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References 345
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14
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Detection and Quantification ofGenome Editing
Events inPreclinical and Clinical Studies
Marina Falaleeva1*, Shengdar Tsai2, Kathleen Meyer1,
and Yanmei Lu
1
Nonclinical Department, Sangamo Therapeutics, Richmond, CA, USA
2
Department of Hematology, St Jude Children’s Research Hospital, Memphis, TN, USA
1
14.1 Introduction
Genome editing technologies, such as engineered nucleases, base editors, or
prime editors, can make permanent genomic modifications to patient cells. These
modifications can result in the disruption of the sequence of a disease‐associated/
mutated gene, correction of a mutated gene, or insertion of a corrective gene into
a precise genomic location. While these technologies remain relatively new and
defining efficacy, durability, and safety in the clinic is ongoing, genome editing
holds enormous potential for treating both inherited and acquired disorders. To
evaluate efficacy and long‐term safety, both on‐ and off‐target editing should be
evaluated and monitored during preclinical and clinical development.
This chapter focuses on the different types of engineered nucleases used in clinical studies and their mechanisms of action leading to induction of double‐strand
breaks (DSBs). A brief overview of US Food and Drug Administration (FDA) and
European Medicines Agency (EMA) regulatory guidance for assessing on‐ and
off‐target nuclease activity is provided, as well as a summary of initial clinical
studies employing engineered nucleases for potential therapeutic benefit. The
workflow to evaluate the efficiency of editing at the intended sites as well as at
potential off‐target sites is described. A detailed overview of methodologies is provided to evaluate the activity of engineered nucleases by quantifying short indels.
347
Drug Development for Gene Therapy: Translational Biomarkers, Bioanalysis, and Companion
Diagnostics, First Edition. Edited by Yanmei Lu and Boris Gorovits.
© 2024 John Wiley & Sons, Inc. Published 2024 by John Wiley & Sons, Inc.
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348
Methods for detecting large genomic rearrangements will also be described. A
special focus will be given to bioanalytical characterization of next‐generation
sequencing (NGS) assays for detection of short indels. The methods described in
this chapter are applicable to both exvivo and invivo genomic editing strategies.
The methodologies used to characterize gene correction, transgene, and vector
integration as well as use of base editors, primer editors, and epigenic regulators
are described elsewhere[1, 2].
14.1.1 Genome Editing Modalities and Molecular Outcomes
There are four main types of engineered nucleases currently available in a drug
developers’ toolbox–meganucleases[3, 4], zinc finger nucleases (ZFNs)[5], transcription activator like effector nucleases (TALENs) [6], and clustered regularly
interspaced short palindromic repeats (CRISPR)‐associated nucleases (Cas)[7, 8].
Meganucleases, ZFNs, and TALENS use DNA‐binding protein domains to recognize
specific DNA sequences while CRISPR‐Cas nuclease employs guide RNA molecules
(gRNA) to target specific DNA sequences through Watson‐Crick base pairing [9].
Regardless of the mechanism of DNA recognition, these technologies are engineered
to create a DSB at a specific site within the genome. Nuclease‐induced DSBs then
trigger one of two main cellular repair mechanisms – homolog‐directed repair
(HDR) and non‐homologous end joining (NHEJ). HDR is a high‐fidelity DNA
repair mechanism that relies upon a donor DNA template containing sequences
homologous to the cleaved ends of the DSB. When supplied together with an engineered nuclease, the donor DNA template can be used to correct disease‐causing
mutations, to insert a therapeutic gene into a genomic safe harbor locus or into its
endogenous locus[10]. NHEJ is error‐prone repair mechanism that can lead to
insertion or deletion of a small number of nucleotides (Figure14.1) or insertion
of larger, nonspecific fragments of the donor DNA template without involvement
of the homology arms[10]. Generation of indels can be leveraged for therapeutic
applications by introducing mutations at a specific site to disrupt the DNA
sequence of a target gene, which can then result in desired clinical outcomes. For
example, editing and subsequent disruption of the BCL11A enhancer gene by
engineered nucleases has shown clinical benefit in β‐thalassemia and sickle cell
anemia (see Section14.1.2). The frequency of indels at the intended site thus can
serve as biomarkers to measure the efficiency of gene editing and as surrogate
potency for the desired biological outcome.
