CRISPR-Directed Gene Editing as a Method to Reduce Chemoresistance in Lung Cancer Cells.

We are advancing a novel strategy for the treatment of solid tumors by employing CRISPR-directed gene editing to reduce levels of standard of care required to halt or reverse the progression of tumor growth. We intend to do this by utilizing a combinatorial approach in which CRISPR-directed gene editing is used to eliminate or significantly reduce the acquired resistance emerging from chemotherapy, radiation therapy, or immunotherapy. We will utilize CRISPR/Cas as a biomolecular tool to disable specific genes involved in the sustainability of resistance to cancer therapy. We have also developed a CRISPR/Cas molecule that can distinguish between the genome of a tumor cell in the genome of a normal cell, thereby conferring target selectivity onto this therapeutic approach. We envision delivering these molecules by direct injection into solid tumors for the treatment of squamous cell carcinomas of the lung, esophageal cancer, and head and neck cancer. We provide experimental details and methodology for utilizing CRISPR/Cas as a supplement to chemotherapy to destroy lung cancer cells.

Homology directed correction, a new pathway model for point mutation repair catalyzed by CRISPR-Cas.

Gene correction is often referred to as the gold standard for precise gene editing and while CRISPR-Cas systems continue to expand the toolbox for clinically relevant genetic repair, mechanistic hurdles still hinder widespread implementation. One of the most prominent challenges to precise CRISPR-directed point mutation repair centers on the prevalence of on-site mutagenesis, wherein insertions and deletions appear at the targeted site following correction. Here, we introduce a pathway model for Homology Directed Correction, specifically point mutation repair, which enables a foundational analysis of genetic tools and factors influencing precise gene editing. To do this, we modified an in vitro gene editing system which utilizes a cell-free extract, CRISPR-Cas RNP and donor DNA template to catalyze point mutation repair. We successfully direct correction of four unique point mutations which include two unique nucleotide mutations at two separate targeted sites and visualize the repair profiles resulting from these reactions. This extension of the cell-free gene editing system to model point mutation repair may provide insight for understanding the factors influencing precise point mutation correction.

Exon skipping induced by CRISPR-directed gene editing regulates the response to chemotherapy in non-small cell lung carcinoma cells.

We have been developing CRISPR-directed gene editing as an augmentative therapy for the treatment of non-small cell lung carcinoma (NSCLC) by genetic disruption of Nuclear Factor Erythroid 2-Related Factor 2 (NRF2). NRF2 promotes tumor cell survival in response to therapeutic intervention and thus its disablement should restore or enhance effective drug action. Here, we report how NRF2 disruption leads to collateral damage in the form of CRISPR-mediated exon skipping. Heterogeneous populations of transcripts and truncated proteins produce a variable response to chemotherapy, dependent on which functional domain is missing. We identify and characterize predicted and unpredicted transcript populations and discover that several types of transcripts arise through exon skipping; wherein one or two NRF2 exons are missing. In one specific case, the presence or absence of a single nucleotide determines whether an exon is skipped or not by reorganizing Exonic Splicing Enhancers (ESEs). We isolate and characterize the diversity of clones induced by CRISPR activity in a NSCLC tumor cell population, a critical and often overlooked genetic byproduct of this exciting technology. Finally, gRNAs must be designed with care to avoid altering gene expression patterns that can account for variable responses to solid tumor therapy.

On the Origins of Homology Directed Repair in Mammalian Cells.

Over the course of the last five years, expectations surrounding our capacity to selectively modify the human genome have never been higher. The reduction to practice site-specific nucleases designed to cleave at a unique site within the DNA is now centerstage in the development of effective molecular therapies. Once viewed as being impossible, this technology now has great potential and, while cellular and molecular barriers persist to clinical implementations, there is little doubt that these barriers will be crossed, and human beings will soon be treated with gene editing tools. The most ambitious of these desires is the correction of genetic mutations resident within the human genome that are responsible for oncogenesis and a wide range of inherited diseases. The process by which gene editing activity could act to reverse these mutations to wild-type and restore normal protein function has been generally categorized as homology directed repair. This is a catch-all basket term that includes the insertion of short fragments of DNA, the replacement of long fragments of DNA, and the surgical exchange of single bases in the correction of point mutations. The foundation of homology directed repair lies in pioneering work that unravel the mystery surrounding genetic exchange using single-stranded DNA oligonucleotides as the sole gene editing agent. Single agent gene editing has provided guidance on how to build combinatorial approaches to human gene editing using the remarkable programmable nuclease complexes known as Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and their closely associated (Cas) nucleases. In this manuscript, we outline the historical pathway that has helped evolve the current molecular toolbox being utilized for the genetic re-engineering of the human genome.

