Context

Gene editing–based therapies are currently in development in the areas of oncology, inherited disease, and infectious disease. These potentially life-altering therapies are derived from decades of research in both academic and industry settings that developed technologies rooted in principles and products of nature. However, with such technologic developments come many important considerations, including adverse risks, high cost, and ethical questions.

Objective

To educate pathologists about gene editing technologies, inform them of potential indications and risks, outline regulatory and practical issues that could affect hospital-based practice and laboratory testing, and advocate that pathologists need to be present at discussions among industry and regulators pertaining to gene editing–based therapies.

Design

A Gene Editing Workgroup, facilitated by the College of American Pathologists Personalized Health Care Committee and consisting of pathologists of various backgrounds, was convened to develop an educational paper to serve as a stimulus to increase pathologist involvement and inquiry in gene editing therapeutic and diagnostic implementation.

Results

Through multiple discussions and literature review, the workgroup identified potential gaps in pathologists' knowledge of gene editing. Additional topics that could impact pathology and laboratory medicine were also identified and summarized in order to facilitate pathologists as stakeholders in gene editing therapy administration and monitoring and potential use in diagnostics.

Conclusions

Gene editing therapy is a complex but potentially transformative area of medicine. This article serves as an introduction to pathologists to assist them in future discussions with colleagues and potentially identify and alter pathology practices that relate to gene editing.

During the last 10 years, the concept and science behind the idea that genes could be modified in order to treat a disease has come to fruition with the recent publication of successful clinical trials in β-thalassemia, sickle cell disease (SCD), and Leber congenital amaurosis.1,2  Historically, this medical revolution is akin to the development of vaccines and antibiotics, but will affect not only infectious disease treatment, but also treatment of life-ending or life-altering inherited diseases, cardiovascular disease, diabetes, and cancer. With such a powerful tool, however, come ethical and regulatory questions that require serious deliberation, education, discussion, and challenging dialogue. Importantly, from a pathologist's perspective, one must also consider how such manipulation will affect preanalytical, analytical, and postanalytical factors in the practice of anatomic and clinical pathology. This article serves as an educational summary for pathologists, outlining the history, development, and technology of gene editing. It also provides an initial point of discussion regarding the rapidly evolving clinical, ethical, and regulatory environment as gene editing–based therapies and diagnostics are introduced into clinical practice. It should be stated that the major focus of this article is on somatic gene editing, that is, editing of a focused cell of interest that is involved in a particular disease. This is opposed to germline gene editing, which results in editing of all or most cells within an individual, which has been the subject of much ethical debate. The ethical issues surrounding gene editing will be discussed at the end of this article.

Gene editing refers to a type of genetic engineering that inserts, deletes, modifies, or replaces DNA at targeted locations within the genome of an organism. Gene editing technologies have evolved over time as newly discovered, naturally occurring molecular factors have been identified. However, there are 4 necessary elements that are required: (1) an engineered system to identify and target the area of modification and serve as an anchor; (2) a nuclease (engineered or recombinant) that can either create the double-stranded DNA breaks (DSDBs) within the targeted sequence or target DNA without cleavage (base editing); (3) inherent cellular processes that alter a mutated base to its desired alternative, repair the break to introduce a deleterious error to inactivate a gene, or introduce an amended functional sequence; (4) a delivery mechanism or vector that allows the system to reach the nucleus of a cell. Most gene editing technology borrows from natural DNA damage and repair processes and uses targeting sequences to achieve precise genetic modification.35 

Zinc finger nucleases (ZFNs) and transcription activator–like effector nucleases (TALENs) were sequentially introduced as gene editing apparatuses that combine sequence-targeting protein polymers that bind the DNA of interest with a FokI nuclease (derived from Flavobacterium okeanokoites) to induce a DSDB (Figure 1).4,5  ZFNs are composed of a combination of zinc finger proteins that are approximately 30 amino acids long and a nuclease domain from FokI. Specific zinc fingers target a unique 3- to 6- nucleotide sequence, and combining zinc fingers allows a design to match approximately 3 to 6 codons (9–18 nucleic acids). The FokI nucleases work as dimers. ZFNs have the disadvantage of high cost, complexity of protein design, and inaccurate cleavage in many settings; however, they are still useful.4  TALENs use the same FokI nuclease but the DNA targeting is more flexible than ZFNs as each monomer (TALE) recognizes a single nucleotide. These single nucleotide–recognizing structures, which are approximately 35 amino acids long, can be assembled as a polymer designed to identify a specific sequence.4  Whereas TALENs are more versatile (owing to mononucleotide versus trinucleotide recognition), the large structural nature of TALENs results in more limitations on the viral vectors that can be used for delivery.

Figure 1

Zinc finger nucleases (ZFNs) and transcription activator–like effector nucleases (TALENs). ZFNs and TALENs use bioengineering of naturally occurring protein domains to bind to specific DNA sequences (both sense and antisense). Cleavage/breakage of double-stranded DNA (DSB) occurs via dimerized FokI nuclease. Cellular repair mechanisms such as homology-directed repair (HDR) or nonhomologous end joining (NHEJ) repair the break. HDR uses an oligonucleotide (oligo), or sister chromatid (SC), to guide a more precise edit. Less-precise NHEJ results in frameshift disruption and gene inactivation. This figure was created with Biorender.com.

Figure 1

Zinc finger nucleases (ZFNs) and transcription activator–like effector nucleases (TALENs). ZFNs and TALENs use bioengineering of naturally occurring protein domains to bind to specific DNA sequences (both sense and antisense). Cleavage/breakage of double-stranded DNA (DSB) occurs via dimerized FokI nuclease. Cellular repair mechanisms such as homology-directed repair (HDR) or nonhomologous end joining (NHEJ) repair the break. HDR uses an oligonucleotide (oligo), or sister chromatid (SC), to guide a more precise edit. Less-precise NHEJ results in frameshift disruption and gene inactivation. This figure was created with Biorender.com.

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CRISPR (clustered regularly interspaced short palindromic repeats)–Cas is perhaps the best recognized system that can be used for gene editing (Figure 2). Starting in 1989, Francisco Mojica and subsequent researchers described unique, approximately 30-base-pair, repetitive, palindromic regions (later termed CRISPR) separated by 30-base-pair spacers in a salt-tolerant microbe.6,7  These repetitive sequences, and those identified in other microbes including Yersinia pestis, Streptococcus pyogenes, and Streptococcus thermophilus, were later theorized, and subsequently proven, to be from foreign (phage or viral) material.810  Further studies identified a Cas protein nuclease that could cleave bacterially derived RNA containing CRISPR repeats to make a guide RNA to target invading phage or viral DNA sequences.11  A protospacer adjacent motif (PAM) was also discovered as a sequence necessary to target the CRISPR-Cas to the sequence to be cleaved 3 nucleotides upstream from the PAM site.8,12  Targeting of the nuclease occurs via a bacterially developed, virally targeted duplex CRISPR guide, crRNA-tracrRNA, thereby cleaving and destroying the invading virus or phage sequence (CRISPR RNA [crRNA] serves as the sequence viral/phage target guide and is hybridized to the transactivating CRISPR RNA [tracrRNA] that serves as a Cas9 anchor).1215  Thus, through the work of many groups, the CRISPR-Cas system was proven to be a system of bacterial adaptive immunity, used to cleave double-stranded DNA (dsDNA) from an invader by a nuclease. In 2012, it was demonstrated that this system could be adapted as a tool for programmable gene editing, monumental work that was awarded the 2020 Nobel Prize Award in Chemistry.16  Furthermore, in 2013, it was shown that CRISPR-Cas could be translated to eukaryotic cells, including human cells, and programmed for gene editing in these cells.17,18 

Figure 2

Clustered regularly interspaced short palindromic repeats–Cas system. This system consists of a Cas nuclease and an engineered single guide RNA (sgRNA) designed to target the gene of interest. The sgRNA binds to the antisense strand of a target gene to open up the DNA molecule. Cleavage occurs upstream of the protospacer adjacent motif (PAM). The Cas protein binds both the guide RNA and the PAM site, allowing for cleavage, resulting in a double-stranded break (DSB). Homology-directed repair (HDR) or nonhomologous end joining (NHEJ) can then occur. This figure was created with Biorender.com. Abbreviations: oligo, oligonucleotide; SC, sister chromatid.

