Research ArticleHEALTH AND MEDICINE

Engineering a far-red light–activated split-Cas9 system for remote-controlled genome editing of internal organs and tumors

See allHide authors and affiliations

Science Advances  10 Jul 2020:
Vol. 6, no. 28, eabb1777
DOI: 10.1126/sciadv.abb1777

Abstract

It is widely understood that CRISPR-Cas9 technology is revolutionary, with well-recognized issues including the potential for off-target edits and the attendant need for spatiotemporal control of editing. Here, we describe a far-red light (FRL)–activated split-Cas9 (FAST) system that can robustly induce gene editing in both mammalian cells and mice. Through light-emitting diode–based FRL illumination, the FAST system can efficiently edit genes, including nonhomologous end joining and homology-directed repair, for multiple loci in human cells. Further, we show that FAST readily achieves FRL-induced editing of internal organs in tdTomato reporter mice. Finally, FAST was demonstrated to achieve FRL-triggered editing of the PLK1 oncogene in a mouse xenograft tumor model. Beyond extending the spectrum of light energies in optogenetic toolbox for CRISPR-Cas9 technologies, this study demonstrates how FAST system can be deployed for programmable deep tissue gene editing in both biological and biomedical contexts toward high precision and spatial specificity.

INTRODUCTION

Many studies have shown that the CRISPR-Cas9 system is a revolutionary technology (1, 2). This relatively easy-to-use technology has provided unprecedented opportunities for scientific research and disease treatments, including applications in high-throughput screening and functional genomics research and treatment of virus infections (3), genetic diseases (4), and cancer (5). Nevertheless, there are now several well-known disadvantages with the CRISPR-Cas9 system, including the fact that single guide RNAs (sgRNAs) can sometimes lead to off-target effects such as double-strand breaks in untargeted genome regions, which can cause unintended adverse consequences such as gene mutations, insertions, deletions, and even tumorigenic events (6). Seeking to overcome these challenges, several strategies have been developed to improve the precision of CRISPR-Cas9 gene editing, including Cas9 modifications (e.g., Cas9 nickase and high-fidelity variants), prime editors, base editors, and selecting sgRNAs with minimal off-target capacity (7, 8). Recently, some inducible Cas9 expression systems have been developed to limit the activity or lifetime of Cas9, thereby lowering the probability of off-target effects by reducing the exposure time of a cell’s genome to the Cas9 nuclease (9).

There are a variety of chemically induced CRISPR-Cas9 systems, including doxycycline-regulated Cas9 (10), trimethoprim (TMP) (11) and 4-hydroxytamoxifen (4-OHT)–controlled Cas9 (12), rapamycin-inducible split-Cas9 (13), 4-OHT–responsive intein–dependent Cas9 (14), and 4-OHT–responsive nuclear receptors split-Cas9 (15), among others. However, a notable adverse effect of these systems is the potential for cytotoxicity from the chemical inducers: Doxycycline can negatively affect cell numbers and colony formation (16), TMP can inhibit uptake of folic acid by the cells (17), 4-OHT can increase cytosolic levels of autophagosomes and cause irregularly clumped chromatin in the nuclei (18), and rapamycin can perturb the endogenous mammalian target of rapamycin pathway (19). Moreover, once these agents are inside the cells or present in an in vivo context, these inducer chemicals can diffuse freely, limiting the spatial resolution of editing induction. In addition, it is difficult to rapidly remove the inducer compounds, so they can persist for a long time, making it difficult to turn Cas9 activity on and off quickly and precisely.

These limitations have helped motivate the development of multiple systems based on the optical control of Cas9 activity because light is a reversible and noninvasive inducer modality that potentially offers fine precise spatiotemporal resolution. The first reported example of a photoactivatable Cas9 system was paCas9 system based on blue light (20). In the paCas9 system, Cas9 nucleases are fragmented into two nonfunctional fragments that can be reconstituted as an active nuclease under blue light illumination based on dimerization of their respective fusion domains, the positive Magnet (pMag) or negative Magnet (nMag) proteins from the filamentous fungus Neurospora crassa (21). Later studies reported the ultraviolet (UV) light–mediated cleavage of a synthesized complementary oligonucleotide element that normally inactivates the editing-guiding function of sgRNAs (22).

