Visualization of RNA virus infection in a marine protist with a universal biomarker.

Half of the marine virosphere is hypothesized to be RNA viruses (kingdom Orthornavirae) that infect abundant micro-eukaryotic hosts (e.g. protists). To test this, quantitative approaches that broadly track infections in situ are needed. Here, we describe a technique-dsRNA-Immunofluorescence (dsRIF)-that uses a double-stranded RNA (dsRNA) targeting monoclonal antibody to assess host infection status based on the presence of dsRNA, a replicative intermediate of all Orthornavirae infections. We show that the dinoflagellate Heterocapsa circularisquama produces dsRIF signal ~ 1000 times above background autofluorescence when infected by the + ssRNA virus HcRNAV. dsRNA-positive virocells were detected across > 50% of the 48-h infection cycle and accumulated to represent at least 63% of the population. Photosynthetic and chromosomal integrity remained intact during peak replication, indicating HcRNAV infection does not interrupt these processes. This work validates the use of dsRIF on marine RNA viruses and their hosts, setting the stage for quantitative environmental applications that will accelerate understanding of virus-driven ecosystem impacts.

Comprehensive analysis and accurate quantification of unintended large gene modifications induced by CRISPR-Cas9 gene editing.

Most genome editing analyses to date are based on quantifying small insertions and deletions. Here, we show that CRISPR-Cas9 genome editing can induce large gene modifications, such as deletions, insertions, and complex local rearrangements in different primary cells and cell lines. We analyzed large deletion events in hematopoietic stem and progenitor cells (HSPCs) using different methods, including clonal genotyping, droplet digital polymerase chain reaction, single-molecule real-time sequencing with unique molecular identifier, and long-amplicon sequencing assay. Our results show that large deletions of up to several thousand bases occur with high frequencies at the Cas9 on-target cut sites on the HBB (11.7 to 35.4%), HBG (14.3%), and BCL11A (13.2%) genes in HSPCs and the PD-1 (15.2%) gene in T cells. Our findings have important implications to advancing genome editing technologies for treating human diseases, because unintended large gene modifications may persist, thus altering the biological functions and reducing the available therapeutic alleles.

Pitfalls in Single Clone CRISPR-Cas9 Mutagenesis to Fine-map Regulatory Intervals.

The majority of genetic variants affecting complex traits map to regulatory regions of genes, and typically lie in credible intervals of 100 or more SNPs. Fine mapping of the causal variant(s) at a locus depends on assays that are able to discriminate the effects of polymorphisms or mutations on gene expression. Here, we evaluated a moderate-throughput CRISPR-Cas9 mutagenesis approach, based on replicated measurement of transcript abundance in single-cell clones, by deleting candidate regulatory SNPs, affecting four genes known to be affected by large-effect expression Quantitative Trait Loci (eQTL) in leukocytes, and using Fluidigm qRT-PCR to monitor gene expression in HL60 pro-myeloid human cells. We concluded that there were multiple constraints that rendered the approach generally infeasible for fine mapping. These included the non-targetability of many regulatory SNPs, clonal variability of single-cell derivatives, and expense. Power calculations based on the measured variance attributable to major sources of experimental error indicated that typical eQTL explaining 10% of the variation in expression of a gene would usually require at least eight biological replicates of each clone. Scanning across credible intervals with this approach is not recommended.

Spatial control of in vivo CRISPR-Cas9 genome editing via nanomagnets.

The potential of clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR associated protein 9 (Cas9)-based therapeutic genome editing is hampered by difficulties in the control of the in vivo activity of CRISPR-Cas9. To minimize any genotoxicity, precise activation of CRISPR-Cas9 in the target tissue is desirable. Here, we show that, by complexing magnetic nanoparticles with recombinant baculoviral vectors (MNP-BVs), CRISPR-Cas9-mediated genome editing can be activated locally in vivo via a magnetic field. The baculoviral vector was chosen for in vivo gene delivery because of its large loading capacity and ability to locally overcome systemic inactivation by the complement system. We demonstrate that a locally applied magnetic field can enhance the cellular entry of MNP-BVs, thereby avoiding baculoviral vector inactivation and causing a transient transgene expression in the target tissue. Because baculoviral vectors are inactivated elsewhere, gene delivery and in vivo genome editing via MNP-BVs are tissue specific.

A Self-Deleting AAV-CRISPR System for In Vivo Genome Editing.

Adeno-associated viral (AAV) vectors packaging the CRISPR-Cas9 system (AAV-CRISPR) can efficiently modify disease-relevant genes in somatic tissues with high efficiency. AAV vectors are a preferred delivery vehicle for tissue-directed gene therapy because of their ability to achieve sustained expression from largely non-integrating episomal genomes. However, for genome editizng applications, permanent expression of non-human proteins such as the bacterially derived Cas9 nuclease is undesirable. Methods are needed to achieve efficient genome editing in vivo, with controlled transient expression of CRISPR-Cas9. Here, we report a self-deleting AAV-CRISPR system that introduces insertion and deletion mutations into AAV episomes. We demonstrate that this system dramatically reduces the level of Staphylococcus aureus Cas9 protein, often greater than 79%, while achieving high rates of on-target editing in the liver. Off-target mutagenesis was not observed for the self-deleting Cas9 guide RNA at any of the predicted potential off-target sites examined. This system is efficient and versatile, as demonstrated by robust knockdown of liver-expressed proteins in vivo. This self-deleting AAV-CRISPR system is an important proof of concept that will help enable translation of liver-directed genome editing in humans.

