Browsing by Author "Gersbach, Charles A"

Duchenne muscular dystrophy (DMD) is a severe, progressive muscle wasting disease that causes loss of ambulation and premature death in affected boys. In the 1980s, the cause of the disease was attributed to mutations in the DMD gene, yet almost 40 years later there is still no cure. The large size of the gene, diversity of patient mutations, and delivery challenge of modifying skeletal muscle systemically have all limited the application of therapies to correct the disease-causing mutations. Gene editing with CRISPR-Cas technology is poised to revolutionize our ability to treat genetic diseases. For DMD, several landmark studies have demonstrated the strong therapeutic potential of using CRISPR to restore dystrophin protein in DMD models. Yet many challenges remain in converting proof-of-concept editing to a safe and effective therapy. Here, we focus on optimizing and developing methods to evaluate the two most promising strategies for using gene editing to treat DMD: exon deletion and exon skipping.In aim 1, we tackle the problem of searching for highly efficient gRNAs in the vast sequence space of DMD introns by applying high-throughput screening techniques to measure the relative deletion efficiency of gRNA pairs. We discover novel gRNA pairs for the deletion of DMD exon 51 and demonstrate an improvement over previous methods of gRNA pair discovery. In aim 2, we evaluate the potential of “CRISPR 2.0” editing tools that provide next-generation control over DNA editing to produce DMD exon skipping with reduced disruption to the genome. We discover a base editor and gRNA design that efficiently skips exon 45 and restores dystrophin expression and apply unbiased characterization methods to interrogate the impact of such editing.

The ends of long bones that articulate with respect to one another are lined with a crucial connective tissue called articular cartilage. This tissue plays an essential biomechanical function in synovial joints, as it serves to both dissipate load and lubricate articulating surfaces. Osteoarthritis is a painful and debilitating disease that drives the deterioration of articular cartilage. Like many chronic diseases, pro-inflammatory cytokines feature prominently in the onset and progression of osteoarthritis. Because cartilage lacks physiologic features critical for regeneration and self-repair, the development of effective strategies to create functional cartilage tissue substitutes remains a priority for the fields of tissue engineering and regenerative medicine. The overall objectives of this dissertation are to (1) develop a bioactive scaffold capable of mediating cell differentiation and formation of extracellular matrix that recapitulates native cartilage tissue and (2) to produce stem cells specifically tailored at the scale of the genome with the ability to resist inflammatory cues that normally lead to degeneration and pain.

Engineered replacements for musculoskeletal tissues generally require extensive ex vivo manipulation of stem cells to achieve controlled differentiation and phenotypic stability. By immobilizing lentivirus driving the expression of transforming growth factor-β3 to a highly structured, three dimensionally woven tissue engineering scaffold, we developed a technique for producing cell-instructive scaffolds that control human mesenchymal stem cell differentiation and possess biomechanical properties approximating those of native tissues. This work represents an important advance, as it establishes a method for generating constructs capable of restoring biological and mechanical function that may circumvent the need for ex vivo conditioning of engineered tissue substitutes.

Any functional cartilage tissue substitute must tolerate the inflammation intrinsic to an arthritic joint. Recently emerging tools from synthetic biology and genome engineering facilitate an unprecedented ability to modify how cells respond to their microenvironments. We exploited these developments to engineer cells that can evade signaling of the pro-inflammatory cytokine interleukin-1 (IL-1). Our study provides proof-of-principle evidence that cartilage derived from such engineered stem cells are resistant to IL-1-mediated degradation.

Extending on this work, we developed a synthetic biology strategy to further customize stem cells to combat inflammatory cues. We commandeered the highly responsive endogenous locus of the chemokine (C-C motif) ligand 2 gene in pluripotent stem cells to impart self-regulated, feedback-controlled production of biologic therapy. We demonstrated that repurposing of degradative signaling pathways induced by IL-1 and tumor necrosis factor toward transient production of cytokine antagonists enabled engineered cartilage tissue to withstand the action of inflammatory cytokines and to serve as a cell-based, auto-regulated drug delivery system.