There is also a recently reported blue light–based anti-CRISPR system comprising AcrIIA4 (23) (a potent Cas9 inhibitor) and the LOV2 blue-light photosensor (24). Without illumination, the AcrIIA4-LOV2 complex remains bound to Cas9, inhibiting its nuclease activity. Under blue light illumination, the AcrIIA4-LOV2 complex is separated from Cas9 and its editing activity can be restored (25). However, neither UV nor blue light is able to penetrate deeply into the body, owing to the strong absorption and scattering of these light energies by biological tissues (26). UV light hardly penetrates the skin and blue light does merely by 1 mm (27, 28). This substantial limitation, viewed alongside the fact that UV and prolonged blue light exposure can cause cytotoxicity (29, 30), highlights the difficulty of applying these light-induced Cas9 systems for in vivo research applications and clinical translation.

We have, for some time, been investigating far-red light (FRL)–inducible genetic systems due to the deep tissue penetration of FRL with above 5 mm beneath the surface of skin (27, 28). We here report our development of an FRL-activated split-Cas9 (FAST) system that can be used to noninvasively induce gene editing activity in cells located deep inside animal tissues. The FAST system relies on two split-Cas9 fusion proteins with high-affinity binding domains: One half of Cas9 is constitutively expressed, while the other is under the FRL-inducible control of the bacterial phytochrome BphS optical controllable system previously established by our group (31). We initially assembled the FAST system components in human embryonic kidney (HEK)–293 cells and used light-emitting diode (LED)–based FRL illumination to demonstrate successful activation of targeted genome editing. Next, after achieving FRL-inducible editing in diverse human cell lines, experiments with implants confirmed that FAST was able to robustly activate editing in cells positioned in subdermal animal tissues. Experiments with the transgenic tdTomato reporter mouse line established FRL-induced FAST–mediated editing of mouse somatic cells (hepatocytes in the liver), and work with cell cycle–inactivating gene edits of cancer cells in xenograft tumor mice demonstrate how FAST can be deployed against disease. Thus, beyond extending the optogenetic toolbox for gene editing of mammalian cells to include induction by the highly in vivo–compatible and deep tissue–penetrating energies of FRL, our study extends this initial technology to demonstrate applications relevant for basic biological and biomedical research.

RESULTS

Design and optimization of the FAST system

To develop an optogenetically controlled device for genome editing with deep tissue–penetrative capacity and with negligible phototoxicity in vivo, first, we constructed an FRL-controlled full-length Cas9 system based on our previously reported orthogonal FRL-triggered optogenetic system (FRL-v2) (31). However, there was serious background leakage in dark state with low-induction performance under illumination. Therefore, we focused on building a FAST system based on split-Cas9 (13) and FRL-v2, which comprises the bacterial FRL-activated cyclic diguanylate monophosphate (c-di-GMP) synthase (BphS) and a c-di-GMP–responsive hybrid transactivator, p65-VP64-BldD. For the FAST system, we then fused the N-terminal Cas9 fragment [Cas9(N)] to the Coh2 domain from Clostridium thermocellum (32) and fused the C-terminal Cas9 fragment [Cas9(C)] to the DocS domain from the same bacterium. Expression of the NLS-Cas9(N)-Coh2 fusion protein is driven by the FRL-v2–specific chimeric promoter (PFRL), while expression of the DocS-Cas9(C)-NES fusion protein is driven by a constitutive promoter (PhCMV). A complete Cas9 protein can be reconstituted upon FRL illumination because of the high-affinity interaction of the Coh2 and DocS domains (Fig. 1). Confirming the editing activity of the reconstituted Cas9, we found that HEK-293 cells cotransfected with pXY137 (PhCMV-p65-VP64-BldD-pA::PhCMV-BphS-P2A-YhjH-pA, 100 ng), pYH20 [PFRL-NLS-Cas9(N)-Linker-Coh2-pA, 50 ng], pYH102 [PhCMV-DocS-Linker-Cas9(C)-NES-pA, 100 ng], and pYW57 [PU6-sgRNA (CCR5)-pA, 50 ng] successfully edited the targeted human CCR5 locus (11.9% indel frequency) upon FRL illumination (1 mW/cm2; from an LED source, 730 nm); no editing was detected for dark control cells (Fig. 2, A and B). These detected edits were analyzed by the mismatch-sensitive T7 endonuclease I (T7E1) assay. We further used Sanger sequencing to confirm that the FRL-induced, FAST-mediated edits (indel mutations) occurred in the targeted region of the human CCR5 locus at a frequency of ~20% using the tracking of indels by decomposition (TIDE) analysis (fig. S1).