Figure 2

Clustered regularly interspaced short palindromic repeats–Cas system. This system consists of a Cas nuclease and an engineered single guide RNA (sgRNA) designed to target the gene of interest. The sgRNA binds to the antisense strand of a target gene to open up the DNA molecule. Cleavage occurs upstream of the protospacer adjacent motif (PAM). The Cas protein binds both the guide RNA and the PAM site, allowing for cleavage, resulting in a double-stranded break (DSB). Homology-directed repair (HDR) or nonhomologous end joining (NHEJ) can then occur. This figure was created with Biorender.com. Abbreviations: oligo, oligonucleotide; SC, sister chromatid.

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CRISPR-Cas systems (Cas9 derived from S pyogenes is the most common but many other Cas systems exist) are designed with a single guide RNA (sgRNA), essentially engineered to reflect crRNA combined with tracrRNA, and a Cas protein nuclease to induce a DSDB and stimulate gene-targeted repair and local mutagenesis.19  DSDBs (induced by ZFNs, TALENs, or CRISPR-Cas) can be repaired either by homology-directed repair, where a donor sequence (from the corresponding sister chromatid or an engineered oligonucleotide) is copied; or by nonhomologous end joining (NHEJ), which joins the broken ends together but occasionally leads to small insertions or deletions (indels) at the break site, often inactivating the gene product (Figures 1 and 2).4  Thus, normal NHEJ repair process would inactivate genes, whereas homology-directed repair can be manipulated to replicate a synthetic sequence at the site, introducing a wild-type sequence in lieu of the mutated sequence or a mutated sequence that could provide resistance to an infectious agent (such as human immunodeficiency virus [HIV]). In vivo, NHEJ repair mechanisms are more common, and therefore, diseases corrected by gene product inactivation have been more commonly investigated.

Whereas the above systems are most useful in disrupting a gene to produce a desired response, many diseases (inherited and somatic) are caused by single-nucleotide variants that potentially could be corrected by base editing (Figure 3). Base editing technologies take advantage of naturally occurring enzymes that can convert 1 nucleotide to a different nucleotide, thereby “repairing” a problematic mutation.20,21  Base editing can manipulate DNA or RNA, but as RNA base editing is currently used for research (it is not a permanent genomic edit to therapeutically correct a disease) it will not be discussed here. DNA base editors include C:G to T:A base editors, based on usage of a cytosine deaminase enzyme that can convert cytosine to uracil, which pairs as a thymidine in DNA. There is also development of A:T to G:C base editors that take advantage of adenosine deamination enzymes to convert adenosine to an intermediate called inosine, which is structurally similar to and is read as a guanosine. Such adenosine editing would result in correction of a G>A (C>T) mismatch, the most common form of DNA mutation in ClinVar, back to a G (or C).20  Such editing technologies have required innovative modification to properly perform the desired effect.20,21  These deaminases are complexed with a Cas nickase enzyme (DNA nicking on the unedited DNA strand encourages base editing of the opposite strand) and a guide RNA to open up the dsDNA and properly target the mutated base to be edited. For cytosine deaminases, a uracil glycosylase inhibitor is also engineered into the construct to reduce unwanted excision of the converted uracil by a natural-occurring uracil DNA glycosylase, allowing the uracil to persist long enough for future incorporation by a thymidine. No such modification is required to inhibit inosine removal in adenosine base editing, as naturally occurring adenosine editors prefer single-stranded RNA (ssRNA) as a target. Base editing is advantageous in that it does not require DSDBs (ideally reducing the chance for undesired structural alterations), and theoretically could correct more than 60% of reported pathogenic single-nucleotide variants.20  Base editing can correct the 4 transition mutations (C>T, G>A, A>G, T>C) but cannot correct the remaining 8 transversion mutations or correct an insertion or deletion of 1 or more nucleotides. Prime editing is another gene editing technology that can correct the alterations, as it identifies a site to be modified and provide instructions for the edit, without a need for DSDBs or donor DNA templates.22  Prime editors use a guide RNA that both targets the DNA of interest and contains the sequence to be copied and a reverse transcriptase fused to a Cas nickase.22  The nickase breaks 1 strand of the DNA at the target area, and the sequence RNA is reverse transcribed onto the nicked DNA.22 

Figure 3

Base editing. A, Cytosine base editing allows for correction of a C:G mutation back to T:A sequence. Targeted machinery consists of a clustered regularly interspaced short palindromic repeats–Cas nuclease engineered with a single guide RNA (sgRNA), a cytosine deaminase (CytD), and a uracil DNA glycosylase inhibitor (UGI). The sgRNA/Cas opens up the target DNA helix, and the CytD deaminates the cytosine to a uracil (which is recognized as a thymidine in replication). The Cas nuclease has been engineered to create a single-stranded nick in the DNA (scissors) to target repair machinery to the site. Cellular repair and replication mechanisms result in a normal/wild-type edited nucleotide of T:A. The UGI enzyme is required to inhibit naturally occurring cellular uracil DNA glycosylase from causing uracil base excision. B, Adenosine base editing functions similarly to correct A:T mutations back to wild-type G:C. After targeting of the machinery and DNA helix opening, adenosine deaminase (AdeD) converts adenine to inosine (subsequently recognized as a guanosine in replication). As there is minimal cellular desire for base excision of inosine (owing to its rarity), an added inhibitor is not necessary. This figure was created with Biorender.com. Abbreviation: PAM, protospacer adjacent motif.

Figure 3

Base editing. A, Cytosine base editing allows for correction of a C:G mutation back to T:A sequence. Targeted machinery consists of a clustered regularly interspaced short palindromic repeats–Cas nuclease engineered with a single guide RNA (sgRNA), a cytosine deaminase (CytD), and a uracil DNA glycosylase inhibitor (UGI). The sgRNA/Cas opens up the target DNA helix, and the CytD deaminates the cytosine to a uracil (which is recognized as a thymidine in replication). The Cas nuclease has been engineered to create a single-stranded nick in the DNA (scissors) to target repair machinery to the site. Cellular repair and replication mechanisms result in a normal/wild-type edited nucleotide of T:A. The UGI enzyme is required to inhibit naturally occurring cellular uracil DNA glycosylase from causing uracil base excision. B, Adenosine base editing functions similarly to correct A:T mutations back to wild-type G:C. After targeting of the machinery and DNA helix opening, adenosine deaminase (AdeD) converts adenine to inosine (subsequently recognized as a guanosine in replication). As there is minimal cellular desire for base excision of inosine (owing to its rarity), an added inhibitor is not necessary. This figure was created with Biorender.com. Abbreviation: PAM, protospacer adjacent motif.

