Systematic evaluation of CrRNA design parameters for optimized Cas13d-mediated RNA targeting in chicken cells.

NEAT1-triggered collateral activity of CRISPR/Cas13 exhibits antileukemic activity

Manipulation of Cell Physiology Enables Gene Silencing in Well-differentiated Airway Epithelia

Modified siRNA Structure With a Single Nucleotide Bulge Overcomes Conventional siRNA-mediated Off-target Silencing

CRISPR-Cas13 exclusively targets RNA. In prokaryotes, Cas13 cleaves both target and non-target RNA indiscriminately upon activation by a specific target RNA, but in eukaryotic cells collateral cleavage activity has been limited. Here we report that LbuCas13a exhibits strong collateral RNA cleavage activity in human cells when delivered as ribonucleoprotein, independent of cell line and targeting both exogenous and endogenous transcripts. Collateral RNA cleavage starts within 50 minutes of ribonucleoprotein delivery resulting in major alterations to the total RNA profile. In response to the collateral RNA cleavage, cells upregulate genes associated with the stress and innate immune response, ultimately leading to apoptotic cell death. This enables us to use LbuCas13a as a flexible and repeatable target-RNA-specific cell elimination tool. Finally, using both total RNA sequencing and Nanopore sequencing, we find that LbuCas13a activation leads to rapid and near-global depletion of cytoplasmic RNAs, and that cleavage occurs at specific nucleotide positions.

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with an attractive off-target profile at both cellular and organismal levels. It is recently reported that the CRISPR/CasRx system can be used to achieve ubiquitous and tissue-specific gene transcript reduction in Drosophila melanogaster. This paper details the methods from the recent work, consisting of three parts: 1) ubiquitousin vivo endogenous RNA targeting using a two-component CasRx system; 2) ubiquitous in vivo exogenous RNA targeting using a three-component CasRx system; and 3) tissue-specific in vivo RNA targeting using a three-component CasRx system. The effects of RNA targeting observed include targeted gene specific phenotypic changes, targeted RNA transcript reduction, and occasional lethality phenotypes associated with high expression of CasRx protein and collateral activity. Overall, these results showed that the CasRx system is capable of target RNA transcript reduction at the organismal level in a programmable and efficient manner, demonstrating that in vivo transcriptome targeting, and engineering is feasible and lays the foundation for future in vivo CRISPR-based RNA targeting technologies.

CasRx, a member of the RNA-targeting Cas13 family, is a promising new addition of the CRISPR/Cas technologies in efficient gene transcript reduction with an attractive off-target profile at both cellular and organismal levels. It is recently reported that the CRISPR/CasRx system can be used to achieve ubiquitous and tissue-specific gene transcript reduction in Drosophila melanogaster. This paper details the methods from the recent work, consisting of three parts: 1) ubiquitous in vivo endogenous RNA targeting using a two-component CasRx system; 2) ubiquitous in vivo exogenous RNA targeting using a three-component CasRx system; and 3) tissue-specific in vivo RNA targeting using a three-component CasRx system. The effects of RNA targeting observed include targeted gene specific phenotypic changes, targeted RNA transcript reduction, and occasional lethality phenotypes associated with high expression of CasRx protein and collateral activity. Overall, these results showed that the CasRx system is capable of target RNA transcript reduction at the organismal level in a programmable and efficient manner, demonstrating that in vivo transcriptome targeting, and engineering is feasible and lays the foundation for future in vivo CRISPR-based RNA targeting technologies.

Escalating vector disease burdens pose significant global health risks, as such innovative tools for targeting mosquitoes are critical. CRISPR-Cas technologies have played a crucial role in developing powerful tools for genome manipulation in various eukaryotic organisms. Although considerable efforts have focused on utilizing class II type II CRISPR-Cas9 systems for DNA targeting, these modalities are unable to target RNA molecules, limiting their utility against RNA viruses. Recently, the Cas13 family has emerged as an efficient tool for RNA targeting; however, the application of this technique in mosquitoes, particularly Aedes aegypti, has yet to be fully realized. In this study, we engineered an antiviral strategy termed REAPER (vRNA Expression Activates Poisonous Effector Ribonuclease) that leverages the programmable RNA-targeting capabilities of CRISPR-Cas13 and its potent collateral activity. REAPER remains concealed within the mosquito until an infectious blood meal is uptaken. Upon target viral RNA infection, REAPER activates, triggering programmed destruction of its target arbovirus such as chikungunya. Consequently, Cas13-mediated RNA targeting significantly reduces viral replication and viral prevalence of infection, and its promiscuous collateral activity can even kill infected mosquitoes within a few days. This innovative REAPER technology adds to an arsenal of effective molecular genetic tools to combat mosquito virus transmission.