Fig. 1 Design of the FAST system.

(A) Schematic of the split-Cas9 fusion protein components of the FAST system. Coh2 and DocS are two C. thermocellum proteins that interact with high affinity. Cas9 is formed from two separate (N- and C-terminal) Cas9 fragments that individually lack nuclease activity. When Cas9’s two fragments Cas9(N) and Cas9(C) are respectively fused with Coh2 and DocS, they readily combine to reconstitute a nuclease-active form of Cas9. (B) Schematic of the FAST system, as deployed in mammalian cells, based on the fragments detailed in (A). FRL (~730 nm) activates the engineered bacterial photoreceptor BphS, which converts guanosine triposphate (GTP) into c-di-GMP. c-di-GMP can bind to BldD (derived from sporulating actinomycete bacteria) and be translocated into the nucleus. This induces dimerization of the synthetic transcriptional activators p65-VP64-BldD [BldD fused with p65 (the nuclear factor κB–transactivating domain) and VP64 (a tetramer of the herpes simplex virus–derived VP16 activation domain)], after which they bind to PFRL to activate expression of the N-terminal fusion fragment of split-Cas9. The other (C-terminal) fusion fragment is constitutively expressed, as driven by the human cytomegalovirus promoter (PhCMV). DNA double-strand breaks are formed by Cas9 after the Coh2-DocS heterodimerization–mediated reconstitution of the two fusion fragments.

Fig. 2 Optimization of the FAST system.

(A) Time schedule of FRL-controlled gene editing in HEK-293 cells. Cells were illuminated (1 mW/cm2; 730 nm) for 4 hours once a day for 2 days and were collected at 48 hours after the first illumination for further analysis. (B) A mismatch-sensitive T7 endonuclease I (T7E1) assay to test HEK-293 cells (6 × 104) transfected with full-length Cas9 (pHP1) or the FAST system (pXY137, pYH20, and pYH102), together with the sgRNA targeting to CCR5 locus (pYW57). FRL-mediated editing (indel deletions) of the human EMX1, CXCR4, and VEGFA loci by FAST was performed using the same experimental procedure as that used when targeting the CCR5 gene. (C) FRL-mediated multiplex editing of the human CCR5 and CXCR4 loci. (D) FAST-mediated DNA insertion via homology-directed repair (HDR), achieved by adding a single-stranded oligodeoxynucleotide (ssODN) template (10 μM), bearing a HindIII restriction endonuclease site. Homologous arms are indicated in red. The target sites of sgRNA (EMX1) are marked in blue. HEK-293 cells (6 × 104) were cotransfected with full-length Cas9 (pHP1) or the FAST system (pXY137, pYH20, and pYH102) and the sgRNA targeting to EMX1 locus (pYH227) via a nucleofection method. In (B) to (D), n = 2 from two independent experiments. Red arrows indicate the expected cleavage bands. Detailed description of genetic components and transfection mixtures are provided in tables S1 and S5. N.D., not detectable.

We next confirmed that the FAST system can cleave different targeted endogenous genomic loci and induce indel mutations via nonhomologous end joining (NHEJ) in an FRL-dependent manner by designing sgRNAs targeting three additional human genes (EMX1, CXCR4, and VEGFA), and these induced indel mutations were detected by T7E1 assay. With each of these sgRNAs, FRL-induced but not dark-induced indel mutations were observed (Fig. 2B). We also confirmed that the FAST system can cleave targeted exogenous d2EYFP reporter efficiently (fig. S2). In addition to single gene targeting, we also tested whether our FAST system can simultaneously edit multiple target sites. Using one sgRNA targeting CCR5 and another sgRNA targeting CXCR4, the FAST system was capable of inducing the desired indel mutations at the two target sites upon FRL illumination (Fig. 2C), demonstrating optogenetic multiplexed control of NHEJ-mediated indel mutations in mammalian cells.