Deconvolution of Complex DNA Repair (DECODR): Establishing a Novel Deconvolution Algorithm for Comprehensive Analysis of CRISPR-Edited Sanger Sequencing Data.

During CRISPR-directed gene editing, multiple gene repair mechanisms interact to produce a wide and largely unpredictable variety of sequence changes across an edited population of cells. Shortcomings inherent to previously available proposal-based insertion and deletion (indel) analysis software necessitated the development of a more comprehensive tool that could detect a larger range and variety of indels while maintaining the ease of use of tools currently available. To that end, we developed Deconvolution of Complex DNA Repair (DECODR). DECODR can detect indels formed from single or multi-guide CRISPR experiments without a limit on indel size. The software is accurate in determining the identities and positions of inserted and deleted bases in DNA extracts from both clonally expanded and bulk cell populations. The accurate identification and output of any potential indel allows for DECODR analysis to be executed in experiments utilizing potentially any configuration of donor DNA sequences, CRISPR-Cas, and endogenous DNA repair pathways.

Precise and error-prone CRISPR-directed gene editing activity in human CD34+ cells varies widely among patient samples.

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and their associated CRISPR-associated nucleases (Cas) are among the most promising technologies for the treatment of hemoglobinopathies including Sickle Cell Disease (SCD). We are only beginning to identify the molecular variables that influence the specificity and the efficiency of CRISPR- directed gene editing, including the position of the cleavage site and the inherent variability among patient samples selected for CRISPR-directed gene editing. Here, we target the beta globin gene in human CD34+ cells to assess the impact of these two variables and find that both contribute to the global diversity of genetic outcomes. Our study demonstrates a unique genetic profile of indels that is generated based on where along the beta globin gene attempts are made to correct the SCD single base mutation. Interestingly, even within the same patient sample, the location of where along the beta globin gene the DNA is cut, HDR activity varies widely. Our data establish a framework upon which realistic protocols inform strategies for gene editing for SCD overcoming the practical hurdles that often impede clinical success.

The Diversity of Genetic Outcomes from CRISPR/Cas Gene Editing is Regulated by the Length of the Symmetrical Donor DNA Template.

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas gene editing systems have enabled molecular geneticists to manipulate prokaryotic and eukaryotic genomes with greater efficiency and precision. CRISPR/Cas provides adaptive immunity in bacterial cells by degrading invading viral genomes. By democratizing this activity into human cells, it is possible to knock out specific genes to disable their function and repair errors. The latter of these activities requires the participation of a single-stranded donor DNA template that provides the genetic information to execute correction in a process referred to as homology directed repair (HDR). Here, we utilized an established cell-free extract system to determine the influence that the donor DNA template length has on the diversity of products from CRISPR-directed gene editing. This model system enables us to view all outcomes of this reaction and reveals that donor template length can influence the efficiency of the reaction and the categories of error-prone products that accompany it. A careful measurement of the products revealed a category of error-prone events that contained the corrected template along with insertions and deletions (indels). Our data provides foundational information for those whose aim is to translate CRISPR/Cas from bench to bedside.

Modeling pediatric AML FLT3 mutations using CRISPR/Cas12a- mediated gene editing.

Clustered regularly interspaced palindromic repeats (CRISPR) with the associated (Cas) nuclease complexes have democratized genetic engineering through their precision and ease-of-use. We have applied a variation of this technology, known as CRISPR-directed mutagenesis (CDM), to reconstruct genetic profiles within the FLT3 gene of AML patients. We took advantage of the versatility of CDM and built expression vectors that, in combination with a specifically designed donor DNA fragment, recapitulate simple and complex mutations within the FLT3 gene. We generate insertions and point mutations including combinations of these mutations originating from individual patient samples. We then analyze how these complex genetic profiles modulate transformation of Ba/F3 cells. Our results show that FLT3 expression plasmids bearing patient-specific single or multiple mutations recapitulate cellular transformation properties induced by FLT3 ITDs and modify their sensitivity or resistance in response to established AML drugs as a function of these complex mutations.

gRNA Sequence Heterology Tolerance Catalyzed by CRISPR/Cas in an In Vitro Homology-Directed Repair Reaction.

