Chemically-inducible CRISPR/Cas9 circuits for ultra-high dynamic range gene perturbation

Domain: biology
Published: 2025-12-12
Editorial outcome: minor_revision

Abstract
CRISPR/Cas9 technologies provide unique capabilities for modeling disease and understanding gene-to-phenotype connections. In cultured cells, chemical-mediated control of Cas9 activity can limit off-target effects and enable mechanistic study of essential genes. However, widely-used Tet-On systems often show leaky Cas9 expression, leading to unintended edits, as well as weak activity upon induction. Leakiness can be problematic in the context of Cas9 nuclease activity, which may result in cumulative DNA damage and degradation of the target cell genome over time. To overcome these deficiencies, we have established transgenic platforms that minimize Cas9 functionality in the OFF-state along with maximized and uncompromised ON-state gene editing efficiency. By combining conditional destabilization and inhibition of Cas9, we have developed an all-in-one (one or multiple guide RNAs and Cas9) ultra-tight, Tet-inducible system with exceptional dynamic range (ON vs. OFF-state) across various cell lines and targets. As an alternative to Tet-mediated induction, we have created a Branaplam-regulated splice switch module for low-baseline and robust Cas9 activity control. Lastly, for circumstances where DNA damage needs to be avoided, we have constructed a dual-control, Tet-inducible CRISPRi module for tight and potent transcriptional silencing. This upgraded suite of inducible CRISPR systems has broad applications for numerous cell types and experimental conditions.

Introduction
CRISPR/Cas9 is a highly adaptable and efficient genome editing tool. Commonly, users design single guide RNAs (sgRNAs) that direct Cas9 to generate double-stranded DNA (dsDNA) breaks at specified genomic target sites. Inaccurate repair of the Cas9-mediated DNA damage results in a permanent genetic modification, which allows for the disruption of individual genes or thousands in the context of functional genomics screens. The power of this technology is further amplified through methods that enable users to manipulate the timing and duration of Cas9 expression. Such control helps to minimize off-target DNA damage and facilitates the study of gene function(s) over a wide range of conditions and cellular states. Inducible gene disruption also enables mechanistic and temporal assessment of cell populations depleted of essential targets, which, due to the lethal phenotypes, present challenges with approaches that rely on constitutive Cas9 expression. However, any inducible Cas9 nuclease system is dependent on low or essentially no background editing in the off state, since this would result in an accumulation of permanent gene edits over time1,2. In addition, the post-induction on state should provide high and uniform targeting efficacy across the desired cell population. Ideally, such a system would function consistently over a wide range of cell types and target loci.

Broadly applied for both in vitro and in vivo experiments, Tetracycline (Tet)/Doxycycline (Dox)-inducible systems have shown promise for timed activation of Cas9 activity. Yet, the dynamic range (e.g., contrast between the on and off state editing) of Tet-based induction is often limited by issues such as leaky background expression or suboptimal gene targeting efficiency after induction3,4. Clonal selection from transgenic cell populations can help mitigate basal Cas9 activity, but this approach is labor-intensive, risks skewing genetic and phenotypic diversity representative of the parental cells, and may fail to fully address variability across cell types. Alternative strategies for induced gene editing have been explored, including integration of a Tet-On/Cas9 cassette with insulators separating the promoter and the reverse Tet transactivator (rtTA) elements into genomic safe harbor sites5, chemical stabilization/destabilization of degron-linked Cas94,6–9, split-Cas10, intein-Cas911, hydroxytamoxifen inducible Cas912,13 Dox-inducible sgRNA14,15 or combination of each4,6,8. Despite showing value, these approaches struggle with challenges related to incomplete induction and/or residual background activity, highlighting the need for technical improvement to achieve peak precision, control, and potency of genome editing.

To address these limitations, we have developed a suite of enhanced, inducible CRISPR/Cas9 systems offering full on-state gene perturbation with near-absent background editing. In addition, we provide vectors that enable streamlined, single-step production of cell populations with optimal Cas9 activity profiles, eliminating the need for single-cell cloning. For general applications, we have engineered an all-in-one, ultra-tight transgene that combines Dox-inducible and degron-destabilized Cas9, a separate degron-linked anti-CRISPR module, and a constitutive sgRNA expression module to achieve ~60-95% editing efficiency and marginal (often unmeasurable) background across multiple genes and cell lines. We have further expanded the use of this system through co-expression of two sgRNAs for induced, population-level excision of a >2 kilobase (kb) genomic segment, facilitating conditional deletion of individual repetitive genomic regions, non-coding loci, and more. For circumstances where Tet/Dox-based induction must be avoided, we have created an all-in-one, chemically controlled anti-CRISPR-integrated splice switch module for Tet/Dox-independent regulation of Cas9 activity. Finally, in recognition of experimental conditions that would benefit from transcriptional silencing (e.g. CRISPR inhibition/CRISPRi) rather than directed DNA damage, we have established a Dox-inducible, degron-tagged Zim3-dCas9 expression vector that provides superior gene suppression compared to the commonly used KRAB-(KOX1) dCas9 and minimal background silencing.

Together, these tools offer a marked contrast between induced and background Cas9 activity in vitro, enabling precise, on-demand CRISPR applications that bolster the flexibility and reliability of gene editing for cell-based experiments.

Results
The Tet-On CRISPR/Cas9 system exhibits significant basal editing
To establish a baseline for background versus drug-induced editing with a foundational Tet-On Cas9 variant, we used piggyBac to stably integrate an all-in-one transgene across a panel of seven human cell lines. Here, S. pyogenes (Sp) Cas9 was placed under the control of the TRE3G Tet-inducible promoter upstream of a U6 promoter for constitutive expression of an sgRNA targeting CD81. A separate constitutive promoter (from human EF1α) was incorporated to drive expression of the reverse tetracycline transactivator (rtTA) and puromycin resistance (PuroR) gene, each separated by a T2A sequence to enable co-expression as distinct polypeptides16. Hereafter, we term this construct, inclusive of the CD81 sgRNA, pTET. Disruption of CD81, encoding a ubiquitously expressed cell surface protein, can be easily detected by a highly specific antibody and quantified through flow cytometry (Fig. 1A, Supplementary Fig. 1A). To confirm antibody specificity, we generated a CD81 knockout HEK293T line (pCAG-Cas9) with constitutive Cas9 and coincident CD81 sgRNA expression from the CAG and U6 promoters, respectively (Supplementary Fig. 1A). Wild-type and knockout cells were stained with anti-CD81 or an IgG isotype control, alongside unstained samples. Minimal signal in the isotype and unstained samples confirmed the specificity of anti-CD81 staining, which was detected in wild-type but not knockout cells. After the generation of stable polyclonal lines with pTET, cell populations were either treated with Dox to induce CD81 knockout or left untreated as a control. To ensure complete Cas9 cutting upon drug treatment and to capture cumulative leaky cutting in the untreated cells, we measured loss of CD81 via flow cytometry ten days after induction. A representative gating strategy is shown for HEK293T (Supplementary Fig. 1B). We observed significant and reproducible Dox-independent loss of CD81 in all tested cell lines (Fig. 1B). Background knockout levels ranged from a low of ~4% in K562 (bone marrow) cells to as high as 60% in SW480 (colorectal) cells. Upon treatment with Dox, robust (≥95%) depletion of CD81 was observed in all but one cell line (HCT116, ~80% depletion). Despite potent target disruption, these data highlight a pervasive problem of leaky Cas9 activity in the context of a conventional Tet-On system.

An ultra-tight TET-CRISPR/Cas9 system with near-absent basal editing and high knockout efficiency
In order to minimize background editing, we applied a series of modifications to pTET. These adopted different strategies to regulate Cas9 in the uninduced state including conditional stabilization of Cas9 protein, modulated anti-CRISPR expression for directed control of Cas9 activity, and drug-regulated alternative splicing to influence translation of the Cas9-containing pre-mRNA (Fig. 2A–D). The various transgenic circuits, unique control modules, and the specific drug(s) required for Cas9 expression/function are defined in Table 1. For each of these modified genetic circuits, we generated stable polyclonal cell lines in contexts that exhibited high (293T), mid (A549) and low (K562) levels of basal editing. Each stable cell population was treated with the appropriate drug or drug combination for ten days to induce CD81 depletion, while an equal number of cells were left untreated as a control. In order to obtain a performance metric for each circuit that encompasses both ON and OFF-state editing, we determined the dynamic range of Cas9 activity (i.e. the fold difference between residual editing in the OFF state and the efficiency achieved in the ON state, providing a quantitative measure of system tightness). This was calculated as the ratio of percentage CD81 knockout comparing treated versus non-treated cells.

Unless otherwise noted, inducer concentrations (2µg/ml doxycycline, 1µM Shield1, 1µM dTAGV-1, 2µM trimethoprim, or 20–150nM Branaplam depending on cell type) were pre-titrated in reporter assays to maximize performance while maintaining viability, and these doses align with ranges previously reported (see Methods).

Broadly, the re-engineered systems provided a substantial decrease in background editing and, consequently, a general improvement in dynamic range (Fig. 2E–H). Across tested lines, both expression of a non-targeting control sgRNA from the pTET-Cas9 vector (pTET-NTC) demonstrated that CD81 levels were unaffected solely by Dox addition (Fig. 2E, G, Supplementary Fig. 2A). For several conditions, basal CD81 disruption in cell lines with high (HEK293T, Fig. 2E) and moderate (A549, Fig. 2G) leakiness was reduced to levels similar to that seen with pTET-NTC. K562 cells, which already demonstrated a low background similar to pTET-NTC, and high Dox-induced editing with pTET, did not benefit from additional vector modification (Supplementary Fig. 2A, B). While there was a general improvement in the dynamic range of editing across the tested conditions, relative to pTET, the peak editing efficiency upon induction was variable; detailed below.

Similar to previous studies, we genetically fused conditional destabilizing degron domains17,18 to Cas9 within the pTET vector context, which would lead to rapid turnover of leaky Cas9 protein in the absence of Dox and the small molecule used to stabilize each degron (Fig. 2B, Table 1)6. Here, DD references a double mutant variant of the FKBP12 protein (FKBP12F36VL106P) that is continuously degraded until treated with Shield117. DHFR is an E. coli dihydrofolate reductase mutant that destabilizes tagged proteins in the absence of Trimethoprim (TMP)18. Although the destabilizing domain-tagged Cas9 variants (pTET-DD-Cas9 and pTET-DHFR-Cas9) displayed comparable percentage knockout in the ON state, background editing was lower with pTET-DD-Cas9 versus pTET-DHFR-Cas9. This resulted in higher dynamic range of basal versus induced activity with the DD-tagged Cas9 module (Fig. 2F, H).

The discovery of anti-CRISPR proteins, several of which can function as natural inhibitors of Cas9 activity, has inspired the development of systems for titratable Cas9 deactivation2,19–21. To investigate the potential application of anti-CRISPRs for reducing basal editing, we modified the pTET vector to constitutively express engineered variants of AcrIIA4, previously identified as a highly effective inhibitor of SpCas919,22. For this, we assessed AcrIIA4 fused to a Nucleoplasmin NLS and, as an additional layer of control, tested C-terminal fusions of FKBP12F36V or ligand-inducible (LID) conditional degrons (Fig. 2C). FKBP12F36V (hereafter referred to as FKBP) can be used for selective degradation upon treatment with an appropriate Tag ligand23,24. LID is an engineered variant of FKBP with a cryptic degron at the C-terminus, and is degraded in response to treatment with Shield125. Co-treatment with Dox and dTag (i.e. dTagV-1) or Shield1 is expected to induce Cas9 expression and in parallel degrade the AcrIIA4, significantly shifting the stoichiometric balance in favor of Cas9 protein/function (Fig. 2C, Table 1).