We further investigated whether FAST can be used for homology-directed repair (HDR)–mediated genome editing. The FAST system components and a donor template (single-stranded oligodeoxynucleotide containing a HindIII site) were electroporated into HEK-293 cells. Assessment of HDR events at the EMX1 locus using restriction endonuclease assays showed that the FAST system induced HindIII site integration at the EMX1 locus at a frequency of 5.7% under FRL illumination; no HDR events were detected in dark controls (Fig. 2D). Together, these results establish that the FAST system can be deployed for optogenetic control of NHEJ-/HDR-mediated indel mutations.

Characterization of the gene editing performance of the FAST system

To demonstrate photoactivatable regulation of gene editing in diverse mammalian cell lines, we introduced the FAST system into four different human cell lines, and it achieved successful FRL-induced gene editing (CCR5 locus) in each of them (Fig. 3A). Next, experiments testing the FRL illumination intensity and duration-dependent activity of the FAST system showed that the frequency of edits (indel mutations at CCR5) increased along with illumination intensity and with illumination time (Fig. 3, B and C), indicating the tunability of the FAST system. We also used a photomask to establish proof of principle for spatially controlled gene editing with the FAST system (Fig. 3, D and E). We also conducted an experiment with two rounds of FRL illumination to verify repeated induction cycles of the FAST system wherein the first round of illumination achieved indel mutations guided by an sgRNA targeting CXCR4 locus, followed by transfection of a second sgRNA targeting the CCR5 locus, which guided successful indel mutations after the second FRL illumination. However, engineered cells shifted to the dark did not have indel mutations in CCR5 locus (fig. S3, A and B). This result indicates that the FAST system is reusable and reversible.

Fig. 3 Characterization of the FAST system.

(A) FAST-mediated gene editing in four human cell lines. (B) Illumination intensity–dependent FAST gene editing. In (A) and (B), cells were collected for mismatch-sensitive T7E1 assays, as indicated in the time schedule of Fig. 2A. (C) Evaluation of exposure time–dependent FAST system gene editing performance. Cells were collected for T7E1 assays at 24 hours after the start of the second illumination. (D) Schematic of the photomask device used to demonstrate the spatial regulation of FAST-mediated gene editing. Cells were illuminated through a photomask containing a 7-mm line pattern. (E) Spatial control of FRL-dependent gene editing mediated by the FAST system. HEK-293 cells (3 × 106) were cotransfected with the FAST system, sgRNA (pYW57), and a frameshift enhanced green fluorescent protein (EGFP) reporter containing a CCR5 locus (pYH244) and were illuminated with FRL (0.5 mW/cm2; 730 nm; 2-min on, 2-min off) for 48 hours. EGFP is not expressed without Cas9 activity because the EGFP sequence is out of frame. Upon double-strand cleavage by Cas9, the frameshifts caused via DNA repair by NHEJ enable EGFP expression. The fluorescence of EGFP was assessed via fluorescence meter ChemiScope 4300 Pro imaging equipment (Clinx) at 48 hours. In (A) to (C), n = 2 from two independent experiments. Red arrows indicate the expected cleavage bands. Detailed description of genetic components and transfection mixtures are provided in tables S1 and S5. SEAP, human placental secreted alkaline phosphatase.

We then evaluated the photocytotoxicity of FRL (730 nm) or blue light (470 nm) illumination on mammalian cells. When HEK-293cells were transfected with human placental secreted alkaline phosphatase (pSEAP2)-control-and then exposed to FRL or blue light for different intensity, the SEAP expression demonstrated that the FRL exposure resulted in negligible cytotoxicity. However, a marked difference was observed from the blue light illumination, which substantially reduced cell viability (fig. S4, A and B). Moreover, we did not observe substantially increased cytotoxicity with FRL illumination of cells engineered with the FAST system (fig. S4, C and D), indicating the inertness and noncytotoxicity of the system constituents. In short, neither FRL illumination nor the ectopic presence of FAST system constituents was verified to influence the gene expression capacity of the engineered cells. In addition, we also compared the controllable gene editing performance of our FAST system with the rapamycin-responsive split-Cas9 system (13) and the blue light–controlled paCas9 system (20) that have been reported. The results showed that the genome editing efficiency of rapamycin-responsive split-Cas9 system was lower than the FAST system (fig. S5, A and B), and the paCas9 system had relative higher background leakage in the dark. Our FAST system showed notable induction of indel mutations under FRL illumination but with negligible background in the dark (fig. S5, C and D). Off-target activity of the FAST system was also assessed simply. We checked a potential off-target site of human BMP1 locus, as reported previously (33). The indel frequencies were determined through T7E1 assay at the on-target and potential off-target sites of BMP1. As a result, no mutations were detected at the potential off-target site after editing by our FAST system (fig. S6, A and B). This is probably due to the FAST-mediated transient expression of split-Cas9 that lowered the probability of off-target effects by reducing the exposure time of a cell’s genome to the Cas9 nuclease (79). However, there might be off-target effects that can still occur in illuminated cells.