CRISPR and associated Cas nucleases are genetic engineering tools revolutionizing innovative approaches to cancer and inherited diseases. CRISPR-directed gene editing relies heavily on proper DNA sequence alignment between the guide RNA (gRNA)/CRISPR complex and its genomic target. Accurate hybridization of complementary DNA initiates gene editing in human cells, but inherent gRNA sequence variation that could influence the gene editing reaction has been clearly established among diverse genetic populations. As this technology advances toward clinical implementation, it will be essential to assess what degree of gRNA variation generates unwanted and erroneous CRISPR activity. With the use of a system in which a cell-free extract catalyzes nonhomologous end joining (NHEJ) and homology-directed repair (HDR), it is possible to observe a more representative population of all forms of gene editing outcomes. In this manuscript, we demonstrate CRISPR/Cas complexation at heterologous binding sites that facilitate precise and error-prone HDR. The tolerance of mispairing between the gRNA and target site of the DNA to enable HDR is surprisingly high and greatly influenced by polarity of the donor DNA strand in the reaction. These results suggest that some collateral genomic activity could occur at unintended sites in CRISPR-directed gene editing in human cells.

Kinetics of Nuclear Uptake and Site-Specific DNA Cleavage during CRISPR-Directed Gene Editing in Solid Tumor Cells.

Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-directed gene editing is approaching clinical implementation in cancer. Thus, it is imperative to define the molecular framework upon which safe and efficacious therapeutic strategies can be built. Two important reaction parameters include the biological time frame within which the CRISPR/Cas complex enters the nucleus and executes gene editing, and the method of discrimination that the CRISPR/Cas complex utilizes to target tumor cell, but not normal cell, genomes. We are developing CRISPR-directed gene editing for the treatment of non-small cell lung carcinoma focusing on disabling Nuclear Factor Erythroid 2-Related Factor-Like (NRF2), a transcription factor that regulates chemoresistance and whose genetic disruption would enhance chemosensitivity. In this report, we define the time frame of cellular events that surround the initialization of CRISPR-directed gene editing as a function of the nuclear penetration and the execution of NRF2 gene disruption. We also identify a unique protospacer adjacent motif that facilitates site-specific cleavage of the NRF2 gene present only in tumor genomes. IMPLICATIONS: Our results begin to set a scientifically meritorious foundation for the exploitation of CRISPR-directed gene editing as an augmentative therapy for lung cancer and other solid tumors. VISUAL OVERVIEW: http://mcr.aacrjournals.org/content/molcanres/18/6/891/F1.large.jpg.

Perspectives on Molecular Diagnostic Testing for the COVID-19 Pandemic in Delaware.

The United States has quickly transitioned into one of the epicenters for the coronavirus pandemic. Limitations for rapid testing for the virus responsible for the pandemic, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), is the single most important barrier for early detection and prevention of future outbreaks. Combining innovative molecular biology techniques, such as clustered regularly interspaced short palindromic repeats (CRISPR)/Cas nuclease systems and next generation sequencing (NGS) may prove to be an effective solution to establish a high-throughput diagnostic and genomic surveillance workflow for COVID-19 in the State of Delaware. Integrating key expertise across the medical institutions in Delaware, including ChristianaCare and Nemours/Alfred I. duPont Hospital for Children, is one potential solution for overcoming current barriers and driving a successful implementation of these techniques.

Understanding the diversity of genetic outcomes from CRISPR-Cas generated homology-directed repair.

As CRISPR-Cas systems advance toward clinical application, it is essential to identify all the outcomes of gene-editing activity in human cells. Reports highlighting the remarkable success of homology-directed repair (HDR) in the treatment of inherited diseases may inadvertently underreport the collateral activity of this remarkable technology. We are utilizing an in vitro gene-editing system in which a CRISPR-Cas complex provides the double-stranded cleavage and a mammalian cell-free extract provides the enzymatic activity to promote non-homologous end joining, micro-homology mediated end joining, and homology-directed repair. Here, we detail the broad spectrum of gene-editing reaction outcomes utilizing Cas9 and Cas12a in combination with single-stranded donor templates of the sense and nonsense polarity. This system offers the opportunity to see the range of outcomes of gene-editing reactions in an unbiased fashion, detailing the distribution of DNA repair outcomes as a function of a set of genetic tools.

CRISPR-Directed Gene Editing Catalyzes Precise Gene Segment Replacement In Vitro Enabling a Novel Method for Multiplex Site-Directed Mutagenesis.

