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.
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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.
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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.
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Our principal executive offices are located 5959 Horton Street, 7th Floor,
Emeryville, California 94608, and our telephone number is (510) 871-4880. Our
website address is www.metagenomi.co.