In this work, we combine principles from synthetic biology, gene therapy, and functional tissue engineering to develop methods for generating constructs with biomimetic molecular and mechanical features of articular cartilage while precisely defining how cells respond to dysfunction in the body’s finely-tuned inflammatory systems. Moreover, our strategy for customizing intrinsic cellular signaling pathways in therapeutic stem cell populations opens innovative possibilities for controlled drug delivery to native tissues, which may provide safer and more effective treatments applicable to a wide variety of chronic diseases and may transform the landscape of regenerative medicine.

Electrostatic interaction is a strong force that attracts positively and negatively charged molecules to each other. Such an interaction is formed between positively charged polycationic polymers and negatively charged nucleic acids. In this dissertation, the electrostatic attraction between polycationic polymers and nucleic acids is exploited for applications in oral gene delivery and nucleic acid scavenging. An enhanced nanoparticle for oral gene delivery of a human Factor IX (hFIX) plasmid is developed using the polycationic polysaccharide, chitosan (Ch), in combination with protamine sulfate (PS) to treat hemophilia B. For nucleic acid scavenging purposes, the development of an effective nucleic acid scavenging nanofiber platform is described for dampening hyper-inflammation and reducing the formation of biofilms.

Non-viral gene therapy may be an attractive alternative to chronic protein replacement therapy. Orally administered non-viral gene vectors have been investigated for more than one decade with little progress made beyond the initial studies. Oral administration has many benefits over intravenous injection including patient compliance and overall cost; however, effective oral gene delivery systems remain elusive. To date, only chitosan carriers have demonstrated successful oral gene delivery due to chitosan’s stability via the oral route. In this study, we increase the transfection efficiency of the chitosan gene carrier by adding protamine sulfate to the nanoparticle formulation. The addition of protamine sulfate to the chitosan nanoparticles results in up to 42x higher in vitro transfection efficiency than chitosan nanoparticles without protamine sulfate. Therapeutic levels of hFIX protein are detected after oral delivery of Ch/PS/phFIX nanoparticles in 5/12 mice in vivo, ranging from 3 -132 ng/mL, as compared to levels below 4 ng/mL in 1/12 mice given Ch/phFIX nanoparticles. These results indicate the protamine sulfate enhances the transfection efficiency of chitosan and should be considered as an effective ternary component for applications in oral gene delivery.

Dying cells release nucleic acids (NA) and NA-complexes that activate the inflammatory pathways of immune cells. Sustained activation of these pathways contributes to chronic inflammation related to autoimmune diseases including systemic lupus erythematosus, rheumatoid arthritis, and inflammatory bowel disease. Studies have shown that certain soluble, cationic polymers can scavenge extracellular nucleic acids and inhibit RNA-and DNA-mediated activation of Toll-like receptors (TLRs) and inflammation. In this study, the cationic polymers are incorporated onto insoluble nanofibers, enabling local scavenging of negatively charged pro-inflammatory species such as damage-associated molecular pattern (DAMP) molecules in the extracellular space, reducing cytotoxicity related to unwanted internalization of soluble cationic polymers. In vitro data show that electrospun nanofibers grafted with cationic polymers, termed nucleic acid scavenging nanofibers (NASFs), can scavenge nucleic acid-based agonists of TLR 3 and TLR 9 directly from serum and prevent the production of NF-ĸB, an immune system activating transcription factor while also demonstrating low cytotoxicity. NASFs formed from poly (styrene-alt-maleic anhydride) conjugated with 1.8 kDa branched polyethylenimine (bPEI) resulted in randomly aligned fibers with diameters of 486±9 nm. NASFs effectively eliminate the immune stimulating response of NA based agonists CpG (TLR 9) and poly (I:C) (TLR 3) while not affecting the activation caused by the non-nucleic acid TLR agonist pam3CSK4. Results in a more biologically relevant context of doxorubicin-induced cell death in RAW cells demonstrates that NASFs block ~25-40% of NF-ĸβ response in Ramos-Blue cells treated with RAW extracellular debris, ie DAMPs, following doxorubicin treatment. Together, these data demonstrate that the formation of cationic NASFs by a simple, replicable, modular technique is effective and that such NASFs are capable of modulating localized inflammatory responses.