SummaryCRISPR-Cas13 systems are widely used in basic and applied sciences. However, its application has recently generated controversy due to collateral activity in mammalian cells and mouse models. Moreover, its efficiency could be improved in vivo. Here, we optimized transient formulations as ribonucleoprotein complexes or mRNA-gRNA combinations to enhance the CRISPR-RfxCas13d system in zebrafish. We i) used chemically modified gRNAs to allow more penetrant loss-of-function phenotypes, ii) improved nuclear RNA-targeting, and iii) compared different computational models and determined the most accurate to predict gRNA activity in vivo. Furthermore, we demonstrated that transient CRISPR-RfxCas13d can effectively deplete endogenous mRNAs in zebrafish embryos without inducing collateral effects, except when targeting extremely abundant and ectopic RNAs. Finally, we implemented alternative RNA-targeting CRISPR-Cas systems with reduced or absent collateral activity. Altogether, these findings contribute to CRISPR-Cas technology optimization for RNA targeting in zebrafish through transient approaches and assist in the progression of in vivo applications.

Clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas) systems, especially type II (Cas9) systems, have been widely developed for DNA targeting and formed a set of mature precision gene-editing systems. However, the basic research and application of the CRISPR-Cas system in RNA is still in its early stages. Recently, the discovery of the CRISPR-Cas13 type VI system has provided the possibility for the expansion of RNA targeting technology, which has broad application prospects. Most type VI Cas13 effectors have dinuclease activity that catalyzes pre-crRNA into mature crRNA and produces strong RNA cleavage activity. Cas13 can specifically recognize targeted RNA fragments to activate the Cas13/crRNA complex for collateral cleavage activity. To date, the Cas13X protein is the smallest effector of the Cas13 family, with 775 amino acids, which is a promising platform for RNA targeting due to its lack of protospacer flanking sequence (PFS) restrictions, ease of packaging, and absence of permanent damage. This study highlighted the latest progress in RNA editing targeted by the CRISPR-Cas13 family, and discussed the application of Cas13 in basic research, nucleic acid diagnosis, nucleic acid tracking, and genetic disease treatment. Furthermore, we clarified the structure of the Cas13 protein family and their molecular mechanism, and proposed a future vision of RNA editing targeted by the CRISPR-Cas13 family.

The clustered regularly interspaced short palindromic repeat (CRISPR)-CRISPR-associated genes (Cas) adaptive immune system defends microbes against foreign genetic elements via DNA or RNA-DNA interference. We characterize the class 2 type VI CRISPR-Cas effector C2c2 and demonstrate its RNA-guided ribonuclease function. C2c2 from the bacterium Leptotrichia shahii provides interference against RNA phage. In vitro biochemical analysis shows that C2c2 is guided by a single CRISPR RNA and can be programmed to cleave single-stranded RNA targets carrying complementary protospacers. In bacteria, C2c2 can be programmed to knock down specific mRNAs. Cleavage is mediated by catalytic residues in the two conserved Higher Eukaryotes and Prokaryotes Nucleotide-binding (HEPN) domains, mutations of which generate catalytically inactive RNA-binding proteins. These results broaden our understanding of CRISPR-Cas systems and suggest that C2c2 can be used to develop new RNA-targeting tools.