Accordingly, in the OFF state, continuous expression of non-degradable AcrIIA4 (pTET;AcrIIA4) virtually abolished basal editing (Fig. 2E, G). However, Dox-induced targeting was compromised with this variant in some cellular contexts (see A549 cells, Fig. 2G). Presumably, the induced levels of Cas9 were insufficient to overcome the suppressive effects of the constitutively expressed anti-CRISPR protein in these contexts. Consistent with this notion, editing efficiency was largely restored and/or consistently high in pTET;AcrIIA4-FKBP and pTET;AcrIIA4-LID, the degron controlled AcrIIA4 lines. In HEK293T and A549 cells, there was no significant difference in drug-induced CD81 disruption between the two degron-tagged AcrIIA4 systems (Fig. 2E, G). In K562 cells, despite a ~10% higher editing efficiency with pTET;AcrIIA4-FKBP compared to pTET;AcrIIA4-LID, the dynamic range was similar due to marginally greater background gene disruption with the AcrIIA4-FKBP fusion (Supplementary Fig. 2A, B). While subtle differences were observed across cellular contexts for pTET-AcrIIA4-FKBP and pTET-AcrIIA4-LID, the directionality was inconsistent, and we determined these systems to be functionally interchangeable with one another and the pTET-DD-Cas9 variant.

Targeted degradation and direct inhibition of Cas9 protein proved to be effective for reducing basal gene editing; we next evaluated whether we could achieve similar results through conditional translation of Cas9. For this, we exploited the Xon alternative splicing switch26 based on a modified, Branaplam-sensitive splicing cassette from SF3B3. Placed within the pTET vector (pTET-Xon-Cas9), immediately downstream of the TRE3G promoter, the modified cassette positions a small exon with an initiator (AUG) codon within an SF3B3 intron segment. In the absence of Branaplam, the exon with the initiator codon will be spliced over (i.e. excluded) and Cas9 translation will be impaired (Fig. 2D). Upon co-treatment with Dox and Branaplam, a switch in the splicing mechanism will retain the AUG-containing exon, allowing in-frame and full-length production of Cas9 protein. Across all tested cell lines, basal editing was reduced with pTET-Xon-Cas9 relative to pTET, but this came at a severe cost to perturbation once activated (Fig. 2E, G, Supplementary Fig. 2A). These results suggested that additional measures were needed to optimize the performance and compatibility of the Xon and pTET systems.

Encouraged by the targeting efficacy after induction with either the degradable Cas9 or anti-CRISPR control modules, and the potential for further reduction in basal editing, we generated a composite pTET vector with both the DD-Cas9 and AcrIIA4-LID systems (pTET-DD-Cas9;AcrIIA4-LID). Using this design, co-treatment with Dox and Shield1 will induce and stabilize Cas9, while simultaneously relieving inhibition of Cas9 activity through depletion of AcrIIA4. We favored this specific combination of control modules, since it restricts the number of inducing agents to two rather than the three required for incorporation of either DHFR-Cas9 or AcrIIA4-FKBP. Application of pTET-DD-Cas9;AcrIIA4-LID resulted in a >100-fold dynamic range when comparing the ON versus OFF states for CD81 depletion in both the HEK293T and A549 backgrounds, which was 2-fold greater than any other system in these contexts, with a trend towards improved dynamic range in K562 cells (Fig. 2F, H, Supplementary Fig. 2A, B). This was due to the low background editing level, which was indistinguishable from the pTET-NTC control. Induced editing remained high (>80% within polyclonal cell populations) with pTET-DD-Cas9;AcrIIA4-LID, but with a marginal (~10%) decrease relative to pTET in both the A549 and K562 cells. We therefore decided to prioritize and expand our evaluation of the pTET-DD-Cas9;AcrIIA4-LID system (hereafter named pTET:Ultra-tight) due to its unique capacity for nearly eliminating background activity with minimal impact on induced editing.

To further validate the function of the control modules, we performed western immunoblotting to assess Cas9 protein expression in pTET and pTET:Ultra-tight systems following drug treatments (Supplementary Fig. 3A). HEK293T cells transfected with each construct were treated with the indicated small molecules for 72h, followed by protein extraction and immunoblotting for Cas9. As expected, Cas9 protein was undetectable in untreated pTET:Ultra-tight cells, reflecting tight regulation. Interestingly, Cas9 protein levels remained below the detection limit in pTET cells despite the significant background editing observed by flow cytometry, suggesting that functional activity can occur at levels undetectable by western blot analysis. As such, target gene depletion is a more sensitive readout of leaky Cas9 activity relative to Cas9 protein detection. Cas9 protein was readily observed upon Dox treatment of both pTET and pTET:Ultra-tight cells. Expression was lower in the pTET:Ultra-tight condition, likely due to degradation via the DD tag in the absence of Shield1. Cas9 protein was not seen when pTET:Ultra-tight cells were treated with Shield1 alone. Co-treatment of pTET:Ultra-tight cells with Dox and Shield1 increased Cas9 protein levels, though to a lesser extent than in pTET cells. However, this expression level was sufficient to drive efficient target gene knockout as indicated by flow cytometry results (Fig. 2E–H, Supplementary Fig. 2A, B).

Attempts to detect Anti-CRISPR protein by immunoblotting were unsuccessful due to limitations of available antibodies. Despite this, changes in CD81 protein levels were consistent with the expected regulation of the degron-tagged Anti-CRISPR in the presence or absence of the specific drugs (Fig. 2E–H).

Time-resolved target depletion demonstrates concordance between protein loss and indel formation
Having established pTET:Ultra-tight as a more tightly regulated system than pTET, we next compared the kinetics of editing for each system over a period of 10 days (Fig. 2I). Untreated pTET cells exhibited ~50% CD81 depletion, which did not increase further over time. In contrast, untreated pTET:Ultra-tight displayed minimal basal editing throughout the time course. In drug treated pTET cells, maximal CD81 depletion occurred by 72h. pTET:Ultra-tight cells took longer than pTET cells to achieve maximal CD81 depletion, but reached equivalent levels by 144-240h.

To confirm that flow cytometry results reflected underlying genomic changes, we performed targeted amplicon sequencing of CD81 on the same samples to quantify indel frequencies over time (Fig. 2J). The amplicon sequencing data closely paralleled target protein levels. Untreated pTET cells displayed ~50% indel generation at the CD81 locus throughout the time course. In pTET:Ultra-tight cells, indels remained minimal without treatment, confirming suppression of leaky Cas9 activity. For both systems, between 6 and 48h after treatment, indel frequencies exceeded the percentage of CD81-depleted cells, suggesting a lag between genomic editing and observable protein loss, likely due to transcriptional/translational rates and/or protein half-life. Similar to flow cytometry results, upon induction, pTET:Ultra-tight cells showed a modest delay compared to pTET cells before achieving maximal editing. Indel frequencies plateaued in both systems by 144h of drug exposure.

A CD81 knockout control line (Supplementary Fig. 1A) was established by stable genomic integration of a constitutive Cas9 and CD81 sgRNA-expressing transgene. This line exhibited complete protein depletion and genomic indel formation by 24h after removal of puromycin (used for selecting stable integrants) and these levels remained constant throughout the duration of our assay (~240h). (Supplementary Fig. 3B, C).

Together, these time-course experiments confirmed the minimal background editing and high efficiency with the pTET:Ultra-tight system as quantified by both indel generation and protein depletion. While our initial assays focused on a 5–10-day window post-induction, the ability to detect robust editing at earlier time points expands the utility of the system, allowing researchers to capture transient phenotypes, resolve early molecular events, or study genes whose prolonged loss impairs cell viability.

Consistent pTET:Ultra-tight functionality across a diverse cell line panel
In order to assess the broad applicability of pTET:Ultra-tight across an expanded set of cell lines, we stably integrated pTET and pTET:Ultra-tight into SW480, HeLa, HAP-1, and HCT116. In our initial evaluation of pTET, these four cell lines exhibited a range of basal editing from ~60% to 8% (Fig. 1B). Consistent with our previous observations, the pTET:Ultra-tight lines displayed substantially reduced basal editing in untreated cells relative to pTET in all cell types (Fig. 3A). Of the tested lines, SW480 cells represented an outlying context, where pTET:Ultra-tight was unable to fully reduce background editing. We therefore decided to compare pTET:Ultra-tight to an expanded set of genetic circuits (Supplementary Fig. 4A), revealing that it remained the superior option for balancing ON versus OFF-state editing (Supplementary Fig. 4B). Upon drug exposure, the pTET:Ultra-tight system maintained a knockout efficiency greater than 90% in all but one cancer cell line; HCT116 (58%). HCT116 cells also displayed the lowest induced knockout rate (~80%) tested across all lines with the pTET circuit, suggesting the Dox-regulated circuit requires further optimization within this particular cellular context (Fig. 1B and Fig. 3A).

Despite a reduction in knockout efficiency within HCT116 cells with the pTET:Ultra-tight system, the gains in dynamic range between the ON and OFF states, due to a near-complete elimination of background editing, were substantial (>50-fold relative to pTET); a trend that extended to the additional lines tested in this study (Fig. 3B).

To confirm that protein-level changes were reflected at the genomic locus, we evaluated indel generation in HAP-1 cells by targeted amplicon sequencing. We observed strong agreement between protein-level and genomic changes (Supplementary Fig. 4C), mirroring the concordance seen in our kinetic analysis.

In addition to immortalized cell lines, we extended our evaluation to include a more sensitive, non-transformed line, KOLF2.1J, an induced pluripotent stem cell (iPSC) line that has been extensively characterized and favored for its genomic stability, efficient CRISPR/Cas9 editing, and robust differentiation capacity27,28. Stable, polyclonal populations were engineered with pTET or pTET:Ultra-tight and treated with either Dox (pTET) or Dox + Shield1 (pTET:Ultra-tight) for 10 days followed by flow cytometry to measure CD81 levels. Both treated and untreated KOLF2.1J stable cell lines exhibited morphological features characteristic of healthy iPSCs, including prominent nuclei, tightly packed colony growth with well-defined borders, and minimal evidence of spontaneous differentiation (Supplementary Fig. 4D). Additionally, SSEA-4, a well-established marker of iPSC pluripotency, was highly expressed across all conditions, confirming that the cells maintained a pluripotent state irrespective of transgene or drug treatment (Supplementary Fig. 4E). While we observed a modest decrease in targeting efficiency when comparing pTET (~93%) to pTET:Ultra-tight (~78%), background editing was absent with pTET:Ultra-tight, resulting in the highest dynamic range of editing for all tested cell types (>200-fold difference between the ON vs. OFF-state). Notably, the ~78% editing efficiency observed with pTET:Ultra-tight in KOLF2.1J represents a substantial improvement over published inducible systems in stem cells, which report efficiencies closer to ~40%5,6. Together, these results suggest pTET:Ultra-tight consistently outperforms pTET across multiple cellular contexts.

The pTET:Ultra-tight system maintains highly controlled regulation at additional target genes
To investigate whether the pTET:Ultra-tight system maintains stringent regulation at target loci other than CD81, we replaced the CD81 sgRNA in pTET and pTET:Ultra-tight constructs with sgRNAs targeting an expanded cohort of non-essential cell surface markers; β2-microglobulin (B2M), CD298, and CD9. As previously, we generated stable, polyclonal HEK293T cell lines with these constructs, treated each line with either Dox or Dox + Shield1, and evaluated target protein levels 10 days later by antibody staining and flow cytometry. Untreated cells were included as a control to determine basal editing. Consistent with our observations for CD81, we detected background editing in untreated cells at all three target loci with the pTET system (Fig. 4A–C). In contrast, pTET:Ultra-tight displayed strikingly diminished basal editing for each target. Importantly, upon drug exposure, population-wide knockout efficiencies >90% were observed for all target genes. Significant improvement in dynamic range was observed across targets with the pTET:Ultra-tight system (Fig. 4D–F). Consistent with prior observations, editing at the B2M locus showed close alignment between indel frequencies and protein loss (Supplementary Fig. 5).

