Determinants of CRISPR Cas12a nuclease activation by DNA and RNA targets

Eric A Nalefski; Remy M Kooistra; Ishira Parikh; Samantha Hedley; Karunya Rajaraman; Damian Madan

doi:10.1093/nar/gkae152

. 2024 Mar 13;52(8):4502–4522. doi: 10.1093/nar/gkae152

Determinants of CRISPR Cas12a nuclease activation by DNA and RNA targets

Eric A Nalefski ^1,^✉, Remy M Kooistra ², Ishira Parikh ³, Samantha Hedley ⁴, Karunya Rajaraman ⁵, Damian Madan ^6,^✉

PMCID: PMC11077072 PMID: 38477377

Abstract

The RNA-guided CRISPR-associated (Cas) enzyme Cas12a cleaves specific double-stranded (ds-) or single-stranded (ss-) DNA targets (in cis), unleashing non-specific ssDNA cleavage (in trans). Though this trans-activity is widely coopted for diagnostics, little is known about target determinants promoting optimal enzyme performance. Using quantitative kinetics, we show formation of activated nuclease proceeds via two steps whereby rapid binding of Cas12a ribonucleoprotein to target is followed by a slower allosteric transition. Activation does not require a canonical protospacer-adjacent motif (PAM), nor is utilization of such PAMs predictive of high trans-activity. We identify several target determinants that can profoundly impact activation times, including bases within the PAM (for ds- but not ssDNA targets) and sequences within and outside those complementary to the spacer, DNA topology, target length, presence of non-specific DNA, and ribose backbone itself, uncovering previously uncharacterized cleavage of and activation by RNA targets. The results provide insight into the mechanism of Cas12a activation, with direct implications on the role of Cas12a in bacterial immunity and for Cas-based diagnostics.

Graphical Abstract

Introduction

Clustered regularly interspaced short palindromic repeats (CRISPR) systems provide bacteria with acquired immunity using enzymes and RNA guides transcribed from CRISPR loci to destroy invading DNA and RNA (1). One CRISPR-associated enzyme, Cas12a, cleaves specific double-stranded DNA (dsDNA) targets downstream of protospacer adjacent motifs (PAM) in cis (2) and non-specific single-stranded DNA (ssDNA) in trans (3–5), which has been coopted to generate detectable signals for diagnostic purposes (6). Though RNA can activate certain Cas12 family members (7–10), DNA but not RNA targets (10–12) have been shown to activate Cas12a from Lachnospiraceae bacterium (LbCas12a) (5,11,13). CRISPR RNA (crRNA) within the Cas12a ribonucleoprotein (RNP) hybridizes with the protospacer target strand (TS), displacing the non-target strand (NTS) (12,14–16). Formation of this R-loop induces a large conformational change within Cas12a, leading to activation of endonucleolytic activity, resulting in sequential cleavage of the NTS and TS and acquisition of trans-activity (17).

Since time to result is an important characteristic of any diagnostic, developers would benefit from a more comprehensive understanding of which target features rapidly activate the ability of Cas12a to generate signal. Besides the protospacer, defined as the TS segment complementary to spacer sequences of the guide RNA, target features responsible for Cas12a activation are poorly understood (18). Much focus has centered on the PAM for detection of dsDNA targets. The canonical TTTV PAM (Figure 1A) is most effectively recognized by LbCas12a (2,19,20), as this AT-rich sequence forms a narrowed minor groove aligning with WED, REC1, and P1 domains to trigger an allosteric transition to the catalytically active state (20). Noncanonical PAMs can activate Cas12a (19–22), but they stimulate less efficient cis-cleavage because structural alterations in both enzyme and target duplex reduce binding stability (20). Hence Cas12a applications commonly target protospacers downstream of canonical PAMs, though RNPs utilizing suboptimal PAMs increase sensitivity for detection of nucleic acid sequences generated by reverse transcription-RPA by reducing amplification interference (21,22). Despite success in characterizing protospacer sequences for Cas-based gene editing (23,24) and general principles of optimal guide design (13,25) relatively little is known about other possible determinants of enzyme activation.

Figure 1. — Wide variability of *trans*-cleavage activities induced by DNA targets. (A) Schematic of LbCas12a RNP bound to dsDNA target. Base-specific interactions provided by side chains of residues in LbCas12a are designated by red arrows (20). PAMs are denoted by the four NTS bases at positions −4* to −1* (5′ to 3′) immediately 5′ to the first protospacer nucleotide (position 1), highlighted in red. For single-stranded targeting, the PAM is denoted by positions −4 to −1 (3′ to 5′) of the TS. (B) Measuring cleavage of C₁₀*trans*-substrates (S) by RNP pre-reacted with unlabeled target (RNP·T) under multiple-turnover conditions (excess of *trans*-substrate over RNP), using FB-C₁₀ tagged at opposite ends with fluorophore (F) and biotin (B) capture probe, enabling removal of uncleaved substrate (Scheme RNP·T + FB-S; (27)), or FQ-C₁₀ where capture tag is replaced with fluorescence quencher (Q) (Scheme RNP·T + FQ-S). (C) *Trans*-substrate cleavage by RNPs targeting protospacer sequences embedded within DNA fragments 858–1378-bp in length. Symbols represent k_app calculated as *trans*-cleavage rates from 1.0 h endpoint data (Supplementary Figure S1a–f) using 1.0 nM RNP, low picomolar concentrations of target, and 100 nM *trans*-substrate at 37°C (Supplementary Table S1). Symbols are color-coded according to PAM sequence at NTS positions −3* and −2* flanking the protospacers recognized. Average coefficient of variation (SE/|value|) over all data was 11%. Significance thresholds, indicated by thick horizontal lines, represents the mean + 3SE of values measured for Cas12a in the absence of crRNA for each target. (D) Representation of how both the rates of RNP activation (k_act) and steady-state *trans*-cleavage (k_ss) can impact measurement of apparent substrate cleavage (k_app). Progress curves (thick solid lines) for three mock reactions (a–c) are displayed on the left, with heat maps of rates on the right. Steady-state components are represented by dashed lines, whose slopes reflect k_ss, which, when extrapolated, intersect the abscissa at the characteristic activation time (1/k_act). In this example the time at which product is measured (circles) results in identical k_app values for reactions b and c. As compared to reference reaction a, the reduction in k_app is solely a result of reduced k_act and k_ss for reactions b and c, respectively.

Wide variability in trans-cleavage activities for different Cas12a RNPs is commonly observed (26–28), suggesting that the field has yet to uncover the rules of target design that govern optimal performance. As some have assumed that target-induced Cas enzyme activation is not rate-limiting (29,30), much attention has been given to understanding how catalytic efficiency impacts enzyme performance (26). However, the extent to which differences in assay performance arise from variation in enzyme activation remains untested. We performed quantitative kinetic analysis of the steps leading to activation of the Cas12a nuclease and observed that acquisition of trans-cleavage capacity is coupled to two activating steps, target binding and an allosteric transition. We show that variability in trans-activity by RNPs arises from both complex determinants of activation and steady-state cleavage of trans-substrate. These results provide a more complete understanding of the role played by Cas12a in bacterial immunity as well as the target features responsible for eliciting the Cas12a assay signal, offering insight into how to optimize the time to result.

Materials and methods

Reagents

crRNA and RNA targets were purchased from Synthego. Single-stranded oligonucleotides and HPLC-purified oligonucleotides duplexes were purchased from Integrated DNA Technologies (IDT; Coralville, IA). Double-stranded (gBlocks Gene Fragments) and single-stranded (Megamer® ssDNA Fragment) DNA fragments were purchased from IDT and GenScript (Piscataway, NJ). Nucleic acids were quantified using UV spectrophotometry based on extinction coefficients provided by the manufacturer or, for dsDNA, calculated elsewhere (http://molbiotools.com/dnacalculator.html). Sequences of short and long synthetic nucleic acids are given in Supplementary Tables S8 and S9. Sources of plasmids are given in Supplementary Table S10. Plasmid DNA was quantified using Qubit™ 1X dsDNA High Sensitivity Kit (cat. no. Q33230) purchased from Thermo Fisher Scientific (Waltham, MA). Recombinant Lba Cas12a (cat. no. M0653T), Nb.BtsI (cat. no. R0707), and 6X Purple Gel Loading Dye (cat. no. B7024S) were purchased from New England BioLabs (Ipswich, MA). Dynabeads™ MyOne™ Streptavidin C1 (cat. no. 65002) were purchased from ThermoFisher.

RNP assembly

200 nM Cas12a was incubated with 100 nM crRNA in Cas12 assay buffer (10 mM Tris–Cl pH 7.5, 10 mM MgCl₂, 1.0 mM Tris(2-carboxyethyl)phosphine hydrochloride, 0.01% Igepal CA-630, 40 μg/ml BSA) for at least 30 min at room temperature and diluted immediately before reaction with substrates. All subsequent cleavage reactions were conducted in Cas12 assay buffer. Herein, [RNP] refers to the nominal concentration of Cas12–crRNA complexes, which is dictated by [crRNA], the limiting reagent in RNP assembly, whereas ‘activated enzyme’ refers to the target-activated RNP.

Trans-substrate cleavage reactions

To measure trans-cleavage using Scheme RNP·T + FB-S (Figure 1B), RNP and targets were pre-reacted at 37°C for 30 min prior to reaction with dual fluorophore-biotin capture tag-labeled trans-substrate FB-C₁₀ in a final volume of 20 μl at 37°C. Time = 0 represents the time at which target-activated RNP was mixed with trans-substrate. For endpoint measurements, at the desired times, unreacted trans-substrate was removed by bead capture using 100 μl of a capture solution (1.0 M NaCl, 10 mM Tris–Cl 7.5, 10 mM EDTA, 0.01% Tween-20) containing 10 μg of streptavidin-conjugated magnetic beads, and products remaining in supernatants were quantified (Figure 3A–D and Supplementary Figure S1a–e, g, i). For Michaelis–Menten analysis, reactants were temperature equilibrated prior to reaction, and signals were corrected for background recorded in cleavage reactions containing RNP without target (Figure 3I–L and Supplementary Figure S4a–c). To measure trans-cleavage using Scheme RNP·T + FQ-S (Figure 1B), RNP and targets were pre-reacted at 37°C for 30 min prior to reaction with dual fluorophore-quencher tag-labeled trans-substrate FQ-C₁₀ in a final volume of 20 μl at 37°C. Time = 0 represents the time at which target-activated RNP was mixed with trans-substrate. For endpoint measurements, at the desired time, 5.0 ul of a stop solution (20 mM Tris 7.5–Cl, 100 mM EDTA, 0.01% Tween-20) was added, and fluorescence was recorded (Figure 7A and Supplementary Figure S1f, h). For Michaelis–Menten analysis, reactants were temperature equilibrated for several minutes prior to mixing, whereupon reactions were continuously monitored, and signals were corrected for background recorded in cleavage reactions containing RNP without target (Supplementary Figure S4d–q). Other reactions assembled via Scheme RNP·T + FQ-S were monitored continuously after assembly (Figure 3G, H and Supplementary Figure S3). To measure trans-substrate cleavage using Scheme RNP + T,FQ-S (Figure 4A), RNP, targets and trans-substrate FQ-C₁₀ were mixed at room temperature prior to incubation at 37°C after which reactions were monitored continuously (Figures 5B, 7B, Supplementary Figure S5e–h, and Supplementary Figure S9).

Figure 3. — DNA and RNA target-activated *trans*-substrate cleavage by crRNA-Cas12a RNP. (A–D) Cleavage of *trans*-substrate FB-C₁₀ using Scheme RNP·T + FB-S and titrated ds- and ssDNA (A, C) or RNA (B, D) targets at 37°C. Cleaved product was determined after 1.0 min using 0.25 nM RNP and 200 nM FB-C₁₀ for DNA targets (A, C). For slower RNA-activated cleavage, product was determined after 2.0 h using 200 nM FB-C₁₀ and 1.0 nM RNP-A2 (B) or after 1.0 h (ssRNA) or 10 min (ssRNA-t) using 100 nM FB-C₁₀ and 0.5 nM RNP-A19 (D). Points represent mean (±SD) of background-corrected replicates (n = 3). Lines show hyperbolic fit with Eq. (2) to obtain k_app (Supplementary Table S2). (E, F) Cleavage of *trans*-substrate FB-C₁₀ by RNPs activated with their cognate ds- and ssDNA and RNA targets from (A–D), where bars represent k_app (±SD) (Supplementary Table S2). (G, H) Cleavage of 100 nM *trans*-substrate FQ-C₁₀ using Scheme RNP·T + FQ-S by 1.0 nM RNPs activated with their cognate ds- and ssDNA-t and RNA-t targets at 37°C. Points represent mean (±SD) of replicates (n = 3) collected after 2.0 h (from Supplementary Figure S3b–g). Inset shows analysis of signal at low target concentrations, where solid line shows fit with linear Eq. (1), the slope of which provides estimates for k_app when normalized to time. LOD and k_app values are summarized in Supplementary Figure S3h. (I–L) Determination of Michaelis–Menten constants for *tran*s-substrate FB-C₁₀ cleavage at 37°C using Scheme RNP·T + FB-S for 1.0 nM RNP-A2 activated by 10 pM DNA target (I, J) or by 1.0 nM RNP-A19 activated by 0.5 nM RNA target (K, L). Representative time courses for the indicated RNP-target combinations (I, K), where lines represent linear fit of individual background-corrected values to obtain V₀ for each concentration of *trans*-substrate (nM). Substrate-dependence of activated enzyme-normalized V₀ (J, L), where points represent V₀/E₀ (±SE), and lines and shaded 95% confidence interval show hyperbolic fit to obtain k_cat/K_M and k_cat (Table 1). Steady-state analysis for all other RNP, target and *trans*-substrate combinations is shown in Supplementary Figure S4.

Figure 7. — Determinants of crRNA-Cas12a RNP activation by ssDNA targets. (A) *Trans*-activities of RNPs in response to long ssDNA targets measured using Scheme RNP·T + FQ-S and 100 nM *trans*-substrate FQ-C₁₀ at 37°C. (Top) Bars represent mean (±SD) of product released after 1.0 h in replicates (n = 3) using 10 pM ssDNA targets ss-E(308) (green) and ss-E(494) (blue) by 1.0 nM RNPs, whose PAM sequences and intended specificity are indicated. (Bottom) Schematic of positions of protospacers (red) in ssDNA corresponding to 308-nt ss-E(308) (green, bottom strand) and 425-nt ss-E(425) (blue, top strand) from the 858-bp Target-E (grey, double strands). Note the lack of ssDNA target-induced activation of RNP-E27 and -E23, which target overlapping protospacers in target ss-E(308), suggesting potential secondary structure interference. (B) Cleavage of 100 nM *trans*-substrate FQ-C₁₀ at 37°C by 1.0 nM RNPs activated with 5.0 pM of their respective ssDNA targets or long dsDNA Target-E, where bars represent k_ss (±SE) (Supplementary Figure S9 and Supplementary Table S6A). RNP-E26 and -27 utilize dipurine PAMs and exhibit little activity when activated by dsDNA targets, whereas RNP-E1 and -E18 utilize dipyrimidine PAMs. (C) Activation time courses for 1.0 nM RNPs induced by 10 pM of their respective short ssDNA targets, employing Scheme RNP + T,FQ-S with 100 nM *trans*-substrate FQ-C₁₀ at 37°C. Points represent mean (±SD) of background-corrected replicates (n = 3). Solid lines show fit to activation Eq. (7) to obtain activation parameters (Supplementary Table S6B) plotted in heat maps. RNP-E18 was activated at a rate too fast to measure and was assigned a lower limit activation rate of 1.0 min⁻¹. Note the lack of requirement for purines in positions −2 and −3 of the TS PAM for ssDNA target-induced activation. (D) Effect of non-specific DNA on activation of RNP-A2 and -A19 as described in (C), yielding values (Supplementary Table S7) plotted in heat maps. 1.0 nM RNPs were reacted with 50 pM 40-bp ssDNA targets (blue symbols), 50 pM 3.87-kb non-specific plasmid DNA (white) or 50 pM of both (red), and 100 nM *trans*-substrate FQ-C₁₀ at 37°C.

