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.

Small-molecule IKKß activation modulator (IKAM) targets MAP3K1 and inhibits pancreatic tumor growth.

Activation of inhibitor of nuclear factor NF-κB kinase subunit-ß (IKKß), characterized by phosphorylation of activation loop serine residues 177 and 181, has been implicated in the early onset of cancer. On the other hand, tissue-specific IKKß knockout in Kras mutation-driven mouse models stalled the disease in the precancerous stage. In this study, we used cell line models, tumor growth studies, and patient samples to assess the role of IKKß and its activation in cancer. We also conducted a hit-to-lead optimization study that led to the identification of 39-100 as a selective mitogen-activated protein kinase kinase kinase (MAP3K) 1 inhibitor. We show that IKKß is not required for growth of Kras mutant pancreatic cancer (PC) cells but is critical for PC tumor growth in mice. We also observed elevated basal levels of activated IKKß in PC cell lines, PC patient-derived tumors, and liver metastases, implicating it in disease onset and progression. Optimization of an ATP noncompetitive IKKß inhibitor resulted in the identification of 39-100, an orally bioavailable inhibitor with improved potency and pharmacokinetic properties. The compound 39-100 did not inhibit IKKß but inhibited the IKKß kinase MAP3K1 with low-micromolar potency. MAP3K1-mediated IKKß phosphorylation was inhibited by 39-100, thus we termed it IKKß activation modulator (IKAM) 1. In PC models, IKAM-1 reduced activated IKKß levels, inhibited tumor growth, and reduced metastasis. Our findings suggests that MAP3K1-mediated IKKß activation contributes to KRAS mutation-associated PC growth and IKAM-1 is a viable pretherapeutic lead that targets this pathway.

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.

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.

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.

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/Cas9-Directed Reassignment of the GATA1 Initiation Codon in K562 Cells to Recapitulate AML in Down Syndrome.

Using a CRISPR/Cas9 system, we have reengineered a translational start site in the GATA1 gene in K562 cells. This mutation accounts largely for the onset of myeloid leukemia in Down syndrome (ML-DS). For this reengineering, we utilized CRISPR/Cas9 to generate mammalian cell lines that express truncated versions of the Gata1s protein similar to that seen in ML-DS, as determined by analyzing specific genetic alterations resulting from CRISPR/Cas9 cleavage. During this work, 73 cell lines were clonally expanded, with allelic variance analyzed. Using Tracking of Indels by DEcomposition (TIDE) and Sanger sequencing, we defined the DNA sequence and variations within each allele. We found significant heterogeneity between alleles in the same clonally expanded cell, as well as among alleles from other clonal expansions. Our data demonstrate and highlight the importance of the randomness of resection promoted by non-homologous end joining after CRISPR/Cas9 cleavage in cells undergoing genetic reengineering. Such heterogeneity must be fully characterized to predict altered functionality inside target tissues and to accurately interpret the associated phenotype. Our data suggest that in cases where the objective is to rearrange specific nucleotides to redirect gene expression in human cells, it is imperative to analyze genetic composition at the individual allelic level.

Insertional Mutagenesis by CRISPR/Cas9 Ribonucleoprotein Gene Editing in Cells Targeted for Point Mutation Repair Directed by Short Single-Stranded DNA Oligonucleotides.

CRISPR/Cas9 and single-stranded DNA oligonucleotides (ssODNs) have been used to direct the repair of a single base mutation in human genes. Here, we examine a method designed to increase the precision of RNA guided genome editing in human cells by utilizing a CRISPR/Cas9 ribonucleoprotein (RNP) complex to initiate DNA cleavage. The RNP is assembled in vitro and induces a double stranded break at a specific site surrounding the mutant base designated for correction by the ssODN. We use an integrated mutant eGFP gene, bearing a single base change rendering the expressed protein nonfunctional, as a single copy target in HCT 116 cells. We observe significant gene correction activity of the mutant base, promoted by the RNP and single-stranded DNA oligonucleotide with validation through genotypic and phenotypic readout. We demonstrate that all individual components must be present to obtain successful gene editing. Importantly, we examine the genotype of individually sorted corrected and uncorrected clonally expanded cell populations for the mutagenic footprint left by the action of these gene editing tools. While the DNA sequence of the corrected population is exact with no adjacent sequence modification, the uncorrected population exhibits heterogeneous mutagenicity with a wide variety of deletions and insertions surrounding the target site. We designate this type of DNA aberration as on-site mutagenicity. Analyses of two clonal populations bearing specific DNA insertions surrounding the target site, indicate that point mutation repair has occurred at the level of the gene. The phenotype, however, is not rescued because a section of the single-stranded oligonucleotide has been inserted altering the reading frame and generating truncated proteins. These data illustrate the importance of analysing mutagenicity in uncorrected cells. Our results also form the basis of a simple model for point mutation repair directed by a short single-stranded DNA oligonucleotides and CRISPR/Cas9 ribonucleoprotein complex.