In general, the robust agreement observed between targeted amplicon sequencing and flow cytometry across cell types, target loci, and experimental contexts supports pTET:Ultra-tight as a superior system for regulating Cas9 activity.

pTET:Ultra-tight system exhibits tight control over essential gene knockout
Large-scale functional genomics screening campaigns have revealed a host of core and condition-specific essential genes29. Detailed mechanistic study of these factors is advantaged with an experimental system that allows precise control over the timing of gene disruption, as constitutive knockout is expected to result in early growth arrest or cell loss. Given the near-absent leakiness and potent induction observed with the pTET:Ultra-tight system, we sought to evaluate its potential for investigating an essential gene phenotype.

We constructed pTET and pTET:Ultra-tight all-in-one plasmids expressing an sgRNA targeting the Polo-like kinase 1 (PLK1) gene. PLK1 is a serine/threonine kinase and a master regulator of cell cycle and mitosis. Disruption of PLK1 results in proliferation defects. PLK1 is overexpressed in several cancers and is considered a potential target for cancer therapy30,31. Each of the PLK1-targeting constructs, as well as a pTET vector expressing a NTC sgRNA, were stably integrated into HEK293T cells to create polyclonal populations. HEK293T cells were chosen based on the relatively high rate of background editing with pTET (Fig. 1B). Each cell population was either treated with Dox (pTET) or Dox + Shield1 (pTET:Ultra-tight) to induce Cas9 activity or left untreated as a control for background (leaky) targeting. Cell growth was monitored by live imaging over five days in the presence/absence of drugs (Fig. 5A). Stable lines were successfully generated for all constructs, but there were notable differences in growth rates in the presence/absence of drug treatment. In the untreated state, the pTET line targeting PLK1 exhibited reduced proliferation, suggesting knockout of the target gene in the absence of induction (Fig. 5B). Conversely, the growth rate of the pTET:Ultra-tight line expressing PLK1 sgRNA was comparable to the pTET-NTC control line (Fig. 5B). Upon drug treatment, growth was almost fully inhibited in both pTET and pTET:Ultra-tight lines, although the latter demonstrated a greater reduction in growth rate. As previously shown with non-essential gene disruption (Fig. 2E–H, Fig. 4A–F), the pTET:Ultra-tight system demonstrated a significantly improved dynamic range between treated and untreated states, as compared to pTET (Fig. 5C) While a population-level phenotype was observed with both pTET and pTET:Ultra-tight, we interpret the difference in dynamic range between the systems to reflect “pruning” of cells with leaky and, likely, the greatest capacity for PLK1 disruption in the pTET context. In that scenario, cells with diminished Cas9 activity would survive but induction would result in a less penetrant phenotype. Our data reinforce the notion that the pTET:Ultra-tight system supersedes pTET for implementing effective, population-level editing control across multiple cell lines and targets including essential genes.

Conditional deletion of a ~2.5 kb genomic locus with paired guide RNAs
Thus far, we have explored the general capability of pTET:Ultra-tight for conditional disruption of genes using individual, coding exon-targeting sgRNAs. The use of paired sgRNAs, where two guides are co-expressed and target positions spaced along the same chromosome, enables specified excision of large regions of DNA, enabling more complex and expanded genomic studies beyond what is capable with single sgRNA targeting. Such edits could include deletion of regulatory DNA elements (i.e. enhancers), non-coding RNAs, repetitive DNA, specific exons, and genes with family members or pseudogenes sharing highly similar exonic sequences32–36. To this end, we assessed the potential for pTET:Ultra-tight and paired sgRNA expression to induce targeted genomic deletions.

As proof-of-concept for conditional genomic excision with pTET:Ultra-tight, we focused on the H3-3A locus, which expresses the non-canonical H3.3A histone variant. H3-3A and the near-identical H3-3B genes present a distinct use case for targeted deletion, since these share 100% nucleotide similarity across their coding sequences, while the intron and untranslated regions are unique37,38. In addition, H3-3A has several pseudogenes with >90% sequence overlap with the H3-3A open reading frame39. These uncommon features preclude the use of single sgRNAs for the disruption of H3-3A via exonic targeting. However, taking advantage of the non-duplicated genomic segments, we designed a pair of sgRNAs targeting intronic sequences flanking exons 2 and 3, which span ~2.5 kilobases (kb), of the H3-3A locus (Fig. 5D).

For expression of the H3-3A-targeting sgRNA pair, we cloned one guide within the original U6 promoter cassette in our pTET and pTET:Ultra-tight vectors, while the second guide of the pair was cloned downstream of a human H1 promoter. Stable, polyclonal lines were generated in HEK293T with both the pTET and pTET:Ultra-tight constructs, and cells were either left untreated or exposed to inducing drug(s) for 72h. After this period, we collected genomic DNA and performed PCR with primers spanning the target sites. An amplicon size of ~2.9kb indicated the presence of the wild-type allele, while successful deletion of the target locus would result in a shorter amplicon of ~0.4kb.

In the absence of drug treatment, the ~2.9kb product corresponding to the wild-type allele was detected in parental, pTET-NTC control, pTET and pTET:Ultra-tight cells (Fig. 5D). However, only within the pTET cells did we observe the ~0.4kb deletion product, indicating leaky editing in this context. Upon drug treatment, the deletion product was observed for both pTET and pTET:Ultra-tight cell lines with a concomitant reduction in signal for the full-length amplicon. Taken together, these results demonstrate the technical feasibility of this approach and validate the use of pTET:Ultra-tight and paired sgRNA expression for precisely controlled deletion of genomic loci.

Development of an enhanced, Dox-independent, conditional CRISPR/Cas9 system
While the Tet/Dox system is commonly employed for conditional transgene expression, it is not uniformly compatible with every cell model or experimental situation. For instance, Dox can disrupt mitochondrial function, making the Tet/Dox system less desirable for studies involving this organelle40,41. Separately, orthogonal control of distinct genetic circuits, to direct or understand complex cellular processes, may be enabled through the use of non-Tet/Dox systems for modulating CRISPR/Cas9 activity42. Furthermore, the ability to orthogonally control two processes in the same cell model is a common need that necessitates exploration of an alternative inducible system for controlling CRISPR activity43,44.

Previous studies have applied constitutive Cas9 expression in combination with conditional degradation or alternative splicing for drug-regulated gene editing8,45,46. To evaluate the potency and background editing of Dox-independent induction of Cas9, we designed all-in-one piggyBac vectors with Cas9 expression driven by a constitutive promoter (CMV) and protein levels controlled by either DD or DHFR fusion domains or a Xon cassette (Supplementary Fig. 6A). In addition, these vectors contained a constitutive CD81-targeting sgRNA expression cassette. Stable, polyclonal HEK293T cell lines were generated for each construct. After antibiotic selection, cell populations were cultured for 10 days with or without respective drug treatments. DD-(N- or C-terminal) and DHFR-fused Cas9 expression resulted in 100% depletion of CD81 in the presence or absence of drug treatment, indicating a complete lack of Cas9 activity control under these conditions (Supplementary Fig. 6B). By contrast, the Xon-Cas9 construct enabled inducible control of Cas9 activity, with ~88% disruption of CD81 in the presence of Branaplam. However, this system displayed nearly 20% basal editing in the untreated HEK293T cell context, which encouraged further optimization (Supplementary Fig. 6B).

Based on our ability to mitigate leaky Cas9 activity within the pTET system, we deployed similar strategies to improve upon the Xon-Cas9 (Fig. 6A). To begin, we added the DD or DHFR domain to the N- or C-terminus of Cas9 under control of Xon. Our data with the optimized TET systems suggested that N-terminal fusion of DD and DHFR would reduce background editing activity (Fig. 2E, G). However, in the context of the Xon switch, an N-terminal degron fusion led to 100% leakiness of Cas9 activity (Supplementary Fig. 6C). We reasoned that this could be due to an internal start codon provided by the DD or DHFR domains, which would enable Cas9 translation without Branaplam-induced exon inclusion (Supplementary Fig. 6D). Consistent with this hypothesis, C-terminal DD or DHFR domain fusions demonstrated a substantial reduction in background effects without impacting ON state editing (Fig. 6B). We next tested the addition of an anti-CRISPR protein AcrIIA4 with or without the FKBP degron. The addition of AcrIIA4 dramatically reduced the leakiness of Xon-Cas9 in HEK293T cells, but at the cost of significantly compromised inducible cutting (Fig. 6B). By contrast, inclusion of the AcrIIA4-FKBP control module not only reduced the leakiness but also maintained full ON state editing, offering the best dynamic range of all conditions tested in this cell type (Fig. 6B, C).

To evaluate editing kinetics in the Xon-Cas9 and Xon-Cas9;AcrIIA4-FKBP systems, we performed a time-course analysis with both flow cytometry (Fig. 6D) and targeted amplicon sequencing (Supplementary Fig. 6E). In the absence of drug treatment, Xon-Cas9 cells exhibited progressive loss of CD81 and increasing indel formation, whereas Xon-Cas9;AcrIIA4-FKBP cells maintained minimal background editing throughout the time course. Upon drug induction, Xon-Cas9 cells showed time-dependent indel formation and depletion of CD81, plateauing by 240h. Drug-treated Xon-Cas9;AcrIIA4-FKBP cells exhibited slower editing relative to Xon-Cas9, but comparable knockout levels by 144h. These time course results confirm that Xon-Cas9;AcrIIA4-FKBP is a superior system for controlling Cas9 activity.

We then tested the Xon-Cas9;AcrIIA4-FKBP combination in K562 cells. Although background editing was nearly absent in this cell line with the pTET system, the unmodified Xon-Cas9 system displayed ~19% leaky target disruption (Fig. 6E). Use of the Xon-Cas9;AcrIIA4-FKBP vector reduced leakiness in this cellular context, but it also resulted in poor induction of Cas9 activity (Fig. 6E, F). Within this vector design, AcrIIA4 expression is under the control of a PGK promoter (Fig. 6A). Separate studies have observed slightly higher or more stable transgene expression from the PGK promoter relative to other constitutive promoters, albeit with context-dependent results47,48. Therefore, we hypothesized that the PGK-driven levels of AcrIIA4 protein were too high, even with dTag-mediated degradation, to fully reverse inhibition of Cas9 function, and that weaker AcrIIA4 expression was needed. To this end, we evaluated two distinct strategies aimed at lowering the baseline expression of AcrIIA4 such that it would maintain suppression of Cas9 in the uninduced state but allow for more complete reversal of this effect when cultured with a dTag molecule. This included exchange of the PGK promoter for the generally less-expressive UBC promoter47,48 or construction of an extended, polycistronic transgene with AcrIIA4 expressed by way of an IRES downstream of the CMV-Xon-Cas9 cassette (Supplementary Fig. 6F). IRES elements are commonly used to express multiple genes simultaneously and have been reported to drive lower expression of the downstream gene compared to the upstream, cap-dependent translated segment49. Both background and induced editing were greatly reduced with the UBC promoter driving the anti-CRISPR module, suggesting that UBC is stronger than PGK in our tested K562 cell line (Fig. 6E). By contrast, IRES-AcrIIA4-FKBP led to significantly reduced leakiness with no compromise in induced cutting and displayed the best dynamic range of editing (Fig. 6E, F). Together, these results demonstrate that non-Dox-inducible Cas9 expression systems can enable high efficiency and low background editing, aided through co-expression of degradable anti-CRISPR proteins, but cell type-dependent variability necessitates empirical testing for optimal performance.