While lipid nanoparticles (LNPs) proved to be effective in delivering siRNA to the liver and hepatocellular carcinoma (HCC) via systemic administration, they showed inferior or no efficacy in the extrahepatic tumor models. To understand the underlying mechanism for LNP activity in delivering siRNA to extrahepatic tumors, we determined siRNA biodistribution and gene-silencing activity of the LNP-siRNA nanoparticles containing different percentages of a polyethylene glycol (PEG) lipid, distinct PEG lipids with a short (C-14) or long (C-18) alkyl chain, or an identical composition with different particle sizes in mouse subcutaneous tumor xenograft models derived from either human HCC (Hep3B) or colon cancer (HT29 or HCT116) cells. We show that the LNPs containing C-18 DSA PEG exhibited a higher retention of siRNA in both Hep3B and HT29 tumors than those with C-14 DMA PEG following an intravenous administration. Intriguingly, LNPs containing 2% of DSA or DMA PEG resulted in similar levels (∼50%) of target knockdown in Hep3B tumors while those containing 6% of DSA or DMA PEG caused no target silencing. Also, none of these LNPs reduced target expression in HT29 tumors despite the retention of siRNA in tumor tissues. When LNPs with an identical composition (2% DMG PEG) but different particle sizes (80 nm vs. 120 nm) were tested, we observed a comparable and moderate suppression of target expression in Hep3B tumors, but no target knockdown in HCT116 tumors. This is despite the fact that the expression of low density lipid receptor, which is shown to be involved in mediating LNP efficacy, is detected in HT29 and HCT116 tumors. Finally, intratumor injection of LNPs only induced minimal target silencing in subcutaneous tumors while provoking significant target knockdown in the liver. Together, enhanced tumor retention of siRNA might not be readily translated to target silencing activity in tumors. Other factors such as the efficiency of transfecting tumor cells and the passive targeting effect through opsonization could impact on LNP efficacy in extrahepatic tumors. Citation Format: {Authors}. {Abstract title} [abstract]. In: Proceedings of the 103rd Annual Meeting of the American Association for Cancer Research; 2012 Mar 31-Apr 4; Chicago, IL. Philadelphia (PA): AACR; Cancer Res 2012;72(8 Suppl):Abstract nr 2888. doi:1538-7445.AM2012-2888