Much of our understanding of eukaryotic genes function comes from studies of the activity of their mutated forms or allelic variability. Mutations have helped elucidate how members of an intricate pathway function in relation to each other and how they operate in the context of the regulatory circuitry that surrounds them. A PCR-based site-directed mutagenesis technique is often used to engineer these variants. While these tools are efficient, they are not without significant limitations, most notably off-site mutagenesis, limited scalability, and lack of multiplexing capabilities. To overcome many of these limitations, we now describe a novel method for the introduction of both simple and complex gene mutations in plasmid DNA by using in vitro DNA editing. A specifically designed pair of CRISPR-Cas12a ribonucleoprotein complexes are used to execute site-specific double-strand breaks on plasmid DNA, enabling the excision of a defined DNA fragment. Donor DNA replacement is catalyzed by a mammalian cell-free extract through microhomology annealing of short regions of single-stranded DNA complementarity; we term this method CRISPR-directed DNA mutagenesis (CDM). The products of CDM are plasmids bearing precise donor fragments with specific modifications and CDM could be used for mutagenesis in larger constructs such as Bacterial Artificial Chromosome (BACs) or Yeast Artificial Chromosome (YACs). We further show that this reaction can be multiplexed so that product molecules with multiple site-specific mutations and site-specific deletions can be generated in the same in vitro reaction mixture. Importantly, the CDM method produces fewer unintended mutations in the target gene as compared to the standard site-directed mutagenesis assay; CDM produces no unintended mutations throughout the plasmid backbone. Lastly, this system recapitulates the multitude of reactions that take place during CRISPR-directed gene editing in mammalian cells and affords the opportunity to study the mechanism of action of CRISPR-directed gene editing in mammalian cells by visualizing a multitude of genetic products.

Functional Gene Knockout of NRF2 Increases Chemosensitivity of Human Lung Cancer A549 Cells In Vitro and in a Xenograft Mouse Model.

Recent studies point to the evolution of drug resistance in lung cancer as being centered, at least in part, on the upregulation of various genes involved in controlling efflux or drug inactivation. Among the most important of these genes is Nuclear Factor Erythroid 2-Related Factor (NRF2), considered the master regulator of 100-200 target genes involved in cellular responses to oxidative and/or electrophilic stress. With increased focus on the development of combinatorial approaches for cancer treatment, we utilized CRISPR/Cas9 to disable the NRF2 gene in lung cancer cells by disrupting the NRF2 nuclear export signal (NES) domain; phenotypically, the protein is largely blocked from transiting into the nucleus after translation. In tissue culture, cells with this gene knockout were found to have a reduced proliferation phenotype and are more sensitive to chemotherapeutic agents, such as cisplatin and carboplatin. These observations were confirmed in xenograft mouse models wherein the homozygous knockout cells proliferate at a slower rate than the wild-type cells, even in the absence of drug treatment. Tumor growth was arrested for a period of 16 days, with a dramatic decrease in tumor volume being observed in samples receiving the combined action of CRISPR-directed gene editing and chemotherapy.

Efficient Delivery and Nuclear Uptake Is Not Sufficient to Detect Gene Editing in CD34+ Cells Directed by a Ribonucleoprotein Complex.

CD34+ cells are prime targets for therapeutic strategies for gene editing, because modified progenitor cells have the capacity to differentiate through an erythropoietic lineage. Although experimental advances have been reported, the associated experimental protocols have largely been less than clear or robust. As such, we evaluated the relationships among cellular delivery; nuclear uptake, often viewed as the benchmark metric of successful gene editing; and single base repair. We took a combinatorial approach using single-stranded oligonucleotide and a CRISPR/Cas9 ribonucleoprotein to convert wild-type HBB into the sickle cell genotype by evaluating conditions for two common delivery strategies of gene editing tools into CD34+ cells. Confocal microscopy data show that the CRISPR/Cas9 ribonucleoprotein tends to accumulate at the outer membrane of the CD34+ cell nucleus when the Neon Transfection System is employed, while the ribonucleoproteins do pass into the cell nucleus when nucleofection is used. Despite the high efficiency of cellular transformation, and the traditional view of success in efficient nuclear uptake, neither delivery methodology enabled gene editing activity. Our results indicate that more stringent criteria must be established to facilitate the clinical translation and scientific robustness of gene editing for sickle cell disease.

CRISPR-Directed In Vitro Gene Editing of Plasmid DNA Catalyzed by Cpf1 (Cas12a) Nuclease and a Mammalian Cell-Free Extract.

Extraordinary efforts are underway to offer greater versatility and broader applications for CRISPR-directed gene editing. Here, we report the establishment of a system for studying this process in a mammalian cell-free extract prepared from HEK-293 human embryonic kidney cells. A ribonucleoprotein (RNP) particle and a mammalian cell-free extract coupled with a genetic readout are used to generate and identify specific deletions or insertions within a plasmid target. A Cpf1 (Cas12a) RNP induces a double-stranded break, and the cell-free extract provides the appropriate enzymatic activities to direct specific deletion through resection and homology directed repair in the presence of single- and double-stranded donor DNA. This cell-free system establishes a foundation to study the heterogeneous products of gene editing, as well as the relationship between nonhomologous end joining and homology directed repair and related regulatory circuitries simultaneously in a controlled environment.

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.