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A gene editing system can be delivered either in vivo or ex vivo and can use various delivery vectors to get the system into the cell and subsequently, into the nucleus.20,23,24  In vivo delivery involves delivery of the gene editing material into the body, without taking cells out of the body. Ex vivo refers to removing cells from the body (eg, T cells or CD34+ stem cells), manipulating those cells with gene editing apparatus, and then returning the cells back to the body. At this time, most gene editing clinical trials involve some sort of ex vivo manipulation, unless it is a therapy directed to a specific organ (eg, eye).

Delivery vectors may be viral, viral-like, or nonviral. Viral vectors can be used for both in vivo and ex vivo delivery and have various pros and cons.2325  Recombinant adeno-associated virus (AAV) is the least pathogenic, with only a mild inflammatory response and low immunogenicity, and is the least likely to integrate into the genome, reducing the risk of insertional mutagenesis. Lentiviruses (derived from HIV-1) have the advantages of a larger insert size (10 kb) versus AAV (4.7 kb), such as needed for TALEN use, with the ability to infect nondividing cells. Integration (through the integrase viral gene of lentivirus into the host DNA) also allows for the transgene to be passed on to cell progeny and potentially persist in its expression. There is a higher risk of insertional mutagenesis and pathology associated with lentiviruses, although manipulation of the lentiviral structure has been repeatedly refined to improve the safety profile, including removal of the integrase genetic component to reduce the risk of insertional mutagenesis.23,25 

Nonviral methods have also been used in the delivery of gene editing apparatus to cells, including plasmid DNA, messenger RNA (mRNA), and ribonucleoproteins (RNPs). RNPs are a newer delivery system by which CRISPR-Cas9, for instance, can be transported into a nucleus.26  In this setting, a recombinant Cas9 protein is complexed with the sgRNA in a delivery particle. This design can potentially reduce complications associated with viral vector, plasmid, or mRNA delivery. For instance, targeted cells would not need a high transcription and translation capacity and there would theoretically be a decreased risk of mutagenesis or off-targeting effects.26  RNP delivery also results in a quicker time to editing but is most often used in the setting of ex vivo cell manipulation. RNPs require a process to cross the cell membrane for efficient and targeted delivery into the cell of interest. The cationic charge of Cas9, somewhat balanced by the negatively charged sgRNA, must be considered in the development and cell delivery of these systems.

There are many methods for delivery of RNPs through the cell membrane including lipid reagents, such as Lipofectamine, cell-penetrating peptides that are covalently or ionically conjugated to Cas9 protein, lipopeptides, or various polymers.26  Such components take advantage of membrane permeability and more so, cellular machinery allowing for endocytosis. Physical microinjection is another mechanism that can directly target a cell population of interest but requires skill so as to not produce physical trauma. Electroporation uses small electrical bursts to briefly disrupt the cell membrane, allowing for influx of RNPs; however, this method can only be used for ex vivo delivery to cells, to be subsequently infused into the patient. Electroporation can potentially cause cell death. Virus-like particles, which package Cas9 protein with other viral proteins and with the sgRNA, have been used, but can elicit an immune response. Other physical and biochemical approaches that have been investigated in eukaryotic, prokaryotic, and plant-based cells have been further reviewed beyond the scope of this article.26 

If an in vivo approach is used, there are many hurdles to delivery of the gene editing system to the cells of interest, including macrophage phagocytosis, various destructive enzymes (DNases, RNases, and proteases), immune activation, and premature excretion.23  One also must consider a targeted approach to cell delivery so as to limit introduction of the gene-altering apparatus into undesired cell types. Additionally, after the gene editing apparatus arrives in the cell (either via in vivo or ex vivo methods), the machinery must escape the endosome. It then must be released and targeted appropriately to the cytoplasm (for mRNA translation of encoded Cas9 protein if need be) and to the nucleus for actual editing to occur.23 

The development of gene editing therapies is similar in its phases and considerations to conventional therapies but also includes unique concerns associated with the action of the gene editing machinery on the genome. The preclinical phase of development seeks to demonstrate feasibility and identify potential toxicity in vitro and in animal models.

Gene editing therapeutics are designed to optimize on-target activity in order to promote efficacy while minimizing off-target activity that may lead to adverse events. As many gene editing approaches rely on making precise DSDBs, there is a risk that off-target editing may contribute to neoplasia owing to the disruption of tumor suppressor genes. Modern laboratory approaches seek to quantify the degree of on-target and off-target gene editing during preclinical development. In silico, computational tools, with the knowledge of the human genome sequence, aid in choosing optimal gene editing therapy designs that maximize on-target activity while minimizing potential off-target effects. In the case of CRISPR-Cas9, the sequence of the sgRNA as well as the selection of the particular Cas9 protein contribute to the targeting of the gene editing machinery. Once selected, the on-target and off-target effects of potential gene editing therapies can be assessed with high-throughput sequencing. Targeted deep sequencing can quantify the proportion of alleles edited at the on-target site. Off-target activity can be assessed by targeted monitoring of sites predicted to be at risk for off-target editing, such as those that resemble, in homology, the on-target sequence. Additionally, specialized library preparation and sequencing methodologies can be used to identify the creation of DSDBs in an unbiased manner. As gene editing therapies involve human DNA sequences, human cells (including human cellular disease models derived from pluripotent stem cells) represent an important tool for assessing gene editing activity on the genome.

The Center for Biologics Evaluation and Research/Office of Cellular, Tissue and Gene Therapies (OCTGT) of the US Food and Drug Administration (FDA) issued a guidance document in November 2013 on preclinical assessment of investigational cellular and gene therapy products.27  This guidance provided recommendations on the general framework for planning and designing preclinical studies for investigational cellular and gene therapies (referred to as CGT products). This document acknowledged the unique scientific challenges posed by CGT products in regulatory review due to the diverse biology and clinical indications, as well as the rapidly evolving research. Therefore, a careful risk-benefit analysis in the context of the particular clinical indication under study is necessary, as is a flexible, science-driven process to incorporate the basic toxicologic principles to address safety issues. The recommendations given by OCTGT for general preclinical program design covered the following major aspects: investigational CGT products used in preclinical studies, animal species selection, animal models of disease/injury selection, proof-of-concept studies, toxicology studies, product delivery considerations, good laboratory practice, product development for later-phase clinical trials, preclinical study reports, and communication with OCTGT pharmacology/toxicology staff.