An enhanced, inducible CRISPRi system for potent, tightly regulated gene silencing
The use of Cas9-mediated dsDNA breaks for gene perturbation may not be suitable for certain applications50. Accordingly, CRISPR interference (CRISPRi) provides a complementary and powerful tool for loss-of-function purposes. Here, a nuclease-dead Cas9 (dCas9) is fused to a Krüppel-associated box (KRAB) repressor or various chromatin and DNA modifying domains to facilitate transcription silencing of an intended target gene through a promoter or enhancer-targeting sgRNA51,52. While dCas9 fused to the KOX1 KRAB domain has commonly been used for CRISPRi, ZIM3 KRAB has emerged as a more potent alternative53,54, in both constitutive and inducible vector contexts53,55,56. Recognizing the utility of inducible CRISPRi for targeting both coding and non-coding regions, we sought to identify key parameters that enable robust, population-wide gene silencing and minimal background through expression of dCas9-KRAB variants.

We began by evaluating a foundational set of TET-inducible CRISPRi systems. For this, KRAB domains from either KOX1 (pTET-KOX1) or ZIM3 (pTET-ZIM3) were fused to dCas9 and this cassette was positioned downstream of a TRE3G promoter within a lentiviral backbone with a constitutively expressed blasticidin resistance and rtTA transgene (Fig. 7A). Lentiviral infection with this vector was used to generate stable, polyclonal DLD1 (colorectal cancer) cell lines. These lines were subsequently transduced with lentivirus expressing an sgRNA targeting the CD81 promoter. Each cell population was treated with Dox or left untreated for 7 days and then assessed for CD81 depletion. The pTET-KOX1 cell line exhibited minimal effect on CD81 levels in the absence of Dox, but also correspondingly low (<50%) activity upon induction (Fig. 7B, Supplementary Fig. 7). Conversely, pTET-ZIM3 demonstrated substantial improvement for silencing CD81 upon Dox induction, although this potency translated to noticeable leakiness, impacting ~40% of the untreated population (Fig. 7B, Supplementary Fig. 7). This prompted us to optimize the ZIM3-based CRISPRi system to reduce its leakiness and maintain its activity via the principles applied for inducible expression of Cas9 nuclease.

To improve dynamic range between the CRISPRi ON and OFF states, we incorporated an N-terminal DD tag to the ZIM3-dCas9 cassette, resulting in pTET-DD-ZIM3. In this instance, a separate anti-CRISPR module was not included to avoid complications with viral particle production related to cargo size limitations with lentiviral backbones. Induction of pTET-DD-ZIM3 cells by combined Dox and dTag treatment reduced CD81 levels similar to pTET-ZIM3 with minimal background gene silencing (Fig. 7B). As a result, dynamic range for the inducible, degron-enabled CRISPRi system increased ~13 fold over pTET-ZIM3 (Fig. 7C). Similar to our evaluation of pTET:Ultra-tight, we compared pTET-ZIM3 with pTET-DD-ZIM3 for disruption of the essential gene, PLK1 (Fig. 7D). As with pTET:Ultra-tight, background editing was largely eliminated with pTET-DD-ZIM3, and this coincided with a more robust phenotype upon induction (Fig. 7E). For CRISPRi, the combination of Dox induction with degron-mediated stabilization of ZIM3-dCas9 offers the most potent tool for gene silencing with minimal background.

Methods
Cell culture and stable line generation
All parental cell lines, except KOLF2.1J, were obtained from Genentech cell bank (gCell), validated by STR profiling, and tested for mycoplasma. The KOLF2.1J iPSC line was obtained from the Jackson Laboratory (https://www.jax.org/jax-mice-and-services/ipsc/cells-collection/JIPSC001000; product code: JIPSC001000), maintained in feeder-free media, and validated by KaryoStat+™ (Thermo Fisher Scientific), PluriTestTM (Thermo Fisher Scientific) and mycoplasma testing. Genome integration was accomplished using the PiggyBac transposition system where each piggyBac transposon plasmid was co-delivered into cells with a plasmid expressing the piggyBac transposase. Cell culture conditions and transfection methods are described in Supplementary Table 1. Briefly, stable integration in KOLF2.1J was achieved by nucleofection using a Lonza 4D-Nucleofector, while immortalized cell lines were seeded as described and transfected in 1 mL Opti-MEM, with the indicated transfection reagents as per manufacturer’s recommendations. Opti-MEM was replaced with growth media 4h post-transfection.

Plasmid constructs were generated by gene synthesis (Genscript) and included either Puromycin or Blasticidin resistance markers to select for stable cells. Unless otherwise indicated, stable lines were enriched with 1 μg/ml Puromycin or 20 μg/ml Blasticidin. Sequences of plasmids used in the study are listed in Supplementary Table 6. Guide RNA sequences for CRISPR/Cas9 nuclease-based editing and cell culture drugs used in this study are listed in Supplementary Tables 2 and 4.

Small molecule dosing rationale
All small molecules used in this study have been characterized in prior publications5,17,18,24–26,76–79 and were further titrated in our laboratory using luciferase or GFP reporter systems. Final working concentrations were chosen to maximize induction or degradation while minimizing cytotoxicity, and they consistently fall within ranges reported in the literature. Unless otherwise noted, experiments were conducted using the following concentrations: Doxycycline (2µg/ml), Shield1 (1µM), dTAGV-1 (1µM), trimethoprim (2µM), and Branaplam (20nM in HEK293T and 150nM in K562). Dose–response assays confirmed that these concentrations achieved near-maximal performance with minimal cytotoxicity.

Flow cytometry
In brief, adherent cells were lifted by TrypLE, or Accutase in the case of KOLF2.1J, at 37°C for 5 min and resuspended in growth media. Dissociated adherent cells or suspension cells were spun down at 500g for 5 min and resuspended in 100 μl FACS buffer (PBS, 0.5% BSA, 0.05% NaN3 sodium azide) with 5 μL of antibody. After a 15-min incubation on ice in the dark, the cells were washed three times with FACS buffer and passed through a 35-μm cell strainer. Data was collected on BD Symphony instruments using FACSDiva acquisition software. Data was analyzed by FlowJo v10.10.0. The gating strategy to isolate live and single cell populations is shown in Supplementary Fig. 1B. The following antibodies were used for flow cytometric analysis. CD81 Monoclonal Antibody (1D6-CD81) APC (Thermo Fisher Scientific, Cat. No. 17-0819-42), mouse IgG1 kappa isotype control antibody APC (Thermo Fisher Scientific, Cat. No. 17-4714-82), SSEA-4 antibody Fluorescein (R&D Systems, Cat. No. FAB1435F), CD81 antibody PE (BioLegend, Cat. No. 349505), β2-microglobulin Antibody FITC (BioLegend, Cat. No. 316304), CD9 Antibody PE/Cyanine7 (BioLegend, Cat. No. 312115) and CD298 Antibody PE/Cyanine7 (BioLegend, Cat. No. 341707).

Western immunoblotting
Whole-cell extracts were prepared by lysing equal cell numbers in 1X Laemmli Sample Buffer (Bio-Rad, Cat. No. #1610737) for 10 min, followed by three cycles of sonication and boiling using a Bioruptor Plus (Diagenode). Proteins were separated on an SDS-PAGE and transferred to PVDF membranes using an iBlot 2 Gel Transfer Device (Fisher Scientific, IB21001), followed by immunoblotting with the following antibodies: Cas9 (NovusBio, Cat. No. 7A9-3A3), α-Tubulin (Sigma Aldrich, Cat. No. CP06), Rabbit Anti-Mouse IgG (H+L)-HRP secondary antibody (SouthernBiotech, Cat. No. 6170-05), and Goat Anti-Rabbit IgG-HRP secondary antibody (SouthernBiotech, Cat. No. 4030-05). Chemiluminescence was performed with Amersham enhanced chemiluminescence (ECL) Prime reagent (Cytiva, RPN2232) and imaged on a ChemiDocTM MP Imaging System (Bio-Rad).

Amplicon sequencing
Genomic DNA was extracted from cells using the Agencourt DNAdvance Kit (Beckman Coulter). PCR primers flanking the target site were designed to amplify 200–300bp amplicon using a 2 stage PCR method. Primers were synthesized at Integrated DNA Technologies (IDT). The PCR includes genomic DNA, i5 and i7 indexed adapters, target-specific primers with universal adapters, and PrimeTime Gene Expression Master Mix (IDT). The first stage of PCR amplifies the target-specific amplicons with target-specific primers flanked by universal adapters at 60°C Tm for 30 cycles. The second stage of PCR incorporates the indexed sequencing adapters in the target-specific amplicons at 55°C for 4 cycles. We use dual-end indexing by incorporating i5 and i7 10bp indexes by IDT to uniquely label amplicons produced from each PCR. The PCRs are assembled using Echo 525 Acoustic Liquid Handler (Beckman Coulter). Following PCR, the amplicons from all of the reactions are pooled together. The amplicon library pool is cleaned twice using KAPA Pure Beads (Kapa) at a 1:1 ratio to remove unused primers and index adapters. The quality and quantity of the clean amplicon library pool is determined using 4200 TapeStation System (Agilent) and High Sensitivity D1000 DNA ScreenTape. The amplicon library pool is diluted to 15 pM with 5% PhiX as per Illumina MiSeq library preparation procedure, and sequenced on Illumina MiSeq, dual read, 150/150 cycles to produce 300bp sequences using the Miseq V2 kit (Illumina). Primer sequences used for amplicon sequencing are listed in Supplementary Table 5.

Sequence analysis
Sequencing reads were demultiplexed using i5 and i7 index barcodes, allowing up to one mismatch per index. Reads were then filtered to retain only those with a minimum Phred quality score of 30. For each reaction, reads were aligned to the expected amplicon sequence using BWA-MEM280. Alignment results were parsed using custom software to extract the CIGAR string (which encodes how each read aligns to the reference, including matches, insertions, and deletions). From this, the total number of aligned reads, the number of exact matches to the wild-type sequence, the number of reads with indels, and the number of partially aligned or non-matching reads were determined. Exact matches were classified as wild-type (unmodified), and reads with indels were classified as edited (reflecting sgRNA activity). Partially matched or non-specific reads were excluded from downstream analysis. Wild-type and indel-containing reads were then reported as a percentage of the total number of high-quality, specific alignments.

Lentivirus production and transduction
HEK293T cells were seeded and grown to 80-90% confluency at the time of transfection. Lentiviral plasmids expressing target genes were co-transfected with packaging plasmids pDelta8.9 and pVSVG into HEK293T cells using Lipofectamine 2000 reagent. 6h after transfection, the medium containing the transfection mix was aspirated and fresh growth media was added to the cells. Lentiviruses were collected twice from the supernatant 24h and 48h post-transfection. Cell debris was removed by passing the crude supernatant through the 45 μm filter. Lentivirus was concentrated by Lenti-X (Takara Bio). DLD1 cells were cultured in RPMI 1640 supplemented with 10% TET-free FBS, 2mM L-Glutamine and pen-strep. Cells were seeded in 6-well Corning plates and grown to 50% confluency on the day of transduction. The lentivirus and 8 μg/ml polybrene were added to the DLD1 cells. Cells were spin infected at 1800 rpm for 45min and incubated for 48h before passage. DLD1 was first transduced with lentivirus expressing inducible CRISPRi systems and selected with 10 μg/ml Blasticidin to obtain the stable cell line, and then transduced with lentivirus expressing sgRNA and selected with 2 μg/ml puromycin.

