Browsing by Subject "CRISPR/Cas9"

Duchenne muscular dystrophy (DMD) is a muscle wasting disease that results from a lack of dystrophin protein, which is an essential musculoskeletal protein. Patients are typically non-ambulatory by their teenage years and suffer prematurely fatal respiratory and/or cardiac complications by the third decade of life. DMD is caused by deleterious mutations in the dystrophin gene, which creates an out-of-frame shift leading to a lack of dystrophin protein and manifestation of DMD. Although scientists have had an understanding of the genetic basis of DMD for decades there has been only modest advancement in improving quality of life for these patients.

Becker muscular dystrophy (BMD) is an allelic disease; BMD is also caused by mutations in the dystrophin gene, although these mutations maintain the translational reading frame and thus a truncated, partially functional dystrophin protein is created. BMD patients have a wide range of symptoms, but BMD typically has much less severe symptoms than DMD. Thus, a common approach to creating a therapy for DMD is to shift the DMD genotype to a BMD genotype. One therapy targeting the genetic cause of the DMD by shifting the messenger RNA (mRNA) and thus protein product to one of BMD has been conditionally approved by the US Food and Drug Administration (FDA), but the treatment is transient and thus far has not demonstrated reliable clinical benefit. DMD presents some unique challenges for developing gene therapies. First, the full-length gene is so large that exogenous delivery in size restricted viral vectors is not an option. Second, popular strategies being explored are transient and would require lifelong administration. The work presented in this dissertation utilized gene editing technology. Building on prior proof-of-concept studies, we show a CRISPR-Cas9 system utilizing Staphylococcus aureus Cas9 (SaCas9) can be used to create permanent changes to the dystrophin gene. This technique overcomes the main challenges presented, as editing the native locus does not require delivering the gene exogenously, and CRISPR-Cas9 mediated DNA double stranded breaks result in permanent changes of the genome. Here we further the proof-of-concept body of work for utilizing CRISPR-Cas9 to treat DMD by targeting exon 51 for excision in a humanized mouse model.

Initially, we recognized the need for a relevant small animal model. A majority of DMD in vivo work is done in the mdx mouse or variants of the mdx mouse, which contains a mutated mouse dystrophin gene such that it does not produce dystrophin protein and displays a mild dystrophic phenotype. While this is a useful research tool, in order to move genome editing closer to the clinic we need to be able to test guide RNAs (gRNAs) that target the human dystrophin gene in a small animal model. As the gRNAs target exact sequences of the genome they must be designed to the human DMD gene. These human DMD targeting gRNAs would not match the mouse Dmd gene, and thus there was a clear need for a preclinical humanized small animal model of DMD. We obtained an hDMD/mdx mouse that contains the full-length, healthy, wild type human DMD gene on mouse chromosome 5. Although this mouse has the human DMD gene, it is ultimately a healthy mouse. Thus, we utilized Streptococcus pyogenes CRISPR-Cas9 (SpCas9) to excise exon 52 of the human DMD gene in the mouse zygotes. We identified a founder mouse that lacked exon 52 in the genomic DNA (gDNA) and bred that mouse with the mdx mouse line. Thus, using genome editing, we created the hDMDΔ52/mdx mouse, which lacks both human and mouse dystrophin protein expression. We confirmed this biochemically by sequencing the gDNA to ensure lack of exon 52 between the gRNA targeted sites, lack of exon 52 in the cDNA, and lack of dystrophin protein by both immunohistochemistry (IHC) staining and Western blot. The hDMDΔ52/mdx mouse also displayed a mild dystrophic phenotype compared to its healthy counterpart, the hDMD/mdx mouse. We have characterized this hDMDΔ52/mdx mouse and shown it lacks dystrophin and has a mild dystrophic phenotype, and this mouse will be a meaningful tool for testing potential DMD therapies.