Another FDA guidance document issued by OCTGT in June 2015 focused on considerations for the design of early-phase clinical trials of cellular and gene therapy products.28  It was mentioned that early experiences indicated that some CGT products might pose substantial risks to subjects, with risk-contributing features including potential for prolonged biological activity after a single administration, a high potential for immunogenicity, or the need for relatively invasive procedures to administer the product. The design of early-phase clinical trials of CGT products involves consideration of clinical safety issues, preclinical issues, along with chemistry, manufacturing, and controls issues. Some characteristics of gene therapy products can influence trial design; for example, expression of a delivered gene may be uncontrolled and interfere with normal function of critical biological process in the recipient, or genomic alteration could cause activation or inactivation of neighboring genes, leading to tumors. Specifically, gene-modified cellular products are considered to have features and potential risks of both cellular and gene therapy products. This document described specific elements of the design of an early-phase trial for a CGT product, focusing on the aspects for CGT products that are often different from other types of products. In the section on monitoring and follow-up, several special considerations were highlighted and the ones relevant to gene editing products included evaluation of a subject's immune response to the products if immunogenicity is a concern, determination of the duration of product persistence and activity, addressing potential for migration from the target site, and monitoring for clonal outgrowths.

In June 2020, the FDA provided a further guidance to industry sponsors developing gene therapy products, relating to the monitoring of long-term effects of gene therapy products.29  These recommendations outlined the risks that gene therapy can present, including off-targeting events or unwanted DNA breaks, prolonged transgene expression, viral integration and latency. It also recommended general elements for long-term follow-up observations and protocol development. Items to be monitored included vector persistence; development of new or exacerbated malignancies; neurologic, infectious, or autoimmune disorders; and other adverse events. A framework to assess the risk of gene therapy–related adverse events was provided through a suggested flowchart. Additionally, preclinical design considerations were outlined to assess biodistribution and persistence of gene therapy, including animal and tissue studies. In this draft, submission of a Pharmacovigilance Plan was recommended with an application to include long-term follow-up. As of this writing another draft guidance is currently underway (March 2022) outlining recommendations for industry sponsors developing human gene therapy products incorporating gene editing of somatic cells.30 

Hereditary disorders were the first group of diseases to be addressed in the early gene replacement therapy trials, so it seems natural that they would also be seen as promising targets for gene editing therapies. The long road to the present juncture has been rocky, with several high-profile fatalities of patient-subjects, resulting in long periods of moratorium during which no gene therapy trials were approved. Most of these deaths were caused by immunologic reactions to the viral vectors used to deliver the transgene to the target organ, or by unintended insertion of the transgene near an oncogenic locus, resulting in iatrogenic malignancy.31,32  Those factors, along with the inherent expense and long development time of gene therapeutics, have made the prospect of treatment by gene editing far more appealing.

The molecular defects in most genetic disorders are localized to particular organs or tissues where the causative gene is expressed—for example, blood for hemoglobinopathies, liver for metabolic disorders, brain for neurodegenerative disorders. Naturally, those tissues that are physically the most accessible to delivery of the editing reagents, either in vivo or ex vivo, have been addressed first, such as bone marrow cells for β-thalassemia and severe combined immunodeficiency, or retinal cells for Leber congenital amaurosis.1,33,34  More recently, correction of a mutant liver gene, TTR, has been achieved by simple infusion into the bloodstream of the CRISPR complex contained in lipid nanoparticles, raising hope the gene editing of other internal organs may be achievable, without the need for direct injection.35 

One important concept to keep in mind for gene editing therapy of hereditary diseases is that the earlier in the course the therapy can be initiated, the better the outcome. This is especially true for neuromuscular disorders such as Duchenne muscular dystrophy, Huntington disease, spinal muscular atrophy—if one waits too long, there will not be a sufficient volume of the target muscle or neuronal cells to correct. As gene editing successes for an increasing array of diseases are reported, our approach to newborn screening, which currently does not include later-onset diseases, may need to be modified.

As the idea that gene editing machinery could be used to potentially cure human disease materialized, it should come as no surprise that hereditary diseases, especially rare and severe, or with an accessible, focused cellular target, were first to be investigated. As not all potential trials can be discussed, a few will be mentioned here to illustrate how these technologies are being used in the area of hereditary disease. A summary of some important trials can be found in Table 1.

Table 1

Selected Clinical Trials for Inherited Diseases Using Gene Editing

Selected Clinical Trials for Inherited Diseases Using Gene Editing
Selected Clinical Trials for Inherited Diseases Using Gene Editing

Significant blood disorders were an obvious focus, owing to the ability to target CD34+ stem cells that could repopulate the hematopoietic system. The first landmark clinical trials using ex vivo gene editing technology to treat inherited blood disorders were conducted by Vertex Pharmaceuticals in collaboration with CRISPR Therapeutics in patients with SCD and transfusion-dependent β-thalassemia (TDT).1  Both diseases are among the most common monogenic inherited diseases worldwide and caused by mutations in the hemoglobin β subunit gene (HBB) with severe manifestations. The biological named “CTX001” was manufactured by ex vivo genetic editing of the autologous CD34+ hematopoietic stem/progenitor cells (HSPCs) by CRISPR-Cas9. Electroporation of CD34+ HSPCs was performed with CRISPR-Cas9 targeting (via inactivation) of erythroid-specific enhancer of BCL11A (an inhibitor of γ-globin synthesis) to restore γ-globin synthesis and reactivate the production of fetal hemoglobin. The delivery of the treatment was through a single infusion via central venous catheter following myeloablative conditioning with busulfan. Frangoul et al1  reported in 2021 the trial results of 2 patients: 1 with SCD and 1 with TDT. With more than a year of follow-up post therapy, both patients retained high levels of allelic editing in bone marrow and blood, increased fetal hemoglobin, transfusion independence, and absence of vaso-occlusive episodes. Both safety and efficacy study clinical trials using CTX001 for SCD and TDT are active at the time of publication of this article (NCT03745287). A multisite, long-term observational follow-up study in subjects who received CTX001 was also opened on ClinicalTrials.gov with the estimated study completion time in September 2039. Besides CTX001, there are also several other ongoing clinical trials applying ex vivo genetic editing technologies (CRISPR-Cas9 or ZFN) targeting the same BCL11A gene in autologous hematopoietic stem/progenitor cells, followed by infusing the product to the patients with SCD or TDT. More safety and efficacy results are expected to be reported from these clinical trials in the coming years.

Transthyretin amyloidosis, also called ATTR amyloidosis, is caused by accumulation of misfolded transthyretin (TTR) protein in various tissues, including the nerves and heart, and can be life-threatening. In 1 clinical trial (NCT04601051) assessing safety and pharmacodynamics, an in vivo gene editing therapeutic agent designed with the CRISPR-Cas9 system was delivered through a lipid nanoparticle encapsulating mRNA for Cas9 protein and an sgRNA targeting TTR, with the goal of reducing the concentration of TTR in serum. Safety and pharmacodynamics of single escalating doses in 6 patients with ATTR amyloidosis with polyneuropathy were assessed. A single dose showed durable knockout of TTR. At day 28, the mean reduction from baseline in serum TTR protein concentration was 52% (range, 47%–56%) and 87% (range, 80%–96%) in the groups that received a dose of 0.1 mg/kg and 0.3 mg/kg, respectively. Only mild adverse events were reported.35 

