We are a precision genetic medicines company committed to developing curative therapeutics for patients using our proprietary, comprehensive metagenomics-derived genome editing toolbox. Genetic diseases are caused by a diverse set of mutations that have been largely inaccessible by genome engineering approaches to date. Genetic mutations are seen in a variety of forms, including deletions, insertions, single-base-pair changes and sequence repeats, and are found throughout the genome and across a variety of different cell types, tissues, and organ systems. Additionally, many diseases lack a genetic origin but have the potential to be effectively and permanently addressed through genome editing. We are harnessing the power of metagenomics, the study of genetic material recovered from the natural environment, to unlock four billion years of microbial evolution to discover and develop a suite of novel editing tools capable of correcting any type of genetic mutation found anywhere in the genome. Our comprehensive genome editing toolbox includes programmable nucleases, base editors, and RNA and DNA-mediated integration systems (including prime editing systems and clustered regularly interspaced short palindromic repeat (“CRISPR”)-associated transposases (“CASTs”)). We believe our diverse and modular toolbox positions us to access the entire genome and select the optimal tool to unlock the full potential of genome editing for patients. The company was founded by pioneers in the field of metagenomics, a powerful science that allows us to tap into the diversity of microbial life on this planet. The metagenomics process starts by collecting samples from microbe-rich ecosystems ranging from simple home gardens to extreme locations such as hydrothermal vents below the ocean. We then extract the DNA from these environmental samples and deeply sequence them to fully reconstruct the genomes of the resident microbes. Each sample may include thousands of distinct genomes from previously unknown organisms revealing novel cellular machinery that we utilize as building blocks for our editing systems. Using high-throughput screening, artificial intelligence (“AI”), and proprietary algorithms, we rapidly mine through billions of novel proteins from our genome-resolved metagenomics database to create genome editing tools. To date, we have analyzed over 460 trillion base pairs, predicted over 7.4 billion proteins, including over 322 million CRISPR-associated (“Cas”) proteins and over 1.75 million CRISPRs, which we estimate has resulted in the identification of over 20,000 novel genome editing systems. Simultaneously, we have assembled extensive libraries of millions of nucleases, deaminases, reverse transcriptases (“RTs”) and over one thousand CASTs. Our platform is designed to enable us to rapidly and effectively find, screen, and select tools with the highest potential targetability, specificity, and efficiency in order to develop them into genetic medicines. The iterative nature of our process, underpinned by AI, allows us to continuously push the boundaries of innovation. Our proprietary toolbox of editing systems We have developed an expansive and modular toolbox of next-generation genome editing systems that will allow us to interact with the human genome in a site-specific manner, where each tool can be matched to specific disease targets. --- Our programmable nucleases are the backbone of our broad set of genome editing tools. These novel nucleases including type II and type V Cas nucleases, of which some are ultra-small systems that we call SMall Arginine-Rich sysTems (“SMART”) nucleases, have unique targeting abilities and can be programmed by guide RNAs (“gRNA”) to target and cut at specific locations in any genome sequence. Targeted genomic breaks trigger DNA repair pathways that can be used for genome editing, for example, to integrate a gene at a target site (knock-in) or for gene inactivation (knockdown). Our toolbox contains thousands of CRISPR nucleases with diverse abilities to target different parts of the genome, allowing us to select the ideal nuclease for targeting any given gene in a site-specific manner and potentially overcome a major limitation of first-generation CRISPR/Cas9 systems. We also modify our nucleases to either nick the genome (i.e., a nickase that cuts one strand of the DNA) or to simply bind to target sites (i.e., a nuclease dead variant). These capabilities (e.g., searching, cutting, nicking, and binding) can be leveraged as a chassis by adding on additional effector enzymes to create base editors for single nucleotide changes and RNA-mediated integration systems (“RIGS”) for both small and large genomic integrations using “Little RIGS” for prime editing and “Big RIGS” for large integrations. Using modular engineering, we match nickases with deaminases and RTs for base editing and RIGS, respectively. Furthermore, nucleases can be engineered by swapping the search modules of the enzyme to expand the targetability of the chassis, which is critical for site-specific genomic modifications. Given the measured targeting density of our toolbox, we believe that essentially any codon in the human genome could be addressed with our gene editing systems. Our highly active nucleases have gone through extensive preclinical evaluation for both in vivo and ex vivo applications, with demonstration of broad potency of these systems across human primary cells, mouse, and nonhuman primate (“NHP”) models. Our base editors, RIGS, and CAST systems have demonstrated activity across various cell-based models. In addition to evaluating system activity, we have undertaken detailed characterization of guide-specific on- and off-target effects. We routinely identify guides that have no or minimal verifiable off-target editing, thus overcoming another limitation of first-generation CRISPR/Cas9 systems. In addition to overcoming the activity, targetability, and specificity limitations of first-generation systems, our nuclease toolbox was designed to have broad compatibility with viral and nonviral delivery technologies. This compatibility is accomplished by having a variety of nuclease and gRNA structures, which range in terms of their size and biochemistry. For example, small guides for some type V Cas systems streamline manufacturing for delivery by lipid nanoparticle (“LNP”) approaches, and SMART nickases can be used to construct base editors that are small enough to fit within the packaging limitations of adeno-associated viruses (“AAV”). SpCas9, which is currently used in most base editing applications, is roughly three times the size of some of our smallest SMART nickases and cannot be efficiently packaged into a single AAV. Combined, we believe these features will facilitate delivery of our genome editing tools to previously inaccessible tissue types and organ systems. While nucleases, base editors, and prime editors can precisely address a wide variety of genomic modifications required to treat disease, the fact that many diseases are caused by a multitude of mutations across a gene means that a diverse set of editing tools are required to fully address these patient populations. The integration of a complete and functional gene through targeted genome editing may provide a way in which every patient with a given disease could potentially be treated by a single genetic medicine. Big RIGS and CASTs are novel genome editing systems that are under development to achieve what has been a major challenge for the genome editing field—large, targeted genomic integrations. Initial preclinical readouts conducted in mammalian cells indicate that these systems could potentially have a major impact on how diseases caused by loss-of-function mutations, the most common cause of genetic diseases, can be addressed through genome editing. Therapeutic translation roadmap and initial programs We are taking a stepwise approach deploying our genome editing toolbox to develop potentially curative therapies for patients. Our lead programs are selected to both address important diseases and to establish new standards in targetability, precision, efficiency, and scope of editing capabilities. Each of these indications were chosen based on our conviction in the underlying biology, existence of validating preclinical and clinical data, availability of pharmacodynamic and translational tools to assess early proof-of-concept, relevant value supporting outcome measures, and ongoing clinical unmet need. While we do not currently have any approved products and all of our product candidates are preclinical, our lead programs capture an ever-growing set of translational learnings and insights that will inform and accelerate future programs. --- Hemophilia A—novel, durable, knock-in approach for expression of Factor VIII Hemophilia A is the most common X-linked inherited bleeding disorder and is caused by mutations in the Factor VIII (“FVIII”) gene leading to loss of functional FVIII protein that impacts the body’s ability to form normal clots in response to injury. FVIII is normally produced in the liver within sinusoidal endothelial cells and is then secreted into the bloodstream where it acts as a cofactor for the catalytic activation of Factor X in the clotting pathway. The lack of functional FVIII disrupts the normal clotting cascade and predisposes patients to increased risk of bleeding, either spontaneously or in response to injury or surgery. Repeated bleeding episodes in joints