Deployment of the FAST system for editing of cells implanted subcutaneously in mice

Having established the basic performance characteristics of the FAST system in human cells, we next conducted experiments with mice to verify the system’s capacity to induce gene editing based on the tissue-penetrating capacity of FRL. Specifically, we conducted an experiment using hollow fiber implantation of HEK-293 cells equipped with the FAST system into the dorsum of mice and exposed to FRL illumination (10 mW/cm2; alternating 2-min on/off for 4 hours) (Fig. 4A). Notably, the FRL illumination of the FAST cell-bearing mice induced notable activation of gene editing (~11.4% of the cells retrieved from the implant fibers was edited at the CCR5 locus versus not detectable for dark control cells) (Fig. 4B). These results demonstrate that the FAST system can be used to activate gene editing inside animal tissues, exploiting the physical properties of FRL as an inducer modality.

Fig. 4 FRL-induced FAST-mediated gene editing (CCR5 locus) of cells residing subcutaneously in mouse implants.

(A) Schematic for the time schedule and experimental procedure for FRL-controlled gene editing in mice harboring hollow fiber implants with HEK-293 cells. Pairs of 2.5-cm hollow fibers containing a total of 5 × 106 transgenic HEK-293 cells (engineered with FAST system) were subcutaneously implanted on the dorsum of wild-type mice and illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off) for 4 hours each day for 2 days. Cells were collected from the hollow fiber implants at 48 hours after the first illumination and assessed with mismatch-sensitive T7E1 assay to assess targeted gene editing efficiency (CCR5 locus). (B) Representative T7E1 assay for FAST-mediated indel mutations. n = 3 mice. The red arrow indicates the expected cleavage bands. Detailed description of genetic components and transfection mixtures are provided in table S1 and S5.

FAST-mediated gene editing of the tdTomato reporter locus in the livers of transgenic mice

We obtained transgenic mice harboring a homozygous Rosa26 CAG [cytomegalovirus (CMV) enhancer fused to the chicken beta-actin] promoter loxP-STOP-loxP-tdTomato cassette present in all cells. In this model, tdTomato is silent because of the stop signal [three repeats of the simian virus 40 (SV40) polyadenylate (polyA) sequence], but the deletion of the stop cassette allows transcription of the tdTomato gene, resulting in fluorescence expression. The Cas9-mediated DNA cleavage of the stop sequence guided by sgRNAs can initiate CAG promoter to drive tdTomato expression (34). Therefore, we used this mouse model to examine the in vivo genome editing performance of the FAST system in mice somatic cells (Fig. 5A). We used hydrodynamic injection to introduce the FAST system components, along with an sgRNA designed to target the deletion of the SV40 polyA stop cassette, which should activate tdTomato reporter protein expression upon successful editing. Note that it is difficult to activate tdTomato expression by Cas9 system as the desired edit requires two cuts on the same allele; we eventually achieved the desired edit, but it required optimization of the delivery mode for the FAST components. Briefly, we chose hydrodynamic injection because it is known to result in enrichment of plasmids (and thus, transgene expression) in liver cells (35). We reduced the overall number of plasmids by combining some constructs (fig. S7, A and B) and explored a number of different injection time and illumination schedules (Fig. 5A), but we only detected weak tdTomato signals in the FRL-illuminated FAST mice (fig. S8).

Fig. 5 FAST-mediated gene editing of the tdTomato reporter locus in Gt(ROSA)26Sortm14(CAG-tdTomato)Hze mice.