PCR genotyping
Genomic DNA was isolated from untreated and treated cells 72h post-treatment using QuickExtract™ DNA Extraction Solution (Lucigen). Both full-length and deleted forms of the target locus were amplified with Q5® High-Fidelity 2X Master Mix (NEB) using primers (forward 5′-GGCTTTGTAGAATGCTCGCTG-3′; reverse 5′-TTTAACGACCCCAACTGCAT-3′) flanking exons 2 and 3 within the H3-3A gene. Primers were synthesized at IDT. Amplicons were resolved on an agarose gel and images were obtained using the ChemiDoc MP imaging system (Bio-Rad).

Cell proliferation assay
An equal number of HEK293T pTET-NTC, pTET or pTET:Ultra-tight stable cells expressing PLK1 sgRNA, were seeded at low density in growth media with or without inducing drug(s). Cell growth was monitored using the IncuCyte S3 Live Cell Imaging and Analysis Instrument (Sartorius). Phase contrast cell images were collected using a 10× objective lens within the instrument every 4h over 5 days and cell proliferation was determined based on confluence.

Cell viability assay
DLD1 stable cells generated by transduction of sgNTC lentivirus or sgPLK1 lentivirus cocktail (PLK1 sgRNA1 + PLK1 sgRNA2) were replated into 96 well format at 1000 cells/well in 80 μl medium. The following day, 20 μl of medium with or without treatment (final concentration: 250 ng/ml dox, 250 ng/ml dox + 1 μM shield1) was added to begin induction. On day 5 after induction, CellTiter-Glo Luminescent Cell Viability Assay (Promega #G7572) was used to read out cell viability on an EnVision plate reader (PerkinElmer/Revvity). Guide RNA sequences are provided in Supplementary Table 3.

Figure production
Flow cytometry plots and histograms were generated in FlowJo v10.10.0. Graphical representations (bar graphs, floating bars, and scatter plots) and statistical analyses were performed in GraphPad Prism v10. Illustration elements were created with BioRender.com.

Statistics and reproducibility
All experiments were conducted in the indicated cell lines under standardized culture and treatment conditions. Experimental groups (stable cell line, treatment, and timepoint) were defined by the study design. Sample sizes were established empirically based on reproducibility across independent experimental replicates, as specified in the figure legends, Methods, or associated text. For quantitative assays, independent biological replicates were performed to confirm reproducibility and enable statistical comparisons. Replicate numbers were chosen according to standard practice for cell-based and molecular assays.

The exact n, statistical test, and data presentation format (e.g., mean ± SD/SEM or floating bars representing min–max with median line) are indicated in the figure legends. Statistical analyses were performed using GraphPad Prism v10.

No statistical method was used to predetermine sample size. No data were excluded from analyses. Experiments were not randomized, and investigators were not blinded to sample allocation or outcome assessment.