Figure 4. — Two-step, target-induced activation of the crRNA-Cas12a RNP *trans*-nuclease. (A) Measuring the time course of target-induced activation of the *trans*-nuclease by Scheme RNP + T,FQ-S under conditions for single-turnover of target and multi-turnover of *trans*-substrate. (B–D) Activation of RNP-A2 *trans*-nuclease by short dsDNA targets, revealing lag and steady-state phases in cleavage of *trans*-substrate FQ-C₁₀. 1.0 nM RNP, 10–40 pM target, and 100 nM *trans*-substrate were reacted by Scheme RNP + T,FQ-S (B) or RNP·T + FQ-S (C) at 25°C. Points represent means (±SD) from background-corrected replicates (n = 3). Lines show fit to activation equation Eq. (7) to obtain rate constants (±SE) for activation of the nuclease k_act and steady-state rate of *trans*-substrate turnover k_ss (B) or fit to linear Eq. (5) (C); bars (D) summarize constants (Supplementary Table S3). Note the lag in the development of *trans*-nuclease activity (B) is eliminated by pre-reacting RNP with target (C). (E–H) Effect of RNP concentration on activation rates using Scheme RNP + T,FQ-S for RNP-A2 reacted with 10 pM short ds- (E, F) or ssDNA (G, H) targets and 100 nM *trans*-substrate FQ-C₁₀ at 25°C. Points in time course (E, G) represent means (±SD) from background-corrected replicates (n = 3) of a representative experiment; lines show fit to activation equation Eq. (7) to obtain k_act. Global fitting from multiple experiments, indicated by lines and shaded 95% confidence intervals, was performed with hyperbolic Eq. (8) over the full RNP concentration range (F, H) to obtain RNP·target dissociation constant (K_d) and net rates of the activation step (k_{act (max)}) from asymptotes, and from ratios of the two, apparent second-order RNP activation rates (k_{on (app)}) representing products of intrinsic RNP–target binding and forward isomerization rates, a lower limit for RNP–target binding. Global analysis over low RNP concentrations (insets) was performed with linear Eq. (9) to yield independent estimates for k_{on (app)} and k_{off (net)} from slopes and y-intercepts, respectively (Table 2A). (I–L) Effect of RNP concentration on activation rates using Scheme RNP + T,FQ-S for RNP-A19 reacted with 100 pM short dsDNA (I, J) or 10 pM ssDNA (K, L) targets and 100 nM *trans*-substrate FQ-C₁₀, carried out as in E–H. Global fitting from multiple experiments, indicated by lines and shaded 95% confidence intervals, was performed with linear Eq. (9) over the full RNP concentration range (J) to obtain k_{on (app)} and k_{off (net)} from slopes and y-intercepts, respectively (Table 2B), with hyperbolic Eq. (8) over the full RNP concentration range (K, L) to obtain K_d and k_{act (max)}), or with linear Eq. (9) over low RNP concentrations (L, inset) to yield independent estimates for k_{on (app)} and k_{off (net)} (Table 2B).

Figure 5. — Low rates of target-induced activation account for poor performance of crRNA-Cas12a RNPs. (A) Activation time courses for RNP-A2 and -A19 induced by their protospacers embedded in a long DNA target (1378-bp Target-A), employing Scheme RNP + T,FQ-S or RNP·T + FQ-S with 1.0 nM RNP, 10 pM target and 100 nM *trans*-substrate FQ-C₁₀ at 37°C. Points represent mean (±SD) of background-corrected replicates (n = 3). Solid lines show fit to activation Eq. (7) to obtain k_act and k_ss for Scheme RNP + T,FQ-S and fit to linear Eq. (5) to obtain k_ss for RNP·T + FQ-S (Supplementary Table S4B) plotted in heat maps along with k_app (Supplementary Table S1). Inset shows the same data plotted to longer times. (B) Cleavage of 100 nM *trans*-substrate FQ-C₁₀ using Scheme RNP + T,FQ-S at 37°C by 1.0 nM RNP-A2 and A19 activated with their cognate short (40-bp) and long (1378-bp Target-A) DNA targets. Points represent mean (±SD) of replicates (n = 3) recorded after 2.0 h (from Supplementary Figure S5e–h). LOD (±SE) was calculated at each time point (insets). (C) Figure of merit (FOM) calculated as LOD × time, from RNPs activated by short DNA targets. Points represent FOM (±SE) calculated at time points (B), indicating rapidly activating RNP-A2 achieves its optimal sensitivity within minutes of reaction, whereas more slowly activating RNP-A19 takes considerably longer to achieve its optimal sensitivity.

Target-cleavage reactions

For cleavage of plasmid DNA targets (Figure 2A, B and Supplementary Figure S2a,b), 5.0 nM RNP and 2.0 nM plasmid were equilibrated at the desired temperature, then mixed to initiate reaction in a final volume of 130 μl. Time = 0 represents the time at which RNP was mixed with target. At desired time points, 20 μl aliquots were removed, and reactions were terminated by mixing with 4.0 μl of 6× Gel Loading Dye to achieve 0.08% SDS and 10 mM EDTA (final). 10 μl was applied to 1.5% agarose/TAE gels containing 7.5 μg/ml ethidium bromide. After electrophoresis for 2.0 h at 75 V, gels were imaged by fluorescence with UV light illumination on a Gel Doc EZ Gel Documentation System (Bio-Rad). Images were processed using Image Lab™ Software. For illustrative purposes, image displays were inverted. Background-subtracted intensities of the linearized bands were quantified using volume tools. Nicked products are not detected in these Cas12a digestion reactions, presumably due to our use of crRNA possessing spacer lengths of 20 nt, which has been shown to promote faster conversion of transient, nicked forms to linearized forms compared to crRNA possessing longer spacer lengths (31).

For cleavage of short ds- and ssDNA targets (Figure 2d,e and Supplementary Figure S2d) using Scheme RNP + FB-T (Figure 2C), RNP and dual fluorophore-biotin capture tag-labeled target were equilibrated to 25°C then mixed to initiate reaction in a final volume of 100 μl, and at desired time points, 10 μl aliquots were removed mixed with 20 μl of stop solution (90% formamide, 20 mM EDTA). Time = 0 represents the time at which RNP was mixed with target. Unreacted targets were removed by bead capture as described above. Signals were corrected for background recorded in target reactions containing enzyme lacking crRNA. For cleavage of RNA target (Figure 2F), RNP and dual fluorophore-biotin-labeled target were incubated for 2.0 h at 37°C prior to removal of unreacted substrate by bead capture.

Trans-nuclease activation reactions

To record the time course of trans-nuclease activation (Figures 4B, E–L, 5A, 6, 7C, D, Supplementary Figure S5a–d, Supplementary Figure S6, and Supplementary Figure S7) via Scheme RNP + T,FQ-S (Figure 4A), temperature-equilibrated RNP and solutions containing target and trans-substrate FQ-C₁₀ were mixed to initiate reaction. Time = 0 represents the time at which RNP was mixed with target and trans-substrate. For the corresponding control experiments to measure trans-cleavage by pre-activated RNP (Figure 4C, 5A, 6A, B, D, Supplementary Figure S5a,b, and Supplementary Figure S7b,c) via Scheme RNP·T + FQ-S (Figure 1B), RNP and targets were pre-reacted at 37°C for 30 min prior to reaction with trans-substrate. Time = 0 represents the time at which target-activated RNP was mixed with trans-substrate. In both schemes, reaction products were monitored continuously at the desired temperature. Where indicated, signals were corrected for background recorded in cleavage reactions containing RNP without target. For testing the effect of DNA topology (Figure 6F), 1.0 μg of plasmid DNA was digested with 10 U Nb.BtsI in NEBuffer2 for 2.0 h at 37°C, then enzyme was heat-inactivated for 20 min at 80°C. Control plasmid treated similarly, except that nuclease was omitted, was used as a source of supercoiled plasmid.

Figure 6. — Determinants of crRNA-Cas12a RNP activation by dsDNA targets. (A) Activation time courses for RNPs induced by their respective protospacers embedded within a fixed target background sequence recognized by RNP-F3 (blue bars), employing Scheme RNP + T,FQ-S and RNP·T + FQ-S with 1.0 nM RNP, 10 pM targets and 100 nM *trans*-substrate FQ-C₁₀ at 25°C. Points represent mean (±SD) of background-corrected replicates (n = 3). Solid lines show fit to activation Eq. (7) to obtain k_act for Scheme RNP + T,FQ-S or to linear Eq. (5) to obtain k_ss for RNP·T + FQ-S (Supplementary Table S5A) plotted in heat maps. (B) Effect of single-base substitutions at each of the four PAM positions on target-induced activation using 1.0 nM RNP, 10 pM targets and 100 nM *trans*-substrate FQ-C₁₀ at 25°C, as described in (A), yielding values (Supplementary Table S5B) plotted in heat maps. (C) Effect of target sequence adjoining the PAM-protospacer on target-induced RNP activation. Chimeric dsDNA targets were constructed from sequences of target rapidly activating RNP-F3 (blue bars) or slowly activating RNP-F25 (red), including native PAMs utilized by RNP-F25 (TTTT) or -F3 (TTTC). Points represent mean (±SD) of background-corrected replicates (n = 3) using 1.0 nM RNP, 10 pM targets and 100 nM *trans*-substrate FQ-C₁₀ at 25°C. Solid lines show fit to activation Eq. (7), yielding values (Supplementary Table S5C) plotted in heat map. See Supplementary Figure S7a for targets containing TTTT PAMs. (D) Effect of target context on RNP activation as described in (A), using 1.0 nM RNP, 20 pM targets and 100 nM *trans*-substrate FQ-C₁₀ at 25°C, yielding values (Supplementary Table S5D) plotted in heat maps, using protospacers embedded with 55-bp duplexes (short) or 1299-bp Target-F (long). See Supplementary Figure S7b for other RNPs. (E) Effect of non-specific DNA on activation of RNP-A2 as described in (C), yielding values (Supplementary Table S5E) plotted in heat maps. 1.0 nM RNPs were reacted with 50 pM 40-bp dsDNA targets (blue symbols), 50 pM 3.87-kb non-specific plasmid DNA (white) or 50 pM of both (red), and 100 nM *trans*-substrate FQ-C₁₀ at 37°C. See Supplementary Figure S7c,d for RNP-A1. (F) Activation time courses for RNPs targeting different protospacers embedded within 3970-bp plasmid DNA encoding Target-A as described in (C), yielding values (Supplementary Table S5F) plotted in heat maps. 1.0 nM RNPs were reacted with 50 pM target, consisting of supercoiled plasmid (S) or relaxed circular plasmid DNA (R) generated by nicking endonuclease digestion, and 100 nM *trans*-substrate FQ-C₁₀ at 37°C.

Fluorescence quantification of reaction products

Alexa488 content of samples was determined on a BioTeK Synergy H1 fluorescence microplate reader using λ_ex 490 nm and λ_em 525 nm. The concentration of product was calculated by converting fluorescence values (RFU) to molarity by interpolation to linear standard curves. Calibration standard concentrations were chosen to span the reactant and product concentrations particular to each experiment. Uncleaved target and trans-substrate FB-C₁₀ were used as standards for reactions employing substrate-capture. Cleaved trans-substrate FQ-C₁₀ standard was prepared by digesting 1.0 μM of FQ-C₁₀ with 1.0 nM of target-activated RNP for 2.0 h at 37°C with cleavage reaction monitoring in real time to confirm reactions had progressed to completion.

Data analysis

Statistical analysis

Data are plotted as mean ± standard deviation (SD) of measurements made on n replicates indicated. Values obtained from curve-fitting, performed using GraphPad Prism (v.8), are expressed as value ± standard error (SE). Errors were propagated through calculations using standard approaches (32). Comparison of k_app for cleavage of trans-substrates FB-C₁₀ using Scheme RNP·T + FB-S and FQ-C₁₀ using Scheme RNP·T + FQ-S by 63 RNPs (Supplementary Figure S1j) was made by calculating the mean absolute errors in measurements, calculated as the sum of absolute errors divided by the sample size.

Trans-substrate cleavage

[Target]-dependence of product formation using a fixed [RNP] (Figure 3G, H(insets) and Supplementary Figure S1a–e,i) was fitted with the 2-parameter linear equation:

(1)

where [Target] represents the total concentration of target nucleic acid, the slope k represents the amount of trans-substrate cleaved per target, and Y₀ is an offset. Since [RNP] was in vast excess of [Target], and stoichiometric analysis (see below) indicates [RNP] are in vast excess of the net K_d for the RNP-target complex, here [Target] represents the concentration of activated RNP, and k_app, the apparent turnover rate, was calculated by dividing k by the assay duration. [Target]-dependence of product formation over stoichiometric ranges of [Target] using fixed [RNP] (Figure 3A–D) was fitted with the 3-parameter hyperbolic equation:

(2)

where parameters are as defined above for Eq. (1), K_1/2 is the midpoint (approximating 1/K_net, the net dissociation constant of the RNP–target complex), and the difference in Y_max and Y₀ represents the net target-normalized product formed, which, when divided by assay duration, represents k_app.

Target cleavage

For cleavage of plasmid targets (Supplementary Figure S2a,b), time (t)-dependence of cleaved product formation (background-corrected band intensity) was fitted with the 3-parameter single-exponential equation:

(3)

where k_obs is the rate constant for target cleavage, Y_f is the final product, and Y₀ is an offset.

For cleavage of ds- and ssDNA targets (Figure 2D, E and Supplementary Figure S2d), time (t)-dependence of product formation from data collected in independent experiments was globally fitted with Eq. (3) utilizing shared parameters. For cleavage of a fixed [RNA target] (Figure 2F), [RNP]-dependence of product formation was fitted with the 3-parameter hyperbolic equation:

(4)

where Y_max is the maximal RNP-normalized product formed at the endpoint, K_1/2 is the midpoint (approximating the net dissociation constant of the RNP–target complex), and Y₀ is an offset; k_obs was calculated by dividing Y_max by the assay duration.

Steady-state analysis of trans-substrate cleavage

Time-dependence of product formation was fitted with the 2-parameter linear equation:

(5)

where the slope v is the reaction velocity and Y₀ is an offset. For Michaelis–Menten analysis (Figure 3I, K and Supplementary Figure S4), v represents rate the initial velocity (V₀). V₀ was normalized to the concentration of activated enzyme E₀ approximated by the [Target], which is justified given the high net affinity of RNP for target as revealed by stoichiometric analysis. The [Substrate] ([S])-dependence of V₀/E₀ (Figure 3J, L and Supplementary Figure S4) was fitted with Johnson's 2-parameter adaptation of the Michaelis–Menten equation (33):

(6)

to directly obtain turnover number (k_cat) and the specificity constant (k_SP) equal to catalytic efficiency (k_cat/K_M), which is treated as a single parameter. The Michaelis–Menten constant (K_M) was calculated by dividing k_cat/K_M by k_cat. This approach provides the same kinetic constants as those obtained by application of the conventional Michaelis–Menten equation but provides more accurate values for catalytic efficiency by avoiding over-estimation of errors when it is calculated from k_cat and K_M (33). Results from all Michaelis–Menten analyses were congruent with consistency criteria (30).