An experimental CRISPR-based treatment has also been designed for the inherited retinal degenerative disease Leber congenital amaurosis 10 (LCA-10), a rare, monogenic cause of early vision loss.36  This therapy aims to correct the mutations in CEP20 that cause LCA-10, thereby restoring normal protein expression. The CRISPR components are packed into nonpathogenic AAV viral vectors that are injected directly into the patient subretinally. Notably, the BRILLIANCE Phase I/II clinical trial (NCT03872479) is designed to enroll both pediatric age (3–17 years) and adult patients. Data from the trial were released in Fall 2020 with some potentially encouraging clinical results in 1, possibly 2, of 5 patients in the mid-dose range and no significant adverse effects (no immune response or AAV effects). Notably, the first pediatric patient was treated in April 2022, the first time a child has been treated with an in vivo CRISPR-based therapeutic.36 

Hereditary angioedema (HAE) is an autosomal dominantly inherited disease caused by a deficiency in C1 inhibitor due to a mutation in SERPING1. This gene encodes a protease inhibitor in the serpin family that regulates proteins in various pathways of complement, coagulation, and contact activation.37  Patients with this disease can have multisystemic swelling leading to severe facial swelling, abdominal pain, and life-threatening airway obstruction. Bradykinin is the mediator of swelling in these patients through activation of the plasma contact system. A CRISPR-Cas9–based knockout of the target gene KLKB1, coding for kallikrein B1, designed to target hepatocytes through systemic administration, was devised to halt the production of bradykinin, thereby preventing HAE attacks (currently under study by Intellia Therapeutics as NTLA-2002; NCT05120830). The ultimate goal is to cure HAE. This editing system is designed to be delivered as Cas9 mRNA and sgRNA via lipid nanoparticles and is currently under phase I/II studies.

These studies give examples of how various gene editing trials are underway in hereditary diseases. Other diseases include Kabuki syndrome, mucopolysaccharidosis II, α1-antitrypsin deficiency, hemophilia A and B, Usher syndrome, and rhodopsin-associated autosomal dominant retinitis pigmentosa, among others. Of note, with the exception of hemoglobinopathies, many of these diseases are rare, and companies need to satisfy investors. These therapies will thus be very expensive and such trials need to demonstrate not only efficacy, but also proof of principle for future therapies in other diseases. Targeting more common diseases, such as cystic fibrosis (CF) as an example, would prove beneficial but many diseases have drawbacks. Compared to other therapeutic strategies for CF, gene editing to correct mutations at the endogenous CFTR locus is still at an early stage.38  Although many proof-of-concept studies in CF cell models have demonstrated the feasibility of CFTR editing, they lack immediate translational potential. One challenge associated with gene editing is related to the large size of the CFTR gene and the wide distribution and variation of mutations, which would likely require different strategies to be developed and tailored to each specific mutation. Another major challenge for clinical translation of gene editing into a therapy for patients with CF lies in the optimal delivery and successful engraftment in the target organ in the presence of extracellular barriers. Despite the current obstacles, gene editing holds promise as a potential therapeutic for CF and more studies are likely to come to address the hurdles.

Despite great potential, the therapeutic application of gene editing to human solid tumors is in its infancy. Currently, most published studies in this area center on the use of gene editing to improve the efficacy of immunotherapy.39  This includes the use of gene editing to engineer T cells to improve the effectiveness of a patient's antitumor T-cell response.39  An additional, and more preliminary, application of gene editing to solid tumor therapy is direct modification of the tumor cell genome. Laboratory testing, particularly molecular approaches such as next-generation sequencing, will likely play a key role in facilitating the transition of solid tumor gene editing into the clinic. In clinical trials, sequencing assays and targeted molecular tests enable assessments of safety and efficacy of gene editing for solid tumor therapies and allow for molecular characterization of the treated tumor. As gene editing therapies for solid tumors emerge from clinical trials and enter oncology practices, molecular assays will likely continue to be used for monitoring of safety and efficacy.

An example of the use of gene editing for solid tumor immunotherapy is the work of Stadtmauer et al.40  In this phase I clinical trial, investigators used gene editing to modify multiple genes in T cells (ex vivo) removed from patients with advanced, refractory cancers (2 myelomas and 1 liposarcoma). Specifically, they used electroporation of RNP CRISPR-Cas9 to inactivate genes encoding the endogenous T-cell receptor (TCR), as well as the gene encoding the programmed death receptor-1 (PD-1). By removing the endogenous TCR, the investigators could use a lentiviral vector to introduce an HLA-A2*0201–restricted TCR specific for NY-ESO-1 and LAGE-1 (cancer-testis antigens expressed by many cancers). Like antibody blockade of PD-1, genetic inactivation of PD-1 is thought to increase T-cell–driven tumor cell killing by “removing the brakes” activated by the programmed death ligand-1 (PD-L1) on tumor cells. To characterize the safety and efficacy of the genetically modified T cells, a wide range of laboratory tests was performed. Molecular testing before infusion of T cells included digital polymerase chain reaction (PCR) to assess the efficiency of gene editing, quantitative PCR to identify unintended translocations, and a sequencing approach called iGUIDE to assess for the presence of off-target effects from gene editing.41  Following infusion, the engraftment of the edited T cells was assessed by quantitative PCR in both tumor and blood. Finally, single-cell RNA sequencing was used to characterize the transcriptome in the patient with liposarcoma and its evolution over time. Ultimately, the patient with liposarcoma had a 50% reduction in tumor burden following treatment and before disease progression.

A separate study in patients with advanced non–small cell lung cancer (PD-L1 positive) used ex vivo manipulation of T cells (via electroporation) and CRISPR-Cas9 (2 sgRNAs targeting exon 2) to knock out PD-1, similarly theorizing that manipulation would improve T-cell cytotoxicity.42  Twenty-two patients were enrolled and 12 received an infusion. All patients demonstrated edited T cells at 4 weeks, and mean editing efficiency was 5.8% (using next-generation sequencing analysis). Median mutational frequency of off-target events was 0.05% (mostly in noncoding regions). No patients had partial response; however, 2 had stable disease (up to 76 weeks). Unfortunately, all patients had disease progression and 11 of 12 had died as of publication.

Unlike immunotherapy, gene editing applications to directly modify the tumor cell genome are largely restricted to preclinical studies. An example of this approach is the editing of the nuclear factor erythroid 2–related factor (NRF2) gene in non–small cell lung carcinoma. NRF2 is frequently upregulated in cancer; this upregulation increases the resistance of cancer cells to chemotherapy agents through a variety of mechanisms.43  Investigators applied CRISPR-Cas9 to knock out NRF2 in chemoresistant A549 lung cancer cells.44  These cells showed significantly increased susceptibility to chemotherapy agents including carboplatin, cisplatin, and vinorelbine both in culture and in a xenograft mouse model.44  The approach of directly targeting NRF2 with CRISPR-Cas9 in non–small cell lung carcinoma has been proposed for a human clinical trial.45  Gene editing has also been applied to editing somatic mutations in tumor cells. For example, researchers used CRISPR-Cas9 to target mutant KRAS in cell lines and showed inhibition of cancer cell proliferation in vitro and in vivo.46  Challenges to directly targeting tumor cell genomes with gene editing include the requirement for high editing efficiency, as edited cancer cells will have decreased fitness relative to unedited cells.47  Additional challenges include difficulty associated with in vivo delivery and the possibility of off-target effects.47 

The therapeutic applications of gene editing in hematologic malignancies have been developed for decades, and there are many clinical trials in both myeloid and lymphoid neoplasms, although most of these trials are still at the early phases, either phase I or phase II. Representative trials in acute myeloid leukemia, acute lymphoid leukemia, B-cell non-Hodgkin lymphoma, and multiple myeloma are listed in Table 2 to illustrate different gene editing methods, types of edits, targeted genes, and delivery methods. Three major gene editing methods are used, including CRISPR-Cas9, TALENs, and meganucleases. The types of editing are either gene knockout or knock-in, and occasionally gene insertion. In acute myeloid leukemia, the target genes include tumor cell surface markers, such as CD33 or CD123, or genes involved in leukemogenesis such as Wilms tumor-1 gene (WT1). In acute lymphoid leukemia, the main target genes are the T-cell receptor α locus and CD19/20, while in multiple myeloma, the targeted genes are the TCR or CD38.