Next, we created a CRISPR-SaCas9 system that would target the human DMD gene for exon 51 excision. While our lab has previously shown efficacy of this method utilizing SpCas9, we switched to the smaller SaCas9 in order to better accommodate the small packaging limit of adeno-associated virus (AAV). gRNAs were designed to target conserved regions in the intronic area flanking exon 51 of dystrophin in both humans and rhesus macaques. gRNAs were tested individually for on-target activity in HEK293T cells and those with on-target activity were assessed for off-target activity in silico. One gRNA upstream of human dystrophin exon 51 and one gRNA downstream of exon 51 were selected based on distance from the exon, percent modification measured by the Surveyor nuclease assay, and potential for off-target activity in humans and rhesus macaques. Those chosen two gRNAs were tested as a deletion pair in both HEK293T cells and immortalized myoblasts from a DMD patient, lacking exons 48 through exon 50 that is correctable by removing exon 51, and shown to create the desired deletion. Currently there is a lack of rules about what makes an effective gRNA, and in particular even the length of the gRNA protospacer sequence for SaCas9 can have effects on on-target activity. Thus, the two chosen gRNAs were tested with protospacer lengths varying from 19 to 23 base pairs (bp) both individually and as deletion pairs in HEK293T cells. The most effective on-target pair was with both gRNA protospacer sequences at 23 bp long. These 23 bp length gRNAs were re-tested in HEK293T cells and DMD patient immortalized myoblasts and shown to be effective at creating deletions in the genome, having that edit carry over in the mRNA of differentiated myoblasts resulting in the loss of exon 51 and the junction of exon 47 to exon 52 when Sanger sequenced, and restored dystrophin protein expression in the differentiated myoblasts by Western blot. Off-target sequences of these 23 bp length protospacers were assessed in silico and ten of the predicted off-target sites for each gRNA were tested in vitro in HEK293T cells by deep sequencing. Although the upstream gRNA did have two off-target sites that had notable small insertion or deletion (indel) rates measured by treated gDNA/untreated gDNA, ultimately all measurable off-target activity was at least two orders of magnitude lower than the on-target rate of indel formation.

Finally, we created a CRISPR-SaCas9 system with gRNAs that target human DMD for exon 51 removal, and these exact gRNAs tested in vitro were tested in vivo in our previously characterized hDMDΔ52/mdx mouse. Initially we did a small proof-of-concept study by packaging our system in AAV8 and performed local injections into the tibialis anterior (TA) muscle of adult hDMDΔ52/mdx mice. 8 weeks after treatment the TA was analyzed. We noted deletion of exon 51 between the gRNA targeted sites in the gDNA, as well as dystrophin protein restoration by IHC and Western blot. While promising, DMD is a systemic disease that affects all skeletal and cardiac muscles. Thus, we next delivered our CRISPR-SaCas9 system using AAV9 systemically by tail vein injections in adult hDMDΔ52/mdx mice or temporal vein injections in neonatal hDMDΔ52/mdx mice. At 16 weeks of age mice were sacrificed for biochemical analysis. Deep sequencing of gDNA at each gRNA target site showed measurable indel formation above the limit of detection in all tissues assayed in mice treated as both adults and neonates. There were a few trends that emerged in this data and hold true throughout analysis of on-target editing: the upstream gRNA is generally more effective at on-target activity than the downstream gRNA, the mice treated as neonates show more on-target activity than mice treated as adults, and there is much more on-target activity in the heart than in the skeletal muscles. Indels are a measure of on-target activity, but we delivered a system to create a deletion and not just individual cuts. Thus, gDNA from the heart and TA of mice treated as adults was assayed by linear amplification sequencing, which revealed approximately 4% deletions of exon 51 in gDNA from the heart and about 1% deletions of exon 51 in gDNA from the TA. Through this method we are also investigated inversions of the targeted sequence and AAV integrations into the targeted cut site, both of which were much more prominently present in the heart gDNA than the TA gDNA. Confident we were able to edit the genome at low, although measurable, levels, we examined changes in mRNA. In the mRNA from hearts of both mice treated as adults and neonates we see clear deletions of exon 51 by endpoint polymerase chain reaction (PCR). Sanger sequencing the deletion band revealed the exact junction of exon 50 to exon 53 as expected. We performed quantitative droplet digital PCR (ddPCR) on cDNA from the heart, TA, diaphragm, and gastrocnemius, and similar to the indel formation we saw the highest amount of exon 51 deletions in the heart cDNA at about 20% in both mice treated as adults and neonates. The deletions in skeletal muscles varied from about 0.15% to about 1.5% and were all measurable above the limit of detection as defined by the average of samples from untreated mice. Lastly, we examined dystrophin protein expression. By Western blot we saw mouse to mouse variability in intensity, but largely some degree of dystrophin protein expression restoration in protein extracted from hearts and gastrocnemius muscles from mice treated as both adults and neonates, although qualitatively the mice treated as neonates have more dystrophin protein expression than those treated as adults. IHC on hearts and TA muscle sections similarly showed variable but nonetheless present dystrophin protein expression restoration in both mice treated as adults and neonates. Consistent with prior data, we saw more dystrophin expression in the heart than in the TA, and this difference is exacerbated in the mice treated as adults.