In other experiments (Supplementary Figure S9), steady-state phases of trans-substrate cleavage reactions were identified and fitted with the 2-parameter linear equation (Eq. 5) to obtain reaction rates, which when normalized to the [Target] approximation of activated enzyme, yielded k_ss.

Trans-nuclease activation reactions

Time-dependence of product formation in experiments utilizing Scheme RNP + T,FQ-S (Figure 4B, E, G, I, K, 5A, 6, 7C, D, Supplementary Figure S5a–d, Supplementary Figure S6, and Supplementary Figure S7) was fitted with a 3-parameter activation equation, described previously as a process leading to a steady-state (34):

(7)

where k is a an apparent steady-state rate of trans-substrate cleavage, k_act is an apparent rate for RNP activation, manifested as a lag in the time course, and Y₀ is an offset; k_ss was calculated by dividing k by the [Target] approximation of activated RNP. This apparent steady-state rate is directly related to k_cat, K_M and substrate concentration via the Michaelis–Menten relationship:

For Scheme RNP·T + FQ-S (Figures 4C, 5A, 6A, B, D, Supplementary Figure S5a, b, and Supplementary Figure S7b, c), where no lag was apparent, data were fitted with Eq. (5), and k_ss was similarly calculated.

For processes displaying saturation kinetics (Figure 4F, H, L, Supplementary Figure S5c, d, and S6), [RNP]-dependence of k_act over wide ranges of [RNP] from multiple independent experiments was globally fitted with the 2-parameter hyperbolic equation:

(8)

with shared parameters from independent experiments, where k_{act (max)} represents the maximal activation rate and K_d is the intrinsic dissociation rate constant for RNP binding to target equal to 1/K, where K is defined in Mechanisms 14 and 16. The bimolecular rate of RNP activation (k_{on (app)}) was calculated from k_{act (max)}/K_d. [RNP]-dependence of k_act over low [RNP] (Figure 4F, H, L(inset), Supplementary Figure S5c, d (insets), and Supplementary Figure S6 (insets)) from independent experiments was likewise globally fitted with shared parameters using the linear equation:

(9)

where k_{on (app)} is the bimolecular rate of RNP activation and k_{off (net)} is the net rate of inactivation. For the slower process showing no signs of saturation kinetics (Figure 4J), [RNP]-dependence of k_act over wide ranges of [RNP] was globally fitted with Eq. (9).

Limits of detection

Analytical limits of detection (LOD) and 95% confidence intervals (Figure 5B and Supplementary Figure S3h) were calculated with four-parameter logistic fit (35) of raw fluorescence values recorded at time intervals over 2.0 h (Supplementary Figure S5e–h and Supplementary Figure S3b–g). Figure of merit (FOM) (Figure 5C) was calculated by multiplying LOD by time (36).

Simulation of trans-nuclease activation reactions

Time-dependence of product formation was simulated (Figure 8b) by joining expressions for enzyme activation (Eq. 7) and steady-state trans-cleavage (Eq. 6):

(10)

The model uses steady-state rates based on parameters of k_cat = 1.0 s⁻¹, k_SP = 10⁸ M⁻¹s⁻¹ and K_M = 10 nM, representing those obtained at higher temperatures, and assumes 100 nM trans-substrate and that [Target] represents the concentration of activated RNP. For each [Target], k_act was varied and the time required to achieve 1% cleavage of trans-substrate (assumed to represent that required for detection) was determined. Steady-state parameters were varied to reflect hypothetical changes brought about by pooling of crRNA (resulting in a 10-fold increase in k_cat), the presence of competitive inhibitors of the trans-cleavage reaction (resulting in a 10-fold increase in K_M), and reduced temperature (resulting in 10-fold decreases in both k_cat and K_M) (Table 1).

Table 1.

Steady-state kinetic constants for C₁₀trans-cleavage by target-activated crRNA-Cas12a RNP

Trans-substrate	Temperature	RNP	Target	k _cat /K_M (10⁸ M⁻¹s⁻¹)	k _cat (s⁻¹)	K _M (nM)	Figure
FB-C₁₀	37°C	F4	dsDNA	0.97 (±0.05)	2.77 (±0.05)	29 (±2)	S4a ^a
		F2	dsDNA	1.43 (±0.19)	3.72 (±0.17)	26 (±4)	S4b ^a
		A2	dsDNA	0.46 (±0.03)	2.81 (±0.09)	61 (±5)	3I, J
			ssDNA	0.87 (±0.08)	2.90 (±0.10)	33 (±3)	3J, S4c
		A19	RNA-t	0.021 (±0.001)	0.61 (±0.01)	285 (±4)	3K, L
FQ-C₁₀	37°C	F4	dsDNA	2.51 (±0.15)	6.27 (±0.16)	25 (±2)	S4d
		F2	dsDNA	3.9 (±0.5)	6.7 (±0.3)	17 (±2)	S4e
		A2	dsDNA	1.05 (±0.09)	3.03 (±0.11)	29 (±3)	S4f
			ssDNA	1.9 (±0.3)	2.43 (±0.14)	13 (±2)	S4g
		A19	dsDNA	4.1 (±0.2)	1.64 (±0.02)	4.0 (±0.2)	S4h
			ssDNA	5.5 (±1.4)	0.91 (±0.04)	1.7 (±0.4)	S4i
		E27	ssDNA	1.58 (±0.10)	8.3 (±0.3)	53 (±4)	S4j
	25°C	F4	dsDNA	1.2 (±0.2)	0.65 (±0.03)	5.3 (±1.0)	S4k
		F2	dsDNA	2.3 (±0.4)	0.80 (±0.04)	3.5 (±0.7)	S4l
		A2	dsDNA	0.64 (±0.05)	0.48 (±0.01)	7.4 (±0.6)	S4m
			ssDNA	1.8 (±0.3)	0.71 (±0.03)	3.9 (±0.6)	S4n
		A19	dsDNA	1.9 (±0.3)	0.59 (±0.03)	3.2 (±0.5)	S4o
			ssDNA	3.7 (±1.6)	0.53 (±0.05)	1.4 (±0.6)	S4p
		A1	dsDNA	2.5 (±0.4)	0.68 (±0.03)	2.7 (±0.4)	S4q

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Values (±SE) obtained from Michaelis–Menten steady-state analysis of 1.0 nM RNPs at the indicated temperatures using short synthetic targets (10 pM DNA or 500 pM RNA) in Scheme RNP·T + FB-S with trans-substrate FB-C₁₀ or Scheme RNP·T + FQ-S with FQ-C₁₀. Initial velocities of linear portions of time courses (5–10 min) determined by fitting with linear Eq. (5) were normalized to the target concentration approximation of activated RNP, plotted against substrate concentrations, and analyzed by hyperbolic Eq. (6) to yield k_cat/K_M and k_cat. K_M calculated as the ratio of k_cat/K_M to k_cat.

^aReanalysis of data from reference (27) using Eqs. (5) and (6).

Results

Canonical PAMs are neither required nor predictive of high trans-nuclease activity

We surveyed the activity of 120 LbCas12a RNPs designed against protospacers in 858–1378-bp DNA targets corresponding to cDNA encoding portions (Targets-A–D) of SARS-CoV-2 (37), E6–E7 oncogenes (Target-E) from human papilloma virus (HPV) (38), and IS2404 (Target-F) of Mycobacterium ulcerans, the causative agent of Buruli ulcer disease (39). RNPs are designated by protospacers against which they are designed: e.g. RNP-A1 recognizes protospacer-1 of Target-A. We used two methods to quantify cleavage of trans-substrates (Figure 1B). In the first, cleavage of trans-substrate FB-S was measured using a magnetic bead pull-down strategy, which can be used to measure cis-cleavage (see below) (27). In the second, cleavage of trans-substrate FQ-S was measured using a fluorescence dequenching strategy that allows for real-time monitoring of enzyme activation (see below). We measured product formation (Supplementary Figure S1a–f), calculated apparent turnover rates (k_app;Supplementary Table S1), and plotted values with color-coding indicating PAM utilization to highlight central dinucleotides of non-canonical PAMs (Figure 1C). Select RNPs that displayed high- and low- target-activated trans-cleavage activity exhibited no cross-reactivity against unrelated targets (Supplementary Figure S1g, h) except for one (RNP-A1). Good qualitative agreement was observed between trans-cleavage of FB-C₁₀ and FQ-C₁₀ by 63 RNPs specific to Targets-E and -F (Supplementary Figure S1i, j).

Most RNPs utilizing canonical TTTV and non-canonical dipyrimidine PAMs displayed higher activity than those utilizing dipurines PAMs, paralleling thymidine preference with cytosine acceptance in central PAM positions. Maximal turnover rates (0.6–2.6 s⁻¹) were achieved with dithymidine PAM-utilizing RNPs, though there was no general rule that RNPs utilizing canonical PAMs displayed the highest activities. Thus, canonical PAM utilization is neither required nor predictive of a high-activity RNP, consistent with numerous studies showing that a canonical PAM is not absolutely required for cleavage of target or trans-substrate interference (2,19–22) and supporting the proposal that NYYN PAM sequences can be effectively used for in vitro applications relying on LbCas12a (19). Besides expanding the repertoire of protospacers amenable for Cas-mediated detection, these results suggest there are structural determinants of RNP activity besides the PAM. Across all targets, RNPs displayed trans-activities that varied by as much as two orders of magnitude, illustrating the complex influence of target sequence on trans-cleavage.

Mechanistic framework to investigate target-activated trans-nuclease activity

To better understand the determinants underlying variability in RNP activity, we begin with a simplified kinetic Mechanism 11 representing apparent trans-substrate (S) cleavage by RNP activated by target (T) depicted byorrect to capitalize:

(11)

where RNP*·T represents target-bound RNP having acquired nuclease activity, i.e. competence for cleaving trans-substrate, and k_app represents an apparent rate of trans-substrate cleavage. This process encompasses both enzyme activation (defined as the conversion of RNP from an inactive to active nuclease) and trans-substrate cleavage, as depicted by an expanded Mechanism 12:

(12)

Here, k_act represents an apparent rate of enzyme activation by target and k_ss represents the apparent steady-state rate constant for trans-substrate cleavage governed by the Michaelis–Menten relationship. Depending on their magnitudes, either may contribute to the overall rate of product formation (Figure 1D). We investigated the molecular determinants of enzyme activation and trans-substrate cleavage in the framework of Mechanism 12, beginning with cis-target binding and cleavage and progressing to activation of the trans-nuclease, expanding upon the mechanism as needed, utilizing illustrative examples of RNPs from the collection screened above (Figure 1C).

Rates of plasmid cis-cleavage parallel trans-activity

Because trans-substrate cleavage is temporally coupled to cis-cleavage of dsDNA targets, we investigated target cleavage, which may be depicted by Mechanism 13:

(13)

where k_obs represents an observed rate of target cleavage (a complex function of the rates of target binding, enzyme activation, and chemical cleavage), T_p represents the PAM-proximal fragment remaining bound to enzyme after cis-cleavage, and T_d represents the released PAM-distal fragment. We tested whether trans-cleavage activities for representative RNPs track with cis-cleavage of supercoiled plasmids (Figure 2A, B). High-activity Target-F RNPs (F3, F5 and F6) cleaved plasmid rapidly and completely within 30 min, whereas lower-activity RNPs (F16 and F25) effected only partial cleavage. Very low-activity dipurine PAM-utilizing RNP (F40) failed to cleave DNA. Similar results were seen for Target-A RNPs, indicating high-activity RNPs cleave targets more rapidly than lower-activity ones. Given target cleavage by RNPs utilizing non-canonical PAMs (A2, F5 and F6) can be on par with that of canonical PAM-utilizing RNPs (F3 and A4), canonical PAMs are not required for rapid target cleavage, as observed elsewhere (22).

Cis-cleavage properties of RNPs activated by DNA and RNA targets

We used short, synthetic targets to further investigate cis-cleavage of targets, focusing on high-activity RNP-A2, which utilizes non-canonical PAM CTTC, comparing results to previous findings on RNPs utilizing canonical PAMs (27). Using Scheme RNP + FB-T (Figure 2C), we measured cleavage of ds- and ssDNA targets by RNP-A2 under single-turnover conditions with dual-labeled targets (FB-T). As before (27), since cleavage was too rapid at 37°C for accurate determination, time courses were recorded at 25°C, yielding a dsDNA TS cleavage rate k_obs (0.69 ± 0.10 min^-1; Figure 2D, G) comparable to that for canonical PAM-utilizing RNP-F4 (0.72 min^–1; (27)). Cleavage of ssDNA TS was indistinguishable (0.72 ± 0.10 s^–1; Figure 2E, G), and similar rates were obtained when reporter and capture probe positions were swapped (Figure 2G).

We then tested whether Cas12a performs cis-cleavage of RNA targets using Scheme RNP + FB-T. Initial experiments indicated little RNA target cleavage at 25°C, so the reaction temperature was raised to 37°C and target was titrated with RNP-A2 (Figure 2F). After 2.0 h, <10% of target was cleaved, revealing a cleavage rate 860-fold slower than for DNA (Figure 2G). The near stoichiometric dependence of target cleavage on RNP concentration suggests that, despite the slow cleavage of RNA targets by Cas12a, this RNP binds target RNA with relatively high affinity. Though RNA targets have been shown to activate Cas12g1 (7,9) and Cas12a2 (8,10), and despite the similarities of DNA–RNA and RNA–RNA helices in solution, no reports have described targeting of RNA by Cas12a RNP. RNA–target activation of trans-cleavage by Cas12a has gone unreported (10–12)—possibly due to the use of shorter (5,11,13), less-efficiently cleaved reporters (27). DNA target-activated LbCas12a cleaves RNA in trans (19,27,40), but this catalytic efficiency is 100-fold lower than that for deoxyribosyl trans-substrates (27), paralleling the diminished rate of RNA target we measure here. Thus, RNA protospacers can activate cis-cleavage by Cas12a RNP, and like the diminished turnover of RNA trans-substrates, the target ribosyl backbone decreases the rate of target cleavage by the activated enzyme, providing further evidence the Cas12a RuvC nuclease is a preferred deoxyribonuclease (5,27,40). We explore trans-substrate cleavage activated by RNA targets below.

Trans-cleavage properties of RNPs activated by DNA and RNA targets

We used short synthetic targets to investigate of trans-cleavage by RNP-A2, comparing its properties to those of low-activity RNP-A19, which utilizes non-canonical PAM ACCA. After confirming target specificity of the RNPs (Supplementary Figure S3a), we performed a quantitative comparison, interpreting target-activated trans-substrate in the framework of Mechanism 14:

(14)

where K_net represents the net affinity of RNP for target, a term comprising K, the intrinsic affinity of RNP for target, and rates corresponding to post-binding events including conformational changes in the RNP-target complex associated with enzyme activation.