Apart from editing at the intended genomic site, unintended nuclease‐induced
DSBs can result in associated genotoxic events such as off‐target indels, inversions, or translocations. Small insertions and deletions can result in frameshift
mutations resulting in lack of production of specific protein or production of a
truncated non‐functional protein. Translocations can occur when the same cell
14.1 Introduction 349
DNA damage activates DNA repair pathways
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Nuclease introduce DSB
1
2
NHEJ introduces indelsHDR introduces specic change
Insertion
Deletion
Figure14.1 Molecular outcomes of gene editing events by engineered nucleases.
Engineered nucleases introduce a double-stranded break (DSB) in genomic DNA (step1).
This DNA damage activates intrinsic DNA repair pathways with a cell (step2). In the
presence of a donor template, homology-directed repair (HDR) can result in integration of
donor transgene into the genome through homologous recombination, at mostly G2 and
S phases of the cell cycle (step3). Non-homologous end joining (NHEJ), error-prone repair,
is active throughout the cell cycle and does not require a repair template. This repair
mechanism introduces small nucleotide insertions and deletions (indels) at the DSB site
(step4). Source: Created with BioRender.com.
contains more than one DSB e.g. one at the intended and one at an off‐target site
or between two off‐target sites. Chromosomal rearrangements were also observed
when only one DSB was detected at the intended site[11, 12].
In addition to off‐target editing due to non‐specific recognition of DNA
sequences, other potential genotoxic events have been described during editing.
This includes nonspecific single‐stranded DNA cleavage by CRISPR‐Cas12a[13]
and large deletions at the site of intended editing[11, 14]. Lastly, all genome editing approaches implementing synthetic DNA repair templates are susceptible to
random integration at DSBs independent of nuclease activity[15].
The health risks related to off‐target editing in the clinic are not well understood, particularly when off‐target editing occurs at a very low frequency or in
intergenic and/or intronic genomic regions.
Due to the potential risk of genotoxic events posed by DSB introduction, the
genome editing field is exploring other means of editing genes or modulating
gene activity. Indeed, in addition to their endonuclease activity, zinc finger proteins (ZFPs) and CRISPR‐Cas9 can be engineered to create other therapeutic
modalities. ZFPs and catalytically inactive Cas9 can be fused to transcription factors that modulate mRNA transcription [16, 17] without inducing DSBs. For
aphaeresis
Ex vivo In vivo
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350
example, ZFP‐transcriptional factors have been used for allele‐selective transcriptional repression of mutant HTT for the treatment of Huntington’s disease.
Further engineering of CRISPR‐Cas proteins has created a diversity of editing
modalities: Cas9 protein was altered to form a nickase that introduces a single‐
stranded break in DNA [7], base editors that can induce transition (and certain
transversion) mutations[18, 19, 20, 21], and prime editors that use Cas9nickase
fused to a reverse transcriptase and prime editing guide RNA (pegRNA) to mediate targeted small insertions, deletions as well as substitutions[22].
14.1.2 Clinical Trials Using Genome Editing Technologies
Clinical therapeutic applications of genome editing comprise exvivo and invivo
gene modifications. There are presently over 50 clinical studies utilizing exvivo
and invivo genome editing strategies listed in clinical trials.gov. For exvivo editing
approach, hematopoietic cells (e.g. autologous hematopoietic stem and progenitor cells (HSPC), autologous CAR‐T cells) are collected from the patients, modified and expanded ex vivo, and then reinfused into patients. The efficiency and
specificity of genomic editing can be assessed in the drug product and samples
collected from patients post infusion. For invivo editing approach, genome editing components are delivered systemically or locally to patients via lipid nanoparticles or recombinant adeno‐associated virus (rAAV), then expressed in target
cells such as hepatocytes (Figure14.2), followed by editing of the genomic target.
Here tissue biopsies are collected, if possible, for assessing editing efficiency.
Cell
Figure14.2 and genome editing for clinical applications. Left: Ex vivo
genome editing. Cells are isolated from a patient, edited, activated/expended, and infused
back into patient. Right: In vivo genome editing. Engineered nucleases are delivered by
viral or nonviral approaches to the patient systemically. Source: Created with
BioRender.com.