Materials availability
Biological materials generated for these studies can be attained via MTA by contacting Andrew Ng (ng.andrew.an1@gene.com). Details related to the MTA process can be found on the Genentech Outgoing MTA Program website: https://www.gene.com/scientists/mta. Vector and guide RNA sequences are provided in Supplementary Tables.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Srinivasan and colleagues developed a collection of improved CRISPR systems to overcome the limitations of exciting approaches. The authors developed an inducible and degron-destabilized Cas9 expression systems, applied degradation approaches to anti-CRISPR proteins, and developed a Branaplam-sensitive splice switch to control the genetic editing machinery. The approaches minimize unintended editing in the off-state while maximizing editing efficiency in the on-state. Overall, this is a valuable toolbox for the community for CRISPR editing experiments. However, I have several comments for the authors to address as the characterization, applications, and specificity of this study are incomplete:\nMajor points:\n1) The dosing and timing of compounds used in this study are unclear. Demonstration of their activity in stabilizing or destabilizing the proteins are also not shown, making it difficult to understand the experimental design in the experiments and the comprehensiveness of evaluation. The authors should show stabilization/destabilization of their control modules and provide data to support the dosing that is selected for experiments in this study.\n2) In Figure 2 ,3 and 5, while the authors show flow cytometry data for their target expression, editing of the genomic locus should also be shown to help verify their observations and efficiencies.\n3) In Figure 4, the characterization of genomic and protein level disruption is insufficient to show PLK1 or H3-3A is altered and to what extent. The consequences of the H3-3A disruption should also be evaluated to show a phenotypic effect. Is there a biological observation the authors can now achieve that was not evident with prior approaches? Most critically, rescue experiments are not provided to show that these effects are on-target and need to be performed with a CRISPR-resistance expression strategy. This will be necessary for the author’s conclusions and is the only biological data shown in the paper.\n4) The authors comment that temporal studies are feasible with these approaches. However, this is not demonstrated, nor does it seem like it would be possible with these approaches. For example, applying the Ultra-tight TET system for KO of CD81 was performed after 10 days but the rationale for this timepoint was not explained. Does it take 10 days of KO to see quantifiable loss of CD81 on the cell surface? How soon after drug treatment(s) are there measurable effects? With these kinetics, how would it be feasible to do a genomics or proteomics study to accurately define protein function?\n5) What are the cell line limitations of these systems? Most of the cell lines used in this study are easy to manipulate. How efficient are these strategies in more difficult to manipulate cancer or normal cells?\n6) The approaches would be valuable if they could improve utility of in vivo studies, which can be a severe limitation of existing approaches due to leakineess. The authors should comment on in vivo potential and provide proof-of-concept support that their advance would be impactful in these settings.\nMinor points:\n1) There is inconsistency throughout the figures where non-significance or significance calculations are noted. The authors more clearly clarify what is significant or non-significant in each panel, either in the legends or figure itself.\n2) For flow based experiments, necessary controls such as IgG or evaluations in non-CD81 lines are not shown and would be helpful for readers to evaluate their results.", "reviewer_2": "In their manuscript, Srinivasan, Sun, and co-authors present a system for chemically controlled expression and activity of CRISPR-Cas9 in mammalian cells. The authors initially employed the conventional reverse tetTA system for Dox-dependent Cas9 expression in standard mammalian cell lines but observed significant leakiness, i.e. Cas9 activity in the non-induced state. To address this issue, they explored various strategies, identifying the most effective approaches as (i) destabilizing Cas9 with a degradation domain and (ii) co-expressing a drug-dependent AcrIIA4 as a conditional Cas9 antagonist.\nFollowing genomic integration of this multi-component system, the authors demonstrated strong drug-mediated control of Cas9 activity, as evidenced by antibody-based detection of a Cas9-targeted cell surface antigen via flow cytometry. They also tested an alternative system based on Branaplam-induced splicing. While generally effective, this approach exhibited strong cell-type dependence. Finally, the authors extended their strategy from catalytically active Cas9 to CRISPRi using dCas9-ZIM3, showcasing regulation of cell growth by targeting the PLK1 promoter.\nOverall, this is a well-executed study and a well-written manuscript. However, I am divided on its suitability for publication in Nature Communications. While the authors successfully integrate multiple existing control strategies into a single system, the conceptual novelty remains overall limited. Furthermore, although the \"ultra-tight\" system has potential applications in genetic screening of essential genes and similar application scenarios, the manuscript does not convincingly demonstrate its superiority over alternative methods, such as chemically inducible split-Cas9 regulators (see extensive literature on chemical CRISPR-Cas9 control).\nIn my view, these limitations currently preclude publication in Nature Communications. However, if the authors can provide compelling evidence that their system offers a substantial performance advantage over the state-of-the-art in chemical CRISPR-Cas9 control and/or demonstrate a practical application, such as screening for gene essentiality in a growing organoid or a developing model organism embryo, I would be fully open to reconsidering my assessment.\nAdditionally, I offer the following specific comments for improvement:\n1. Time-course experiment: The authors typically assess CRISPR-mediated knockdown only after 5-10 days of drug treatment, i.e. when editing is saturated. It is crucial to include a time-course experiment with samples taken at multiple time points (e.g., days 1, 2, 3, 5, and 10) to compare the genome editing kinetics of the ultra-tight CRISPR-Cas9 system against the conventional pTet CRISPR-Cas9. Given the nature of the ultra-tight system, its kinetics could be slower than the conventional pTet due to an overall diminished Cas9 activity, which could limit its utility for precisely timed gene knockouts.\n2. A direct comparison between the ultra-tight system and leading strategies for chemical Cas9 control, e.g. inducible split-Cas9 regulators, would be highly informative.\n3. The authors predominantly assess genome editing indirectly, i.e. through cell surface protein expression. To exclude potential artifacts, they should validate editing at the genomic level as well (e.g. via amplicon sequencing of the Cas9-targeted locus) for at least a subset of experiments, e.g. using 3–4 sgRNAs across 2–3 cell lines.\n4. Including a constitutively expressed Cas9 (i.e. not Dox/drug-dependent) as a control would be important, particularly for the time-course experiment mentioned in point 1, so assess effects the drugs could have on Cas9 function/genome editing in general (i.e. irrespective of the author’s regulatory modules).\nMinor points to address:\n* Page 3: \"separated by a T2A linker\" → be more precise, i.e. T2A mediates ribosome skipping, resulting in the expression of two separate polypeptides\n* Last paragraph of the introduction: The statement beginning with \"Together, these tools offer…\" should be moderated, especially given the lack of direct comparison with state-of-the-art methods.\n* Figure 2 legend title: The phrase “ultra-tight pTET system that is tight” should be revised for clarity.\nFinally, while I have reservations about its suitability for Nature Communications, I acknowledge the strengths of this manuscript and the potential use of the ultra-tight system. If not accepted here, I am confident it would be well-suited for a well-recognized, more specialized journal."}, "author_response": "General responses to all reviewers:\nWe sincerely thank the editor and reviewers for their thorough evaluation of our\nmanuscript and for the constructive feedback provided. The comments raised important\npoints that prompted several additional experiments, which we believe have substantially\nimproved the quality and rigor of the manuscript while also broadening the applicability of\nour new technology platforms. Notable new results now included are as follows: (i)\nMeasurement of gene editing at the genomic level by NGS for two different cell lines and\ntwo different target loci highlighting the reliability of flow cytometry assays to assess Cas9\nfunctionality (Fig. 2J, S3B-C, S4C, S5, S6E) (ii) Time course evaluation of editing kinetics\nat protein and genomic level for both the pTET:Ultra-tight and Xon-Cas9;AcrIIA4-FKBP\nsystems (Fig. 2I-J, 6D, S6E), (iii) Evaluation of pTET:Ultra-tight in an iPSC line\ndemonstrating expanded applicability in complex cell types (Fig. 3A-B, S4D-E), and (iv)\nWestern immunoblotting experiments demonstrating stabilization/destabilization of Cas9\nin pTET vs. pTET:Ultra-tight (Fig. S3A). Below we provide a point-by-point response to\neach of the reviewers’ comments.\nPoint-by-point responses to each Reviewer:\nReviewer #1 (Remarks to the Author):\nSrinivasan and colleagues developed a collection of improved CRISPR systems to\novercome the limitations of exciting approaches. The authors developed an inducible\nand degron-destabilized Cas9 expression systems, applied degradation approaches to\nanti-CRISPR proteins, and developed a Branaplam-sensitive splice switch to control the\ngenetic editing machinery. The approaches minimize unintended editing in the off-state\nwhile maximizing editing efficiency in the on-state. Overall, this is a valuable toolbox for\nthe community for CRISPR editing experiments. However, I have several comments for\nthe authors to address as the characterization, applications, and specificity of this study\nare incomplete:\nMajor points:\n1) The dosing and timing of compounds used in this study are unclear. Demonstration\nof their activity in stabilizing or destabilizing the proteins are also not shown, making it\ndifficult to understand the experimental design in the experiments and the\ncomprehensiveness of evaluation. The authors should show stabilization/destabilization\nof their control modules and provide data to support the dosing that is selected for\nexperiments in this study.\nWe appreciate the reviewer’s comments. In response to the question about dosing:\na) All of the small molecules used in this study for inducing expression or for\nmodulating protein stability have been extensively characterized and published\npreviously (PMID: 31101683, PMID: 31959800, PMID: 32994412,\nhttps://www.takarabio.com/learning-centers/gene-function/gene-\nediting/crisprcas9-delivery-methods/tet-inducible-cas9-for-gene-\nediting?srsltid=AfmBOoomRdgFVpxsgPgmWSfopv7wnlgm2LcYqXE5_p4UGyhP\nHNefkSfY, PMID: 32948771, PMID: 39504960, PMID: 21725303, PMID:\n20851347, PMID: 16959577, PMID: 33795428, PMID: 34321659). In general, we\nreferred to the published data for our dosage choice. Additionally, we performed\ndose titration experiments using various reporter cell lines (generated by our lab)\nto confirm or fine-tune published concentrations. The resulting data, combined\nwith information from published studies, were then applied across the broader\npanel of cell lines used in all experiments shown for the current study. We detail\nour optimization process and the results in the following section.\nb) To determine an optimal concentration for doxycycline (Dox), we performed a\ndose-response analysis using a stable, dual-reporter system in HEK293 cells\n(Reviewer Figure 1). In this context, Firefly luciferase (FLuc) is expressed from\nthe doxycycline-inducible pTET promoter, serving as a readout for transcriptional\nactivation, while Nanoluciferase (NLuc) is constitutively expressed from the\nEF1s promoter, serving as an internal normalization control. Cells were treated\nwith the indicated concentrations of Dox for 48 hours, and fold induction of\nnormalized FLuc was determined. A concentration of 2000 ng/mL Dox yielded\nmaximal induction. This dosage aligns with concentrations used in previous\nstudies across various cell lines (PMID: 31959800, PMID: 31101683 and PMID:\n32994412) and was therefore selected for subsequent experiments.\nTo assess stabilization/destabilization via the FKBP degron tag, we generated\nstable DLD1 and HCT116 cell lines expressing a FLuc reporter either untagged\nor fused to the FKBP degron, rendering it degradable by addition of the dTAGV-1\nligand (Reviewer Figure 2). NLuc was co-expressed from the same transcript via\nan IRES element to serve as an internal control. Cells were treated with the\nindicated concentrations of dTAGV-1 for 24 hours, and normalized FLuc activity\nrelative to vehicle control cells was determined. Treatment with 1000 nM dTAGV-\n1 resulted in maximal reporter degradation in both cell lines. This concentration\nwas consistent with doses used in prior studies across different cell types (PMID:\n32948771, PMID: 39504960, PMID: 33795428) and was therefore selected for\ndownstream experiments. FLuc activity in untagged cells remained relatively\nstable at all concentrations.\nTo evaluate the functionality of destabilizing domains (DD and DHFR), we\ngenerated stable DLD1 cell lines expressing FLuc fused to either the DD or\nDHFR tag, resulting in constitutive degradation under basal conditions (Reviewer\nFigure 3). Untagged FLuc constructs were included as controls. FLuc was co-\nexpressed with NLuc via an IRES element to enable internal normalization. Cells\nwere treated for 24 hours with the indicated concentrations of TMP or Shield1,\nand normalized FLuc activity was determined relative to untagged vehicle-treated\ncontrol samples. Ligand addition resulted in dose-dependent stabilization of the\ntagged reporters as evident from increasing FLuc activity. Based on these results\n2 µM TMP and 1000 nM Shield1 were selected for use in subsequent\nexperiments, and these doses were within the concentration range reported in\nprevious publications in various cell lines (PMID: 21725303, PMID: 20851347,\nPMID: 1695957).\nIn order to determine appropriate Branaplam concentration for inducibility\nassays, stable K562 cells were generated where a GFP reporter was placed\nunder control of the Xon splice switch (Reviewer Figure 4). A dosage titration of\nBranaplam was performed in the K562 cell line. GFP induction and cell viability\n(via staining of 7-AAD, Thermo Fisher Scientific) were analyzed by flow\ncytometry. A range of 120-250nM of Branaplam demonstrated high induction with\nminimal impact on cell viability. This dosage range was comparable to\nconcentrations used in prior studies with this drug (PMID: 34321659, PMID:\n35241644), and was selected for subsequent experiments in K562 cells. We\nselected a lower concentration (20nM) based on the previous study (PMID:\n34321659), for HEK293T treatment.\nIn summary, we referred to both previously published literature and our pilot\nreporter assay to select the dose of each inducer drug.\nc) Please see our detailed response in point 4, which addresses a point raised by\nthe reviewer about the timing of treatment.\nd) To address the comment regarding stabilization and destabilization of the control\nmodules, we performed immunoblotting to detect Cas9 protein in treated and\nuntreated HEK293T cells expressing either the pTET or pTET:Ultra-tight systems\n(Fig. S3A). In pTET-expressing cells, Cas9 was undetectable in the absence of\ndoxycycline but was strongly induced following treatment. Notably, untreated\npTET cells still exhibited ~50% CD81 knockout by flow cytometry (Fig. 2E),\ndespite the absence of a detectable Cas9 band. This suggests that Cas9 may\nhave been expressed at levels below the detection threshold of our assay, or in a\ntransient or heterogeneous manner, yet still sufficient to drive substantial genome\nediting. This is plausible given the potency of Cas9 when programmed with our\nCD81 guide RNA and the low copy number of the CD81 locus (~3 copies per cell\ngiven HEK293T is hypotriploid). These observations highlight a limitation of\nimmunoblotting in this context: low-level Cas9 expression may evade detection\nwhile remaining functionally significant. A functional assay that measures\ndepletion of the target protein, as we demonstrated in the manuscript, would\ntherefore be a more stringent approach.