An understandable way to clinically apply the NASF is as a wound bandage. Chronic wounds are a serious clinical problem that is attributed to an extended period of inflammation as well as the presence of biofilms. An NASF bandage can potentially have two benefits in the treatment of chronic wounds by reducing the inflammation and preventing biofilm formation. NASF can prevent biofilm formation by reducing the NA present in the wound bed, therefore removing large components of what the bacteria use to develop their biofilm matrix, the extracellular polymeric substance, without which the biofilm cannot develop. The NASF described above is used to show the effect of the nucleic acid scavenging technology on in vitro and in vivo biofilm formation of P. aeruginosa, S. aureus, and S. epidermidis biofilms. The in vitro studies demonstrated that the NASFs were able to significantly reduce the biofilm formation in all three bacterial strains. In vivo studies of the NASF on mouse wounds infected with biofilm show that the NASF retain their functionality and are able to scavenge DNA, RNA, and protein from the wound bed. The NASF remove DNA that are maintaining the inflammatory state of the open wound and contributing to the extracellular polymeric substance (EPS), such as mtDNA, and also removing proteins that are required for bacteria/biofilm formation and maintenance such as chaperonin, ribosomal proteins, succinyl CoA-ligase, and polymerases. However, the NASF are not successful at decreasing the wound healing time because their repeated application and removal disrupts the wound bed and removes proteins required for wound healing such as fibronectin, vibronectin, keratin, and plasminogen. Further optimization of NASF treatment duration and potential combination treatments should be tested to reduce the unwanted side effects of increased wound healing time.

The eukaryotic epigenome has an instrumental role in determining and maintaining cell identity and function. Epigenetic components such as DNA methylation, histone tail modifications, chromatin accessibility, and DNA architecture are tightly correlated to central cellular processes, while their dysregulation manifests in aberrant gene expression and disease. The ability to specifically edit the epigenome holds the promise of enhancing understanding how epigenetic modifications function and enabling manipulation of cell phenotype for scientific or therapeutic purposes. Genome targeting technologies, such as the CRISPR/Cas9 system, have successfully been harnessed to create epigenome editing tools to alter gene expression. Prominently, two leading CRISPR-based technologies, CRISPRa and CRISPRi, were shown to be highly specific and effective in controlling gene transcription levels. These tools, however, often lead to formation of complexes that affect a multitude of endogenous factors, thus mitigating our ability to elucidate the role of individual epigenetic marks. Moreover, changes in epigenetic marks are associated with numerous health conditions, therefore the development of tools that can modify specific marks may help in creating disease models, or the restoration of a “healthy” epigenome. We first created a suite of CRISPR-based epigenome modifiers (CRISPR-GEMs) that were aimed to catalyze the removal or addition of specific histone tail marks. Next, we tested a few promising CRISPR-GEMs on multiple target genes to characterize their effect on gene expression and chromatin marks. Furthermore, we utilized these tools to deepen our insights into the relationship of individual histone marks and gene expression in different contexts and to better our understanding of the kinetics and dynamics of several of these novel tools alongside existing ones. Additionally, we decided to use the CRISPRa platform to explore senescence, a cellular process that is at the epicenter of aging and has been shown to play a key role in various age-related diseases. Using the CRISPRa platform in an inducible-senescence cell model, we found and validated multiple transcription factors (TFs) that regulate senescence-associated growth arrest (SAGA). Lastly, we characterized genetic pathways that are pivotal to successful inhibition of SAGA, thereby demonstrating a new application of epigenome editing in a senescence model that enhanced our understanding of the pathways that govern SAGA.