Genome
editing
Edited cells
infused back
into patient
Lipid nano particles
or viral vectors with
genome editing
therapeutic product
14.1 Introduction 351
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ZFNs pioneered the genome editing field[5] with the first exvivo and invivo
clinical studies. In initial preclinical efforts, autologous T cells were modified
exvivo to disrupt the C–C motif chemokine receptor 5 (CCR5) gene, with the aim
to induce resistance to HIV infection[23] (NCT03617198) as disruption of CCR5
restricts the ability of HIV to enter CD4+ T cells through the cell surface expressed
CCR5 receptor. This was followed by several gene‐edited cell therapy clinical studies with HIV patients (add references). Both ZFN and CRISPR‐Cas9 technologies
were used to disrupt the BCL11A erythroid‐specific enhancer in autologous
HSPCs to reactivate fetal hemoglobin (HbF) expression. This disruption consisted
of NHEJ derived indels resulting from the nuclease‐derived targeted DSB [24].
The expression of HbF was expected to ameliorate the symptoms of both beta
thalassemia and sickle cell anemia. In these studies, cells were collected from
individual patients, edited exvivo and then reinfused into patients after myeloablation (ClinicalTrials.gov Identifier: NCT03655678; NCT03745287; NCT03653247)
(Figure 14.2). Further examples of such ex vivo editing include CRISPR‐Cas9,
TALENs and meganucleases to engineer autologous and allogenic chimeric antigen receptor T lymphocytes (CAR‐T) for antitumor immunity (ClinicalTrials
.gov Identifier: NCT02735083; NCT02808442; NCT02746952; NCT03081715;
NCT02793856; NCT04244656; NCT04035434; NCT04142619; NCT03190278;
NCT04150497; NCT04649112).
The first‐in‐human invivo editing studies utilized ZFNs and corrective transgene
components packaged in recombinant AAVs and delivered intravenously to permanently modify patient hepatocytes. Expression of the ZFNs was driven by a
liver‐specific promoter. The ZFNs targeted the albumin intron 1locus in hepatocytes, a safe harbor site, and following induction of a DSB resulted in the insertion
of a corrective transgene at the albumin locus (Figure 14.2). This strategy
was applied for the treatment of mucopolysaccharidosis type I (MPS I; Clinical
Trials.gov Identifier NCT02702115), mucopolysaccharidosis type II (MPS II;
ClinicalTrials.gov Identifier NCT03041324), and hemophilia B (ClinicalTrials
.gov Identifier NCT02695160). In the MPS II clinical study, one patient showed
transient plasma transgene protein at therapeutic levels, but expression was
diminished due to a suspected immune response, indicated by elevated levels of
alanine transaminase and aspartate transferase. Protein expression did not reach
therapeutic levels in other patients in the studies[25]. In another study, CRISPR‐
Cas9mRNA and single guide RNA were targeted to the liver using apolipoprotein
E‐modified lipid nanoparticles to address transthyretin amyloidosis through permanent disruption of the transthyretin gene (TTR)[26]. The study showed a 96%
reduction of TTR in patient serum. If this treatment is proven to be durable, it is
expected to improve disease symptoms.
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352
14.2 Regulatory Guidance onEngineered Nuclease
On- and Off-target Assessment
As interest is exponentially increasing for the development of new therapies
derived from engineered nucleases, harmonized regulatory guidance is needed to
ensure the appropriate characterization of safety profiles and risk assessments of
these genomic medicines. The FDA issued a draft guidance in 2022 to provide
recommendations for assessment of safety and quality of products incorporating
genome editing in human somatic cells to support Investigational New Drug
(IND) applications[27]. The aim of this guidance is to assist in translation of gene
editing products from laboratory bench to clinical studies by providing recommendations for assessing the safety and quality as well as addressing the potential
risks for these products. Some of the risks associated with genome editing include
off‐target editing, unintended consequences as well as the unknown long‐term
effects of on‐ and off‐target editing.