(A) Schematic showing the time schedule and experimental procedure for assessing in vivo gene editing. The minicircle iteration of the FAST system pYH412, pYH413, and pYH414 at a 7:15:4 (w/w/w) ratio were injected hydrodynamically via tail vein. Twenty-four hours after injection, mice were illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off) for 4 hours per day for 3 days. A second injection of the minicircle-based FAST system components was performed on the fifth day, followed by 4 hours daily illumination for three additional days. In our design, the tdTomato reporter protein was expressed after a stop cassette was destroyed by Cas9 editing. (B) Fluorescence IVIS image of mouse livers. (C) The frequency of edits (targeting the aforementioned stop cassette) by monitoring fluorescence intensity of the tdTomato reporter in Gt(ROSA)26Sortm14(CAG-tdTomato)Hze mice. (D) Representative fluorescence microscopy images of tdTomato and tdTomato+ hepatocytes present in frozen liver sections from FRL-illuminated mice. Blue indicates 4′,6-diamidino-2-phenylindole (DAPI) staining nuclei; red indicates endogenous tdTomato expression. The images represent typical results from three independent measurements. Scale bar, 100 μm. Data in (C) are means ± SEM; n = 3 mice. P values were calculated by Student’s t test. ****P < 0.0001 versus control.

We speculated that this apparently weak induction of editing activity may result from rapid degradation of the plasmids, so we constructed minicircle (36) iterations of our FAST system. Minicircle DNA vectors without the bacterial backbone of the plasmid, markedly reducing the possibility of random integration of bacterial DNA sequences into the genome, have been shown to maintain gene expression in cells for long durations because these molecules are resistant to degradation (37). We delivered the minicircle iterations of the FAST via hydrodynamic injection and used FRL illumination schedules as follows: alternating 2-min on/off for 4 hours, once each day for 3 days; we then monitored the fluorescence signal intensity in livers. FRL illumination of the mice bearing the FAST system resulted in strong editing and thus, tdTomato reporter expression (Fig. 5, B and C). We also detected strong tdTomato expression in liver sections prepared from the FRL-illuminated FAST mice (Fig. 5D), and Sanger sequencing of genomic DNA extracted from the livers verified the success of the targeted excision of the SV40 polyA stop cassette in the FRL-induced FAST mice (fig. S9). Collectively, these results demonstrate that the FAST system can be used for in vivo editing of the genomes of somatic cells located in the internal organs of mice.

Gene editing–based inhibition of tumor growth mediated by the FAST system in xenograft nude mouse model

We further investigated the optogenetic activation of the FAST system in tumor models as proof-of-concept examples for therapeutic genome editing. The polo-like kinase (PLK1) protein is a highly conserved serine-threonine kinase that promotes cell division, and strong PLK1 expression is a marker in various types of tumor (38). Extensive work has established that inhibition or depletion of PLK1 leads to cell-cycle arrest, apoptosis, and a so-called “mitotic catastrophe” in cancer cells, which provides a promising modality for anticancer therapy (39, 40). After initially confirming that the FAST system can edit the PLK1 locus (indel mutations and extensive apoptosis) in the FRL-illuminated human lung cancer A549 cells in vitro (fig. S10, A to D), we then evaluated the tumor therapy application of our FAST system by testing the in-tumor editing performance of the FAST system for the disruption of the PLK1 locus in mice bearing A549 xenograft tumors.

We first delivered the minicircle iterations of the FAST system alongside a PLK1-targeting sgRNA minicircle vector when the tumors had reached 80 to 100 mm3; note that we also injected transfection reagent, a cationic polymer-coated nanoparticle (APC), (41) to facilitate the transfection of tumor cells in situ. Subsequently, FRL illumination was delivered to the xenograft-bearing mice via LED for 4 hours each day for 7 days (Fig. 6A), and tumor development was monitored by measuring the sizes of the tumors every 2 days. Notable inhibition of tumor growth was observed for the FAST mice that received FRL illumination; no such inhibition was observed for the dark control FAST or FRL-illuminated vehicle control mice (Fig. 6, B to D). Mismatch-sensitive T7E1 assays confirmed that the FRL-induced FAST system achieved the desired genome disruption of PLK1 gene in the tumor tissue (Fig. 6E) at a frequency of ~21.5% detected by TIDE analysis (Fig. 6F). Moreover, quantitative real-time polymerase chain reaction (qRT-PCR) verified the expected reductions in tumor PLK1 mRNA expression upon FRL illumination (Fig. 6G). Consistent with the observed antitumor efficacy, subsequent histologic analysis of tumor sections revealed extensive cancer cell necrosis (Fig. 6H) and very extensive cell apoptosis [via both terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick end labeling (TUNEL) and caspase-3–labeling assays; Fig. 6, I and J]. Thus, FRL-triggered FAST-mediated gene editing can inhibit cancer cell growth in xenograft tumors in mice. These results further indicate that our FAST system could be deployed for deep tissue gene editing.