or soft tissues can lead to progressive joint damage, inflammation, pain, and mobility impairment. Intracranial bleeding is of greatest concern as this can be rapidly fatal or lead to major morbidity. Rather than provide the FVIII gene in an episomal location, which risks dilution from cell division or cell death as well as episomal transcriptional silencing, our approach is to insert a FVIII DNA cassette into a "safe harbor location," within an intron of the albumin gene that is not expected to have deleterious effects. FVIII expression is then driven by the strong native albumin promoter. This approach has previously been demonstrated in preclinical studies to lead to therapeutically relevant expression of a different clotting factor (Factor IX) with negligible impact to systemic circulating albumin levels. Our FVIII knock-in approach is designed to provide stable expression and clinically relevant circulating levels of FVIII, even at low integration rates because of the strength of the albumin promoter. We have demonstrated the feasibility of the FVIII gene knock-in approach in mice with several mouse specific guides and different FVIII DNA donor cassettes, with integration of the FVIII gene leading to FVIII mRNA expression and therapeutically relevant levels of FVIII protein in the blood. In an ongoing NHP study we demonstrated integration of a surrogate cynomolgus-FVIII cassette (used to avoid immune response that would occur with a foreign human FVIII protein) and observed therapeutically relevant levels of the cyno-FVIII protein encoded by the integrated cassette in all 3 treated animals that has extended for 4.5 months following a single dose of the AAV-cFVIII virus followed five weeks later by a liver trophic LNP encapsulating the mRNA encoding MG29-1 and guide 2 at a dose of 1mg/kg body weight. We intend to continue measuring FVIII levels in these monkeys up to the 12 month time point to generate a robust data set on durability. Evaluation of different human FVIII donor DNA cassettes has been completed in mice resulting in the selection of 2 lead cassettes that will be compared in another NHP study, potentially leading to a development candidate selection anticipated in Q2 2024. In parallel, we are manufacturing mRNA, gRNA, AAV and LNP to support future investigational new drug (“IND”) enabling studies. Primary Hyperoxaluria, Type 1 (“PH1”)—a durable knockdown of HAO1 for substrate reduction therapy PH1 is a rare autosomal recessive metabolic disease arising from loss of function mutations in the alanine-glyoxylate aminotransferase (“AGXT”) gene that encodes alanine glyoxylate aminotransferase. This enzyme is found in peroxisomes of the liver where it catalyzes the conversion of glyoxylate to glycine and pyruvate. Lack of functional AGXT leads to an accumulation of glyoxylate substrate, which is then converted to oxalate and excreted in the kidney. The excess urinary oxalate forms an insoluble complex with urinary calcium that leads to the production of calcium oxalate crystal precipitates. This pathologic process results in the formation of repeated calcium oxalate urolithiasis and nephrolithiasis, which in turn leads to obstructive uropathy, inflammation, fibrosis, tubular toxicity, and progressive loss of kidney function. PH1 is a serious disease that causes kidney failure. More than 70% of individuals with PH1 mutations will develop end-stage renal disease, with a median age in young adulthood. The goal of our genome editing approach is to durably knock down HAO1 resulting in stable and permanent reduction of oxalate levels to effect a lifelong benefit. We have performed nuclease and guide screening to select an optimal nuclease and gRNA combination. Along with our partner ModernaTX, Inc. (“Moderna”), we have achieved preclinical proof-of-concept in an AGXT knock-out mouse which is an accepted disease model of PH1. We are in the final stages of confirming the candidate to take into NHP studies and expect to have NHP data in 2024 to support final development candidate selection. Transthyretin Amyloidosis—a single treatment to knockdown TTR gene expression Transthyretin amyloidosis is a disease of misfolded and aggregated transthyretin (“TTR”) protein that can deposit in tissues causing organ dysfunction, primarily in the heart and/or peripheral nerves. The TTR protein is normally produced in the liver and circulates in a homotetramer (four copies of the same TTR protein bound together) where it serves as a carrier protein for vitamin A and thyroxine. Certain mutations have been identified that can cause TTR homotetramers to fall apart, misfold, and aggregate into insoluble fibrils that deposit in cardiac tissue and peripheral nerves. However, more commonly, the