In sum, the objective of this dissertation was to create a clinically relevant CRISPR-SaCas9 system and test it in vitro and in vivo in a diseased humanized mouse model. This work is an incremental step to propel forward methods to permanently correct the dystrophin gene by gene editing technology to treat DMD. We created a useful mouse model for the field to test preclinical therapies in vivo and make the most of the rapidly advancing gene editing tools. Collectively this work is significant in extending early proof-of-principle studies to a translational strategy for gene editing as a potential treatment for DMD.

Duchenne muscular dystrophy (DMD) is a severe hereditary disorder caused by a loss of dystrophin, an essential musculoskeletal protein. Decades of promising research have yielded only modest gains in survival and quality of life for these patients and there have been no approved gene therapies for DMD to date. There are two significant hurdles to creating effective gene therapies for DMD; it is difficult to deliver a replacement dystrophin gene due to its large size and current strategies to restore the native dystrophin gene likely require life-long administration of a gene-modifying drug. This thesis presents a novel method to address these challenges through restoring dystrophin expression by genetically correcting the native dystrophin gene using engineered nucleases that target one or more exons in a mutational hotspot in exons 45-55 of the dystrophin gene. Importantly, this hotspot mutational region collectively represents approximately 62% of all DMD mutations. In this work, we utilize various engineered nuclease platforms to create genetic modifications that can correct a variety of DMD patient mutations.

Initially, we demonstrate that genome editing can efficiently correct the dystrophin reading frame and restore protein expression by introducing micro-frameshifts in exon 51, which is adjacent to a hotspot mutational region in the dystrophin gene. Transcription activator-like effector nucleases (TALENs) were engineered to mediate highly efficient gene editing after introducing a single TALEN pair targeted to exon 51 of the dystrophin gene. This led to restoration of dystrophin protein expression in cells from DMD patients, including skeletal myoblasts and dermal fibroblasts that were reprogrammed to the myogenic lineage by MyoD. We show that our engineered TALENs have minimal cytotoxicity and exome sequencing of cells with targeted modifications of the dystrophin locus showed no TALEN-mediated off-target changes to the protein coding regions of the genome, as predicted by in silico target site analysis.

In an alternative approach, we capitalized on the recent advances in genome editing to generate permanent exclusion of exons by using zinc-finger nucleases (ZFNs) to selectively remove sequences important in specific exon recognition. This strategy has the advantage of creating predictable frame restoration and protein expression, although it relies on simultaneous nuclease activity to generate genomic deletions. ZFNs were designed to remove essential splicing sequences in exon 51 of the dystrophin gene and thereby exclude exon 51 from the resulting dystrophin transcript, a method that can potentially restore the dystrophin reading frame in up to 13% of DMD patients. Nucleases were assembled by extended modular assembly and context-dependent assembly methods and screened for activity in human cells. Selected ZFNs had moderate observable cytotoxicity and one ZFN showed off-target activity at two chromosomal loci. Two active ZFN pairs flanking the exon 51 splice acceptor site were transfected into DMD patient cells and a clonal population was isolated with this region deleted from the genome. Deletion of the genomic sequence containing the splice acceptor resulted in the loss of exon 51 from the dystrophin mRNA transcript and restoration of dystrophin expression in vitro. Furthermore, transplantation of corrected cells into the hind limb of immunodeficient mice resulted in efficient human dystrophin expression localized to the sarcolemma.