Article Figures and data Abstract eLife digest Introduction Results Discussion Materials and methods References Decision letter Author response Article and author information Metrics Abstract Transcription is an inherently stochastic, noisy, and multi-step process, in which fluctuations at every step can cause variations in RNA synthesis, and affect physiology and differentiation decisions in otherwise identical cells. However, it has been an experimental challenge to directly link the stochastic events at the promoter to transcript production. Here we established a fast fluorescence in situ hybridization (fastFISH) method that takes advantage of intrinsically unstructured nucleic acid sequences to achieve exceptionally fast rates of specific hybridization (∼10e7 M−1s−1), and allows deterministic detection of single nascent transcripts. Using a prototypical RNA polymerase, we demonstrated the use of fastFISH to measure the kinetic rates of promoter escape, elongation, and termination in one assay at the single-molecule level, at sub-second temporal resolution. The principles of fastFISH design can be used to study stochasticity in gene regulation, to select targets for gene silencing, and to design nucleic acid nanostructures. https://doi.org/10.7554/eLife.01775.001 eLife digest The body produces proteins by transcribing DNA (genes) to make messenger RNA, which is then translated to make a protein. Transcription begins when an enzyme called RNA polymerase binds to the DNA and catalyzes the process by which genetic information from the double helix is copied to a complementary RNA transcript, which subsequently becomes the messenger RNA. Because a living cell usually contains only one or a few copies (alleles) of a given gene, molecular fluctuations play a crucial role in cellular transcription. Therefore, studying transcription kinetics at the level of single molecules may provide critical insights into how cells deal with—or even take advantage of—molecular fluctuations. A number of different single-molecule techniques can be used to follow transcription, but these techniques are often relatively slow compared to transcription in living cells, or they suffer from other problems such as only being able to study one step in the transcription process. Now, Zhang, Revyakin et al. have systematically devised a technique called ‘fastFISH’ that is fast enough to track the production of single RNA molecules directly and instantaneously. FastFISH builds on an existing technique called FISH—short for fluorescence in situ hybridization—in which fluorescent molecules are attached to single strands of DNA or RNA. These single strands pair with specific regions of complementary DNA or RNA molecules, and they can be visualized with a fluorescence microscope. However, conventional FISH is a ‘snap-shot’ technique that is not suitable for making real-time observations under physiological conditions. FastFISH relies on single strands of fluorescently labeled DNA and RNA that bind to complementary strands of DNA or RNA extremely quickly, even under physiological conditions, because they contain only three of the four ‘regular’ nucleotides that make up DNA or RNA. As a proof of principle, Zhang, Revyakin et al. used fastFISH to study the kinetics of transcription by the bacteriophage T7 RNA polymerase and were able to measure multiple stages of the transcription cycle in a single-molecule experimental setup. By allowing each stage of transcription to be tracked in real-time at the level of single-molecules, fastFISH will permit a more in-depth analysis of the factors that regulate how genes are expressed as proteins in our cells. Moreover, the ability to design single-strand probes that bind rapidly to DNA and RNA targets could have many additional applications, including new strategies for more efficient gene silencing. https://doi.org/10.7554/eLife.01775.002 Introduction Transcription is the first, and frequently the most regulated step in the flow of genetic information from DNA to protein. Transcription is a dynamic, multi-step process in which RNA polymerase (RNAP) (i) binds to the promoter to form the closed complex (RPc); (ii) melts the promoter to form the open complex (RPo); (iii) performs several abortive cycles of synthesis and release of short 2–12 nt RNA products as the initial transcribing complex (RPitc); (iv) escapes the promoter; (v) undergoes some promoter-proximal pausing; (vi) forms an elongating complex (RDe) whose processivity can either be interrupted by more pauses or stimulated by trailing RNAPs and elongation factors; and, finally (vii) terminates at the end of the transcription unit (reviewed in DeHaseth et al., 1998; Murakami and Darst, 2003; Cheung and Cramer, 2012). Due to the low copy number of genes in a cell (usually one in prokaryotes and two in eukaryotes), molecular fluctuations at any of the above steps may cause large cell-to-cell variability in the amount of the final RNA transcript produced in populations of otherwise genetically identical cells grown under identical conditions, and thus can affect gene expression, and cell physiology (Ogawa, 1993; Raj and Van Oudenaarden, 2009; Yamanaka, 2009; Gupta et al., 2011; Lionnet and Singer, 2012). Therefore, understanding how molecular fluctuations at different steps of the transcription cycle alter transcriptional outcomes is required to dissect the mechanism of gene regulation. Single-molecule techniques are ideally suited to directly monitor molecular fluctuations in multi-step reactions in real-time, without averaging out their inherent stochasticity (Weiss, 1999), and have provided important insights into the dynamics of transcription, unattainable by conventional ensemble methods (Bai et al., 2006). For instance, single-molecule assays based on optical nanomanipulation have revealed pausing and backtracking of RDe (Neuman et al., 2003; Shaevitz et al., 2003; Galburt et al., 2007) and measured the force and the torque generated by RDe (Wang et al., 1998; Ma et al., 2013). Methods based on magnetic nanomanipulation and single-molecule fluorescence resonance energy transfer have probed conformational changes in DNA and RNAP within RPo, and RPitc (Kapanidis et al., 2006; Revyakin et al., 2006; Tang et al., 2009; Chakraborty et al., 