For RNP-A2, trans-activity increased stoichiometrically with titrated target and approached an asymptote (Figure 3A, B), indicating high-affinity binding to both DNA and RNA targets. Owing to the high affinity of the interaction, accurate values for K_net could not be determined, so approximations (K_1/2) were calculated based on the concentration of target activating half-maximal trans-substrate cleavage (Supplementary Table S2), indicating sub-nanomolar apparent dissociation constants. These affinities are considerably larger than the intrinsic affinities of RNP for target (see below). Additionally, this confirms that under conditions of nanomolar RNP in vast excess of target, formation of activated enzyme is achieved at levels dictated by target concentrations, indicating that target concentration may serve as a surrogate for the concentration of activated RNP. Both ds- and ssDNA targets induced similar trans-activities by this RNP (1.4–2.5 s⁻¹; Figure 3E), higher than observed for protospacer embedded within a long dsDNA target (0.68 ± 0.03 s⁻¹; Supplementary Table S1). Further, removal of four nucleotides at the 5′ PAM-distal end to mimic end products of target cleavage (2) had no effect on ssDNA target potency. RNA target-induced trans-activities were considerably lower (Figure 3E), but 5′-truncated RNA targets activated higher rates of trans-cleavage than longer counterparts. Similar analysis performed using RNP-A19 (Figure 3C, D, F) revealed despite exhibiting low trans-activity in response to long dsDNA target (0.0126 ± 0.0001 s⁻¹; Supplementary Table S1), it exhibits 160-fold greater activity in response to short ds- or ssDNA targets (2.0–2.2 s⁻¹) and higher RNA target-induced activity than RNP-A2. Again 5′-truncated RNA targets effected considerably higher rates of trans-cleavage than longer ones. Together these results demonstrate protospacer context and target ribose backbone contribute to activating RNP trans-cleavage. Further, activating RNP with RNA targets possessing bases beyond presumptive cleavage sites diminishes trans-activity, suggesting unprocessed target interferes with trans-cleavage.

We measured DNA and RNA target-detection sensitivity using trans-activity by RNP-A2 and -A19 (Figure 3G, H). As for FB-C₁₀trans-cleavage (27), LODs for synthetic DNA targets were well below picomolar concentrations (Supplementary Figure S3h), 16 and 18 fM (RNP-A2) and 161 and 116 fM (RNP-A19) for ds- and ssDNA targets. Consistent with stoichiometric analysis, LOD for target RNA-t by RNP-A19 (880 fM) was considerably lower than that for RNP-A2 (12 pM) owing to 18-fold higher rate of trans-cleavage (Supplementary Figure S3h). These results confirm that low femtomolar LODs for ds- and ssDNA targets are achievable, consistent with previous findings (27), and indicate that even RNA targets may be detected by LbCas12a at sub-picomolar levels using trans-substrate cleavage reporters.

dsDNA targets containing non-canonical PAMs and ssDNA can trigger formation of high-activity trans-nucleases

We then directed our attention to the steady-state trans-substrate cleavage properties of the target-activated enzyme. Estimation of steady-state constants for trans-substrate cleavage by Michaelis–Menten analysis relies on pre-formation of RNP-target complexes prior to reaction with substrate, whereupon trans-cleavage commences, as depicted by Mechanism 15:

(15)

We performed Michaelis–Menten analysis to compare kinetic properties of DNA and RNA target-activated nucleases (Figure 3I–L and Table 1). First, using Scheme RNP·T + FB-S, we measured FB-C₁₀trans-cleavage by RNP-A2. Consistent with apparent turnover values (Figure 3E, F), and observed elsewhere (26,30), ds- and ssDNA-activated RNP exhibited similar turnover (2.8–2.9 s⁻¹) and catalytic efficiency (4.6–8.7 × 10⁷ M⁻¹s⁻¹), higher than reported in some but not other studies (41) but comparable to those for canonical PAM utilizing RNP-F4 and -F2 (Table 1). For RNA target-activated RNP-A19, >20-fold lower catalytic efficiency was measured, confirming the negative influence of the target ribose backbone on trans-cleavage.

Using Scheme RNP·T + FQ-S, we then measured trans-substrate FQ-C₁₀ cleavage by various target-activated RNPs (Table 1) at both 37°C and 25°C for comparison with FB-C₁₀ and DNA target cleavage rates. For dsDNA target-activated RNPs, though catalytic efficiency was somewhat higher for FQ-C₁₀ than for FB-C₁₀ at 37°C, the difference was small. Consistency between these values indicates that the higher catalytic efficiencies we previously measured (relative to others in the literature) were not due to use of the substrate-capture approach (27). As observed for FB-C₁₀, catalytic constants for ssDNA target-activated RNP-A2 and -A19 were indistinguishable from those for their respective dsDNA target-activated RNPs. Though reduction of temperature to 25°C had little effect on catalytic efficiency, turnover and K_M decreased for all RNP tested, indicating the impact on assay performance near room temperatures.

These results further confirm that for RNPs utilizing non-canonical PAMs, including those composed of dicytosines, ds- and ssDNA targets can induce formation of equally high-activity trans-nucleases, indicating their catalytically active enzymes possess similar conformations and active sites of comparable chemical architectures. RNPs activated by short or long DNA targets, as exemplified by RNP-A19, may exhibit widely disparate levels of trans-activity, indicating targets possess complex determinants for RNP activation.

Defining activation and steady-state phases of the trans-nuclease

Having characterized steady-state properties of Cas12a trans-nuclease activated by ds- and ssDNA targets, we used Scheme RNP + T,FQ-S (Figure 4A) to investigate activation kinetics. Reaction of the high-activity RNP-A2 resulted in a cleavage time course composed of an exponential lag phase followed by a linear phase (Figure 4B). The lag phase, composed of a characteristic time on the order of minutes and independent of target concentrations over the target range tested, disappeared with prolonged pre-reaction of RNP-A2 and prior to incubation with trans-substrate via Scheme RNP·T + FQ-S (Figure 4C, D). Linear phase slopes were directly proportional to target concentration and, when normalized to target concentrations serving as proxies for activated RNP, yielded steady-state rates (k_ss) indistinguishable from each other and those of RNPs pre-reacted with target (Figure 4D). This steady-state rate (0.6 s⁻¹; Supplementary Table S3) agrees closely with that (0.5 s⁻¹) predicted by Michaelis–Menten analysis (Table 1). Together, these results indicate the lag phase represents conversion of RNP to an active nuclease initiated by target binding, which precedes steady-state trans-cleavage, as depicted by Mechanism 12. The timescale of activation (on the order of minutes) is considerably longer than the timescale of trans-substrate turnover (on the order of seconds), indicating RNP activation by target, not trans-substrate cleavage, can be rate-limiting and that that trans-product cleavage may be used to characterize rates of target-induced activation.

Defining two steps in target-induced enzyme activation

For the high-activity RNP-A2, holding dsDNA target concentration constant and varying RNP concentration resulted in a hyperbolic increase in activation rate that approached a maximum, k_{act (max)} (Figure 4E, F). This observation suggests that formation of activated nuclease, the first step in Mechanism 12, proceeds via two steps (42), as depicted in Mechanism 16:

(16)

Here, step 1 represents rapidly equilibrating, readily reversible RNP-target binding mediated by PAM contacts with a dissociation constant K_d equal to 1/K, defined previously for Mechanism 14. As such, the intrinsic affinity of RNP for target is considerably smaller than the net affinities (K_net) owing to the post-binding steps that extend lifetimes of RNP-target complexes. Step 2 represents a slower first-order conversion, described as an allosteric transition (17), of enzyme to an activated state, comprising R-loop formation, conformational changes in the enzyme that activate the nuclease site, cis-cleavage, and release of the PAM-distal fragment. Owing to the essentially ‘single-turnover’ nature of Cas12a activation by dsDNA targets, we depict the allosteric step in Mechanism 16 as largely irreversible over the time frame of measurements. Fitting the RNP concentration dependence of k_act with Eq. (8) enables estimation of the rate of the allosteric transition (k_{act (max)}) from the asymptotic value and K_d, and the ratio of the two represents a second-order RNP forward activation rate (k_{on (app)}). At low RNP concentrations (i.e. [RNP] << K_d), k_act increases linearly, and curve fitting with linear Eq. (9), provides estimation of k_{on (app)} from the slope and the net rate of enzyme inactivation (k_{off (app)}) from the y-intercept.

For RNP-A2, affinity of dsDNA target binding was in the low nanomolar range and its activation plateaued at 0.60 ± 0.09 min⁻¹ (Table 2A), comparable to dsDNA TS cis-cleavage (0.69 ± 0.10 min⁻¹; Figure 2G), consistent with our previous determination that activation of trans-nucleolytic potential is directly coupled to slow cleavage of dsDNA target by Cas12a (27). Based on these values, the apparent RNP activation rate (k_{on (app)}) was estimated at 0.30 ± 0.11 × 10⁷M⁻¹s⁻¹, and a similar value (0.17 ± 0.07 × 10⁷M⁻¹s⁻¹) was obtained from analysis at low RNP (see inset to Figure 4F). These values represent a lower limit for RNP-target binding and is comparable to those for LbCas12a (1.3 × 10⁷ M⁻¹s⁻¹; (43)) and AsCas12a (∼10⁸ M⁻¹s⁻¹; (44)). However, as these values fall well below the diffusion limit (∼10⁹–10¹⁰ M⁻¹s⁻¹; (45)), they suggest that post-binding steps diminish productivity of the RNP-target encounter by two-orders of magnitude. The inactivation rate (k_{off (net)}) from extrapolated y-intercepts could not be determined with high precision: visual inspection reveals that it is low, considerably smaller than k_{act (max)}, which is consistent with single-target turnover behavior of Cas12a (5,43).

Table 2.

Activation parameters of crRNA-Cas12a RNP reacted with DNA targets

A. Activation rates that approach an asymptotic maximum^a
	Full RNP range^b				Limiting RNP range^c
RNP	DNA target	k _{act (max)} (min⁻¹)	K _d (nM)	k _{on (app)} (10⁷ M⁻¹s⁻¹)	k _{on (app)} (10⁷ M⁻¹s⁻¹)	k _{off (net)} (min⁻¹)	n	Figure
A2	ds	0.60 (±0.09)	3.4 (±1.1)	0.30 (±0.11)	0.17 (±0.07)	0.01 (±0.02)	5	4E, F
	ss	1.1 (±0.2)	1.1 (±0.4)	1.6 (±0.7)	1.6 (±1.0)	−0.04 (±0.15)	5	4G, H
	ss-t	1.1 (±0.2)	1.4 (±0.7)	1.3 (±0.7)	0.8 (±0.5)	0.01 (±0.07)	3	S5c
A19	ss	1.1 (±0.2)	0.36 (±0.26)	5 (±4)	2.6 (±2.0)	−0.03 (±0.29)	5	4K, L
	ss-t	0.9 (±0.2)	0.30 (±0.25)	5 (±4)	2.5 (±1.7)	−0.05 (±0.24)	4	S5d
F2	ds	0.87 (±0.11)	0.27 (±0.13)	5.3 (±2.6)	3.1 (±0.5)	−0.01 (±0.03)	4	S6a
F3	ds	0.74 (±0.08)	0.47 (±0.16)	2.7 (±1.0)	1.55 (±0.18)	−0.01 (±0.02)	3	S6b
F4	ds	0.81 (±0.14)	5.0 (±1.8)	0.27 (±0.11)	0.19 (±0.02)	−0.010 (±0.007)	3	S6c
F23	ds	0.83 (±0.08)	0.7 (±0.2)	1.9 (±0.6)	1.06 (±0.16)	0.01 (±0.04)	3	S6d
F25	ds	0.079 (±0.008)	2.8 (±0.7)	0.047 (±0.012)	0.041 (±0.006)	−0.006 (±0.002)	3	S6e
B. Activation rates that do not approach an asymptotic maximum^d
RNP	DNA target	k _{on (app)} (10⁷ M⁻¹s⁻¹)			k _{off (net)} (min⁻¹)		n	Figure
A19	ds	0.0051 (±0.0001)			0.043 (±0.001)		2	4I, J

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^a Values (±SE) obtained from fit of time courses using Scheme RNP + T,FQ-S with activation Eq. (7). Varying RNP were reacted with 10–20 pM short targets and 100 nM trans-substrate FQ-C₁₀ at 25°C. Representative plots provided in the indicated figures.

^b Maximation activation rate (k_{act (max)}) and dissociation rate constant for RNP binding to target (K_d) obtained from global fit of k_act over the full range of RNP concentrations from n pooled experiments with hyperbolic Eq. (8). Bimolecular rate of RNP activation (k_{on (app)}) was calculated from k_{act (max)}/K_d.

^c Bimolecular rate of RNP activation (k_{on (app)}) obtained from the slope from global fit of k_act at limiting RNP concentrations with linear Eq. (9). Net rate of inactivation (k_{off (net)}) estimated from y-intercept.

^d Values (±SE) obtained from fit of time courses using Scheme RNP + T,FQ-S with activation Eq. (7). Varying RNP-A19 were reacted with 100 pM short target and 100 nM trans-substrate FQ-C₁₀ at 25°C. Representative plot provided in the indicated figure. Bimolecular rate of RNP activation (k_{on (app)}) obtained from the slope from global fit of k_act over the full range of RNP concentrations from n pooled experiments with linear Eq. (9). Net rate of inactivation (k_{off (net)}) estimated from y-intercept.

Rapid activation of trans-activity arises from high-affinity RNP-target binding and rapid conversion to activated enzyme

Activation of RNP-A2 with ssDNA targets resulted in a biphasic time courses whose activation rates rose hyperbolically with RNP concentration (Figure 4G, H), suggesting that RNP activation by ssDNA targets also proceeds via two-step Mechanism 16. Using the analysis described above, we observed ssDNA target induces conversion of RNP-A2 to activated enzyme at a rate (1.1 ± 0.2 min⁻¹) greater than that observed for dsDNA target (0.60 ± 0.09 min⁻¹) (Table 2A) and slightly higher than ssDNA target cleavage itself (0.72 ± 0.10 min⁻¹; Figure 2G). Removal of PAM-distal nucleotides at 5′ ends of ssDNA targets had no effect on the activation rate. The affinity of binding to ssDNA targets was somewhat higher (K_d 1.1 ± 0.4 and 1.4 ± 0.7 nM) than that of dsDNA targets (3.4 ± 1.1 nM), and, together with higher allosteric transition rates, hastened activation (k_{on (app)} 1.3 ± 0.7 and 1.6 ± 0.7 × 10⁷ M⁻¹s⁻¹) above that for dsDNA target (0.30 ± 0.11 × 10⁷ M⁻¹s⁻¹). Since trans-cleavage appears slightly faster than ssDNA TS cis-cleavage, for at least some ssDNA targets, trans-cleavage might commence prior to TS cleavage, which would be consistent with previous observation that RNPs activated by non-cleavable ssDNA targets retain high trans-activity (17). This might explain why, for RNA targets, which are cleaved 860-fold more slowly than DNA counterparts, the removal of four nucleotides from 5′ ends to mimic cleaved TS fragments greatly enhances trans-cleavage, suggesting intact portions of longer targets interfere with trans-substrate accessibility. For dsDNA targets, trans-activity appears at the same rates as TS cleavage, suggesting both events are tightly coupled and that full trans-activity is unleashed only upon target cleavage and, by implication, dissociation of the PAM-distal segment of the dsDNA target (17).