Finally, we exploited the increased versatility, efficiency, and multiplexing capabilities of the CRISPR/Cas9 system to enable a variety of otherwise challenging gene correction strategies for DMD. Single or multiplexed sgRNAs were designed to restore the dystrophin reading frame by targeting the mutational hotspot at exons 45-55 and introducing either intraexonic small insertions and deletions, or large deletions of one or more exons. Significantly, we generated a large deletion of 336 kb across the entire exon 45-55 region that is applicable to correction of approximately 62% of DMD patient mutations. We show that, for selected sgRNAs, CRISPR/Cas9 gene editing displays minimal cytotoxicity and limited aberrant mutagenesis at off-target chromosomal loci. Following treatment with Cas9 nuclease and one or more sgRNAs, dystrophin expression was restored in Duchenne patient muscle cells in vitro. Human dystrophin was detected in vivo following transplantation of genetically corrected patient cells into immunodeficient mice.

In summary, the objective of this work was to develop methods to genetically correct the native dystrophin as a potential therapy for DMD. These studies integrate the rapid advances in gene editing technologies to create targeted frameshifts that restore the dystrophin gene around patient mutations in non-essential coding regions. Collectively, this thesis presents several gene editing methods that can correct patient mutations by modification of specific exons or by deletion of one or more exons that results in restoration of the dystrophin reading frame. Importantly, the gene correction methods described here are compatible with leading cell-based therapies and in vivo gene delivery strategies for DMD, providing an avenue towards a cure for this devastating disease.

Skeletal muscle has the innate ability to robustly regenerate in a highly orchestrated fashion that is initiated by satellite cells, the resident stem cell population. These cells are defined by their uniform expression of the transcription factor, PAX7, which plays a key role in myogenesis through specification and maintenance of satellite cells, as well as regulation of myogenic differentiation. In conditions of skeletal muscle wasting such as cachexia, sarcopenia, and muscular dystrophies, the deterioration of muscle overwhelms the regenerative capabilities of satellite cells, which are believed to undergo early senescence due to exhaustive proliferation. There is significant potential for harnessing satellite cells for gene and cell therapies for such diseases; however, satellite cell specification and regulation is still poorly understood.

The CRISPR/Cas9 system has been established as a multifaceted tool that can be used as a platform for a variety of applications, including sequence-specific genome and epigenome editing for cell differentiation and treatment of genetic diseases. The objective of my research proposal was to use CRISPR/Cas9-based genome engineering technologies toward applications for skeletal muscle regeneration. First, I used a CRISPR/Cas9-based transcriptional activator to direct differentiation of human pluripotent stem cells into functional skeletal muscle progenitor cells. Next, I conducted a high-throughput CRISPR activation screen to identify novel upstream regulators of myogenic progenitor cell differentiation. Lastly, I demonstrated that satellite cells can be targeted in vivo with AAV and subsequently gene-edited to correct the dystrophin reading frame in a mouse model for Duchenne muscular dystrophy. Together, this work provides novel contributions to the field of satellite cell biology and highlights the utility of CRISPR/Cas9 genome engineering in stem cells for skeletal muscle regeneration.