Cellular fate specifications and functions are controlled through precise and complex patterns in gene expression. Although all cells in the human body share the same DNA sequence, the expression of RNA and proteins is diverse across tissue and cell types. This orchestrated control of thousands of genes is performed by transcription factors which bind to DNA regulatory elements and modulate gene expression through assembly of transcriptional complexes or interactions with other transcriptional proteins. These regulatory elements harbor sequence variants that can impact their ability to bind transcription factors and thus influence gene expression to ultimately manifest in various traits and diseases. Our ability to interpret the functional consequences of sequence variation relies on understanding the function of the sequence the variation lies in. However, our knowledge of regulatory element function and to which genes they regulate is still limited to a small number of individually validated sequences or predicted regions from large-scale genomic dataset correlations. Recent advancements in massively parallel reporter assays have enabled the testing of millions of DNA sequences and their regulatory activity but are limited to testing outside of their native contexts. Thus, our goal was to develop scalable methods based on the CRISPR/Cas9 system to facilitate the screening of regulatory element function in their endogenous state in high throughput. We first sought to adapt CRISPR/Cas9 based epigenetic repressors and activators to perturb hundreds of putative regulatory elements in their endogenous context and screen their contribution to a single gene’s expression in single pooled experiments. Our results demonstrated the capability of the system to accurately assign function to both known and novel regulatory elements. Finally, we sought to apply the system to screen more than 100,000 putative regulatory elements for essentiality and their contribution towards cellular fitness. Overall, these studies demonstrate novel methods for decoding regulatory networks and furthering our ability to assign function to regulatory sequence influencing disease.

Technologies for engineering new functions into proteins are advancing biological research, biotechnology, and medicine at an astounding rate. Building on fundamental research of natural protein structure and function, scientists are identifying new protein domains with previously undescribed properties and engineering new proteins with expanded functionalities. Such tools are enabling the precise study of fundamental aspects of cellular behavior and the development of a new class of gene therapies that manipulate the expression of endogenous genes. The applications of these gene regulation technologies include but are not limited to controlling cell fate decisions, reprogramming cell lineage commitment, monitoring cellular states, and stimulating expression of therapeutic factors.

While the field has come a long way in the past 20 years, there are still many limitations. Historically, gene therapy and gene replacement therapies have relied on over-expression of natural transcription factors that activate specific endogenous gene networks. However, natural transcription factors are often inadequate for generating efficient, fast, and homogenous cellular responses. Furthermore, most natural transcription factors have complex structures and functions that are difficult to improve or alter by rational design. This thesis presents three novel and widely applicable methods for engineering transcription factors for programming cell fate decisions in primary human cells. MyoD is the master transcription factor defining the myogenic lineage. Expression of MyoD in certain non-myogenic lineages induces a coordinated change in differentiation state. We use MyoD as a model for developing our protein engineering techniques because myogenesis is a well-studied pathway that is characterized by an easily detected change in phenotype from mono-nucleated to multinucleated cells. Furthermore, efficient generation of myocytes in vitro presents an attractive patient-specific method by which to treat muscle-wasting diseases such as muscular dystrophy.

We first demonstrate that we can improve the ability of MyoD to convert human dermal fibroblasts and human adipose-derived stem cells into myocyte-like cells. By fusing potent modular activation domains to the MyoD protein, we increased myogenic gene expression, myofiber formation, cell fusion, and global reprogramming of the myogenic gene network. The engineered MyoD transcription factor induced myogenisis in a little as ten days, a process that takes three or more weeks with the natural MyoD protein.

While increasing the potency of transcriptional activation is one mechanism by which to improve transcription factor function, there are many other possible routes such as increasing DNA-binding affinity, increasing protein stability, altering interactions with co-factors, or inducing post-translational modifications. Endogenous regulatory pathways are complex, and it is difficult to predict specific amino acid changes that will produce the desired outcome. Therefore, we designed and implemented a high-throughput directed evolution system in mammalian cells that allowed us to enrich for MyoD variants that are successful at inducing expression of the myogenic gene network. Directed evolution presents a well-established and currently unexplored approach for uncovering amino acid substitutions that improve the intrinsic properties of transcription factors themselves without any prior knowledge. After ten rounds of selection, we identified amino acid substitutions in MyoD that increase expression of a subset of myogenic gene markers in primary human cells.

Rather than guide cell fate decisions by expressing an exogenous factor, it may be beneficial to activate expression of the endogenous gene locus. In comparison to delivering the transcription factor cDNA, expression from the endogenous locus may induce chromatin remodeling and activation of positive feedback loops to stimulate autologous expression more quickly. Recent discoveries of the principles of protein-DNA interactions in various species and systems has guided the development of methods for engineering designer enzymes that can be targeted to any DNA target site. We make use of the RNA-guided Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 system to induce expression of the endogenous MyoD gene in human induced pluripotent stem cells (iPSCs). Through complementary base pairing, chimeric guide RNAs (gRNAs) direct a Cas9 transcriptional activator to a target DNA sequence, leading to endogenous gene expression. A current limitation of CRISPR/Cas9-based gene regulation is the potency of transcriptional activation and delivery of the CRISPR/Cas9 components. To address these limitations, we first developed a platform to express Cas9 and up to four gRNAs from a single lentiviral vector. We then optimized the gRNAs and Cas9 transcriptional activator to induce endogenous MyoD expression and differentiate iPSCs into myocyte-like cells.