Analyses of point mutation repair and allelic heterogeneity generated by CRISPR/Cas9 and single-stranded DNA oligonucleotides.

The repair of a point mutation can be facilitated by combined activity of a single-stranded oligonucleotide and a CRISPR/Cas9 system. While the mechanism of action of combinatorial gene editing remains to be elucidated, the regulatory circuitry of nucleotide exchange executed by oligonucleotides alone has been largely defined. The presence of the appropriate CRISPR/Cas9 system leads to an enhancement in the frequency of gene editing directed by single-stranded DNA oligonucleotides. While CRISPR/Cas9 executes double-stranded DNA cleavage efficiently, closure of the broken chromosomes is dynamic, as varying degrees of heterogeneity of the cleavage products appear to accompany the emergence of the corrected base pair. We provide a detailed analysis of allelic variance at and surrounding the target site. In one particular case, we report sequence alteration directed by a distinct member of the same gene family. Our data suggests that single-stranded DNA molecules may influence DNA junction heterogeneity created by CRISPR/Cas9.

Regulation of Gene Editing Activity Directed by Single-Stranded Oligonucleotides and CRISPR/Cas9 Systems.

Single-stranded DNA oligonucleotides (ssODNs) can direct the repair of a single base mutation in human genes. While the regulation of this gene editing reaction has been partially elucidated, the low frequency with which repair occurs has hampered development toward clinical application. In this work a CRISPR/Cas9 complex is employed to induce double strand DNA breakage at specific sites surrounding the nucleotide designated for exchange. The result is a significant elevation in ssODN-directed gene repair, validated by a phenotypic readout. By analysing reaction parameters, we have uncovered restrictions on gene editing activity involving CRISPR/Cas9 complexes. First, ssODNs that hybridize to the non-transcribed strand direct a higher level of gene repair than those that hybridize to the transcribed strand. Second, cleavage must be proximal to the targeted mutant base to enable higher levels of gene editing. Third, DNA cleavage enables a higher level of gene editing activity as compared to single-stranded DNA nicks, created by modified Cas9 (Nickases). Fourth, we calculated the hybridization potential and free energy levels of ssODNs that are complementary to the guide RNA sequences of CRISPRs used in this study. We find a correlation between free energy potential and the capacity of single-stranded oligonucleotides to inhibit specific DNA cleavage activity, thereby indirectly reducing gene editing activity. Our data provide novel information that might be taken into consideration in the design and usage of CRISPR/Cas9 systems with ssODNs for gene editing.

The position of DNA cleavage by TALENs and cell synchronization influences the frequency of gene editing directed by single-stranded oligonucleotides.

With recent technological advances that enable DNA cleavage at specific sites in the human genome, it may now be possible to reverse inborn errors, thereby correcting a mutation, at levels that could have an impact in a clinical setting. We have been developing gene editing, using single-stranded DNA oligonucleotides (ssODNs), as a tool to direct site specific single base changes. Successful application of this technique has been demonstrated in many systems ranging from bacteria to human (ES and somatic) cells. While the frequency of gene editing can vary widely, it is often at a level that does not enable clinical application. As such, a number of stimulatory factors such as double-stranded breaks are known to elevate the frequency significantly. The majority of these results have been discovered using a validated HCT116 mammalian cell model system where credible genetic and biochemical readouts are available. Here, we couple TAL-Effector Nucleases (TALENs) that execute specific ds DNA breaks with ssODNs, designed specifically to repair a missense mutation, in an integrated single copy eGFP gene. We find that proximal cleavage, relative to the mutant base, is key for enabling high frequencies of editing. A directionality of correction is also observed with TALEN activity upstream from the target base being more effective in promoting gene editing than activity downstream. We also find that cells progressing through S phase are more amenable to combinatorial gene editing activity. Thus, we identify novel aspects of gene editing that will help in the design of more effective protocols for genome modification and gene therapy in natural genes.

Combinatorial gene editing in mammalian cells using ssODNs and TALENs.