Likewise, activation time courses of low-activity RNP-A19 were biphasic (Supplementary Figure S5a,b). However, unlike high-activity RNP-A2, activation of RNP-A19 by dsDNA targets was inefficient, necessitating use of higher target and RNP concentrations for further characterization. For RNP-A19, activation rates increased linearly, remaining low without reaching saturation (Figure 4I, J), indicative of relatively low-affinity binding (K_d > 20 nM). Though estimates for k_{act (max)} could not be determined, the apparent rate for activating RNP-A19 by its dsDNA target was very low (k_{on (app)} 5.1 ± 0.1 × 10⁴ M⁻¹s⁻¹), 60-fold smaller than that of RNP-A2 for its dsDNA target. Estimates for rates of enzyme inactivation could be determined (k_off 0.043 ± 0.001 min⁻¹; Table 2B), suggesting even these low-activity RNP–target complexes are relatively long-lived, with lifetimes of 23 minutes. As expected from rapid, efficient activation by ssDNA targets (Supplementary Figure S5b), RNP-A19 showed high affinity for ssDNA targets (K_d 0.30 ± 0.25 and 0.36 ± 0.26 nM) and high allosteric transition rates (k_{act (max)} 0.9 ± 0.2 and 1.1 ± 0.2 min^–1), both contributing to its high activation rates (k_{on (app)} 5 ± 4 × 10⁷ M⁻¹s⁻¹ for both)(Figure 4K, L and Table 2A). Together, these results demonstrate activation kinetics, specifically reduction in the forward rate of enzyme activation, arising from reduced affinity of RNP-A19 for ds-, but not ssDNA targets, accounts for the low dsDNA-induced trans-cleavage activity observed for this RNP (Figure 1C).

We compared time courses for activation of RNP-A2 and -A19 by long DNA targets (Figure 5A and Supplementary Table S4B). As expected, high-activity RNP-A2 displayed rapid and efficient activation by its protospacer embedded within the 1379-bp target, revealing a k_ss value similar to that measured using the same target and FB-S (Supplementary Table S1). By contrast, activation of low-activity RNP-A19 with the long DNA target was slow and inefficient, as was the case for its short DNA target, but even prolonged target pre-activation failed to generate high-activity RNP. Thus, though both high-activity RNP-A2 and low-activity RNP-A19 exhibit capacity for formation of high catalytic efficiency trans-nucleases (Table 1), rates at which these RNPs are activated by their targets differ considerably (Table 2), confirming at least some variation in RNP performance may be explained by timescales of enzyme activation.

Activation kinetics impact assay performance

We tested how differences in activation kinetics, as exemplified by RNP-A2 and -A19, impact assay performance, specifically sensitivity for detection of short and long DNA targets under Scheme RNP + T,FQ-S (Figure 5B), which involves assembly steps resembling those likely employed in a diagnostic assay. For both RNPs, LODs for both targets evolved over 2 hr or more (see insets to Figure 5B), and as expected, RNP-A2 displayed high sensitivity detection for both targets, with LODs of 76 ± 9 and 160 ± 20 fM, respectively, at 2.0 h, whereas RNP-A19 displayed a considerably higher sensitivity for the short target (130 ± 9 fM) compared to the long target (7500 ± 70 fM). These results illustrate how poor kinetics of enzyme activation (Figure 5a) can profoundly impair RNP sensitivity for target detection (Figure 5b).

To better understand the time-dependence of assay performance, we applied the figure of merit (FOM), a recently developed metric for assessing performance of CRISPR sensing systems, which is calculated as LOD x time (36). Underpinning this metric is the assumption that the speed of product formation is dictated by trans-substrate turnover, with the implied assumption that enzyme activation is fast and not rate-limiting; hence FOM should indirectly depend on the catalytic constants for trans-substrate cleavage (k_cat and K_M). For its short DNA target, high-activity RNP-A2 approached optimal performance within minutes of encounter with target and trans-substrate, achieving a FOM of ∼10 pM min that remained nearly constant for 2 h or more (Figure 5C). In contrast, performance of low-activity RNP-A19 developed only slowly, maturing with a rate constant of approximately 30 min, which we interpret corresponds to the rate of target-induced activation under these conditions. Together these results indicate that rapidly activating RNPs displaying high affinity for DNA targets (such as A2) achieve high sensitivity for target detection quickly after assay assembly, but slower-activating RNPs with lower target affinity (such as A19) may require tens of minutes for maturation of full performance. Consequently, the FOM metric (as derived) incompletely describes enzyme performance, as it overlooks the timescale of enzyme activation, which, as we show here, may be extensive, even for RNPs of relatively high catalytic potential.

Determinants of enzyme activation by ds- and ssDNA targets

The results presented so far indicate two parameters define assay performance of a given Cas12a RNP, as illustrated by Mechanism 12: first, activation time, which depends upon RNP-target affinity and a first-order transition that ultimately limits activation, as depicted by Mechanism 16; and second, steady-state trans-cleavage rate, depicted by Mechanism 15, which depends upon Michaelis-Menten rate constants for activated enzyme. Previous assumptions that activation is fast and the trans-cleavage step is rate-limiting (29,30) justified focus on steady-state properties of activated RNPs, measured using short synthetic dsDNA targets, as the sole determinant of assay sensitivity (26). Having developed methodologies for measuring rates of target-induced RNP activation, and highlighting its previously unappreciated importance in assay performance, we embarked on a survey to uncover determinants that underly enzyme activation.

Protospacer

We determined the RNP concentration-dependence of five additional RNPs activated by protospacers within short, 55-bp dsDNA targets (Table 2), including four (F3, F47, F41 and F23) utilizing canonical PAMs and one (F25) utilizing non-canonical PAM TTTT. As before, activation rates rose hyperbolically with RNP concentration and net inactivation rates were very small in comparison to maximal activation rates, which for all RNPs except F25 were comparable to RNP-A2 activation by short dsDNA. That these rates fall somewhat below those for ssDNA targets (1.1 min⁻¹) suggests the latter represents an upper limit for the enzyme at 25°C. As for RNP-A2 (see above), dsDNA k_{act (max)} for RNP-F4 (0.81 ± 0.14 min⁻¹) was similar to target cleavage (k_obs 0.72 min¹; see Figure 4C,D in ref. (27)), again confirming that appearance of trans-activity is coupled to dsDNA target cleavage. In contrast, k_{act (max)} for RNP-F25 was at least 9-fold lower than the others, paralleling its slow cleavage of plasmid DNA target and low trans-activity (Figure 2A), though as will be shown, utilization of a non-canonical PAM only partly accounts for its slower activation. These results indicate that even for RNPs utilizing dithymidine PAMs, apparent rates of activation by short dsDNA targets can vary widely, mainly due to differences in allosteric transition rates. All RNPs analyzed in Table 2 are capable of performing high activity trans-cleavage (Table 1 and Supplementary Table S5D), indicating that variation in RNP-target dissociation constants (75-fold, from 0.27 to >20 nM) and allosteric transition rates (11-fold, 0.079 to 0.87 min⁻¹) may together contribute to an even wider range of target-induced activation rates (1000-fold, 0.0051 × 10⁷ to 5.3 × 10⁷ M⁻¹s⁻¹).

To assess protospacer contribution to RNP activation and trans-cleavage, we measured activation of several RNPs by their protospacers embedded within a backbone of the target recognized by canonical PAM-utilizing RNP-F3 (Figure 6A), since this target rapidly activates RNP trans-activity (Table 2). We chose protospacers for RNPs (F20, F25, F27, and F32) weakly activated by long DNA targets, with all but RNP-F25 utilizing canonical PAMs. Though steady-state rates of trans-cleavage activated by chimeric targets were within a factor of three of that for RNP-F3, rates for activating these RNPs were 5- to 20-fold lower (Supplementary Table S5A), indicating protospacer sequences predominantly influence activation rates. Additionally, we observed RNP-F25 is more rapidly activated by its protospacer in this chimeric target (k_act 0.22 ± 0.01 min⁻¹) than in its native context (0.079 ± 0.008 min⁻¹; Table 2), suggesting sequences immediately outside the protospacer also influence RNP activation, an effect we investigate below. Since Cas12a itself makes no direct sequence-specific protospacer contacts except through the spacer, either sequence may indirectly (as secondary structure formation within crRNA (25) or directly influence multiple steps in activation, including R-loop initiation and propagation and target cleavage. Since all R-loop positions contribute to the rate of target binding to AsCas12a, and reversible propagation of R-loop formation must pass through a late transition state for target cleavage (44), differences in transition state height or timing might modulate net rates of activation demonstrated by different protospacers. Although progress has been made in predicting efficient protospacers for DNA targeting in cells (reviewed in ref. (41)), further investigation into the molecular basis for the effect of protospacer sequence on activation of Cas12a will be required.

Sequence of the PAM

We probed the influence of PAM sequence on RNP activation and trans-cleavage by testing individual substitutions of the four nucleotides in the canonical TTTC PAM of RNP-F3 (Figure 6B). Substitution at positions −1*, −2* and −4* reduced the activation rate 9- to 67-fold but had little effect on steady-state rates of target-induced trans-cleavage, resulting in k_ss comparable to the original target-RNP combination. In contrast, substitution at position −3* slowed the activation rate 18-fold and resulted in reduced trans-cleavage activity. Even after prolonged pre-incubation of RNP with this target, steady-state trans-cleavage was reduced 20-fold, suggesting this substitution might affect both rates of enzyme activation and trans-cleavage. Together, these results confirm each of the four PAM bases contributes to the determinants of RNP activation by dsDNA targets as observed elsewhere for cleavage of target and activation of trans-cleavage (19–22). The striking preference of LbCas12a for the canonical sequence TTTV sequence, both for cleavage of targets in vitro and for in vivo gene editing (2), arises from the high stability of the LbCas12a-target complex: changes in PAM sequences decrease RNP affinity due to alterations in shape- and sequence-specific interactions driving enzyme conformational changes (20).

Target sequence outside the PAM-distal end of protospacers

Having observed that RNP-F25 is more rapidly activated by its protospacer in a chimeric target than in its native context, we probed the influence of target sequences immediately outside the protospacer and PAM (Figure 6C). Chimeric dsDNA targets containing the protospacer recognized by RNP-F25 were constructed from flanking sequences of targets for RNP-F3 and -F25. Converting non-canonical TTTT PAM to a canonical TTTC sequence had little or no effect on the rate of RNP-F25 activation. However, for chimeric targets, the presence of PAM-distal sequences outside the protospacer, specifically the first 10-bp, of target-F3 promoted more rapid target-induced activation of RNP-F25 than those containing native sequences of target-F25. Rate enhancement by this F3-sequence were 4- to 5-fold for targets containing canonical PAMs (Figure 6C) and 2- to 4-fold for non-canonical PAMs (Supplementary Figure S7a), indicating determinants of target-induced RNP activation reside within the first 10-bp immediately adjacent to the protospacer PAM-distal end. Thus, the relatively slow activation of the low-activity RNP-F25 by its native target (Table 2) arises from the combined determinants of protospacer and sequences immediately adjacent to its PAM-distal end, not by its utilization of a non-canonical PAM (Figure 6A, C). The importance of determinants adjacent to the PAM-distal end are likely related to previous observations that the stability of base pairs, and subsequent DNA distortions, flanking the PAM-distal region of the R-loop is linked to TS cleavage (46). Since after NTS cleavage, TS cleavage commences only upon formation of a transient but stable clamped state involving DNA sequences downstream of the R-loop that serve to reorient the TS for cleavage (31), formation of the intermediate state may be affected by this 10-bp sequence.

Protospacer context

We explored the importance of protospacer context by testing the effect on RNP activation by non-specific DNA, provided in cis (on the same fragment as protospacers) or in trans (elsewhere), as would likely be encountered in a diagnostic context. First, we compared several RNPs activated by protospacers embedded within 1299-bp DNA fragments or 55-bp duplexes (Figure 6D). Activation rates were slowed when protospacers were embedded within long DNA targets, as were steady-state rates of trans-substrate cleavage, for all but RNP-25. Second, we observed that the addition of non-specific plasmid DNA slowed rates of RNP activation by short dsDNA targets and reduced steady-state trans-cleavage (Figure 6E). These results suggest non-specific DNA inhibits enzyme activity, as observed elsewhere (19), by either of two mechanisms. First, it slows activation rates, possibly by providing sites for non-productive encounters, resulting in an apparent reduced affinity of RNP for target. Though such non-productive, transient RNP encounters with off-target sites, which have been directly visualized (43,47), may be effectively overcome with non-limiting RNP concentrations, the net rate remains limited by the first-order conversion to activated enzyme. Second, non-specific DNA might additionally reduce trans-cleavage by competing for the binding of ssDNA trans-substrates, consistent with previous findings that Cas12a can cleave dsDNA trans-substrates, albeit poorly (27,40).

Target DNA supercoiling

Target context was further tested by examining the effect of plasmid supercoiling on activation for RNPs of varying trans-activities and PAM utilization (Figure 6F). Paralleling the cleavage of plasmid DNA (Figure 2A, B), RNPs exhibiting high trans-activity (A1, A2, A4 and A6) are more rapidly activated by plasmid DNA than RNPs of lower activity (A8 and A19) (Figure 6F). As for activation by long DNA targets (Figure 5A, B), low-activity RNP-A19 was activated by plasmid considerably more slowly and less efficiently than high-activity RNP-A2. Canonical PAM-utilizing RNP-A6 and -A4 were more rapidly activated than all other RNPs, and supercoil relaxation by site-specific nicking had little effect on their activation, decreasing rates by less than 2-fold, but decreasing activation rates for non-canonical PAM-utilizing RNPs by as much as 8-fold. These results indicate target-induced RNP activation rates are influenced by supercoiling of the DNA in which cognate protospacers are embedded, recapitulating the enhancement of LbCas12a target cleavage by negative DNA supercoiling brought on by faster R-loop formation (48). Since upon DNA binding, LbCas12a undergoes a structural rearrangement that is responsive to the degree of helix distortion within the PAM duplex (20), supercoiling may accelerate RNP activation by stabilizing favorable conformations in either enzyme or PAM that increase binding affinity or accelerate DNA unwinding (31,44,48).

Determinants in ssDNA targets

We surveyed trans-activity of Target-E RNPs activated by synthetic 308- and 494-nt ssDNA targets (Figure 7A). Control RNPs targeting protospacers absent from both ssDNA targets failed to be activated by either ssDNA, and seven RNPs were activated by correct, but not incorrect targets. However, two RNPs (E23 and E27) failed to be activated by their expected target. These two RNPs target overlapping protospacers within the long ssDNA (see Supplementary Figure S8a,b): RNP-E23 utilizes a dipyrimidine PAM and is activated by dsDNA, whereas RNP-E27 utilizes a dipurine PAM and is not activated by dsDNA but can be activated by 40-nt ssDNA (Figure 7B). Given the capacity of these two RNPs to form high-activity trans-nucleases, we speculate that formation of secondary structure within or proximal to their protospacers in long ssDNA interferes with activation. In addition, since three RNPs (E24, E25 and E29) activated by ssDNA-E(494) target protospacers flanked by dipyrimidines in positions −2 and −3 of the TS PAM and exhibit trans-cleavage activities comparable to RNPs (E21 and E18) targeting protospacers flanked by TS PAMs containing dipurines (Figure 7A), there is no requirement for dipurines in central positions of the TS PAM. This is as expected if activation by ssDNA targets is PAM-indifferent (5). Since high activation rates and similar steady-state trans-activities are exhibited by dipurine- and dipyrimidine PAM-utilizing RNPs (Figure 7C), these results provide further evidence that dipurines in central positions of the PAM (TS positions −2 and −3) are not required for rapid activation by ssDNA, perhaps indicating PAM-indifferent activation by ssDNA arises from greater target flexibility to effect the allosteric transition. Further, Michaelis-Menten analysis reveals that ssDNA-activated RNP-E27 exhibits a catalytic efficiency on par with other RNPs (Table 1). Finally, the addition of non-specific plasmid DNA slowed rates of RNP activation by short ssDNA targets and reduced steady-state trans-cleavage (Figure 7D), as was observed for activation of RNPs by dsDNA targets (Figure 6E). However, the extent of inhibition was lesser for ssDNA targets, likely due to the higher affinity of the RNPs for these targets (Table 2). Together these results indicate that target context plays a role in determining rapid, PAM-indifferent activation of Cas12a by ssDNA targets.