In summary, the objective of this work is to develop protein engineering techniques to improve both natural and synthetic transcription factor function for programming cell fate decisions in primary human cells. While we focus on myogenesis, each method can be easily adapted to other transcription factors and gene networks. Engineered transcription factors that induce fast and efficient remodeling of gene networks have widespread applications in the fields of biotechnology and regenerative medicine. Continuing to develop these tools for modulating gene expression will lead to an expanded number of disease models and eventually the efficient generation of patient-specific cellular therapies.

The cell-type diversity in a multicellular organism is precisely defined through epigenetic regulation of an underlying DNA sequence. However, cell fate can be reprogrammed artificially with the proper extrinsic and intrinsic cues. This ability to manipulate cell-type specification has revolutionized the fields of regenerative medicine and disease modeling that aim to program cell phenotypes to better understand and provide therapy for human disease. Strategies for cell reprogramming commonly involve the overexpression of naturally occurring transcription factors to instruct the transcriptional codes of a new cell identity. However, this approach often results in a low efficiency of conversion with poor maturation and limited functional integration in vivo. Thus, our objective was to utilize tools in synthetic biology based on the CRISPR/Cas9 system to more precisely and accurately control endogenous gene expression towards applications in cell reprogramming. We first applied CRISPR/Cas9- based transcription factors to convert fibroblasts to neurons. This strategy entailed activating endogenous proneural genes within fibroblasts, rewriting the epigenetic signatures at the target loci and enabling stable autonomous expression of the target genes. Next, we systematically profiled the proneural activity of every human transcription factor with CRISPR activation pooled gRNA screens. Through these unbiased screens, we uncovered transcription factor combinations that increase conversion efficiency, influence subtype specification and improve synaptic maturation of in vitro-derived neurons. Finally, we applied the same CRISPR-based transcription factors to study mechanisms of gene regulation at the 15q11-13 imprinted locus implicated in the development of Prader-Will syndrome (PWS), a genetic neurobehavioral disorder. Through this work, we identified allele-specific regulatory elements that serve as candidate target sites for epigenetic therapy in PWS. Overall, we've developed a novel approach using tools in synthetic biology to improve the specification of neuronal cell types and to elucidate gene regulatory mechanisms in a neurobehavioral disorder.

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.

Over the past several years genome and epigenome engineering has been propelled forward by CRISPR-Cas technologies. These prokaryotic defense systems work well in mammalian cells in a manner that is remarkably robust: they are non-toxic, fold into a catalytically active state, localize to targeted cellular compartments, and act on the eukaryotic genome, which is heavily compacted in chromatin. While all these are true, CRISPR-cas nucleases did not evolve to function as highly specific genome engineering tools. Thus, the major goals of the work presented herein are to i) refine the specificity of CRISPR-Cas enzymes, ii) develop methods that facilitate genome engineering in human cells, and iii) apply these technologies toward outstanding problems in human gene regulation. With regard to the first goal, we set out to develop a method that could be easily applied to increase the specificity of diverse CRISPR systems. Adopting RNA-engineering to achieve this goal, we modulate the kinetics of DNA strand invasion to increase the specificity of Cas enzymes. Since the guide RNA is a feature that is common across all CRISPR systems, we expect that this new method to tune the activity and specificity of Cas enzymes will be broadly useful. To address the second goal, we set out to develop an experimental pipeline for the high throughput, precise modification of mammalian genomes. Specifically, we modify the C-termini of genes to include an epitope tag for the genome-wide profiling of transcription factor binding sites. We apply this method to over 30 genes, encoding a variety of transcription factors, chromatin modifying enzymes, and gene regulatory proteins. Out of the large number of genes we focus particularly on members of the AP-1 transcription factor family and nuclear receptor co-activator and co-repressor families. Using this ChIP-seq data, which profiles genome wide binding, and integrating a variety of other genomic information, including chromatin modifications, chromatin accessibility, other TF binding, and inherent regulatory activity, we investigate the dimerization preferences of AP-1 subunits, their genomic binding patterns, and the regulatory potential of theses subunits. Toward addressing the third goal, we decided to focus on the glucocorticoid receptor (GR). The dual activating and repressive function of the GR is incompletely understood, and this duality is a property of many other stimuli responsive transcriptional responses (e.g. NFKB signaling). Thus, how one transcription factor is biochemically endowed with the ability to both activate and repress gene expression is an outstanding problem in gene regulation. It is hypothesized that the GR recruits a variety of distinct protein complexes in order to mediate its diverse function. We used CRISPR based loss of function screening in order to discover new GR cofactors. Using this method, we find a number of cofactors, both canonical and novel, that regulate this response in A549 cells. Ongoing work investigates how general these cofactors are across the transcriptome and whether they provide an avenue to decouple GR’s dual function, which has been a major goal in drug development. Through these studies we have found a way to make CRISPR systems more specific, developed and applied CRISPR based method to define AP-1 binding and function, and used unbiased CRISPR based screens to discover novel regulators of the glucocorticoid drug response.