Glycogen storage disease type Ia (GSD Ia) is an autosomal recessive metabolicdisorder caused deficiency of glucose-6-phosphatase (G6Pase) resulting from pathogenic variants in the G6PC gene. G6Pase catalyzes the hydrolysis of glucose-6-phosphate to release glucose which can then enter the bloodstream. GSD Ia patients have excess glycogen accumulation mainly in the liver and kidneys and suffer from life threatening hypoglycemia. The current treatment for GSD Ia is dietary therapy that requires patients to frequently consume uncooked cornstarch on a strict schedule. Cornstarch provides a complex carbohydrate that slowly releases glucose to prevent hypoglycemia. This treatment fails to prevent long-term complications associated with GSD Ia including renal failure and the development of hepatocellular adenomas and carcinomas. This lab and others have developed adeno-associated virus (AAV) vector based gene therapies to deliver and therapeutic G6PC transgene to affected tissues in GSD Ia animal models. However, the therapeutic effect is limited as AAV vector genomes are rapidly lost and the biochemical correction declines. Currently no treatment for GSD Ia exists that provides stable, robust expression of G6Pase that can clear glycogen and prevent hypoglycemia. This study employed a novel genome editing approach designed to insert the therapeutic G6PC into the endogenous locus in canine and murine models of GSD Ia. Integration of the transgene into the genome will promote stable expression of G6Pase and prevent the decline of vector genomes and the therapeutic benefit. This genome editing approach utilizes the CRISPR/Cas9 system to generated targeted double stranded DNA breaks at a targeted site in the genome. The G6PC transgene is present in a Donor template with homology to the DNA break to drive homology directed repair (HDR) resulting in the integration of the transgene into the genome. In a canine model of GSD Ia, editing and incorporation of the transgene was achieved in both adult dogs and puppies. Up to 1.0% of alleles were edited in the dog livers and contained the transgene. G6Pase production from the integrated transgene was detected, which correlated with prevention of hypoglycemia during fasting. This demonstrated genome editing in the liver of a large animal model for an inherited metabolic disorder using HDR to insert a therapeutic transgene. A subsequent study in GSD Ia mice also showed incorporation of a G6PC transgene in the mouse genome. Mice were treated with either the Donor transgene vector alone or with both the Donor and a CRISPR/Cas9 vector to assess to role of nuclease activity on integration. Mice treated with both vectors demonstrated improved blood glucose concentrations during fasting, decreased liver glycogen, and increased vector genome copies. Treatment with the pan PPAR agonist bezafibrate increased the efficiency of genome editing. Mice treated with bezafibrate that received both editing vectors had 5.9% of alleles that contained the integrated transgene, whereas only 3.1% of alleles contained the transgene in mice not treated with the drug. This work showed that integration of a therapeutic transgene using CRISPR/Cas9 based genome editing is possible in murine and canine models of GSD Ia. Editing resulted in biochemical correction and sustained transgene expression. These data support the further development of genome editing technologies for GSD Ia and other inherited metabolic disorders.

Synthetic biology facilitates both the design and fabrication of biological components and systems that do not already exist in the natural world. From an engineering point of view, synthetic biology is akin to building a complex machine by assembling simpler parts. Complex genetic machines can also be built by a modular and rational assembly of simpler biological parts. These biological machines can profoundly affect various cellular processes including the transcriptional machinery. In this thesis I demonstrate the utilization of biological parts according to synthetic biology principles to solve three distinct transcription-level problems: 1) How to efficiently select for transgene excision in induced pluripotent stem cells (iPSCs)? 2) How to eliminate transposase expression following piggyBac-mediated transgenesis? 3) How to reprogram cell lineage specification by the dCas9/gRNA transactivator-induced expression of endogenous transcription factors?

Viral vectors remain the most efficient and popular in deriving induced pluripotent stem cells (iPSCs). For translation, it is important to silence or remove the reprogramming factors after induction of pluripotency. In the first study, we design an excisable loxP-flanked lentiviral construct that a) includes all the reprogramming elements in a single lentiviral vector expressed by a strong EF-1α promoter; b) enables easy determination of lentiviral titer; c) enables transgene removal and cell enrichment using LoxP-site-specific Cre-recombinase excision and Herpes Simplex Virus-thymidine kinase/ganciclovir (HSV-tk/gan) negative selection; and d) allows for transgene excision in a colony format. With our design, a reprogramming efficiency comparable to that reported in the literature without boosting molecules can be consistently obtained. To further demonstrate the utility of this Cre-loxP/HSV-tk/gan strategy, we incorporate a non-viral therapeutic transgene (human blood coagulation Factor IX) in the iPSCs, whose expression can be controlled by a temporal pulse of Cre recombinase. The robustness of this platform enables the implementation of an efficacious and cost-effective protocol for iPSC generation and their subsequent transgenesis for downstream studies.