Discussion

The results presented provide mechanistic insight into Cas12a activation, indicating that in addition to steady-state properties of the mature enzyme, the timescale of enzyme activation is a critical determinant of RNP performance. Our data agree with a model where the first step in RNP activation involves a rapid, reversible interrogation of targets mediated though PAM contacts followed by, for dsDNA targets, a second, slower step involving an allosteric transition accompanying R-loop formation resulting in formation of the nucleolytic site (17). Different RNP-target combinations exhibit varying activation profiles, and ds- and ssDNA targets possess different structural determinants that govern the rapidity of RNP activation. These determinants reside within the protospacer, include bases within the PAM (for ds- but not ssDNA targets), consist of sequences within and outside those complementary to the spacer, and are shaped by target context, including DNA topology, target length, and presence of non-specific DNA (Figure 8A). RNA targets may be cleaved by the crRNA-Cas12a RNP and such targets also elicit trans-cleavage of ssDNA, albeit at low potency, indicating the ribose backbone of target sequences impacts enzyme activation. These findings have implications for both function of Cas12a in bacterial immunity as well as utilization of the enzyme in diagnostic assays.

Implications for bacterial systems

To date the available data suggest that Cas12a provides bacterial defense against mobile genetic elements (MGE) mainly through specific cis-cleavage of invading dsDNA, but not ssDNA targets (49), without any role played by non-specific trans-substrate cleavage (49,50). Among our findings are several that appear paradoxical to this mechanism, namely that ssDNA may activate Cas12a RNPs more rapidly than dsDNA targets do and that both targets activate RNPs to the same high level of catalytic activity for trans-cleavage, which is considerably higher against ssDNA than dsDNA trans-substrates (27). However, other results help resolve this apparent paradox, suggesting deeper insight into the mechanism of action of Cas12a in host immunity. First, we show non-specific DNA slows enzyme activation by ssDNA (or dsDNA) targets, and in bacterial cells the presence of a vast excess of genomic DNA over ssDNA invaders may kinetically focus the enzyme on non-productive encounters with host dsDNA during early stages of infection, allowing ssDNA invaders to evade detection and rapidly undergo conversion into dsDNA replicative forms (49). The low availability of free ssDNA due to masking by DNA-binding proteins (51) would further kinetically disfavor cis-cleavage of transient ssDNA appearing during replication. Second, we show that despite rapid activation of Cas12a by ssDNA targets, secondary structure in ssDNA may interfere with target search, and Cas12a may not possess the ability to target protospacers within regions of secondary structure except for those within double-stranded helices tagged with recognizable PAM-sequences, likely formed when ssDNA invaders are converted to dsDNA replicative forms. Third, we show that non-specific DNA, provided in cis- or trans-, also slows trans-substrate cleavage, suggesting any potential for collateral cleavage unleashed by target engagement might be attenuated by host DNA. Finally, despite the relatively high catalytic efficiency for trans-substrate cleavage, on the order of 10⁸ M⁻¹ s⁻¹, turnover of Cas12a for ssDNA trans-substrates is low (single digits/s) and even lower for RNA and dsDNA trans-substrates (27). This low turnover contrasts with the considerably higher turnover of Cas13a (hundreds/s) for trans-RNA (27,41), suggesting collateral destruction of endogenous substrates by trans–cleavage of activated Cas12a might be slow enough to be effectively offset by resynthesis to minimize the deleterious effects of collateral cleavage (49) perhaps also explaining successful use of Cas12a in gene editing (2). The mechanism of action by Cas12a in bacterial cells contrasts with the protection against MGE afforded by Cas12a2, which induces abortive infection though RNA target-induced trans-cleavage of cytoplasmic dsDNA and small RNAs (8), inducing cellular dormancy though its collateral activity, reminiscent of that mediated by Cas13a (52). Whether the specific crRNA-guided RNA target cleavage activity by Cas12a we report plays a physiological role in bacterial immunity remains to be tested.

Implications for diagnostics

Our findings provide strategic guidance for the design of Cas-based diagnostics, since time to result is a critical parameter of any diagnostic assay, especially those used at the point of care. We found that both activation time and steady-state trans cleavage rates directly impact Cas12a signal generation (Figure 1D), an effect we illustrate by modeling how variations in both parameters influence assay time (Figure 8B). As realistic values can lead to unworkable assay times, developers would benefit from measuring both parameters using the methodologies we describe, consider how either are impacted by assay conditions or determinants contributed by targets and sample composition, or apply selection criteria for RNPs employed in different assay formats (see below). Some findings may be specific to LbCas12a, but general observations and the experimental approaches developed are likely widely applicable to other orthologs.

Regardless of assay format, the highest catalytic activity (high turnover, low K_M) for trans-substrate cleavage by activated RNPs is generally desirable in diagnostic contexts. A modest 10-fold improvement in catalytic efficiency is predicted to profoundly shorten the time to detect targets in the sub-pM range (Figure 8B). Increased catalytic activity may be achieved through protein engineering, as suggested by several studies describing engineered Cas12a variants exhibiting improved editing efficiency (53–55), though to date no detailed examination of effects on catalytic activity by Cas12a engineering has been reported. Alternatively, net catalytic activity can be increased using pools of RNPs formed from multiple, distinct crRNA guides specifying different protospacers on the same target, a strategy successful for detection of dsDNA by Cas12a and for viral RNA by Cas13 (27,28), improving sensitivity by increasing rates of target-activated trans-cleavage (27). For instance, pooling 20 crRNA guides increased the apparent turnover of reporter 22-fold over that of single guides, simultaneously increasing target detection sensitivity 180-fold (27). Because of the scarcity of canonical PAMs, maximizing protospacer coverage across a target might necessitate inclusion of RNPs utilizing non-canonical PAMs.

For assays relying on trans-cleavage at ambient temperatures below 37°C, as might be done in a disposable point-of-care diagnostics aiming to decrease reliance on capital equipment, the effect of temperature reduction on both performance parameters is likely to profoundly impact assay outcome. Decreasing temperature to 25°C decreases rates of enzyme activation (compare k_act at 37°C Supplementary Table S5E versus 25°C Supplementary Table S4A), though it has little effect on catalytic efficiency (Table 1). However, the observed decrease in turnover would reduce rates of trans-substrate cleavage almost 10-fold under conditions likely employed to capitalize on substrate turnover ([S] >> K_M), and such reduction would profoundly delay time to result for targets even in the low-pM range (Figure 8B). Performing assays on targets embedded within long DNA fragments and in matrices composed of complex nucleic acid mixtures, also likely encountered in diagnostic assays, decreases both parameters (Supplementary Table S5E). Assuming at least some of the decrease in catalytic activity due to non-specific DNA is competitive in nature, it is worth considering that a 10-fold increase in apparent K_M for trans-substrate has a modest effect on time to result (Figure 8B).

Direct target detection

For diagnostics using amplification-free detection of targets, RNPs with rapid dsDNA activation profiles are desirable since these RNPs would produce the fastest accumulation of cleavage products above detection thresholds. Though RNPs utilizing consensus PAMs may be able to achieve high activation rates owing to high-affinity target binding (K_d < 1 nM) and rapid R-loop formation, unfavorable determinants, including those within the protospacer and PAM-distal segment, may suppress rates. Hence not all such RNPs may succeed and other RNPs utilizing non-canonical PAMs may be considered, especially when applying the crRNA pooling approach described above. Some deviations from the consensus PAM are tolerable, enabling fast activation owing to encounters with favorable determinants, though others slow activation by reducing target-binding affinity. For all RNPs, activation may be further modulated by other target-imposed constraints, such as protospacer context, competing non-specific DNA, and DNA topology, each of which must be considered when testing impure samples of biological origin.

Coupling detection to exponential target amplification

For assays that indirectly detect targets via exponential target amplification in one-pot formats, RNPs with slowed dsDNA activation profiles may instead be desirable. RNPs utilizing non-optimal PAMs are likely more slowly activated by their targets but allow faster accumulation of amplicons by diminishing the rate of template destruction during the amplification phase of the reaction, with the increase in amplicon production at shorter times offsetting the slower appearance of trans-nuclease activity (22). The high concentration of non-specific dsDNA relative to specific targets early in amplification might also reduce the rate of target search, sparing intended targets ultimately undergoing amplification. At later points in amplification, copies of specific targets outnumber non-specific sites, thus promoting trans-cleavage resulting from amplicon-induced activation of RNPs. Assay sensitivity may be improved by generating short target amplicons, to reduce the negative effect of nonspecific, cis DNA on trans activity. The acceptability of non-canonical PAM sites for this approach allows greater flexibility in assay design, and in principle, crRNA pooling should increase assay sensitivity of this format. The time to result will represent a complex function of the amplification time as well as the time for enzyme activation.

Future assays

Given the rapid activation of RNP trans-activity by ssDNA targets, and a resulting catalytic efficiency that matches or exceeds that induced by dsDNA targets, assay formats incorporating amplification strategies that generate ssDNA from targets may be envisioned. In contrast to both assay formats described above, RNPs utilizing dipurine PAMs, which are slowly activated by dsDNA targets but rapidly activated by ssDNA targets, would avoid destruction of dsDNA targets but promote generation of potent signals upon synthesis of ssDNA products.

Finally, our results suggest potential use for Cas12a as a tool for amplification-free detection of specific RNA. Besides assays in which Cas12a activity is coupled to conversion of RNA targets to cDNA by reverse transcriptase (6), other strategies for direct RNA detection by Cas12a have been devised (11), although sensitivity is not as great as observed here. We demonstrated that sub-picomolar LOD for direct detection of RNA targets can be achieved, sensitivity that outperforms that for Cas12a2 (8,10), and is likely to be improved with future optimization of assay conditions and reagents, as has been the case for Cas12a detection of dsDNA targets. It is worth considering that in the original report of collateral cleavage by LbCas12a, the LOD for amplification-free detection of HPV DNA was ∼10 pM (Figure S10 in (5)), but LODs for dsDNA are often low picomolar (6,28) and as shown before (27) and in the present report, LODs of low fM are achievable. The LOD for direct detection of RNA targets by Cas12a we report here lies at the middle range of values obtained using target-activated trans-cleavage of RNA reporters by Cas13a, which span 10–1000 fM (28,39) to 1–92 pM (26). Other orthologs of Cas12a may perform better for detection of RNA targets than LbCas12a used here, and alternative assay strategies, such as pooling of crRNA targeting multiple protospacers within RNA targets, may provide additional boosts in cleavage of trans-substrate reporters. Despite the potential for high sensitivity, our results suggest LbCas12a may not be well-suited for direct detection of long RNA targets, such as mRNA, without modification owing to RNA secondary structure and suppression of trans-cleavage by target 5′ extensions, though shorter RNA targets, such as microRNAs might be amenable for detection.

Conclusions

We identify performance characteristics of Cas12a targets that should be considered when developing rapid diagnostic assays. We show that the rate of RNP activation can dramatically affect assay time and is a function of both affinity of the RNP for target and the rate of a slower post-binding allosteric transition in the RNP-target complex associated with cis-target cleavage. Among RNPs capable of high activity trans-cleavage, measured affinities vary by as much as 75-fold and rates of allosteric transitions vary at least 11-fold, together contributing to target-induced activation rates that span a 1000-fold range. Together with kinetic constants for trans-substrate cleavage, rates of RNP activation play an important role in determining the sensitivity and timing of target detection. Finally, we identify several structural determinants of enzyme activation and trans-substrate cleavage that contribute to Cas12a activity in vitro and suggest that these determinants may also shape the role played by the enzyme in bacterial immunity.

Supplementary Material

gkae152_Supplemental_File

gkae152_supplemental_file.pdf^{(1.5MB, pdf)}

Acknowledgements

We thank Dr Phillip Leung for technical assistance with simulations and Nidi Patel and Zeba Islam for guide RNA characterization. We also thank Dr Ilya Finkelstein for helpful discussions and critical comments on the manuscript. Some cartoons were created with BioRender.com.

Notes

Present address: Samantha Hedley, Dartmouth College, Hanover, NH 03755 USA.

Present address: Karunya Rajaraman, Northeastern University, Boston, MA 02115, USA.

Contributor Information

Eric A Nalefski, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Remy M Kooistra, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Ishira Parikh, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Samantha Hedley, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Karunya Rajaraman, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Damian Madan, Global Health Labs, Inc, Bellevue, WA 98007, USA.

Data availability

All data are contained within the manuscript and/or supplementary files.

Supplementary data

Supplementary Data are available at NAR Online.

Funding

Funding for open access charge: Global Health Labs, Inc.

Conflict of interest statement. Some methods described herein have patents pending on which E.A.N., R.M.K. and D.M. are inventors.