Chapter 1 broadly introduces this work, its motivations, and aims of research presented herein.

Chapter 2 provides an introduction to both genome engineering and gene regulation. Specifically, it describes the development and application of CRISPR-cas tools and details outstanding problems in gene regulation through the lens of nuclear receptors.

Chapter 3 describes the purification of Cas9 protein and its characterization biochemically. Specifically, we use AFM to determine the DNA binding properties of Cas9 in vitro.

Chapter 4 introduces a new method to modulate the specificity of CRISPR systems in human cells. Therein we show that RNA secondary structure can be applied to diverse CRISPR systems to tune their activity.

Chapter 5 details a method for the high throughput tagging of transcription factors. It specifically investigates members of the AP-1 transcription factor complex.

Chapter 6 is an investigation of the glucocorticoid receptor and its cofactors. We apply a variety of genome engineering and genomic methods to characterize known co-factors and discover new ones.

Chapter 7 is an outlook on the fields of genome-engineering and gene regulation. It describes key questions that are still unanswered and possible lines of attack to address them.

The growing complexity of scientific research demands further development of advanced gene regulation systems. For instance, the ultimate goal of tissue engineering is to develop constructs that functionally and morphologically resemble the native tissue they are expected to replace. This requires patterning of gene expression and control of cellular phenotype within the tissue engineered construct. In the field of synthetic biology, gene circuits are engineered to elucidate mechanisms of gene regulation and predict the behavior of more complex systems. Such systems require robust gene switches that can quickly turn gene expression on or off. Similarly, basic science requires precise genetic control to perturb genetic pathways or understand gene function. Additionally, gene therapy strives to replace or repair genes that are responsible for disease. The safety and efficacy of such therapies require control of when and where the delivered gene is expressed in vivo.

Unfortunately, these fields are limited by the lack of gene regulation systems that enable both robust and flexible cellular control. Most current gene regulation systems do not allow for the manipulation of gene expression that is spatially defined, temporally controlled, reversible, and repeatable. Rather, they provide incomplete control that forces the user to choose to control gene expression in either space or time, and whether the system will be reversible or irreversible.

The recent emergence of the field of optogenetics--the ability to control gene expression using light--has made it possible to regulate gene expression with spatial, temporal, and dynamic control. Light-inducible systems provide the tools necessary to overcome the limitations of other gene regulation systems, which can be slow, imprecise, or cumbersome to work with. However, emerging light-inducible systems require further optimization to increase their efficiency, reliability, and ease of use.