Transgene insertion plays an important role in gene therapy and in biological studies. Transposon-based systems that integrate transgenes by transposase-catalyzed "cut-and-paste" mechanism have emerged as an attractive system for transgenesis. Hyperactive piggyBac transposon is particularly promising due to its ability to integrate large transgenes with high efficiency. However, prolonged expression of transposase can become a potential source of genotoxic effects due to uncontrolled transposition of the integrated transgene from one chromosomal locus to another. In the second study we propose a vector design to decrease post-transposition expression of transposase and to eliminate the cells that have residual transposase expression. We design a single plasmid construct that combines the transposase and the transpositioning transgene element to share a single polyA sequence for termination. Consequently, the transposase element is deactivated after transposition. We also co-express Herpes Simplex Virus thymidine kinase (HSV-tk) with the transposase. Therefore, cells having residual transposase expression can be eliminated by the administration of ganciclovir. We demonstrate the utility of this combination transposon system by integrating and expressing a model therapeutic gene, human coagulation Factor IX, in HEK293T cells.

Genome editing by the efficient CRISPR/Cas9 system shows tremendous promise with ease of customization and the capability to multiplex distinguishing it from other such technologies. Endogenous gene activation is another aspect of CRISPR/Cas9 technology particularly attractive for biotechnology and medicine. However, the CRISPR/Cas9 technology for gene activation leaves much room for improvement. In the final study of this thesis we show that the fusion of two transactivation (VP64) domains to Cas9 dramatically enhances gene activation to a level that is sufficient to achieve direct cell reprogramming. Targeted activation of the endogenous Myod1 gene locus with this system leads to stable and sustained reprogramming of mouse embryonic fibroblasts into skeletal myocytes.

In conclusion, this dissertation demonstrates the power of utilizing biological parts in a rational and systematic way to rectify problems associated with cell fate reprogramming and transposon-based gene delivery. Through design of genetic constructs aided by synthetic biology principles, I aspire to make contributions to the related fields of cellular reprogramming, stem cell differentiation, genomics, epigenetics, cell-based disease models, gene therapy, and regenerative medicine.

Gene regulation is a complex and tightly controlled process that defines cell function in physiological and abnormal states. Programmable gene repression technologies enable loss-of-function studies for dissecting gene regulation mechanisms and represent an exciting avenue for gene therapy. Established and recently developed methods now exist to modulate gene sequence, epigenetic marks, transcriptional activity, and post-transcriptional processes, providing unprecedented genetic control over cell phenotype. Our objective was to apply and develop targeted repression technologies for regenerative medicine, genomics, and gene therapy applications. We used RNA interference to control cell cycle regulation in myogenic differentiation and enhance the proliferative capacity of tissue engineered cartilage constructs. These studies demonstrate how modulation of a single gene can be used to guide cell differentiation for regenerative medicine strategies. RNA-guided gene regulation with the CRISPR/Cas9 system has rapidly expanded the targeted repression repertoire from silencing single protein-coding genes to modulation of genes, promoters, and other distal regulatory elements. In order to facilitate its adaptation for basic research and translational applications, we demonstrated the high degree of specificity for gene targeting, gene silencing, and chromatin modification possible with Cas9 repressors. The specificity and effectiveness of RNA-guided transcriptional repressors for silencing endogenous genes are promising characteristics for mechanistic studies of gene regulation and cell phenotype. Furthermore, our results support the use of Cas9-based repressors as a platform for novel gene therapy strategies. We developed an in vivo AAV-based gene repression system for silencing endogenous genes in a mouse model. Together, these studies demonstrate the utility of gene repression tools for guiding cell phenotype and the potential of the RNA-guided CRISPR/Cas9 platform for applications such as causal studies of gene regulatory mechanisms and gene therapy.