References

1. Wang J.Y., Pausch P., Doudna J.A.. Structural biology of CRISPR–Cas immunity and genome editing enzymes. Nat. Rev. Micro. 2022; 20:641–656. [DOI] [PubMed] [Google Scholar]
2. Zetsche B., Gootenberg J.S., Abudayyeh O.O., Slaymaker I.M., Makarova K.S., Essletzbichler P., Volz S.E., Joung J., Van Der Oost J., Regev A.et al.. Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell. 2015; 163:759–771. [DOI] [PMC free article] [PubMed] [Google Scholar]
3. Gootenberg J.S., Abudayyeh O.O., Kellner M.J., Joung J., Collins J.J., Zhang F.. Multiplexed and portable nucleic acid detection platform with Cas13, Cas12a, and Csm6. Science. 2018; 360:439–444. [DOI] [PMC free article] [PubMed] [Google Scholar]
4. Li S.Y., Cheng Q.X., Liu J.K., Nie X.Q., Zhao G.P., Wang J.. CRISPR-Cas12a has both cis- and trans-cleavage activities on single-stranded DNA. Cell Res. 2018; 28:491–493. [DOI] [PMC free article] [PubMed] [Google Scholar]
5. Chen J.S., Ma E., Harrington L.B., Tian X., Doudna J.A., Costa M.Da, Tian X., Palefsky J.M., Doudna J.A. CRISPR-Cas12a target binding unleashes single-stranded DNase activity. Science. 2018; 360:436–439. [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Kaminski M.M., Abudayyeh O.O., Gootenberg J.S., Zhang F., Collins J.J.. CRISPR-based diagnostics. Nat. Biomed. Eng. 2021; 5:643–656. [DOI] [PubMed] [Google Scholar]
7. Li Z., Zhang H., Xiao R., Han R., Chang L.. Cryo-EM structure of the RNA-guided ribonuclease Cas12g. Nat. Chem. Biol. 2021; 17:387–393. [DOI] [PMC free article] [PubMed] [Google Scholar]
8. Dmytrenko O., Neumann G.C., Hallmark T., Keiser D.J., Crowley V.M., Vialetto E., Mougiakos I., Wandera K.G., Domgaard H., Weber J.et al.. Cas12a2 elicits abortive infection through RNA-triggered destruction of dsDNA. Nature. 2023; 613:588–594. [DOI] [PMC free article] [PubMed] [Google Scholar]
9. Yan W.X., Hunnewell P., Alfonse L.E., Carte J.M., Keston-Smith E., Sothiselvam S., Garrity A.J., Chong S., Makarova K.S., Koonin E.V.et al.. Functionally diverse type V CRISPR-Cas systems. Science. 2019; 363:88–91. [DOI] [PMC free article] [PubMed] [Google Scholar]
10. Bravo J.P.K., Hallmark T., Naegle B., Beisel C.L., Jackson R.N., Taylor D.W.. RNA targeting unleashes indiscriminate nuclease activity of CRISPR–Cas12a2. Nature. 2023; 613:582–587. [DOI] [PMC free article] [PubMed] [Google Scholar]
11. Rananaware S.R., Vesco E.K., Shoemaker G.M., Anekar S.S., Sandoval L.S.W., Meister K.S., Macaluso N.C., Nguyen L.T., Jain P.K.. Programmable RNA detection with CRISPR-Cas12a. 2023; bioRxiv doi:30 January 2023, preprint: not peer reviewed 10.1101/2023.01.29.525716. [DOI] [PMC free article] [PubMed]
12. Yamano T., Nishimasu H., Zetsche B., Hirano H., Slaymaker I.M., Li Y., Fedorova I., Nakane T., Makarova K.S., Koonin E.V.et al.. Crystal structure of Cpf1 in complex with guide RNA and target DNA. Cell. 2016; 165:949–962. [DOI] [PMC free article] [PubMed] [Google Scholar]
13. Nguyen L.T., Smith B.M., Jain P.K.. Enhancement of trans-cleavage activity of Cas12a with engineered crRNA enables amplified nucleic acid detection. Nat. Commun. 2020; 11:4906. [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Stella S., Alcón P., Montoya G.. Structure of the Cpf1 endonuclease R-loop complex after target DNA cleavage. Nature. 2017; 546:559–563. [DOI] [PubMed] [Google Scholar]
15. Stella S., Mesa P., Thomsen J., Paul B., Alcón P., Jensen S.B., Saligram B., Moses M.E., Hatzakis N.S., Montoya G.. Conformational activation promotes CRISPR-Cas12a catalysis and resetting of the endonuclease activity. Cell. 2018; 175:1856–1871. [DOI] [PubMed] [Google Scholar]
16. Swarts D.C., van der Oost J., Jinek M.. Structural basis for guide RNA processing and seed-dependent DNA targeting by CRISPR-Cas12a. Mol. Cell. 2017; 66:221–233. [DOI] [PMC free article] [PubMed] [Google Scholar]
17. Swarts D.C., Jinek M.. Mechanistic insights into the cis- and trans-acting DNase activities of Cas12a. Mol. Cell. 2019; 73:589–600. [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Kellogg E.H., Gootenberg J., Abudayyeh O.O., Wong A.S.L., Dahlman J.E., Lapinaite A., Myhrvold C.. Cell Systems what are the current bottlenecks in developing and applying CRISPR technologies?. Cell Syst. 2022; 13:589–593. [DOI] [PubMed] [Google Scholar]
19. Fuchs R.T., Curcuru J.L., Mabuchi M., Noireterre A., Weigele P.R., Sun Z., Robb G.B.. Characterization of Cme and Yme thermostable Cas12a orthologs. Commun. Biol. 2022; 5:325. [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Yamano T., Zetsche B., Ishitani R., Zhang F., Nishimasu H., Nureki O.. Structural basis for the canonical and non-canonical PAM recognition by CRISPR-Cpf1. Mol. Cell. 2017; 67:633–645. [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Lin K., Guo J., Guo X., Li Q., Li X., Sun Z., Zhao Z., Weng J., Wu J., Zhang R.et al.. Fast and visual detection of nucleic acids using a one-step RPA-CRISPR detection (ORCD) system unrestricted by the PAM. Anal. Chim. Acta. 2023; 1248:340938. [DOI] [PubMed] [Google Scholar]
22. Lu S., Tong X., Han Y., Zhang K., Zhang Y., Chen Q., Duan J., Lei X., Huang M., Qiu Y.et al.. Fast and sensitive detection of SARS-CoV-2 RNA using suboptimal protospacer adjacent motifs for Cas12a. Nat. Biomed. Eng. 2022; 6:286–297. [DOI] [PubMed] [Google Scholar]
23. Kim D., Kim J., Hur J.K., Been K.W., Yoon S.H., Kim J.S. Genome-wide analysis reveals specificities of Cpf1 endonucleases in human cells. Nat. Biotechnol. 2016; 34:863–868. [DOI] [PubMed] [Google Scholar]
24. Kim H.K., Song M., Lee J., Menon A.V., Jung S., Kang Y.M., Choi J.W., Woo E., Koh H.C., Nam J.W.et al.. In vivo high-throughput profiling of CRISPR-Cpf1 activity. Nat. Methods. 2017; 14:153–159. [DOI] [PubMed] [Google Scholar]
25. Creutzburg S.C.A., Wu W.Y., Mohanraju P., Swartjes T., Alkan F., Gorodkin J., Staals R.H.J., der Oost J.van.. Good guide, bad guide: spacer sequence-dependent cleavage efficiency of Cas12a. Nucleic Acids Res. 2020; 48:3228–3243. [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Huyke D.A., Ramachandran A., Bashkirov V.I., Kotseroglou E.K., Kotseroglou T., Santiago J.G.. Enzyme kinetics and detector sensitivity determine limits of detection of amplification-free CRISPR-Cas12 and CRISPR-Cas13 diagnostics. Anal. Chem. 2022; 94:9826–9834. [DOI] [PubMed] [Google Scholar]
27. Nalefski E.A., Patel N., Leung P.J.Y., Islam Z., Kooistra R.M., Parikh I., Marion E., Knott G.J., Doudna J.A., Le Ny A.-L.M.et al.. Kinetic analysis of Cas12a and Cas13a RNA-guided nucleases for development of improved CRISPR-based diagnostics. iScience. 2021; 24:102996. [DOI] [PMC free article] [PubMed] [Google Scholar]
28. Fozouni P., Son S., Doudna J.A., Fletcher D.A., Díaz de León Derby M., Knott G.J., Gray C.N., D’Ambrosio M.V., Zhao C., Switz N.A.et al.. Amplification-free detection of SARS-CoV-2 with CRISPR-Cas13a and mobile phone microscopy. Cell. 2021; 184:323–333. [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Santiago J.G. Inconsistent treatments of the kinetics of Clustered regularly Interspaced Short Palindromic Repeats (CRISPR) impair assessment of its diagnostic potential. QRB Discov. 2022; 3:e9. [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Ramachandran A., Santiago J.G.. CRISPR enzyme kinetics for Molecular Diagnostics. Anal. Chem. 2021; 93:7456–7464. [DOI] [PubMed] [Google Scholar]
31. Naqvi M.M., Lee L., Montaguth O.E.T., Diffin F.M., Szczelkun M.D.. CRISPR–Cas12a-mediated DNA clamping triggers target-strand cleavage. Nat. Chem. Biol. 2022; 18:1014–1022. [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Taylor J.R. An Introduction to Error Analysis: The Study of Uncertainties in Physical Measurements. 1996; 2nd edn.University Science Book. [Google Scholar]
33. Johnson K.A. New standards for collecting and fitting steady state kinetic data. Beilstein J. Org. Chem. 2019; 15:16–29. [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Easterby J.S. Coupled enzyme assays: a general expression for the transient. Biochim. Biophys. Acta. 1973; 293:552–558. [DOI] [PubMed] [Google Scholar]
35. Holstein C.A., Griffin M., Hong J., Sampson P.D.. Statistical method for determining and comparing limits of detection of bioassays. Anal. Chem. 2015; 87:9795–9801. [DOI] [PubMed] [Google Scholar]
36. Nouri R., Dong M., Politza A.J., Guan W.. The figure of merit for CRISPR-based nucleic acid-sensing system: improvement strategies and performance comparison. ACS Sens. 2022; 7:900–911. [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Wu F., Zhao S., Yu B., Chen Y.M., Wang W., Song Z.G., Hu Y., Tao Z.W., Tian J.H., Pei Y.Y.et al.. A new coronavirus associated with human respiratory disease in China. Nature. 2020; 579:265–269. [DOI] [PMC free article] [PubMed] [Google Scholar]
38. Münger K., Baldwin A., Edwards K.M., Hayakawa H., Nguyen C.L., Owens M., Grace M., Huh K.. Mechanisms of Human papillomavirus-induced oncogenesis. J. Virol. 2004; 78:11451–11460. [DOI] [PMC free article] [PubMed] [Google Scholar]
39. Johnson P.D.R., Stinear T., Small P.L.C., Pluschke G., Merritt R.W., Portaels F., Huygen K., Hayman J.A., Asiedu K.. Buruli Ulcer (M. ulcerans Infection): new insights, new hope for disease control. PLoS Med. 2005; 2:e108. [DOI] [PMC free article] [PubMed] [Google Scholar]
40. Fuchs R.T., Curcuru J., Mabuchi M., Yourik P., Robb G.B.. Cas12a trans-cleavage can be modulated in vitro and is active on ssDNA, dsDNA, and RNA. 2019; bioRxiv doi: 08 April 2019, preprint: not peer reviewed 10.1101/600890. [DOI]
41. Feng W., Zhang H., Le X.C.. Signal amplification by the trans-cleavage activity of CRISPR-Cas systems: kinetics and performance. Anal. Chem. 2023; 95:206–217. [DOI] [PMC free article] [PubMed] [Google Scholar]
42. Johnson K.A. Sigman D.S. Transient-state kinetic analysis of enzyme reaction pathways. The Enzymes. 1992; 20:Academic Press, Inc; 1–61. [Google Scholar]
43. Singh D., Mallon J., Poddar A., Wang Y., Tippana R., Yang O., Bailey S., Ha T.. Real-time observation of DNA target interrogation and product release by the RNA-guided endonuclease CRISPR Cpf1 (Cas12a). Proc. Natl. Acad. Sci. U.S.A. 2018; 115:5444–5449. [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Strohkendl I., Saifuddin F.A., Rybarski J.R., Finkelstein I.J., Russell R.. Kinetic basis for DNA target specificity of CRISPR-Cas12a. Mol. Cell. 2018; 71:816–824. [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Bar-Even A., Milo R., Noor E., Tawfik D.S.. The moderately efficient enzyme: futile encounters and enzyme floppiness. Biochemistry. 2015; 54:4969–4977. [DOI] [PubMed] [Google Scholar]
46. Cofsky J.C., Karandur D., Huang C.J., Witte I.P., Kuriyan J., Doudna J.A.. CRISPR-Cas12a exploits R-loop asymmetry to form double-strand breaks. eLife. 2020; 9:e55143. [DOI] [PMC free article] [PubMed] [Google Scholar]
47. Jeon Y., Choi Y.H., Jang Y., Yu J., Goo J., Lee G., Jeong Y.K., Lee S.H., Kim I.-S., Kim J.-S.et al.. Direct observation of DNA target searching and cleavage by CRISPR-Cas12a. Nat. Commun. 2018; 9:2777. [DOI] [PMC free article] [PubMed] [Google Scholar]
48. van Aelst K., Martínez-Santiago C., Cross S., Szczelkun M.. The effect of DNA topology on observed rates of R-loop formation and DNA strand cleavage by CRISPR Cas12a. Genes (Basel). 2019; 10:169. [DOI] [PMC free article] [PubMed] [Google Scholar]
49. Marino N.D., Pinilla-Redondo R., Bondy-Denomy J.. CRISPR-Cas12a targeting of ssDNA plays no detectable role in immunity. Nucleic Acids Res. 2022; 50:6414–6422. [DOI] [PMC free article] [PubMed] [Google Scholar]
50. Liu S., Rao X., Zhao R., Han W.. The trans DNA cleavage activity of Cas12a provides no detectable immunity against plasmid or phage. Front Genome Ed. 2022; 4:929929. [DOI] [PMC free article] [PubMed] [Google Scholar]
51. Meyer R.R., Laine P.S.. The single-stranded DNA-binding protein of Escherichia coli. Microbiol. Rev. 1990; 54:342–380. [DOI] [PMC free article] [PubMed] [Google Scholar]
52. Varble A., Marraffini L.A.. Three new Cs for CRISPR: collateral, communicate, cooperate. Trends Genet. 2019; 35:446–456. [DOI] [PMC free article] [PubMed] [Google Scholar]
53. Merker L., Schindele P., Huang T.K., Wolter F., Puchta H.. Enhancing in planta gene targeting efficiencies in Arabidopsis using temperature-tolerant CRISPR/LbCas12a. Plant Biotechnol. J. 2020; 18:2382–2384. [DOI] [PMC free article] [PubMed] [Google Scholar]
54. Schindele P., Puchta H.. Engineering CRISPR/LbCas12a for highly efficient, temperature-tolerant plant gene editing. Plant Biotechnol. J. 2020; 18:1118–1120. [DOI] [PMC free article] [PubMed] [Google Scholar]
55. Huang T.K., Armstrong B., Schindele P., Puchta H.. Efficient gene targeting in Nicotiana tabacum using CRISPR/SaCas9 and temperature tolerant LbCas12a. Plant Biotechnol. J. 2021; 19:1314–1324. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

gkae152_Supplemental_File

gkae152_supplemental_file.pdf^{(1.5MB, pdf)}

Data Availability Statement

All data are contained within the manuscript and/or supplementary files.