Initially, we engineered a light-inducible gene regulation system that combines zinc finger protein technology and the light-inducible interaction between Arabidopsis thaliana plant proteins GIGANTEA (GI) and the light oxygen voltage (LOV) domain of FKF1. Zinc finger proteins (ZFPs) can be engineered to target almost any DNA sequence through tandem assembly of individual zinc finger domains that recognize a specific three base-pair DNA sequence. Fusion of three different ZFPs to GI (GI-ZFP) successfully targeted the fusion protein to the specific DNA target sequence of the ZFP. Due to the interaction between GI and LOV, co-expression of GI-ZFP with a fusion protein consisting of LOV fused to three copies of the VP16 transactivation domain (LOV-VP16) enabled blue-light dependent recruitment of LOV-VP16 to the ZFP target sequence. We showed that placement of three to nine copies of a ZFP target sequence upstream of a luciferase or eGFP transgene enabled expression of the transgene in response to blue-light. Gene activation was both reversible and tunable based on duration of light exposure, illumination intensity, and the number of ZFP binding sites upstream of the transgene. Gene expression could also be spatially patterned by illuminating the cell culture through photomasks containing various patterns.

Although this system was useful for controlling the expression of a transgene, for many applications it is useful to control the expression of a gene in its natural chromosomal position. Therefore we capitalized on recent advances in programmed gene activation to engineer an optogenetic tool that could easily be targeted to new, endogenous DNA sequences without re-engineering the light inducible proteins. This approach took advantage of CRISPR/Cas9 technology, which uses a gene-specific guide RNA (gRNA) to facilitate Cas9 targeting and binding to a desired sequence, and the light-inducible heterodimerizers CRY2 and CIB1 from Arabidopsis thaliana to engineer a light-activated CRISPR/Cas9 effector (LACE) system. We fused the full-length (FL) CRY2 to the transcriptional activator VP64 (CRY2FL-VP64) and the N-terminal fragment of CIB1 to the N-, C-, or N- and C- terminus of a catalytically inactive Cas9. When CRY2-VP64 and one of the CIBN/dCas9 fusion proteins are expressed with a gRNA, the CIBN/dCas9 fusion protein localizes to the gRNA target. In the presence of blue light, CRY2FL binds to CIBN, which translocates CRY2FL-VP64 to the gene target and activates transcription. Unlike other optogenetic systems, the LACE system can be targeted to new endogenous loci by solely manipulating the specificity of the gRNA without having to re-engineer the light-inducible proteins. We achieved light-dependent activation of the IL1RN, HBG1/2, or ASCL1 genes by delivery of the LACE system and four gene-specific gRNAs per promoter region. For some gene targets, we achieved equivalent activation levels to cells that were transfected with the same gRNAs and the synthetic transcription factor dCas9-VP64. Gene activation was also shown to be reversible and repeatable through modulation of the duration of blue light exposure, and spatial patterning of gene expression was achieved using an eGFP reporter and a photomask.

Finally, we engineered a light-activated genetic "on" switch (LAGOS) that provides permanent gene expression in response to an initial dose of blue light illumination. LAGOS is a lentiviral vector that expresses a transgene only upon Cre recombinase-mediated DNA recombination. We showed that this vector, when used in conjunction with a light-inducible Cre recombinase system,1 could be used to express MyoD or the synthetic transcription factor VP64-MyoD2 in response to light in multiple mammalian cell lines, including primary mouse embryonic fibroblasts. We achieved light-mediated upregulation of downstream myogenic markers myogenin, desmin, troponin T, and myosin heavy chains I and II as well as fusion of C3H10T½ cells into myotubes that resembled a skeletal muscle cell phenotype. We also demonstrated LAGOS functionality in vivo by engineering the vector to express human VEGF165 and human ANG1 in response to light. HEK 293T cells stably expressing the LAGOS vector and transiently expressing the light-inducible Cre recombinase proteins were implanted into mouse dorsal window chambers. Mice that were illuminated with blue light had increased microvessel density compared to mice that were not illuminated. Analysis of human VEGF and human ANG1 levels by enzyme-linked immunosorbent assay (ELISA) revealed statistically higher levels of VEGF and ANG1 in illuminated mice compared to non-illuminated mice.

In summary, the objective of this work was to engineer robust light-inducible gene regulation systems that can control genes and cellular fate in a spatial and temporal manner. These studies combine the rapid advances in gene targeting and activation technology with natural light-inducible plant protein interactions. Collectively, this thesis presents several optogenetic systems that are expected to facilitate the development of multicellular cell and tissue constructs for use in tissue engineering, synthetic biology, gene therapy, and basic science both in vitro and in vivo.