Preclinical studies are recommended to identify and characterize the risk of
genome editing at on‐ and off‐target loci, including identification of off‐target
editing activity, including type, frequency, and location of all off‐target editing
events. Although no specific methods for assessment of off‐target editing are
noted, instead multiple orthogonal methods are recommended for identification
of potential off‐target sites, including an unbiased genome‐wide analysis. Potential
off‐target sites should be verified using methods with adequate sensitivity to
detect low‐frequency events. The acceptable sensitivity for detecting low‐
frequency events was not specified by the agency and would likely depend on
specific genomic location and risk posed by the editing. The analytical methodologies used for off‐target evaluation need to be well described in regulatory submissions, including bioanalytical parameters such as sensitivity, specificity, accuracy,
precision, and description of the reference materials. An assessment of genomic
integrity is also advised, including evaluation of potential chromosome rearrangements, large insertions and deletions, integration of exogenous DNA, and potential
oncogenicity or insertional mutagenesis. Evaluation of the biological consequences
associated with on‐ and off‐target editing is also necessary, as feasible. In addition,
characterization is needed for the kinetic profile of genome editing components
expression and editing activity. For clinical studies, consideration should be given
for adequate monitoring of any off‐target editing and adequate assessment of the
outcomes of unintended consequences of on‐ and off‐target editing.
The 2020 European Medicines Agency (EMA) guidance also emphasizes the
importance of characterizing on‐ and off‐target editing in the genetically modified
cells [28]. Since genome editing is a rapidly evolving field, EMA recommends
using current scientific knowledge for selecting a strategy for evaluating on‐ and
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off‐target activity. Additionally, on‐target genome editing should be characterized
to establish that the target site is correctly edited and no unintended changes have
occurred. To evaluate off‐target activity, in silico prediction and at least one sensitive and well‐characterized experimental assay should be used in the cell type that
will be used therapeutically, and an unbiased genome‐wide evaluation of off‐target
activity invitro is also necessary. The chosen assay strategy should be justified,
and the sensitivity of the methods should be indicated. In addition, the possibility
of large genomic rearrangements needs to be evaluated based on the actual profile
of on‐ and off‐target edits. The risk associated with off‐target activity and large
genomic rearrangements should be assessed in the therapeutic cell type. EMA
and FDA guidance documents do not recommend specific methods or acceptable
levels of assay sensitivity, most likely due to the evolving nature of the methods to
assess off‐target activity.
With multiple clinical trials ongoing and forthcoming, sensitive and specific
methods are needed to assess genome editing outcomes and potential risks to
patients. Best practices for indel assessment are being discussed in the genome
editing, bioanalytical, and safety assessment field, yet the number of publications
and white papers remain limited[29], while other genetic outcomes such as long
rearrangements are yet to be discussed in the bioanalytical literature.
The National Institute of Standards (NIST)‐led Genome Editing Consortium
was recently organized[30], and encompasses collaboration between NIST, industry experts, academia, and other government agencies. One of the aims of this
consortium is to establish tools for understanding reproducibility, performance,
and comparability of the assays used for detecting genome editing outcomes. This
work is currently ongoing, and its results are highly anticipated by the field.
This review discusses the current and evolving methods for these assessments
of on‐ and off‐target activity and genomic integrity. Some of these methods are
more focused on invitro preclinical assessments and others for assessing editing
in invivo preclinical studies as well as in clinical studies.
14.3 Strategies and Methodologies toEvaluate
On-target and Off-target Activities
14.3.1 Strategies toEvaluate Off-target Sites inPreclinical and
Clinical Studies
An approach to select the therapeutic genome editing lead candidate with the
least potential off‐target sites can be divided into several phases. First, in the discovery phase, a broad range of methods are utilized in combination to identify the
genome‐wide activity of engineered nucleases and nominate a list of candidate
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off‐target sites. Next, in the validation phase, targeted high‐throughput sequencing or other comparable approaches can be used to measure editing frequencies in
relevant cell types or samples from preclinical in vivo studies. Finally, the validated sites can then be quantitatively measured using well‐characterized or qualified bioanalytical assays in patient samples from clinical studies (Figure14.3).
Broadly, methods for discovering the unintended genome‐wide off‐target activity of genome editors can be divided into in silico, cellular, and biochemical categories. In silico nomination methods are easy and inexpensive, but their ability to
accurately rank off‐target activity remains unreliable[31]. Cellular methods are
the most direct but have limitations in terms of sensitivity. Biochemical methods
are the most sensitive but may nominate sites that are not modified in cells at
frequencies above assay limits of detection and may lack influences of chromatin
on genomic DNA structure and accessibility. To identify off‐target sites as comprehensively as possible, multiple orthogonal methods in multiple donors are
recommended[27].