[B1] 1. Wang J.Y., Pausch P., Doudna J.A.. Structural biology of CRISPR–Cas immunity and genome editing enzymes. Nat. Rev. Micro. 2022; 20:641–656. [DOI] [PubMed] [Google Scholar]

[B2] 2. Zetsche B., Gootenberg J.S., Abudayyeh O.O., Slaymaker I.M., Makarova K.S., Essletzbichler P., Volz S.E., Joung J., Van Der Oost J., Regev A.et al.. Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell. 2015; 163:759–771. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B3] 3. Gootenberg J.S., Abudayyeh O.O., Kellner M.J., Joung J., Collins J.J., Zhang F.. Multiplexed and portable nucleic acid detection platform with Cas13, Cas12a, and Csm6. Science. 2018; 360:439–444. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B4] 4. Li S.Y., Cheng Q.X., Liu J.K., Nie X.Q., Zhao G.P., Wang J.. CRISPR-Cas12a has both cis- and trans-cleavage activities on single-stranded DNA. Cell Res. 2018; 28:491–493. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B5] 5. Chen J.S., Ma E., Harrington L.B., Tian X., Doudna J.A., Costa M.Da, Tian X., Palefsky J.M., Doudna J.A. CRISPR-Cas12a target binding unleashes single-stranded DNase activity. Science. 2018; 360:436–439. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B6] 6. Kaminski M.M., Abudayyeh O.O., Gootenberg J.S., Zhang F., Collins J.J.. CRISPR-based diagnostics. Nat. Biomed. Eng. 2021; 5:643–656. [DOI] [PubMed] [Google Scholar]

[B7] 7. Li Z., Zhang H., Xiao R., Han R., Chang L.. Cryo-EM structure of the RNA-guided ribonuclease Cas12g. Nat. Chem. Biol. 2021; 17:387–393. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B8] 8. Dmytrenko O., Neumann G.C., Hallmark T., Keiser D.J., Crowley V.M., Vialetto E., Mougiakos I., Wandera K.G., Domgaard H., Weber J.et al.. Cas12a2 elicits abortive infection through RNA-triggered destruction of dsDNA. Nature. 2023; 613:588–594. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B9] 9. Yan W.X., Hunnewell P., Alfonse L.E., Carte J.M., Keston-Smith E., Sothiselvam S., Garrity A.J., Chong S., Makarova K.S., Koonin E.V.et al.. Functionally diverse type V CRISPR-Cas systems. Science. 2019; 363:88–91. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B10] 10. Bravo J.P.K., Hallmark T., Naegle B., Beisel C.L., Jackson R.N., Taylor D.W.. RNA targeting unleashes indiscriminate nuclease activity of CRISPR–Cas12a2. Nature. 2023; 613:582–587. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B11] 11. Rananaware S.R., Vesco E.K., Shoemaker G.M., Anekar S.S., Sandoval L.S.W., Meister K.S., Macaluso N.C., Nguyen L.T., Jain P.K.. Programmable RNA detection with CRISPR-Cas12a. 2023; bioRxiv doi:30 January 2023, preprint: not peer reviewed 10.1101/2023.01.29.525716. [DOI] [PMC free article] [PubMed]

[B12] 12. Yamano T., Nishimasu H., Zetsche B., Hirano H., Slaymaker I.M., Li Y., Fedorova I., Nakane T., Makarova K.S., Koonin E.V.et al.. Crystal structure of Cpf1 in complex with guide RNA and target DNA. Cell. 2016; 165:949–962. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B13] 13. Nguyen L.T., Smith B.M., Jain P.K.. Enhancement of trans-cleavage activity of Cas12a with engineered crRNA enables amplified nucleic acid detection. Nat. Commun. 2020; 11:4906. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B14] 14. Stella S., Alcón P., Montoya G.. Structure of the Cpf1 endonuclease R-loop complex after target DNA cleavage. Nature. 2017; 546:559–563. [DOI] [PubMed] [Google Scholar]

[B15] 15. Stella S., Mesa P., Thomsen J., Paul B., Alcón P., Jensen S.B., Saligram B., Moses M.E., Hatzakis N.S., Montoya G.. Conformational activation promotes CRISPR-Cas12a catalysis and resetting of the endonuclease activity. Cell. 2018; 175:1856–1871. [DOI] [PubMed] [Google Scholar]

[B16] 16. Swarts D.C., van der Oost J., Jinek M.. Structural basis for guide RNA processing and seed-dependent DNA targeting by CRISPR-Cas12a. Mol. Cell. 2017; 66:221–233. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B17] 17. Swarts D.C., Jinek M.. Mechanistic insights into the cis- and trans-acting DNase activities of Cas12a. Mol. Cell. 2019; 73:589–600. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B18] 18. Kellogg E.H., Gootenberg J., Abudayyeh O.O., Wong A.S.L., Dahlman J.E., Lapinaite A., Myhrvold C.. Cell Systems what are the current bottlenecks in developing and applying CRISPR technologies?. Cell Syst. 2022; 13:589–593. [DOI] [PubMed] [Google Scholar]

[B19] 19. Fuchs R.T., Curcuru J.L., Mabuchi M., Noireterre A., Weigele P.R., Sun Z., Robb G.B.. Characterization of Cme and Yme thermostable Cas12a orthologs. Commun. Biol. 2022; 5:325. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B20] 20. Yamano T., Zetsche B., Ishitani R., Zhang F., Nishimasu H., Nureki O.. Structural basis for the canonical and non-canonical PAM recognition by CRISPR-Cpf1. Mol. Cell. 2017; 67:633–645. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21. Lin K., Guo J., Guo X., Li Q., Li X., Sun Z., Zhao Z., Weng J., Wu J., Zhang R.et al.. Fast and visual detection of nucleic acids using a one-step RPA-CRISPR detection (ORCD) system unrestricted by the PAM. Anal. Chim. Acta. 2023; 1248:340938. [DOI] [PubMed] [Google Scholar]

[B22] 22. Lu S., Tong X., Han Y., Zhang K., Zhang Y., Chen Q., Duan J., Lei X., Huang M., Qiu Y.et al.. Fast and sensitive detection of SARS-CoV-2 RNA using suboptimal protospacer adjacent motifs for Cas12a. Nat. Biomed. Eng. 2022; 6:286–297. [DOI] [PubMed] [Google Scholar]

[B23] 23. Kim D., Kim J., Hur J.K., Been K.W., Yoon S.H., Kim J.S. Genome-wide analysis reveals specificities of Cpf1 endonucleases in human cells. Nat. Biotechnol. 2016; 34:863–868. [DOI] [PubMed] [Google Scholar]

[B24] 24. Kim H.K., Song M., Lee J., Menon A.V., Jung S., Kang Y.M., Choi J.W., Woo E., Koh H.C., Nam J.W.et al.. In vivo high-throughput profiling of CRISPR-Cpf1 activity. Nat. Methods. 2017; 14:153–159. [DOI] [PubMed] [Google Scholar]

[B25] 25. Creutzburg S.C.A., Wu W.Y., Mohanraju P., Swartjes T., Alkan F., Gorodkin J., Staals R.H.J., der Oost J.van.. Good guide, bad guide: spacer sequence-dependent cleavage efficiency of Cas12a. Nucleic Acids Res. 2020; 48:3228–3243. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26. Huyke D.A., Ramachandran A., Bashkirov V.I., Kotseroglou E.K., Kotseroglou T., Santiago J.G.. Enzyme kinetics and detector sensitivity determine limits of detection of amplification-free CRISPR-Cas12 and CRISPR-Cas13 diagnostics. Anal. Chem. 2022; 94:9826–9834. [DOI] [PubMed] [Google Scholar]

[B27] 27. Nalefski E.A., Patel N., Leung P.J.Y., Islam Z., Kooistra R.M., Parikh I., Marion E., Knott G.J., Doudna J.A., Le Ny A.-L.M.et al.. Kinetic analysis of Cas12a and Cas13a RNA-guided nucleases for development of improved CRISPR-based diagnostics. iScience. 2021; 24:102996. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B28] 28. Fozouni P., Son S., Doudna J.A., Fletcher D.A., Díaz de León Derby M., Knott G.J., Gray C.N., D’Ambrosio M.V., Zhao C., Switz N.A.et al.. Amplification-free detection of SARS-CoV-2 with CRISPR-Cas13a and mobile phone microscopy. Cell. 2021; 184:323–333. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B29] 29. Santiago J.G. Inconsistent treatments of the kinetics of Clustered regularly Interspaced Short Palindromic Repeats (CRISPR) impair assessment of its diagnostic potential. QRB Discov. 2022; 3:e9. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B30] 30. Ramachandran A., Santiago J.G.. CRISPR enzyme kinetics for Molecular Diagnostics. Anal. Chem. 2021; 93:7456–7464. [DOI] [PubMed] [Google Scholar]

[B31] 31. Naqvi M.M., Lee L., Montaguth O.E.T., Diffin F.M., Szczelkun M.D.. CRISPR–Cas12a-mediated DNA clamping triggers target-strand cleavage. Nat. Chem. Biol. 2022; 18:1014–1022. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B32] 32. Taylor J.R. An Introduction to Error Analysis: The Study of Uncertainties in Physical Measurements. 1996; 2nd edn.University Science Book. [Google Scholar]

[B33] 33. Johnson K.A. New standards for collecting and fitting steady state kinetic data. Beilstein J. Org. Chem. 2019; 15:16–29. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B34] 34. Easterby J.S. Coupled enzyme assays: a general expression for the transient. Biochim. Biophys. Acta. 1973; 293:552–558. [DOI] [PubMed] [Google Scholar]

[B35] 35. Holstein C.A., Griffin M., Hong J., Sampson P.D.. Statistical method for determining and comparing limits of detection of bioassays. Anal. Chem. 2015; 87:9795–9801. [DOI] [PubMed] [Google Scholar]

[B36] 36. Nouri R., Dong M., Politza A.J., Guan W.. The figure of merit for CRISPR-based nucleic acid-sensing system: improvement strategies and performance comparison. ACS Sens. 2022; 7:900–911. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B37] 37. Wu F., Zhao S., Yu B., Chen Y.M., Wang W., Song Z.G., Hu Y., Tao Z.W., Tian J.H., Pei Y.Y.et al.. A new coronavirus associated with human respiratory disease in China. Nature. 2020; 579:265–269. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B38] 38. Münger K., Baldwin A., Edwards K.M., Hayakawa H., Nguyen C.L., Owens M., Grace M., Huh K.. Mechanisms of Human papillomavirus-induced oncogenesis. J. Virol. 2004; 78:11451–11460. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B39] 39. Johnson P.D.R., Stinear T., Small P.L.C., Pluschke G., Merritt R.W., Portaels F., Huygen K., Hayman J.A., Asiedu K.. Buruli Ulcer (M. ulcerans Infection): new insights, new hope for disease control. PLoS Med. 2005; 2:e108. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B40] 40. Fuchs R.T., Curcuru J., Mabuchi M., Yourik P., Robb G.B.. Cas12a trans-cleavage can be modulated in vitro and is active on ssDNA, dsDNA, and RNA. 2019; bioRxiv doi: 08 April 2019, preprint: not peer reviewed 10.1101/600890. [DOI]

[B41] 41. Feng W., Zhang H., Le X.C.. Signal amplification by the trans-cleavage activity of CRISPR-Cas systems: kinetics and performance. Anal. Chem. 2023; 95:206–217. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B42] 42. Johnson K.A. Sigman D.S. Transient-state kinetic analysis of enzyme reaction pathways. The Enzymes. 1992; 20:Academic Press, Inc; 1–61. [Google Scholar]

[B43] 43. Singh D., Mallon J., Poddar A., Wang Y., Tippana R., Yang O., Bailey S., Ha T.. Real-time observation of DNA target interrogation and product release by the RNA-guided endonuclease CRISPR Cpf1 (Cas12a). Proc. Natl. Acad. Sci. U.S.A. 2018; 115:5444–5449. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B44] 44. Strohkendl I., Saifuddin F.A., Rybarski J.R., Finkelstein I.J., Russell R.. Kinetic basis for DNA target specificity of CRISPR-Cas12a. Mol. Cell. 2018; 71:816–824. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B45] 45. Bar-Even A., Milo R., Noor E., Tawfik D.S.. The moderately efficient enzyme: futile encounters and enzyme floppiness. Biochemistry. 2015; 54:4969–4977. [DOI] [PubMed] [Google Scholar]

[B46] 46. Cofsky J.C., Karandur D., Huang C.J., Witte I.P., Kuriyan J., Doudna J.A.. CRISPR-Cas12a exploits R-loop asymmetry to form double-strand breaks. eLife. 2020; 9:e55143. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B47] 47. Jeon Y., Choi Y.H., Jang Y., Yu J., Goo J., Lee G., Jeong Y.K., Lee S.H., Kim I.-S., Kim J.-S.et al.. Direct observation of DNA target searching and cleavage by CRISPR-Cas12a. Nat. Commun. 2018; 9:2777. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B48] 48. van Aelst K., Martínez-Santiago C., Cross S., Szczelkun M.. The effect of DNA topology on observed rates of R-loop formation and DNA strand cleavage by CRISPR Cas12a. Genes (Basel). 2019; 10:169. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B49] 49. Marino N.D., Pinilla-Redondo R., Bondy-Denomy J.. CRISPR-Cas12a targeting of ssDNA plays no detectable role in immunity. Nucleic Acids Res. 2022; 50:6414–6422. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B50] 50. Liu S., Rao X., Zhao R., Han W.. The trans DNA cleavage activity of Cas12a provides no detectable immunity against plasmid or phage. Front Genome Ed. 2022; 4:929929. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B51] 51. Meyer R.R., Laine P.S.. The single-stranded DNA-binding protein of Escherichia coli. Microbiol. Rev. 1990; 54:342–380. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B52] 52. Varble A., Marraffini L.A.. Three new Cs for CRISPR: collateral, communicate, cooperate. Trends Genet. 2019; 35:446–456. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B53] 53. Merker L., Schindele P., Huang T.K., Wolter F., Puchta H.. Enhancing in planta gene targeting efficiencies in Arabidopsis using temperature-tolerant CRISPR/LbCas12a. Plant Biotechnol. J. 2020; 18:2382–2384. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B54] 54. Schindele P., Puchta H.. Engineering CRISPR/LbCas12a for highly efficient, temperature-tolerant plant gene editing. Plant Biotechnol. J. 2020; 18:1118–1120. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B55] 55. Huang T.K., Armstrong B., Schindele P., Puchta H.. Efficient gene targeting in Nicotiana tabacum using CRISPR/SaCas9 and temperature tolerant LbCas12a. Plant Biotechnol. J. 2021; 19:1314–1324. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Determinants of CRISPR Cas12a nuclease activation by DNA and RNA targets

Eric A Nalefski

Remy M Kooistra

Ishira Parikh

Samantha Hedley

Karunya Rajaraman

Damian Madan

Abstract

Graphical Abstract

Graphical Abstract.

Introduction

Figure 1.

Materials and methods

Reagents

RNP assembly

Trans-substrate cleavage reactions

Figure 3.

Figure 7.

Figure 4.

Figure 5.

Target-cleavage reactions

Figure 2.

Trans-nuclease activation reactions

Figure 6.

Fluorescence quantification of reaction products

Data analysis

Statistical analysis

Trans-substrate cleavage

Target cleavage

Steady-state analysis of trans-substrate cleavage

Trans-nuclease activation reactions

Limits of detection

Simulation of trans-nuclease activation reactions

Figure 8.

Table 1.

Results

Canonical PAMs are neither required nor predictive of high trans-nuclease activity

Mechanistic framework to investigate target-activated trans-nuclease activity

Rates of plasmid cis-cleavage parallel trans-activity

Cis-cleavage properties of RNPs activated by DNA and RNA targets

Trans-cleavage properties of RNPs activated by DNA and RNA targets

dsDNA targets containing non-canonical PAMs and ssDNA can trigger formation of high-activity trans-nucleases

Defining activation and steady-state phases of the trans-nuclease

Defining two steps in target-induced enzyme activation

Table 2.

Rapid activation of trans-activity arises from high-affinity RNP-target binding and rapid conversion to activated enzyme

Activation kinetics impact assay performance

Determinants of enzyme activation by ds- and ssDNA targets

Protospacer

Sequence of the PAM

Target sequence outside the PAM-distal end of protospacers

Protospacer context

Target DNA supercoiling

Determinants in ssDNA targets

Discussion

Implications for bacterial systems

Implications for diagnostics

Direct target detection

Coupling detection to exponential target amplification

Future assays

Conclusions

Supplementary Material

Acknowledgements

Notes

Contributor Information

Data availability

Supplementary data

Funding

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

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