2012; Robb et al., 2013). Finally, single-molecule localization studies have probed the dynamics of the initial promoter search by RNAP (Wang et al., 2012; Friedman et al., 2013). However, most current single-molecule methods focus on only one step of transcription, and are not well suited to relate protein-DNA complex assembly and dynamics during multiple stages of the transcription cycle to RNA production. In addition, most methods do not measure RNA production directly, but rather infer it from changes in DNA conformation or movement of RNAP along the DNA. Most recently, single-molecule tracking of key protein-DNA interactions coupled with detection of the RNA production has been demonstrated in the bacterial (Friedman and Gelles, 2012) and human transcription systems (Revyakin et al., 2012). The former study achieved time resolution for RNA detection on the order of ∼10 s, and thus provided a dynamic, quantitative view of the full transcription cycle of bacterial RNAP. However, time scales of many events in transcription are on the order of ∼1 s, particularly under physiological conditions (for instance, the residence time of transcription factors on DNA (reviewed in Hager et al., 2009), the rate of initiation and promoter clearance by RNAPs (Revyakin et al., 2006; Tang et al., 2009), and the expected time delay between cooperatively elongating RNAPs molecules (Epshtein and Nudler, 2003)). Thus, a real-time method for nascent transcript detection at ∼1 s time scales would significantly enhance our ability to dissect the dynamics of the transcription process, and to correlate stochastic fluctuations at different steps with transcriptional outcomes. Currently, the most sensitive and specific methods for detecting RNA transcripts continue to rely on complementary nucleic acid hybridization. However, oligonucleotide probes typically hybridize several orders of magnitude slower than diffusion under physiological conditions (effective rate constant kon less than 105 M−1 s−1, [Chan et al., 1995 and references therein]) and, as a result, do not permit real-time nascent RNA detection with common single-molecule setups, such as total internal reflection fluorescence (TIRF) microscopy. Here we present a fast fluorescence in situ hybridization method (fastFISH) to detect synthesis of nascent RNA transcripts at sub-second time resolution, at the single-molecule level. The method takes advantage of our finding that single-stranded nucleic acid probes with sequences comprised of just three of the four bases (A, U and C for RNA probes, and A, T, and G for the complementary DNA targets) are intrinsically unstructured and, as a result, hybridize at exceptionally fast rates (∼107 M−1s−1) without compromising sequence specificity. As a proof of concept, we applied fastFISH to probe the production of nascent RNA transcripts by the speedy bacteriophage T7 RNA Polymerase (T7 RNAP) in vitro. Furthermore, by using fastFISH in combination with fluorescently labeled RNAP, we dissected the full T7 RNAP transcription cycle (promoter binding, promoter escape, elongation, and termination). FastFISH can be used to study transcription by multi-subunit prokaryotic and eukaryotic RNA polymerases at the level of stochastic molecular interactions. The rules for generating fastFISH probe-target pairs can also be used for gene silencing, gene profiling and bottom-up assembly of nucleic acid nanostructures (Rothemund, 2006). Results Design and characterization of fastFISH hybridization probes To achieve real-time nascent RNA detection, we set out to find a general rule for designing RNA-probe pairs that hybridize at the fastest possible rates. Hybridization of two single-stranded nucleic acid fragments requires unfolding of the fragments into unstructured coils, which then anneal to form an intermolecular base paired helix (Lima et al., 1992 and references therein). In support of this notion, nucleic acid probes with less stable secondary structures show faster hybridization rates to complementary nucleic acid targets (Lima et al., 1992; Kushon et al., 2001; Wang and Drlica, 2004; Gao et al., 2006; Yilmaz et al., 2006). Likewise, decreasing the length of hairpins in conventional molecular beacons increases their hybridization rates (Tsourkas et al., 2003). Thus, we reasoned that intrinsically unstructured nucleic acid oligonucleotide pairs of optimal lengths should have the fastest hybridization kinetics under physiological conditions without compromising specificity. To systematically examine the propensity of RNA sequences to form base-paired secondary structures, we used the nucleic acid structure prediction tool Mfold (Zuker, 2003) to calculate the free energy of self-folding, ΔG37°C, of randomly selected short RNA sequences (N = 338,417). We chose RNA sequence with length of 19 nt and GC content between 0.4 and 0.6 as representative of a typical oligonucleotide primer. We found that ΔG37°C values were mostly negative and widely distributed (−1.7 ± 1.7 kCal mol−1), indicating that an average random 19-mer RNA sequence is mostly structured (Figure 1A). Next, we closely examined a subset of 19-mer RNA sequences that had positive ΔG37°C (>+2 kCal mol−1, N = 2,768, <1% of the total pool of random sequences), and found that these mostly unstructured sequences were composed predominantly of only three bases A, U, and C (∼84% had only 2 G residues or less, significantly less than the 4.75 G residues expected on average, Figure 1—figure supplement 1). The bias towards the lower G-content in the unstructured sequences was not surprising, because guanine is the most potent in base-paring interactions: it forms a triple hydrogen bond with cytosine and a wobble pair with uracil. We then calculated ΔG37°C for randomly picked 19-mers composed of only A, U, and C (N = 99,777, GC content between 0.4 and 0.6, Figure 1A), and found that these AUC-sequences had mostly positive ΔG37°C, with a much narrower distribution (+2.5 ± 0.6 kCal mol−1, Figure 1A). Analysis of other three-base-derived RNA sequences (AUG, CAG, CUG) indicated that AUC-sequences were unique in their largely positive ΔG37°C (Figure 1A). Calculation of ΔG37°C for random DNA 19-mers (that could be used as complementary probes for RNA targets) indicated that ATG-based and ATC-based DNA 