Table 2

Clinical Trials for Hematologic Malignancies Using Gene Editing

Clinical Trials for Hematologic Malignancies Using Gene Editing
Clinical Trials for Hematologic Malignancies Using Gene Editing

In addition to the transformative impact that the CRISPR-Cas system is making in the treatment of genetic disorders and cancer, it also stands to reshape the diagnosis and management of a number of infectious diseases. To date, there have been 2 broad applications of CRISPR in the infectious disease space: (1) point-of-care diagnostics and (2) more effective antiviral, antibacterial, and antiparasitic treatments.

Diagnostics

The gold standard method for sensitive and specific nucleic acid–based (DNA and RNA) diagnostics in microbiology is PCR. However, as recent experiences with the SARS-CoV-2 epidemic have demonstrated, the challenges associated with developing and running PCR-based assays for emerging and re-emerging pathogens, namely the high cost, complexity, and turnaround time, have motivated the development of rapid point-of-care diagnostics. In the past few years, the CRISPR-Cas system has been engineered to enable sensitive and rapid point-of-care diagnostics for human pathogens.

As described above, the CRISPR-Cas system consists of 2 parts: a variable sgRNA that localizes the RNP complex to a specific nucleic acid sequence—dsDNA, single-stranded DNA (ssDNA), or ssRNA—and a Cas nuclease that is activated upon sgRNA binding, cleaving dsDNA either in cis or in trans. The general strategy for harnessing the power of this 2-part system for pathogen purposes is designing an sgRNA that is specific for a virus or bacterial species, and converting the Cas protein into a “reporter,” whereby binding to the pathogen-specific nucleic acid sequence triggers the activation of a detectable signal reported as fluorescence, biotin-based lateral flow readouts, or electrochemical signal.

The most broadly studied and widely deployed Cas nuclease is Cas9, which, when bound to a dsDNA fragment targeted by its complexed sgRNA, cleaves the dsDNA near the sgRNA-specific PAM. However, it was the subsequent discovery of other classes of Cas nucleases, namely Cas12 and Cas13, that greatly accelerated the development of Cas-based infectious disease assays. Unlike Cas9, which only targets dsDNA, Cas12 recognizes both dsDNA and ssDNA, and Cas13 recognizes ssRNA. Furthermore, Cas12 and Cas13 share the ability to, when bound, cleave nontarget nucleic acids in trans,48  enabling the construction of nucleic acid–cleavage reporting methods. In short, Cas12 and Cas13 exhibit a built-in nuclease reporter, a powerful feature that is the basis of a growing number of infectious disease assays.49 

The list of diagnostic platforms that leverage the CRISPR-Cas system to detect pathogens is expanding daily, and the detailed strategies of these new methods have been reviewed comprehensively elsewhere.5053  Generally speaking, these platforms can be organized according to (1) the type of Cas effector protein (eg, Cas12 or Cas13), which determines what type of nucleic acid target can be detected; (2) the type of reporting or readout mechanism (eg, fluorescence, lateral flow); (3) the mechanism by which the target nucleic acid is amplified before detection, if at all; and (4) the clinical application (ie, analyte). Platforms using Cas12 include DETECTR, which detects human papillomavirus type 16 (HPV16) and 18 (HPV18), and more recently, SARS-CoV-2; HOLMES, which detects pseudorabies virus and Japanese encephalitis virus; E-CRISPR, which targets HPV16 and parvovirus B19; and a number of others.5357  Cas13 has been the biochemical basis of a number of recent innovative methods, the most prominent of which is specific high-sensitivity enzymatic reporter unLOCKing or SHERLOCK, which has been shown to successfully detect several bacteria and viruses, including SARS-CoV-2, for which it received emergency use authorization in 2020.5860 

Treatments

The natural function of the CRISPR-Cas system is to disrupt viral genes in bacteria. As has been highlighted above, this feature can be harnessed to manipulate the genetic activity with tremendous precision in mammalian cells, but much like ZFNs and TALENs, it can also be exploited to disrupt critical genes in pathogenic viruses, bacteria, and parasites.61,62 

One of the first applications of CRISPR to prevent or treat an infectious disease was the prevention of HIV/AIDS. One of the earliest attempts to target the virus using a CRISPR-Cas system consisted of cleaving latent HIV provirus in T cells; however, in subsequent years, 1 of the most promising strategies, now approaching the clinic, is the targeting of chemokine receptors (eg, CCR5) that are necessary for HIV entry into T cells.6368  In addition to HIV, the CRISPR-Cas system is being explored as an experimental therapy in preclinical models for other pathogenic viruses, including hepatitis B, hepatitis C, and most recently, SARS-CoV-2.6972 

The CRISPR system is also being deployed to address pathogenic bacteria, specifically those that have acquired resistance to antibiotics. Bacteria typically acquire resistance to antibacterial drugs via the transfer and production of numerous copies of plasmids that encode resistance genes, and investigators are adapting the CRISPR-Cas system to disrupt these clinically problematic genes.7376  For example, in a recent proof-of-concept study, investigators demonstrated the feasibility of restoring sensitivity to β-lactams in clinically isolated, multidrug-resistant strains of Escherichia coli, Enterobacter hormaechei, and Klebsiella variicola.76 

Finally, much like it is revolutionizing the treatment of viral and bacterial diseases, CRISPR is beginning to make a transformative impact in parasitology, both with respect to advancing our understanding of the basic biology of specific organisms (including their interactions with insect vectors and hosts) and the development of new treatments for diseases such as malaria, leishmaniasis, trypanosomiasis, and many others. For malaria, a promising strategy and major focus of research recently has been deploying CRISPR in Anopheles mosquitos to block Plasmodium transmission and suppress the mosquito population.7780  However, perturbing the genetics of mosquito populations raises a number of ethical and ecological concerns that will require careful consideration.81 

Gene editing technologies hold great promise to treat and even cure disease in novel ways. At the same time, they carry the potential for formidable adverse effects, which represent a barrier to clinical use.29,82  Toxicities that will be discussed here include genotoxicity and immune effects.

Genotoxicity refers to effects that result from unintended modification of the genome, which can occur both at and away from the site targeted for gene editing. Off-target activity is generally due to tolerance of the target-recognizing domain of the gene editing apparatus to mismatches with host DNA. The resultant nonspecific binding can lead to cleavage at sites other than the intended site. When host cell repair mechanisms attempt to fix the break, mutations (typically point mutations and small indels) may occur.83  Both on- and off-target activity may lead to the generation of multiple DSDBs in the same cell. In this scenario, large chromosomal alterations, such as deletions, inversions, and translocations, can result, potentially leading to the loss of significant amounts of genetic material and to genomic instability.83,84  Some references investigating potential genotoxic events caused by CRISPR technology in the research setting are outlined in Table 3.