The regulation of gene editing is being elucidated in mammalian cells and its potential as well as its limitations are becoming evident. ssODNs carry out gene editing by annealing to their complimentary sequence at the target site and acting as primers for replication fork extension. To effect a genetic change, a large amount of ssODN molecules must be introduced into cells and as such induce a Reduced Proliferation Phenotype (RPP), a phenomenon in which corrected cells do not proliferate. To overcome this limitation, we have used TAL-Effector Nucleases (TALENs) to increase the frequency, while reducing the amount of ssODN required to direct gene correction. This strategy resolves the problem and averts the serious effects of RPP. The efficiency of gene editing can be increased significantly if cells are targeted while they progress through S phase. Our studies define new reaction parameters that will help guide experimental strategies of gene editing.

Electrospun fiber membranes enable proliferation of genetically modified cells.

Polycaprolactone (PCL) and its blended composites (chitosan, gelatin, and lecithin) are well-established biomaterials that can enrich cell growth and enable tissue engineering. However, their application in the recovery and proliferation of genetically modified cells has not been studied. In the study reported here, we fabricated PCL-biomaterial blended fiber membranes, characterized them using physicochemical techniques, and used them as templates for the growth of genetically modified HCT116-19 colon cancer cells. Our data show that the blended polymers are highly miscible and form homogenous electrospun fiber membranes of uniform texture. The aligned PCL nanofibers support robust cell growth, yielding a 2.5-fold higher proliferation rate than cells plated on standard plastic plate surfaces. PCL-lecithin fiber membranes yielded a 2.7-fold higher rate of proliferation, while PCL-chitosan supported a more modest growth rate (1.5-fold higher). Surprisingly, PCL-gelatin did not enhance cell proliferation when compared to the rate of cell growth on plastic surfaces.

Proliferation of genetically modified human cells on electrospun nanofiber scaffolds.

Gene editing is a process by which single base mutations can be corrected, in the context of the chromosome, using single-stranded oligodeoxynucleotides (ssODNs). The survival and proliferation of the corrected cells bearing modified genes, however, are impeded by a phenomenon known as reduced proliferation phenotype (RPP); this is a barrier to practical implementation. To overcome the RPP problem, we utilized nanofiber scaffolds as templates on which modified cells were allowed to recover, grow, and expand after gene editing. Here, we present evidence that some HCT116-19, bearing an integrated, mutated enhanced green fluorescent protein (eGFP) gene and corrected by gene editing, proliferate on polylysine or fibronectin-coated polycaprolactone (PCL) nanofiber scaffolds. In contrast, no cells from the same reaction protocol plated on both regular dish surfaces and polylysine (or fibronectin)-coated dish surfaces proliferate. Therefore, growing genetically modified (edited) cells on electrospun nanofiber scaffolds promotes the reversal of the RPP and increases the potential of gene editing as an ex vivo gene therapy application.Molecular Therapy - Nucleic Acids (2012) 1, e59; doi:10.1038/mtna.2012.51; published online 4 December 2012.

Oligonucleotide delivery by nucleofection does not rescue the reduced proliferation phenotype of gene-edited cells.

Gene editing using single-stranded oligonucleotides (ODNs) can be used to reverse or create a single base mutation in mammalian cells. This approach could be used to treat genetic diseases caused, at least in part, by a nucleotide substitution. The technique could also be used as a tool to establish single base polymorphisms at multiple sites and thus aid in creating cell lines that can be used to define the basis for drug resistance in human cells. A troubling outcome of the gene-editing reaction is the effect on normal growth of cells that have undergone nucleotide exchange. In this work, we attempt to overcome this reduced proliferation phenotype by changing the method by which the ODN is introduced into the target cell. Using a series of assays that measure gene editing, DNA damage response, and cell viability, we report that chemically modified ODNs, the most active form of ODN for gene editing, can be used successfully if introduced into the cell by the method of nucleofection. Unlike electroporation, which has been used as the standard mode of ODN delivery, one new result is that nucleofection does not induce a dramatic loss of viability within the first 24 hours after the start of gene editing. In addition, and importantly, ODNs introduced to the cell by nucleofection do not activate the DNA damage response pathway as dramatically as ODNs introduced by electroporation. These 2 novel findings are encouraging for the application of gene editing in other systems. However, reduced proliferation phenotype is still observed when the population of corrected cells is monitored out to 8 days, and thus, delivery by nucleofection does not solve the proliferation problem encountered by cells bearing an edited gene.