\nFor the pTET:Ultra-tight system, we analyzed Cas9 expression across untreated,\nsingle-drug-treated, and double-drug-treated conditions. As expected, Cas9 was\nnot detected in untreated cells due to active degradation by the DD tag. Cas9\nalso remained undetectable in cells treated with Shield1 alone, which may reflect\npartial stabilization of leaky Cas9 expression that remains below the detection\nthreshold—similar to what was observed in untreated pTET cells. A weak Cas9\nband was observed with doxycycline alone, while a stronger signal was detected\nwith combined doxycycline and Shield1 treatment. This result is consistent with\ndox-induced expression of Cas9-DD and Shield1-mediated stabilization of the\nprotein.\nWe also attempted to detect the anti-CRISPR protein AcrIIA4, which is regulated\nby the LID degron in the pTET:Ultra-tight system. Despite testing both\ncommercially available antibodies (Abcam, Diagenode), we were unable to\ndetect AcrIIA4 by immunoblotting. However, flow cytometry data for target\ndepletion aligned with the expected behavior of the LID-degron-tagged or the\nFKBP degron-tagged AcrIIA4 in response to Shield1 or dTagV-1 treatment,\nrespectively (Fig. 2G, 6B). Specifically, AcrIIA4 expression reduces background\nediting in untreated cells but also limits the extent of CD81 knockout. Addition of\nthe appropriate degron ligand restores maximal editing, consistent with the\nmodule functioning as designed.\n2) In Figure 2 ,3 and 5, while the authors show flow cytometry data for their target\nexpression, editing of the genomic locus should also be shown to help verify their\nobservations and efficiencies.\nTo address the reviewer’s comment, we performed amplicon sequencing by NGS for\ntwo different targets (CD81 and B2M; Fig. 2J. S3C and S5 ), before and after induction\nof Cas9 activity, in two different cell lines (HEK293T and HAP1; Fig. 2J, S3C and S4C).\nWe found that the KO efficiency as measured by NGS/indel rate mirrored KO efficiency\nmeasured by flow cytometry. In addition, we also performed targeted genomic\nsequencing as part of the newly-included time course experiments for pTET,\npTET:Ultra-tight, Xon-Cas9, Xon-Cas9;AcrIIA4-FKBP (Fig. 2J, S6E). We observed strong\nagreement in the kinetic profiles between the protein-level (via flow cytometry) and indel\ngeneration dynamics. Interestingly, we found that indel formation preceded detectable\nloss of CD81 protein. This corresponds to the expected delay between genomic\ndisruption and mRNA/protein turnover. Collectively, these new results support using\nflow cytometry as a quantifiable and robust readout for editing efficiency.\n3) In Figure 4, the characterization of genomic and protein level disruption is insufficient\nto show PLK1 or H3-3A is altered and to what extent. The consequences of the H3-3A\ndisruption should also be evaluated to show a phenotypic effect. Is there a biological\nobservation the authors can now achieve that was not evident with prior approaches?\nMost critically, rescue experiments are not provided to show that these effects are on-\ntarget and need to be performed with a CRISPR-resistance expression strategy. This\nwill be necessary for the author’s conclusions and is the only biological data shown in\nthe paper.\nWe thank the reviewer for this thoughtful comment. The experiments presented in\nFigure 4 were designed to evaluate the performance of the ultra-tight inducible CRISPR\nsystem in scenarios where conventional systems are limited by leaky effector\nexpression—specifically, in the context of essential gene targeting (PLK1) and\nmultiplexed guide dropout strategies (H3-3A). Our aim was to demonstrate the system’s\ncapacity to prevent premature selection and maintain balanced multi-guide activity,\nrather than to characterize the downstream biological consequences of gene disruption.\nWe agree that comprehensive phenotypic analyses and rescue experiments are critical\nfor establishing gene-specific mechanisms. However, such studies are not required to\nsupport the conclusions of this work, which focus on the development and validation of\nan improved inducible genome editing platform. The observed editing at both the\ngenomic and protein levels, in conjunction with the functional outcomes shown,\nsufficiently illustrate the system's utility for enabling these types of experiments.\nWhile we appreciate the reviewer’s interest in further biological investigation, we\nrespectfully consider this to be beyond the scope of the current study, which is centered\non methodological advancement. We believe the presented data provide a robust\ndemonstration of the system’s intended function and its potential to support future\nmechanistic studies where temporal control is essential.\n4) The authors comment that temporal studies are feasible with these approaches.\nHowever, this is not demonstrated, nor does it seem like it would be possible with these\napproaches. For example, applying the Ultra-tight TET system for KO of CD81 was\nperformed after 10 days but the rationale for this timepoint was not explained. Does it\ntake 10 days of KO to see quantifiable loss of CD81 on the cell surface? How soon after\ndrug treatment(s) are there measurable effects? With these kinetics, how would it be\nfeasible to do a genomics or proteomics study to accurately define protein function?\nWe thank the reviewer for raising this important point. Our initial rationale for using a 10-\nday endpoint was to ensure maximal gene editing and protein depletion, while also\ncapturing the cumulative effects of any leaky Cas9 expression; particularly relevant\nwhen comparing inducible systems. This rationale has now been clarified in the revised\nmanuscript.\nIn response to the reviewer’s comment, we performed more extensive time-course\nexperiments to assess the kinetics of CD81 disruption using the pTET, pTET:Ultra-tight,\nXon-Cas9, and Xon-Cas9;AcrIIA4-FKBP systems (Fig. 2I–J, 6D, S6E). With the pTET\nsystem, ~50% CD81 loss was detected throughout the time course, consistent with\nsubstantial basal Cas9 activity. By contrast, CD81 knockout in the pTET:Ultra-tight\nsystem was only observed after inducing expression of and stabilizing Cas9, with\nminimal editing at early time points and progressive loss of CD81 between 24 and 72\nhours. With this vector system and target gene, editing and protein loss plateaued by\nday 6.\nAmplicon sequencing confirmed that indel formation preceded protein depletion, as\nexpected due to mRNA/protein turnover kinetics. Importantly, our NGS results\ncorrelated well with the flow cytometry data, supporting the reliability and consistency\nbetween both genomic and protein-level readouts (Fig. 2J, S6E).\nIn the Xon-Cas9 systems, editing began within the first few days and increased through\nday 10. In this context, the AcrIIA4-FKBP variant demonstrated substantially reduced\nbackground editing albeit with slower activation kinetics, but still reached near-maximal\nediting by the final timepoint (Fig. 6D, S6E).\nTo summarize, while 10 days of treatment ensures complete (or near complete) editing\nand may be suitable for endpoint ‘omics experiments, earlier timepoints (e.g., 48–72\nhours) were sufficient to observe strong target protein depletion in many contexts. The\noptimal duration will depend on the target gene, protein half-life, and experimental\nobjectives. Notably, the pTET:Ultra-tight system enabled controlled gene disruption with\nnear-absent background editing, making it well suited for applications that require\nprecise and exquisitely regulated temporal resolution. We are appreciative of the\nreviewer for encouraging this line of experimentation, since the ability to measure loss-\nof-function phenotypes before a day 10 time point would open up broader use cases for\nour technology platform.\n5) What are the cell line limitations of these systems? Most of the cell lines used in this\nstudy are easy to manipulate. How efficient are these strategies in more difficult to\nmanipulate cancer or normal cells?\nWe appreciate the reviewer’s question and the broader comment regarding the\nimportance of testing these systems in more technically demanding cell types. However,\nwe respectfully note that the ease or difficulty of manipulating a given cell line is highly\ncontext-dependent. What is routine in one lab may be considerably more challenging in\nanother, depending on available protocols, infrastructure, and expertise.\nTo more directly address this concern, we evaluated both pTET and pTET:Ultra-tight in\nthe KOLF2.1J iPSC line. While increasingly used as a reference line due to its well-\ncharacterized behavior and differentiation capacity (PMID: 36459969), KOLF2.1J iPSCs\nare generally more sensitive to transduction and genome editing than transformed lines.\nIn this context, pTET:Ultra-tight maintained excellent performance (in fact, it\ndemonstrated the highest on vs. off-target editing efficiency of any cell type in our\npanel): background editing observed with pTET (~10%) was fully eliminated, and ~80%\nknockout efficiency was achieved upon induction (Fig. 3A,B). Importantly, transduction\nand treatment did not disrupt the undifferentiated state, as evidenced by maintained\niPSC morphology and high SSEA-4 expression (Fig. S4D,E).\nThese results demonstrate that the pTET:Ultra-tight system offers strong temporal\ncontrol and robust activity even in sensitive and difficult-to-engineer models, supporting\nits broader applicability across diverse research settings.\n6) The approaches would be valuable if they could improve utility of in vivo studies,\nwhich can be a severe limitation of existing approaches due to leakiness. The authors\nshould comment on in vivo potential and provide proof-of-concept support that their\nadvance would be impactful in these settings.\nWe thank the reviewer for this thoughtful comment. We agree that in vivo studies could\nsignificantly benefit from the enhanced control offered by our ultra-tight systems,\nparticularly in overcoming the challenges posed by leaky gene expression. However,\nconducting in vivo proof-of-concept experiments (especially in transgenic mouse\nmodels) would require extensive optimization and significant resource investment,\nwhich is beyond the scope of this current study.\nThat said, there is strong precedent supporting the in vivo potential of our systems. The\nmajority of the regulatory modules incorporated into our ultra-tight pTET and Xon-\nCas9;AcrIIA4-FKBP circuits have been successfully deployed in mouse models. For\nexample, the Tet-ON system has been widely used in vivo and is reviewed in Godecke\net al. (PMID: 19394373). Similarly, the control modules - DD/Shield1, LID/Shield1,\nFKBP/dTAGv1, and Xon/Branaplam - have all demonstrated functional efficacy in murine\nsystems (e.g., PMID: 26757616, 18836461, 32948771, 32559430, 36097386,\n34321659, and 18836461). Importantly, the pharmacological inducers, Dox and\nShield1, used within the pTET:Ultra-tight system, as well as Branaplam, are bioavailable\nand have been shown to penetrate the blood–brain barrier (PMID: 30007162,\n26757616, and 35241644).\nCollectively, these provide strong rationale for the potential applicability of our ultra-tight\ninducible systems in vivo, and we view this as an exciting area for future investigation.\nMinor points:\n1) There is inconsistency throughout the figures where non-significance or significance\ncalculations are noted. The authors more clearly clarify what is significant or non-\nsignificant in each panel, either in the legends or figure itself.\nWe appreciate the reviewer’s attention to clarity in figure presentation. In instances\nwhere multiple groupings are shown, we focused statistical comparisons on the most\nrelevant and informative conditions to maintain figure readability and highlight key\ndifferences. For these selected comparisons, significance has been explicitly indicated\nin the appropriate figure panels and legends. If the reviewer has specific panels where\nadditional comparisons would be helpful, we would be happy to incorporate them.\n2) For flow based experiments, necessary controls such as IgG or evaluations in non-\nCD81 lines are not shown and would be helpful for readers to evaluate their results.\nWe appreciate the reviewer’s suggestion and added the IgG control as well as a CD81-\nnull cell line for the flow cytometry data. The result is in Figure S1A.\nReviewer #2 (Remarks to the Author):\nIn their manuscript, Srinivasan, Sun, and co-authors present a system for chemically\ncontrolled expression and activity of CRISPR-Cas9 in mammalian cells. The authors\ninitially employed the conventional reverse tetTA system for Dox-dependent Cas9\nexpression in standard mammalian cell lines but observed significant leakiness, i.e.\nCas9 activity in the non-induced state. To address this issue, they explored various\nstrategies, identifying the most effective approaches as (i) destabilizing Cas9 with a\ndegradation domain and (ii) co-expressing a drug-dependent AcrIIA4 as a conditional\nCas9 antagonist.\nFollowing genomic integration of this multi-component system, the authors\ndemonstrated strong drug-mediated control of Cas9 activity, as evidenced by antibody-\nbased detection of a Cas9-targeted cell surface antigen via flow cytometry. They also\ntested an alternative system based on Branaplam-induced splicing. While generally\neffective, this approach exhibited strong cell-type dependence. Finally, the authors\nextended their strategy from catalytically active Cas9 to CRISPRi using dCas9-ZIM3,\nshowcasing regulation of cell growth by targeting the PLK1 promoter.\nOverall, this is a well-executed study and a well-written manuscript. However, I am\ndivided on its suitability for publication in Nature Communications. While the authors\nsuccessfully integrate multiple existing control strategies into a single system, the\nconceptual novelty remains overall limited. Furthermore, although the \"ultra-tight\"\nsystem has potential applications in genetic screening of essential genes and similar\napplication scenarios, the manuscript does not convincingly demonstrate its superiority\nover alternative methods, such as chemically inducible split-Cas9 regulators (see\nextensive literature on chemical CRISPR-Cas9 control).\nIn my view, these limitations currently preclude publication in Nature Communications.\nHowever, if the authors can provide compelling evidence that their system offers a\nsubstantial performance advantage over the state-of-the-art in chemical CRISPR-Cas9\ncontrol and/or demonstrate a practical application, such as screening for gene\nessentiality in a growing organoid or a developing model organism embryo, I would be\nfully open to reconsidering my assessment.\nWe thank the reviewer for this thoughtful critique and the opportunity to clarify what we\nbelieve to be the core novelty and significance of our study. While we agree that\nindividual regulatory components used in our system have been described previously,\nwe respectfully submit that the design, integration, and functional optimization of these\nelements into a modular and highly tunable system represent a novel contribution to the\nfield.\nA key innovation lies in the incorporation of a drug-regulated anti-CRISPR protein\n(AcrIIA4) as a repressive module that works in parallel with inducible Cas9 expression.\nWhile anti-CRISPRs (AcrIIA4) themselves are not new, their use as conditionally\nregulated antagonists within a multi-tiered control system (with separate, tunable\nexpression from Cas9) is, to our knowledge, unique. This unique module is essential for\nachieving both undetectable background nuclease activity and robust induced editing,\nand provides advantages over approaches that rely solely on inducible Cas9\nexpression/stabilization.