Table 3

Selected Reports of Potentially Adverse or Genotoxic Events With CRISPR in Preclinical Research Studies

Selected Reports of Potentially Adverse or Genotoxic Events With CRISPR in Preclinical Research Studies
Selected Reports of Potentially Adverse or Genotoxic Events With CRISPR in Preclinical Research Studies

If unintended DNA alterations impact a site critical to cell or organism function or survival, biologic harm can result. Of particular concern is the potential for gene editing systems to alter tumor suppressor genes or proto-oncogenes, thereby promoting oncogenesis. A related phenomenon was reported with earlier iterations of gene therapy, when retroviral vectors—used to transduce cells with genetic material intended to replace dysfunctional genes—integrated in or near genes involved in key cellular processes, leading to the development of hematopoietic neoplasms in several patients.85,86  Notably, more recent reports of myeloid neoplasms in 2 patients who underwent lentiviral-based gene therapy for SCD have determined that, for these 2 patients, leukemogenesis was not directly related to insertional oncogenesis.87,88  In theory and in practice, the specificity of gene editing systems appears to exceed that of earlier forms of gene therapy, reducing this risk. However, genotoxicity remains a significant concern.

On a more generic level, the generation of DSDBs by CRISPR-Cas9 has been reported to induce a DNA damage response mediated by p53, leading to cytotoxicity. This observation has generated concern that gene editing may select for cells with deficient DNA repair pathways, which in turn may be more vulnerable to oncogenic events.89,90 

Although all gene editing systems are capable of potentially genotoxic events, several parameters impact the rate of off-target activity. In general, TALENs appear to be more specific than ZFNs, and ZFNs in turn are more specific than CRISPR systems.91,92  With regard to CRISPR-Cas9 specifically, in vitro and cellular assessments have demonstrated that off-target mutations occur with significant frequency that, at least in 1 human-cell-based study, rivaled on-target activity.93  Notably, with current gene editing systems, genotoxicity is related to the generation of DSDBs. However, a CRISPR-Cas system known as CRISPR Cas12a has been shown to cleave ssDNA in a nonspecific manner.83 

The site of cell manipulation—in vivo or ex vivo—influences the risk of adverse effects. Generally, ex vivo engineering permits a degree of quality control over the editing process, because the altered cells can be assessed for flaws before delivery to the patient. Currently, many gene editing trials in humans target blood diseases, in part because hematopoietic stem cells can be harvested, manipulated in the laboratory, and then returned to the patient. In contrast, with in vivo editing, accurate and efficient delivery to target tissues and cells becomes a significant consideration. Consequently, some current trials aim to treat conditions that manifest in a single accessible organ, such as the eye, thereby improving the chance that gene editing machinery is taken up by targeted tissues, and potentially limiting the anatomic extent of possible side effects.

Both in vivo and ex vivo methods expose the host to novel materials, and hence carry the potential to engage the host immune system. Gene editing technologies rely on components derived from microbes, which can trigger both the innate and adaptive immune systems. Viral vectors, for example, represent a key tool used to deliver gene editing nucleases into host cells. AAV has proven to be less immunogenic than other viral vectors, leading to its broad adoption as a delivery vector.94  Electroporation is a nonviral delivery method that avoids the immunogenicity of viruses but—as noted earlier—is primarily useful in ex vivo gene editing techniques.84,94  Gene editing nucleases themselves—specifically, TALEN and CRISPR—originate from bacteria, and consequently pose an immune risk. Studies of human populations have identified varying levels of antibodies against Cas9, and in mice, Cas9 proteins have been shown to evoke a humoral response.9496 

By enabling patients to produce proteins that they are congenitally unable to create, gene therapy may cure certain genetic diseases. However, the host immune system may perceive the protein products of corrected genes as foreign, leading to an immune response against the protein itself or against edited cells. Such a reaction would limit therapeutic efficacy and could lead to inflammatory consequences for the patient.94  Fortunately, therapies designed to eliminate or reduce expression of a defective gene product or to enhance expression of a desired gene product would not result in the production of immunologically foreign antigens and thus would be unlikely to prompt an immune response.94  For therapies that yield a novel protein or antigen, approaches to monitor or modify the host immune response could be required.97  Tremendous efforts are underway to improve the safety and efficacy of gene editing. Strategies to enhance the fidelity of programmable nucleases, predict off-target effects, and minimize the immune consequences of gene editing represent important areas of research.94  Alternative methods that do not rely on DNA cleavage, such as base editing and prime editing, are under exploration.98  These efforts will undoubtedly advance the promise of gene editing therapies.

As clinical applications of gene editing expand, patients will need to be monitored for adverse effects. Although specialized monitoring will likely remain the responsibility of the manufacturers and vendors of these therapies, a myriad of tasks may eventually fall within the purview of pathologists. For example, hematopathologists may help monitor bone marrow function, survey patients for hematologic malignancies, and provide follow-up for patients with hematolymphoid neoplasms treated with gene editing modalities. They may even be asked to discern the likelihood of relatedness of a neoplasm to receipt of a gene-altering therapy, depending on the situation. For instance, there have been 3 reported cases of myelodysplastic syndrome in patients receiving otherwise efficacious and FDA-approved elivaldogene autotemcel (eli-cel) gene therapy (autologous CD34+ cells transduced with a Lenti-D lentiviral vector carrying ABCD1 cDNA) for cerebral adrenoleukodystrophy.99  It is possible hematopathologists could be asked their opinion as to etiology in similar situations. Molecular pathologists, surgical pathologists, cytogeneticists, and flow cytometrists could conceivably evaluate patients for development of clonal and neoplastic cell populations and similarly be asked of relatedness to such therapies. Clinical chemists, microbiologists, and pathologists who serve as laboratory medical directors may supervise measurement of inflammatory markers, monitor the levels of immunosuppressant drugs, and test for posttherapy infections. While the precise responsibilities of pathologists are yet to be determined, it is likely that the practice of pathology will soon require familiarity with the intent and effects of gene editing therapies.

The regulatory aspects of gene editing/drug product manufacturing, including those needed for an investigational new drug application as part of a clinical trial, are expected to be performed by a third-party facility separate from the hospital and are outside the scope of this document.30  This section focuses solely on potential regulatory and practical issues that would be encountered in a hospital-based laboratory service.