In the last decade, the paradigm for cancer therapy has incrementally transitioned away from non-specific cytotoxic therapies (radiation, chemotherapy) and targeted therapies (small molecules, biologics) and towards immune cell-based therapies. Immune cell-based therapies such as adoptive T cell therapy (ACT) harness the intrinsic ‘sense and respond’ functions of immune cells to selectively target and eliminate cancer cells. Nevertheless, more than half of cancer patients either do not respond or relapse to existing ACTs. Several studies have defined specific transcriptional and epigenetic signatures of the infused T cell product associated with clinical response, indicating that T cell state and fitness is linked to ACT efficacy. Thus, epigenetically reprogramming T cells with enhanced potency and durability has the potential to improve ACT. However, this potential has yet to be fully realized due to technical challenges of adapting CRISPR-based epigenome editing technologies for applications in primary human T cells. To overcome these challenges, we developed and rigorously characterized compact and robust CRISPR repressors and activators for endogenous gene regulation. Next, we leveraged these technologies to systematically interrogate the effects of >100 transcriptional and epigenetic regulators on human CD8+ T cell state and function through complementary CRISPR interference (CRISPRi) and activation (CRISPRa) screens. These CRISPRi/a screens converged on basic leucine zipper ATF-like transcription factor (BATF3). Subsequent assays revealed that BATF3 overexpression promotes specific features of memory T cells (such as increased expression of IL7R and glycolysis), counters T cell exhaustion, and enhances CAR T cell potency in both in vitro and in vivo tumor models. In addition, BATF3 programs a transcriptional profile strongly associated with positive clinical response to CD19 CAR T cell therapy. Given that BATF3 is a compact transcription factor (TF) without any transactivation or epigenetic domains, we speculated that BATF3 achieves its widespread effects by interacting with other TFs. To identify these factors, we conducted parallel CRISPR knockout screens targeting all TFs with or without BATF3 overexpression. Using IL7R expression as a proxy for BATF3 activity, we identified both BATF3-independent and dependent transcriptional regulators of IL7R expression. For example, JUNB and IRF4 were uniquely enriched in the low IL7R population in the screen with BATF3 overexpression, suggesting BATF3 heterodimerizes with JUNB and interacts with IRF4 to regulate gene expression. Finally, these CRISPR knockout screens illuminated other candidate therapeutic targets for future exploration and characterization. Overall, we have developed a widely applicable synthetic biology toolkit of orthogonal epigenome editors, which we used to systematically identify regulators of human CD8+ T cell state and function. This catalogue of regulators could serve as the basis for engineering next generation T cell therapies for cancer.

Cell differentiation is the foundation for tissue development and regeneration, disease modeling, and cell-based therapies. The differentiation of skeletal myoblasts has been well-studied, and expression of the transcription factor myogenin is recognized as an early indicator of a cell committed to the myogenic differentiation. Not as much is known regarding how individual cells activate myogenin, the dynamics with which this happens, or the genomic regions that regulate this activation beyond the promoter. This research tested the following hypotheses: 1) The single-cell expression profile of myogenin could reveal distinct subpopulations of myogenic cells and indicate unique cell states. 2) The dynamic evolution of myogenin expression is indicative of the processes regulating differentiation. 3) There exist other genomic loci beyond the immediate promoter of myogenin that regulate the expression of myogenin. The primary conclusions of this dissertation are as follows: 1) Differentiating or reprogrammed cells activating myogenin show a bimodal distribution, with cells either expressing low or high levels of myogenin, and there is a critical dose of MyoD required to transition to differentiate. 2) Myogenic lineage commitment can be delayed but not prevented by serum, and myogenic reprogramming can be accelerated by increasing the forced expression of MyoD. 3) In addition to the promoter, there are additional enhancer sites that regulate the expression of myogenin in differentiating myoblasts.

Recent advances in synthetic transcription factor design have given scientists new tools for probing genome biology and treating human disease. However, the effects of these designer proteins on the cellular physiology have not been fully characterized. Here we describe the effects of synthetic activators on the DNA methylation status of targeted promoters. Through clonal bisulfite analysis we found that these activators induce a statistically significant decrease in DNA methylation in the targeted region. This study uncovers an important epigenetic modification that synthetic transctiptional activators can induce.

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.