normal aging process is associated with an increased propensity for TTR misfolding and aggregation in the heart without any known genetic sequence variation. These distinctions lead to TTR amyloidosis being characterized as either hereditary transthyretin amyloidosis (“ATTRv”) caused by mutations in TTR, or wild-type ATTR amyloidosis (“ATTRwt”). It is estimated that globally there are approximately 50,000 patients with ATTRv and between 300,000 and 500,000 patients with ATTRwt. Among the larger ATTRwt patient population, the most common presentation is a rapidly progressive, restrictive, and hypertrophic cardiomyopathy due to progressive deposition of insoluble TTR fibrils, which result in thickening of the myocardium and stiffening of the ventricles. These pathologic processes lead to impaired diastolic function and progressive cardiomyopathy that typically leads to progressive heart failure and often death within three to five years from disease onset. Although cardiac manifestations are more common and severe, patients with neurologic manifestations also experience significant morbidity, loss of functionality, and impaired quality of life. Using our novel nucleases, we aim to provide efficient TTR knockdown and halt further deposition of amyloid fibrils. Previous experience suggests a clinical correlation between the degree of TTR knockdown and potential for benefit in familial forms of the disease, which are expected to translate similarly to wild type forms. The high degree of in vivo editing efficiency and specificity of our nuclease platform suggest the potential for a single treatment to knockdown TTR gene expression and remove the requirement for life-long therapy. Along with our partner Ionis Pharmaceuticals, Inc. (“Ionis”), we are currently in advanced stages of nuclease and guide selection, having achieved more than 90% knockdown of human TTR protein after a single dose in a humanized TTR mouse model, and expect to move into NHP studies in 2024. Further areas of therapeutic activity and interest In parallel with our translation efforts in our lead programs using our novel programmable nucleases to knock-in or knockdown gene expression in liver-associated targets, we are developing more complex editing systems for liver associated targets as well as moving beyond the liver. Given that our genome editing toolbox contains small editing systems designed to be amenable to viral vector delivery, and given the progress established in targeting the central nervous system and muscle with established AAV capsids, our first extrahepatic indications will be neurodegenerative and neuromuscular diseases. Building on our experience delivering our nucleases to the liver via LNP systems, we are extending that experience delivering novel RIGS to the liver to potentially correct ATP7B mutations in Wilson’s disease and PiZ mutations in alpha-1-antitrypsin deficiency (“A1AT deficiency”). We are also exploring addressing A1AT deficiency via a base editor approach given the predominant mutation involves a single base pair. Both of these liver diseases have well-defined biology, readily available translational biomarkers for early proof-of-concept, established development pathways based on prior drug approvals, and important unmet medical needs. Building on our experience with our novel type II and type V programmable nucleases, we are extending that experience by working to deliver these nucleases via AAV to the central nervous system to potentially knockdown genetic targets important for both spontaneous and familial amyotrophic lateral sclerosis (SOD1, ATXN2) and Charcot-Marie-Tooth Type 1a (PMP22). In addition, we are working to address a series of mutations common in Duchenne Muscular Dystrophy with our programmable nucleases through exon skipping approaches. In diseases outside of the liver, we intend to initially leverage known biology and clinical validation achieved with RNA-targeted approaches like antisense and small interfering RNA (“siRNA”) to advance more potent and definitive one-time genome editing treatments. Building on our experience with both knock-in gene expression and smaller gene corrections with RIGS, we are progressing our larger RNA- and DNA-mediated integration systems to potentially provide a single curative approach to cystic fibrosis. As opposed to currently-available therapies limited to subsets of patients with individual mutations, we intend to deliver a full copy of a functional cystic fibrosis transmembrane conductance regulator (“CFTR”) gene. This approach can similarly be pursued across many other diseases characterized by loss of function mutations. Our principal executive offices are located 5959 Horton Street, 7th Floor, Emeryville, California.