with the selection of appropriate markers during critical manufacturing steps for assuring a defined stage of differentiation that is intended for therapeutic use.

For any cell-based product that has undergone a substantial *ex vivo* manipulation, a robust process validation process is therefore paramount and should include a combination of genetic stability, tumourigenicity, and phenotypic profile assessments of both wanted and unwanted cell populations at all critical stages of manufacturing to ensure safety requirements are met. The mosaic nature of cell-based medicinal products often complicate their identity and purityrelated characterisation. Most often, it is reasonable to accept that purity does not always equate homogeneity. Truly selective markers that could accurately map and distinguish different cell types and differentiation stages are yet to be identified, which often render product characterisation a challenging task for most developers. Nevertheless, one should not underestimate the importance of a thorough demonstration of product consistency as a minimum requirement for characterisation purposes.

The ability to track any cell-based therapy following administration in patients is crucial for clinical monitoring purposes. However, limitations in current medical methodologies do not allow a full biodistribution profile to be drawn from human studies. Hence, the importance of thoroughly designed biodistribution studies in nonclinical models should take into account the multistep biodistribution characteristics of cell products, including migration, niche, engraftment, differentiation, and persistence, together with reliable *in vivo* tracking methods such as the use of marker genes or labelled cells. Current European regulatory requirements do not give exemptions when the risk profile of the cell-based product under investigation is subject to a safety concern or when its route of administration (such as intravenous delivery) warrants a special attention. For this, noninvasive methods based on clinically accepted tracers should be considered and their use should be justified when conducting biodistribution studies in human trials.

#### **7. Conclusion**

For diseases caused by genetic defects, means to correct the defect are attractive routes to cure the disease at its source. Medical scientists working in the field of muscular dystrophies are therefore excited by the opportunities that recent developments in gene therapeutics bring to the field. Technologies do now exist for manipulating the human genome in *ex vivo* cultured cells with considerable specificity. However, there are further technical challenges to solve before such technologies translate into viable medical treatments for affected individuals. The medical need is evident from the severity and incidence of these conditions, which are frequently severely debilitating. It should also be noted that muscular dystrophies are not a single disease but cover a range of conditions caused by different mutations, and it is to be expected that, for specific gene-based therapeutics, a different medicinal product will generally be required according to the different mutation that has caused the disease.

The increasing number of investigational medicinal products entering clinical trials involving patients with DMD and BMD has prompted the EMA to publish its guidelines on the clinical investigation of medicinal products for the treatment of DMD and BMD (EMA/CHMP/ 236981/2011, Corr. 1) [91], expected to come into effect in July 2016. This gives a general guidance to be taken into account during the clinical development and evaluation of currently investigated therapies for DMD and BMD [91].

As described in this chapter, major recent advances for the enablement of gene-based treat‐ ments of muscular dystrophies include the following:

#### **Author details**

Houria Bachtarzi\* and Tim Farries

\*Address all correspondence to<EMAIL_ADDRESS>
ERA Consulting (UK) Ltd. (European Regulatory Affairs), London Gas Museum, Twelvetrees Crescent, London, E3 3JG, United Kingdom

#### **References**

## *Edited by Michael S.D. Kormann*

Site-specific endonucleases create double-strand breaks within the genome and can be targeted to literally any genetic mutation. Together with a repair template, a correction of the defective locus becomes possible. This book offers insight into the modern tools of genome editing, their hurdles and their huge potential. A new era of in vivo genetic engineering has begun.

Photo by Sergey Nivens / AdobeStock

Modern Tools for Genetic Engineering

Modern Tools for

Genetic Engineering

*Edited by Michael S.D. Kormann*