19-mers were also significantly less structured than their four-base counterparts (Figure 1A). Therefore, we hypothesized that the use of the AUC alphabet for RNA targets, and ATG alphabet for DNA probes should allow the fastest annealing reactions under physiological temperature of 37°C. Figure 1 with 3 supplements see all Download asset Open asset Design of fastFISH probe-target pairs. (A) Probability distribution of Mfold-calculated free energies of self-folding of randomly selected, single-stranded 19-mer RNA (left) and DNA (right) oligonucleotides, composed of three or four bases. Results of analysis of three independent sets are shown as ‘+’, ‘x’, and ‘○’. About 100,000 three-letter sequences, and about 300,000 four-letter sequences were analyzed in each set. (B) Lempel–Ziv complexity analysis of three-letter 19-mer oligonucleotides (one set of ∼100,000 AUC sequences), four-letter 19-mer oligonucleotides (one set of ∼300,000 AUGC sequences), and all tiling 19-mers from the exome of the human chromosome 22. (C) Single-molecule measurements of hybridization rates of fastFISH probe-target pairs, and the effect of G-residues in the targets on the rates. Left: schematic of experiment. Cy3-labeled 90-base RNA oligonucleotides containing a single target sequence were immobilized on a glass surface through a biotin moiety at the 3′ end. Atto633-labeled DNA probes were injected into the imaging flow cell, and their hybridization was detected using TIRF/CoSMoS to obtain the probe arrival time Twait. Right: table of RNA target sequences, Mfold-calculated free energies of self-folding of RNA targets (ΔGtarget), DNA probes (ΔGprobe), combined energies of targets and probes, and on-rates calculated based on probe Twait and concentrations. (D) Self-quenching approach to reduce fluorescence background from unbound DNA probes in TIRF imaging of probe-target hybridization. Left: schematic of experiment. A quencher (e.g., Iowa Black FQ) is placed on one end of a DNA probe, and a fluorophore (e.g., Cy3) is placed on the opposite end of the DNA probe. The short persistence length of single-stranded DNA (lo ∼0.8 nm, Smith et al., 1996; Dessinges et al., 2002) enables quenching of Cy3. Upon hybridization to the target, the distance between Cy3 and Iowa Black FQ increases due to the larger lo ∼50 nm of the DNA-RNA duplex, leading to an increase of Cy3 fluorescence. Middle: representative TIRF image and a corresponding three-dimensional plot of target-hybridized F1 probes acquired in the presence of unbound, self-quenched F1 probe at 100 nM. Right: same set of molecules imaged in the presence of unbound, unquenched F1 probe at 100 nM. https://doi.org/10.7554/eLife.01775.003 To ensure that the use of the three-base alphabet did not compromise the specificity of probe-target hybridization, we calculated the textual complexity of randomly picked three-letter 19-mers using the algorithm of Lempel and Ziv (Ziv and Lempel, 1976; Kaspar and Schuster, 1987; Orlov and Potapov, 2004). This algorithm, commonly used in lossless data compression programs, calculates the minimal number of operations required to reconstruct a sequence of symbols by copying and inserting segments of an existing sub-sequence. Thus, nucleic acid sequences that contain repetitive elements would have a lower LZ complexity (and would be less specific in hybridization) (Wright and Church, 2002). We found that three-letter 19-mer sequences had complexity indices of 8.1 ± 0.8 (N = 99,777), while four-letter 19-mers had complexity indices of 9.3 ± 0.8 (N = 338,417, Figure 1B). As a comparative reference, we calculated complexity indices for tiling 19-mers in all exons of human chromosome 22 to be 8.9 ± 1.0 (Figure 1B). Importantly, a significant fraction of three-letter 19-mers (∼31%) had complexity indices of 9 and higher, which matched or exceeded the average complexity of human exons. These calculations indicate that a significant fraction of the random probe sequences composed of only three bases nevertheless retain sequence complexity and specificity that match their physiological, four-base derived counterparts. We applied the AUC rule and the complexity filter to generate two candidate probe-target pairs (referred to as F1 and F2). The calculated ΔG37°C for the F1 and F2 pairs were +1.5 and +2.2 kCal mol−1, respectively, for the AUC-based RNA targets, and +1.0 and +0.9 kcal mol−1 for their ATG-based complementary DNA probes (Figure 1C), suggesting that the pairs are mostly unstructured and likely to be fast-hybridizing. Indeed, the F1 and F2 probes annealed to their surface-immobilized RNA targets at exceptionally fast rates—6 × 106 M−1s−1 and 4 × 106 M−1s−1, respectively, as measured by TIRF-based Colocalization Single-Molecule Spectroscopy (CoSMoS, Friedman et al., 2006) (Figure 1C, Figure 1—figure supplement 2). These on-rates were at least 100-fold faster than typical four-base nucleic acid probes of similar lengths reported in literature (Lima et al., 1992; Kushon et al., 2001; Wang and Drlica, 2004; Friedman et al., 2006; Gao et al., 2006; Yilmaz et al., 2006). Consistent with the AUC rule, introducing back one or more G residues into the F2 RNA target sequence decreased the rate of its hybridization to the complementary DNA probe (10-fold reduction for one G residue and 300-fold reduction for four G residues), which correlated with the progressively lower free energies of self-folding (Figure 1C, Figure 1—figure supplement 2D). We conclude that the target sequences designed using the AUC rule, combined with the complexity filter, achieved our goal of generating superior hybridization kinetics for fast transcript detection. Although the F1 and F2 probes were exceptionally fast, detection of RNA on the sub-second time scales would require the use of fluorescent probe concentrations above 100 nM (Figure 1—figure supplement 2D). Such concentrations are generally incompatible with common single-molecule detection setups such as TIRF microscopy, due to high fluorescence background from the freely diffusing, unbound probe molecules. We overcame this problem by attaching a quencher to the single-stranded probe at the end opposite of the fluorophore (Marras et al., 2002). This self-quenching strategy enabled the use of free probes at concentrations up to 