Here, we describe cellular and biochemical methods for discovering the
genome‐wide off‐target activity of editors with an emphasis on those that have
been more frequently used to characterize therapeutic genome editing candidates.
Biochemical methods
CIRCLE-seq
gDNA
CHANGE-seq
treated by
Digenome-seq
nuclease
SITE-seq, etc.
in vitro
Discovery
In silico tools Cellular methods
Homology based
computational prediction
by sequence alignment
with human genomes
Nomination
Validation
Monitoring
Candidate off-targets list and ranking
Targeted NGS in cell type of
interest and/or in vivo samples
Bioanalytical assay to measure
validated off-targets in patients
Cells
transfected
with
nuclease
GUIDE-seq
Discover-seq
BLESS/
BLISS
IDVL capture
HTGTS, etc.
Figure14.3 Workflows for evaluating off-target activities during genome editor drug
development. Characterization of off-target activities is divided into different phases.
Discovery can include in silico prediction, cellular and biochemical methods that result in
candidate off-target list followed by validation of candidate off-target sites using
targeted NGS in the cell type of interest. The validated sites are then quantitatively
measured during the clinical studies. Source: Created with BioRender.com.
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14.3.2 Techniques toIdentify Genome Wide Off-target Sites
Cellular methods to define the genome‐wide off‐target activity of genome editing
nucleases have become broadly adopted to characterize the specificity of editors
for therapeutic treatment of diseases such as inherited childhood blindness and
sickle cell disease.
There are now a number of cellular methods to define genome‐wide off‐target
activity of genome editing nucleases that include: GUIDE‐seq[32–34], Discover‐
seq [35], Discover‐seq+ [36], BLESS/BLISS [37, 38], IDLV capture [39], and
HTGTS[40].
Of these cellular methods, DNA end‐capture methods like GUIDE‐seq have
become the most widely used. GUIDE‐seq is based on the principle of efficient
integration of end‐protected short DNA tags into the sites of nuclease‐induced
DNA DSBs followed by tag‐specific amplification and high‐throughput sequencing of flanking genomic DNA (gDNA). Its advantages are that it is fairly sensitive,
with the capability of detecting off‐target sites with mutation rates as low as 0.1%,
although it is not as sensitive as some biochemical assays (Figure 14.4A).
It works well in some therapeutically relevant cell types, such as T cells[33, 41]
and fibroblasts. Some limitations are that there may be dsDNA‐related toxicity or
variable integration rates in some cell types, such as human hematopoietic stem
cells or iPS cells. Quantitative tag integration proportional to indel mutation frequencies also implies that linear increases in number of input genomes and
sequencing would be required to scale GUIDE‐seq to higher sensitivity. GUIDE‐
seq and related end‐capture‐based methods have been used to analyze the specificity of ZFNs and TALENS as well[42, 43].
A number of biochemical methods to define the genome‐wide off‐target activity
of genome editors have also been developed. The advantages of biochemical
methods are that they have the potential to be more sensitive and scalable to many
target sites.
Biochemical methods for defining the genome‐wide activity of therapeutic genome
editors include CIRCLE‐seq[44], CHANGE‐seq[45], Digenome‐seq[46], and SITE‐
seq[47]. Digenome‐seq was one of the first biochemical methods to be developed
and is based on the principle of whole‐genome sequencing of nuclease‐modified
gDNA and bioinformatic of reads that have a signature of editing, such as uniform
start positions. An advantage is that it is simple to practice and PCR‐free; limitations
are that it requires large amounts of sequencing and it may be challenging to distinguish background reads that line up by chance from true signal (Figure 14.4B).
CIRCLE‐seq, CHANGE‐seq, and SITE‐seq are all methods for selectively
sequencing nuclease‐modified gDNA. CIRCLE‐seq and CHANGE‐seq achieve
this by generation of libraries of highly purified, circularized gDNA followed by
treatment with Cas9 ribonucleoprotein complex. Only gDNA circles that have
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