Trans-spliced Cas9 allows cleavage of HBB and CCR5 genes in human cells using compact expression cassettes.

CRISPR/Cas9 systems have been used in a wide variety of biological studies; however, the large size of CRISPR/Cas9 presents challenges in packaging it within adeno-associated viruses (AAVs) for clinical applications. We identified a two-cassette system expressing pieces of the S. pyogenes Cas9 (SpCas9) protein which splice together in cellula to form a functional protein capable of site-specific DNA cleavage. With specific CRISPR guide strands, we demonstrated the efficacy of this system in cleaving the HBB and CCR5 genes in human HEK-293T cells as a single Cas9 and as a pair of Cas9 nickases. The trans-spliced SpCas9 (tsSpCas9) displayed ~35% of the nuclease activity compared with the wild-type SpCas9 (wtSpCas9) at standard transfection doses, but had substantially decreased activity at lower dosing levels. The greatly reduced open reading frame length of the tsSpCas9 relative to wtSpCas9 potentially allows for more complex and longer genetic elements to be packaged into an AAV vector including tissue-specific promoters, multiplexed guide RNA expression, and effector domain fusions to SpCas9. For unknown reasons, the tsSpCas9 system did not work in all cell types tested. The use of protein trans-splicing may help facilitate exciting new avenues of research and therapeutic applications through AAV-based delivery of CRISPR/Cas9 systems.

Efficient fdCas9 Synthetic Endonuclease with Improved Specificity for Precise Genome Engineering.

The Cas9 endonuclease is used for genome editing applications in diverse eukaryotic species. A high frequency of off-target activity has been reported in many cell types, limiting its applications to genome engineering, especially in genomic medicine. Here, we generated a synthetic chimeric protein between the catalytic domain of the FokI endonuclease and the catalytically inactive Cas9 protein (fdCas9). A pair of guide RNAs (gRNAs) that bind to sense and antisense strands with a defined spacer sequence range can be used to form a catalytically active dimeric fdCas9 protein and generate double-strand breaks (DSBs) within the spacer sequence. Our data demonstrate an improved catalytic activity of the fdCas9 endonuclease, with a spacer range of 15-39 nucleotides, on surrogate reporters and genomic targets. Furthermore, we observed no detectable fdCas9 activity at known Cas9 off-target sites. Taken together, our data suggest that the fdCas9 endonuclease variant is a superior platform for genome editing applications in eukaryotic systems including mammalian cells.

CRISPR/Cas9 systems have off-target activity with insertions or deletions between target DNA and guide RNA sequences.

CRISPR/Cas9 systems are a versatile tool for genome editing due to the highly efficient targeting of DNA sequences complementary to their RNA guide strands. However, it has been shown that RNA-guided Cas9 nuclease cleaves genomic DNA sequences containing mismatches to the guide strand. A better understanding of the CRISPR/Cas9 specificity is needed to minimize off-target cleavage in large mammalian genomes. Here we show that genomic sites could be cleaved by CRISPR/Cas9 systems when DNA sequences contain insertions ('DNA bulge') or deletions ('RNA bulge') compared to the RNA guide strand, and Cas9 nickases used for paired nicking can also tolerate bulges in one of the guide strands. Variants of single-guide RNAs (sgRNAs) for four endogenous loci were used as model systems, and their cleavage activities were quantified at different positions with 1- to 5-bp bulges. We further investigated 114 putative genomic off-target loci of 27 different sgRNAs and confirmed 15 off-target sites, each harboring a single-base bulge and one to three mismatches to the guide strand. Our results strongly indicate the need to perform comprehensive off-target analysis related to DNA and sgRNA bulges in addition to base mismatches, and suggest specific guidelines for reducing potential off-target cleavage.

Gold nanoparticles electroporation enhanced polyplex delivery to mammalian cells.

Nonviral methods have been explored as the replacement of viral systems for their low toxicity and immunogenicity. However, they have yet to reach levels competitive to their viral counterparts. In this paper, we combined physical and chemical methods to improve the performance of polyplex delivery of DNA and small interfering RNA. Specifically, gold nanoparticles (AuNPs) were used to carry polyplex (a chemical approach) while electroporation (a physical approach) was applied for fast and direct cytosolic delivery. In this hybrid approach, cationic polymer molecules condense and/or protect genetic probes as usual while AuNPs help fix polycations to reduce their cytotoxicity and promote the transfection efficiency of electroporation. AuNPs of various sizes were first coated with polyethylenimine, which were further conjugated with DNA plasmids or small interfering RNA molecules to form AuNPs-polyplex. The hybrid nanoparticles were then mixed with cells and introduced into cell cytosol by electroporation. The delivery efficiency was evaluated with both model anchor cells (i.e., NIH/3T3) and suspension cells (i.e., K562), together with their impact on cell viability. We found that AuNP-polyplex showed 1.5∼2 folds improvement on the transfection efficiency with no significant increase of toxicity when compared to free plasmid delivery by electroporation alone. Such a combination of physical and chemical delivery concept may stimulate further exploration in the delivery of various therapeutic materials for both in vitro and in vivo applications.