1 μM (Figure 1D, Figure 1—figure supplement 3). We believe that, in the absence of the secondary structure, the self-quenching effect was likely mediated by random polymer motion and/or contact quenching (Johansson et al., 2002; Marras et al., 2002). We refer to our method for fast nucleic acid detection using unstructured, sequence specific, self-quenched fluorescent probes as ‘fastFISH’. Real-time single-molecule detection of transcription with fastFISH To demonstrate that fastFISH can detect nascent transcripts in real time at single-molecule resolution, we used the bacteriophage T7 RNAP. This single-subunit RNAP is an excellent test case for fastFISH: at physiological temperature (37°C) it initiates transcripts at an effective rate of at least 1 s−1 (Martin and Coleman, 1987), and elongates transcripts at ∼240 nt s−1 (Golomb and Chamberlin, 1974; Bonner et al., 1994). As a model template, we used a fluorescently labeled linear DNA fragment containing the consensus promoter for T7 RNAP (Milligan et al., 1987) and the F1 target sequence between and from the transcription and at We surface-immobilized the Cy3-labeled DNA a biotin moiety attached to the end of the (Figure In this RNAP molecules were expected to transcription at the the nascent RNA in the from the and the end at with their nascent RNA Figure 2 Download asset Open asset Real-time single-molecule detection of transcription by T7 RNAP using (A) of experiment. DNA containing a single consensus promoter for T7 RNAP or a single promoter were immobilized on a with the promoter transcription towards the free end The the F1 fastFISH target sequence from the promoter to which was expected to for hybridization in the nascent RNA the RNAP (B) analysis of F1 interactions in a representative experiment. Left: Right: (C) and (D) as (A) and but for the F2 target and probe. The the F2 fastFISH target sequence to which was expected to for hybridization in the nascent RNA the RNAP We RNAP, and the self-quenched F1 probe to the imaging and the interactions of the F1 probe with the DNA by TIRF/CoSMoS (Friedman et al., 2006; Revyakin et al., 2012). We reasoned that the F1 target sequence in the nascent RNA would for hybridization the of elongating RNAP (RDe) into the bases of nascent RNA by RNAP and The RDe complex would then on the DNA it at the expected average RNAP elongation rate of nt s−1, the F1 probe would have a ∼1 s of to the nascent transcript at the DNA of hybridization to the RNA would then be as a fluorescent with the DNA Figure the analysis of interactions in a typical single-molecule fastFISH experiment. This analysis all DNA molecules that with a probe for more than 0.4 during the and the between the probe and DNA molecules as a (Revyakin et al., 2012). interactions were as indicated by a at of the In a typical of DNA in a of view with a probe. This was not due to probe hybridization Figure 4 and was likely by the of the DNA due to surface interactions were in a with a DNA containing a promoter et al., indicating that the was due to transcription. We also real-time single-molecule transcription using the other unstructured probe, and identical (Figure these that fastFISH can detect production of nascent transcripts in Single-molecule dynamics of T7 interactions To measure the of real-time detection of nascent transcripts by we a to the and end of each single-molecule transcription We reasoned that the interactions between RNAP molecules and the DNA during promoter and can this Therefore, we fluorescently labeled RNAP with using (Figure supplement and to an imaging flow cell containing immobilized DNA and interactions with the DNA molecules (Figure 3). We RNAP events that between and s, and were (Figure The events were comprised of two events whose well to a single distribution time and a towards events time The of interactions were not by the of the (Figure supplement 2). We the stochastic interactions to be but and RPo, because interactions were with containing a promoter sequence rates of Figure and similar interactions were in the absence of s, Figure Figure 3 with 2 supplements see all Download asset Open asset Single-molecule dynamics of T7 interactions. (A) of experiment. (B) analysis of interactions. Left: promoter DNA in the presence of promoter DNA in the presence of Right: promoter DNA in the absence of (C) of RNAP interactions with containing consensus promoter for a × of at a DNA = nm, imaged at fluorescence time corresponding to the shown on of Left: out in the presence of Right: out in the absence of indicate the of RNAP (D) time of Left: out in the presence of to a of single and is shown in Right: out in the absence of to a single is shown in of the time of interactions on the length of the DNA schematic of experiment. DNA containing segments from to or were and interactions of labeled RNAP were at time of interactions for the three DNA shown in N = for the N = for the and N = for the DNA The were calculated by the to a of single and of the time of interactions the length of the DNA of of RNAP fluorescence during interactions as an of elongation by RNAP. interactions than 0.8 s in were by RNAP = and by RNAP and of RNAP and were for DNA segments of different lengths between RNAP and were set at for the DNA measured in We the of events to be transcription because (i) the events were not in the absence of (Figure (ii) the time of the with the length of the DNA s, s, and s for DNA segments with = and respectively, Figure and (iii) the RNAP fluorescence on average, towards the end of events (Figure with RNAP transcription at the promoter to the surface or and then elongating towards the DNA the of the TIRF We conclude that the time of interactions transcription events which can be used as a to the of nascent RNA detection by and to dissect the full transcription cycle by RNAP. We that the of the plot of RNAP DNA ± × s (Figure an of RNAP elongation rate of nt The of the plot with the time = an of the time that RNAP on the DNA without = ± s, which the of promoter and abortive and and, the time RNAP at the free DNA end Single-molecule of the T7 RNAP transcription cycle with fastFISH To the of nascent RNA detection by and to demonstrate the use of fastFISH in the kinetics of the full transcription we interactions and the production