Once FDA-approved and no longer under clinical trials, laboratory testing supporting gene editing–associated therapies has the potential to be subject to accreditation (Association for the Advancement of Blood and Biotherapies, College of American Pathologists [CAP], Foundation for the Accreditation of Cellular Therapy) and regulatory oversight (Centers for Medicare and Medicaid Services and the FDA). As such, there is anticipated involvement of pathologists or clinical laboratory staff in the hospital or clinic setting. Even with FDA approval, many hospitals will be responsible for the procurement of autologous donor cells to be shipped to a manufacturing facility and receipt of the final product for use. This would be analogous to novel cellular therapies such as for CAR (chimeric antigen receptor)–T cells. The following areas that laboratories should consider from a regulatory and practical perspective, including having quality reviews and documentation of proper performance at each of these steps, include (1) precollection assessment of the autologous donor for autologously derived products including infectious disease testing and precollection treatment of the donor where appropriate; (2) pre–gene editing processing, storage, and shipping of cells to the manufacturing facility; (3) gene editing process control (dependent on the platform and whether on-site or off-site); (4) receipt, inspection, and storage of gene-edited cells returned from the manufacturing facility; (5) confirming the minimum percentage of cells with the change; (6) confirming that there are no off-target changes (may be supplied by manufacturer); (7) mechanism to confirm no intact viral vectors (may be supplied by manufacturer); (8) poststorage processing (thawing or washing or dilution if needed); (9) issuing of the therapeutic from the laboratory area; (10) infusion of the product; (11) postinfusion monitoring—laboratory testing or adverse event reporting; and (12) longitudinal and aggregate review of the safety of the product and mechanism development to capture any/all unintended events.

Unknown at this point is whether these changes to accommodate this new form of therapy will be incorporated into expanded sections of existing standards and checklists for cellular therapy and laboratory testing, or if they would instead be separate stand-alone documents. It would be anticipated, at minimum, that they would initially be worded to match existing applicable standards and checklist items. Similarly, inclusive or exclusive changes to existing laboratory quality plans would need to be made to address the presence of precursor materials or gene-edited cells. This would include the use of proficiency testing, quality control review, audits, and competency assessments. Depending on the assays used to monitor the impact of gene editing, they may require a combination of existing chemical, enzymatic, molecular, or immunophenotypic assays or the use of proprietary genetic sequences to monitor for viral vectors or percentage of cells with the change over time. Similar to existing cellular therapy products in clinical use, different degrees of rigor and regulations may apply for the testing or handling of the gene-edited products versus testing or handling of patient specimens.

Given the enormous amount of attention afforded to the CRISPR technique in the lay press, it is no surprise that concerns about potential ethical abuses have risen to the top of the international conversation. Indeed, there are significant ethical questions to confront, but the first thing to recognize is that these issues are not new and did not suddenly appear with the advent of CRISPR or its recognition by the Nobel Prize in Chemistry in 2020. Gene editing by CRISPR-Cas was preceded by many other gene editing techniques, going back several decades—and by gene replacement strategies even before that. The fact that CRISPR has proven to be more precise, more efficient, and less expensive than those earlier techniques does not fundamentally change the nature of the ethical concerns. Besides the usual ethical concerns that surround any investigational therapies—with all the attendant requirements for institutional review board approval, appropriate informed consent, benefit versus risk calculations, equitable access, among others—both gene therapy and gene editing raise the specter of eugenics: the deliberate alteration of the human gene pool for either aspirational or nefarious purposes. The latter have a long history, albeit in far cruder form, of discrimination, racial superiority theory, forced sterilizations, and genocide.100  The former, while ostensibly benign, also raise concerns about potential harms due to off-target or otherwise unpredictable effects.101  This, too, is nothing new, as we only have to acknowledge the small but tragic series of patient deaths in more traditional gene replacement trials.32 

Most of the ethical objections at present center around germline, as opposed to somatic, gene editing.102  This is because eugenic effects, by definition, can only occur by influencing the genetic traits of offspring, and if done by genetic manipulation, that can only occur if the germ cells (sperm or oocyte) or the early embryo are the targets of gene editing. For that reason, most of the ethical debates are focused on gene editing as treatment for hereditary diseases, where (except in rare cases of mosaicism) every nucleated cell in the body contains the disease-causing mutation(s). As such, the treatment might be directed at embryonic stem cells or germ plasm (as opposed to targeting just the affected tissue, such as liver, eye, or hematopoietic stem cells), opening up the possibility for the reproductive organs and gametes to be so altered, and thus passed on to successive generations. In contrast, most gene editing approaches for cancer treatment (somatic) would be expected to target the neoplastic tissue only.

In fact, the only known instance of such an approach, which led to international outrage and arrest of the investigator, involved the editing of early twin embryos by He Jiankui in China in 2018.102  The optics of this incident were made worse by the fact that these twins did not have, nor were they at any risk for, a genetic disorder. Rather, Dr Jiankui performed an ostensibly innocuous “proof-of-principle” experiment in which he introduced the common 32-bp deletion in the CCR5 gene, which is known to provide relative resistance to HIV infection (not that the babies were at any real risk of contracting AIDS). Thus, the gene edit would not provide any particular benefit, and at this early stage we cannot assume it is entirely benign either, since there could be unanticipated off-target effects that might cause serious disease later in life. As a result of this incident, many governments and professional medical and scientific organizations issued statements to the effect that there should be an immediate moratorium on germline gene editing that leads to a human pregnancy.103,104  However, most have also stated that editing studies of gametes and early embryos can still be permitted (with appropriate approval and oversight), as long as those cells are not implanted into a woman's uterus.104 

While most observers agree with these concerns, some arguments could, in fact, be made in favor of germline gene editing, as a means to potentially wipe out devastating genetic diseases, to lower the number of abortions of affected fetuses, and to keep the field moving forward in an open way, in view of the very low incidence of, but still concerning, observed harmful off-target effects in vitro and in animal models.105109 

In some ways, the current debates over applications of gene editing are not so different from those that occurred in the mid-1970s over the advent of recombinant DNA technology and gene cloning. There were ethicists at that time who demanded that all research using such techniques be shut down completely.110  Fortunately, more pragmatic views prevailed, and the work was allowed to continue with appropriate containment and ethical precautions.111,112  We should expect much the same for human gene editing, and it is crucial that pathologists, who will be essential players in any patient therapies as discussed throughout this document, have a seat at the table.

In conclusion, the development of gene editing capabilities is an example of how small steps in basic science research can lead to potentially major shifts in the practice of clinical medicine. These technologies hold promise in significantly altering the course of, or potentially curing, life-threatening diseases in the cancer, infectious disease, and inherited disease settings. However, with such monumental prospects comes also the need for caution and thoughtful, responsible assessment. Long-term adverse effects and costs are just 2 of many considerations in the area of somatic gene editing. Germline gene editing has presented an even greater set of ethical questions and concerns that will need to be addressed in both the research and clinical setting. How such therapies may affect laboratory testing in molecular diagnostics, flow cytometry, hematology, chemistry, transfusion medicine, and anatomic pathology is not, of course, fully known and requires an educated field of pathologists and laboratorians to identify potential issues. Even more, pathologists need to be at the table with industry, regulators, and professional medical societies in order to impart their knowledge of clinical regulation of laboratory testing and how these therapies could affect hospital-based protocols and procedures and laboratory reimbursement. The CAP's recent addition to the National Institute of Standards and Technology (NIST) Genome Editing Consortium will hopefully provide some pathology insight on the existence of such considerations. It is imperative that pathologists' perspectives are heard as gene editing–based therapies are adopted, and the development of these perspectives require education and a knowledge base in the area of gene editing.

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Author notes

Hiemenz is an employee of Foundation Medicine, a wholly owned subsidiary of Roche, and has an equity interest in Roche. The other authors have no relevant financial interest in the products or companies described in this article.

This paper was presented at the College of American Pathologists 2021 and 2022 annual meetings; September 26, 2021; Chicago, Illinois; and October 9, 2022; New Orleans, Louisiana.