\nWe also designed the system to exploit the opposing effects of Shield-1 on two\nregulatory domains (DD and LID), enabling simultaneous stabilization of Cas9 and\ndestabilization of AcrIIA4. This coordinated use of a single small molecule to exert dual,\nopposing control over key effectors adds another layer of functionality that has not been\ndemonstrated in other Cas9 regulatory systems.\nOur experiments further show that not all combinations of known components yield\ndesirable outcomes. For example, pairing the Branaplam-inducible Xon system with\ndegradation domains resulted in either minimal improvement or substantial leakiness,\nand combinations of the Xon module with a Tet-responsive promoter significantly\nreduced inducibility. These observations reinforce the point that system performance\ndepends not only on component selection but on thoughtful architectural design, timing,\nand orthogonality between modules.\nWith respect to the reviewer’s reference to split-Cas9 strategies, we agree that such\nsystems have been successfully deployed, particularly in contexts where transgene size\nis limiting (e.g., AAV delivery) or in specialized applications such as photo-inducible\ngene editing. However, split-Cas9 approaches are less widely used for general in vitro\napplications (and to our knowledge are rarely used for high-throughput or screening\ncontexts) where simplicity, high efficiency, and compatibility with standard delivery\nformats are essential. Our system was specifically designed with these broader, cell-\nbased applications in mind.\nIn summary, the overall novelty of our system lies in the rational integration of\ncomplementary regulatory modules, Cas9, anti-CRISPR (new from our study), and dual-\nuse degradation domains, into a unified, high-performance platform with consistent\nresults across all tested contexts. We hope this perspective clarifies the unique\ncontribution of our work and its relevance to a broad range of experimental and\ntherapeutic applications.\nAdditionally, I offer the following specific comments for improvement:\n1. Time-course experiment: The authors typically assess CRISPR-mediated knockdown\nonly after 5-10 days of drug treatment, i.e. when editing is saturated. It is crucial to\ninclude a time-course experiment with samples taken at multiple time points (e.g., days\n1, 2, 3, 5, and 10) to compare the genome editing kinetics of the ultra-tight CRISPR-\nCas9 system against the conventional pTet CRISPR-Cas9. Given the nature of the\nultra-tight system, its kinetics could be slower than the conventional pTet due to an\noverall diminished Cas9 activity, which could limit its utility for precisely timed gene\nknockouts.\nWe appreciate the reviewer’s suggestion on the time-course experiment. We picked 5-\n10 days of treatment to ensure drug-induced Cas9 activity and target depletion are\ncomplete based on our empirical results and to avoid underestimating leaky editing at\nearlier time points.\nWe agree with the reviewer that a more detailed time course comparison across\nsystems will benefit users of this technology. To further understand the kinetics of\ninducible Cas9, we performed a time course experiment to measure the rate of CD81\ntarget editing before day 10 (6, 24, 48, 72, 144hrs) for pTET, pTET:Ultra-tight, Xon-Cas9,\nand Xon-Cas9;AcrIIA4-FKBP in HEK293T (Fig. 2I, 6D). In addition to flow cytometry to\nmonitor loss of CD81 protein, we also measured the CD81 indel rate by amplicon\nsequencing (Fig. 2J, S6E).\nThe conventional pTET system achieves saturating indel levels faster than pTET:Ultra-\ntight (24 vs. 48hrs), however, this is due to the significant leaky editing rate of pTET.\nLoss of CD81 as measured by flow cytometry lags behind indel formation as expected.\nThere is no sacrifice in editing rate for pTET:Ultra-tight. In fact, due to the reduction in\nleaky editing, the induced target disruption appears faster for pTET:Ultra-tight than\npTET. By contrast, the Xon-Cas9;AcrIIA4-FKBP system had a slower overall editing rate\nand when compared to Xon (Fig. 6D, S6E).\nTo summarize, marginal differences in editing efficiency (indel formation and/or protein\ndepletion) were observed between pTET and pTET:Ultra-tight. However, a substantial\ndegree of editing was observed with pTET at the earliest measured time points, and\nsubtraction of this population showed that pTET:Ultra-tight was perhaps more active\nupon induction. A plausible explanation for this is that the non-leaky pTET cells\nintegrated the transgene into a less-expressive site, which both minimizes background\nand induced expression. Separately, the Xon-Cas9;AcrIIA4-FKBP system, despite near-\nabsent background, did present slower kinetics compared to Xon. Given that the actual\nkinetics could vary between cell types and targets, we recommend researchers perform\na similar time course experiment for their specific context.\n2. A direct comparison between the ultra-tight system and leading strategies for\nchemical Cas9 control, e.g. inducible split-Cas9 regulators, would be highly informative.\nWe thank the reviewer for this comment. We agree that direct comparisons across\ntechnologies are important and we have included performance benchmarking against\nthe most commonly-applied chemically-inducible Cas9 systems, specifically, dox-\ninducible and degron-modified Cas9, throughout the manuscript.\nThere are indeed several chemically-inducible Cas9 systems previously described\nincluding split Cas9. As noted above, split Cas9 is more conventionally used where\ntransgene size is limiting (as in the case of AAV vector–based delivery, which we have\npublished on previously; PMID: 36419469) or for niche applications like those requiring\nphoto-inducible control of gene editing. That said, we were able to identify studies that\nuse chemical-inducible control of one or both split Cas9 elements. While these\napproaches showed reduced background editing, the reported knockout efficiencies\nrange from ~20% to ~60% at best, whereas we observe ~80-95% KO efficiency with our\nultra-tight systems in those same cell lines (see table below for a summary).\nReferences: PMID: 27363581, PMID: 25643054, PMID: 35615005, PMID: 32994412,\nPMID: 25848930, PMID: 27618190.\nStudy Strategy Chemical Cell Line Best\ninducer %editing\nPMID: 27363581 Split Cas9 4OHT HEK293 ~25%\nPMID: 25643054 Split Cas9 rapamycin HEK293FT ~40%\nPMID: 35615005 Dual control Dox + Shield1 H9 ESC ~40%\nPMID: 32994412 Safe harbor Dox Multiple ~60%\nintegration\nPMID: 25848930 Intein-Cas9 4OHT HEK293 ~15%\nPMID: 27618190 ERT2-Cas9 4OHT HEK293 ~25%\nGiven this reviewer’s interest in split Cas9, in particular, they are likely well aware of\ndifferences in overall Cas9 efficiency based on expression levels and their control\nmechanisms for each half of the split Cas9, amino acid position of the split, intein\nsequences, chemical dimerization strategies, etc. We feel this is an exciting area of\nresearch, but a direct comparison of all of the split Cas9 methods, and optimization of\nany one of them for use across multiple cell contexts, represents a full study on its own\nand is beyond the scope of this manuscript. We have updated the Discussion section to\nsummarize the above information, including split Cas9.\n3. The authors predominantly assess genome editing indirectly, i.e. through cell surface\nprotein expression. To exclude potential artifacts, they should validate editing at the\ngenomic level as well (e.g. via amplicon sequencing of the Cas9-targeted locus) for at\nleast a subset of experiments, e.g. using 3–4 sgRNAs across 2–3 cell lines.\nWe thank the reviewer for this suggestion. Reviewer 1 raised a similar point, and we\naddressed this in response to their comment above. To summarize here, we performed\ntargeted genome sequencing before and after chemical induction of Cas9 of two\ndifferent targets CD81 and B2M and for two different cell types HEK293T and HAP1\n(Fig. 2J, S3C, S4C, S5). In both cell types as well as at both target loci, we found strong\nagreement between KO efficiency measured by indel rate and flow cytometry.\nIn addition, we now include targeted genomic sequencing results for the newly-included\ntime course experiments with pTET, pTET:Ultra-tight, Xon-Cas9, Xon-Cas9;AcrIIA4-\nFKBP (Fig. 2J, S6E). We found that KO efficiency by indel rate and flow cytometry were\nwell-aligned at the measured time points. As discussed in more detail above in\nresponse to the comment form Reviewer 1, we observed that indel formation preceded\nthe loss of CD81 protein.\n4. Including a constitutively expressed Cas9 (i.e. not Dox/drug-dependent) as a control\nwould be important, particularly for the time-course experiment mentioned in point 1, so\nassess effects the drugs could have on Cas9 function/genome editing in general (i.e.\nirrespective of the author’s regulatory modules).\nTo this reviewer’s point, we now include an experiment with constitutively expressed\nCas9 as a positive control for gene disruption. In brief, to be consistent with the rest of\nour designs, we generated a piggyBac construct with a CAG promoter-driven Cas9, a\nsgRNA targeting CD81 and a puromycin selective marker (Fig. S1A). We generated the\ncell line via piggyBac transposition and selected stable cells with puromycin for 7 days.\n24 hours after puro removal, we observed that CD81 KO reached near 100%, which\nstayed the same until day 10 (Fig. S3B-C). Stable line generation and puromycin\nselection take several days. Given that Cas9 and sgRNA were co-expressed in the\ncells, it is not surprising that KO reached completion by the time the constitutive Cas9\nline was established.\nMinor points to address:\n* Page 3: \"separated by a T2A linker\" → be more precise, i.e. T2A mediates ribosome\nskipping, resulting in the expression of two separate polypeptides\nThank you. For clarity we have modified the text to “each separated by a T2A sequence\nto enable co-expression as distinct polypeptides”\n* Last paragraph of the introduction: The statement beginning with \"Together, these\ntools offer…\" should be moderated, especially given the lack of direct comparison with\nstate-of-the-art methods.\nWe thank the reviewer for flagging this and have modified the text accordingly to\n“Together, these tools offer a marked contrast between induced and background Cas9\nactivity…”\n* Figure 2 legend title: The phrase “ultra-tight pTET system that is tight” should be\nrevised for clarity.\nWe appreciate this and agree with the reviewer. The Figure 2 legend title has been\nchanged to “Development of an “ultra-tight” pTET system”."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors addressed several of my concerns during their revision. I appreciated the information that they provided on their rationale for dosing selection and experimental design. My suggestion would be to provide a summary of rationale for potential users, such that there is some detail then included in the manuscript or methods, to help rationalize for others that may be interested in these approaches. I also appreciated the time-course experiments that are now included.\nAdditionally, several major points were not addressed that would elevate the study such as biological application, rescues, in vivo applications, or clear comparisons to existing approaches (pointed out by Reviewer 2). At the very least, the authors should discuss some of these points as limitations of their study in the discussion.", "reviewer_2": "The authors have satisfactorily addressed my comments. Therefore, I now gladly recommend this manuscript for publication in Nature Communications. Congratulations to the authors on this excellent piece of work.\nMinor comment:\nOn page 4: “Similar to flow cytometry results, upon induction, As before, pTET:Ultra-tight cells look longer than pTET cells to achieve maximal editing” Revise as appropriate"}, "author_response": "Point-by-point response to reviewers:\nWe thank the reviewers for their thoughtful evaluations and constructive feedback. Below we\nprovide a point-by-point response, with reviewer comments in bold and our responses in plain\ntext. All textual changes in the manuscript are highlighted in the revised version.\nReviewer #1\nThe authors addressed several of my concerns during their revision. I appreciated the\ninformation that they provided on their rationale for dosing selection and experimental\ndesign. My suggestion would be to provide a summary of rationale for potential users, such\nthat there is some detail then included in the manuscript or methods, to help rationalize for\nothers that may be interested in these approaches. I also appreciated the time-course\nexperiments that are now included.\nWe thank the reviewer for this feedback. In response, we have added a new Methods subsection\n(“Small molecule dosing rationale”) that summarizes the inducer concentrations used\nthroughout (Dox 2 µg/mL, Shield1 1 µM, dTAGv-1 1 µM, trimethoprim 2 µM, and Branaplam\n20–150 nM depending on cell type). We explain that these doses were selected from internal\ntitration assays to maximize induction or degradation while minimizing cytotoxicity, and that\nthey align with previously reported ranges.\nIn addition, we have inserted a clarifying statement in the Results (Fig. 2 introduction) that\nreferences these concentrations and explicitly defines “dynamic range” as the fold difference\nbetween residual editing in the OFF state and the efficiency achieved in the ON state.\nAdditionally, several major points were not addressed that would elevate the study such as\nbiological application, rescues, in vivo applications, or clear comparisons to existing\napproaches (pointed out by Reviewer 2). At the very least, the authors should discuss some\nof these points as limitations of their study in the discussion.\nWe agree with the reviewer that these aspects represent important and potential future extensions\nof the present work. In response, we have expanded the Discussion to include:\n1. A new comparisons section that discusses how pTET:Ultra-tight relates to split/intein-\nCas9 and ERT2-Cas9 systems, highlighting differences in editing efficiency, background\nactivity, and compatibility with high-throughput applications.\n2. A new Supplementary Table S7 that summarizes reported efficiencies of representative\nsplit/intein Cas9 systems side-by-side with those achieved by pTET:Ultra-tight.\n3. A new Limitations and future directions paragraph within the Discussion section,\nwhich explicitly acknowledges that:\no We did not include phenotypic characterization or rescue experiments, which will\nbe important for future biological studies.\n1\no Although the modules employed (Tet/Dox, DD/Shield1, FKBP/dTAG,\nXon/Branaplam) have demonstrated functionality in vivo, we did not conduct\ndirect animal studies in this manuscript.\no While we discussed and summarized representative comparisons with split-Cas9\nand related inducible systems, we did not perform head-to-head benchmarking\nagainst these platforms in our own experiments, which we view as an important\nfuture direction.\nTogether, these additions provide context for users of our technology, and frame these aspects as\nopportunities for extending the platform.\nReviewer #2\nThe authors have satisfactorily addressed my comments. Therefore, I now gladly\nrecommend this manuscript for publication in Nature Communications. Congratulations\nto the authors on this excellent piece of work.\nWe are very grateful for the reviewer’s positive evaluation and their recommendation for\nacceptance. We appreciate the recognition of the technical advances and conceptual\ncontributions of this work.\nMinor comment: On page 4: “Similar to flow cytometry results, upon induction, As before,\npTET:Ultra-tight cells look longer than pTET cells to achieve maximal editing” Revise as\nappropriate.\nWe have corrected this sentence. It now reads:\n“Similar to flow cytometry results, upon induction, pTET:Ultra-tight cells showed a modest\ndelay compared to pTET cells before achieving maximal editing.”\nGeneral manuscript improvements\n● Added a standardized statistics line to all figure legends:\n“Bars show mean ± SD (n as indicated). Significance by [test] with [correction]; ns, not\nsignificant; *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.”\n● Inserted Supplementary Table S7 comparing published inducible Cas9 systems to\npTET:Ultra-tight.\nWe believe these revisions address all outstanding reviewer comments and improve the clarity,\nbalance, and long-term value of the manuscript."}]

Decision letter