Purification and in vivo, cell-free, and in vitro characterization of CRISPR-Cas12a2.

Escalating vector disease burdens pose significant global health risks, so innovative tools for targeting mosquitoes are critical. We engineered an antiviral strategy termed REAPER (vRNA Expression Activates Poisonous Effector Ribonuclease) that leverages the programmable RNA-targeting capabilities of CRISPR Cas13 and its potent collateral activity. Akin to a stealthy Trojan Horse hiding in stealth awaiting the presence of its enemy, REAPER remains concealed within the mosquito until an infectious blood meal is up taken. Upon target viral RNA infection, REAPER activates, triggering programmed destruction of its target arbovirus such as chikungunya. Consequently, Cas13 mediated RNA targeting significantly reduces viral replication and its promiscuous collateral activity can even kill infected mosquitoes. This innovative REAPER technology adds to an arsenal of effective molecular genetic tools to combat mosquito virus transmission.

The approach to characterize preparations of recombinant Cas13a-nuclease in terms of specific collateral activity has been proposed for standardization of enzyme preparations. The standardization of Cas13a preparations by the specific activity may benefit both the development of assays employing Cas13a collateral ribonuclease activity and the optimization of ribonuclease expression, purification, and storage. The approach is based on measurement of the initial rate of a cleavage of specially designed commercially available RNA molecules (�reporters� labelled with a fluorophore and a quencher) by a preparation of recombinant Cas13a-nuclease and commercial RNase A of known activity. This requires the optimization of a molar ratio for the formation of Cas13a complexes with guide RNA as well as the optimization of amount of the RNA target. The use of a synthetic RNA target appears preferable compared with total RNA preparations.