Fig. 6 In-tumor gene editing (PLK1 locus) with the FAST system in xenograft (A549 cell) mice.

(A) Schematic showing the time schedule and experimental procedure for the in-tumor FAST-mediated gene editing. The minicircle iteration of the FAST system targeting to PLK1 locus pYH412, pYH420, and pYH414 at a 7:15:4 (w/w/w) ratio were injected intratumorally. Twenty-four hours after per injection, mice were illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off) for 4 hours per day totally for 7 days. (B) Images of tumor tissues from the different treatments. (C) Tumor growth curves for the different treatments. (D) The weight of tumor tissues after the different treatments. (E) Indel mutations in the tumor tissues detected via mismatch-sensitive T7E1 assays. Red arrows indicate the expected cleavage bands. (F) The gene editing efficacy quantified by the TIDE analysis. (G) Relative mRNA expression levels of the PLK1 gene quantified by quantitative real-time polymerase chain reaction (qRT-PCR). The data are means ± SEM; n = 5 mice. P values were calculated by Student’s t test. ****P < 0.0001 versus control. (H) Representative fluorescence microscopy images of hematoxylin and eosin (H&E) staining of tumor tissues. The images represent typical results from three independent measurements. Scale bar, 100 μm. Representative fluorescence microscopy images of TUNEL staining (I) and caspase-3 (J) staining of tumor tissues. The images represent typical results from three independent measurements. Scale bars, 100 μm. Photo credit: Yuanhuan Yu, East China Normal University.

DISCUSSION

CRISPR-Cas9 is an undeniably revolutionary technology that is changing biological and medical research (4, 5, 42), and several innovative extensions of the basic CRISPR-Cas9 concept have enabled a new era of conditional genome editing activation iterations with chemical (1015) and UV/blue light inducers (20, 22, 25). Nevertheless, limitations with these systems warrant the development of alternatives that exploit different induction sources. The FAST system we developed in the present study opens the door for spatiotemporally selective induction of Cas9 genome editing deep inside animal tissues. It bears emphasis that our induction uses LED lights rather than lasers or optical fibers, highlighting that FAST should be very easy to deploy in a wide range of experimental contexts. Although we did face initial hurdles with induction efficiency for in vivo applications, our development of a minicircle-based iteration of the FAST system easily overcame this and permitted robust editing in mouse livers. The deep tissue–penetrating utility of the FAST system was applied to achieve anticancer therapy by disrupting PLK1 gene in mice bearing A549 xenograft tumors. In this way, we could greatly reduce side effects of the anticancer drugs and promote the precision treatment of cancers. We also envision that the FAST system can be used to study the function of cancer-associated genes during tumor development process by controlling gene knockout or interference in specific tissues at different time nodes.

While we do demonstrate FAST system applications for biological research and the treatment of disease, the present paper merely reports the initial proof-of-principle study. Given that FAST is a fully genetically encoded system, a variety of vectors, alternative plasmids, and tissue-specific promoters could be used to selectively deliver FAST system components to diverse tissues, and we fully anticipate that adeno-associated virus vectors will become a popular modality for this task. Moreover, there is no obvious factor to prevent the deployment of FAST as a genome-integrated stable system, which should enable researchers to selectively activate targeted editing anywhere that they are able to supply sgRNAs and FRL illumination from an LED.

We anticipate that the combination of precise temporal control and deep tissue penetration will enable rapid-uptake FAST in a variety of research communities. Chemical inducers can cause adverse effects in cells and can diffuse freely, and the complexity of cellular and organismal metabolism makes it exceedingly difficult to precisely control the spatiotemporal dynamics of inducible gene editing systems (1619). In this light, perhaps researchers can deploy FAST and FRL induction strategies to explore the development, basic biology, or etiopathological basis of diverse processes that occur in animal internal organs such as the heart, lungs, liver, kidneys, etc., and in tissues, including muscles and bone marrow. In theory, the FAST system should give researchers previously unattainable precise control of conditional genetic knockout and knock-in experiments. A huge variety of temporal illumination schemes should be feasible with FAST because FRL has low phototoxicity, representing a clear advantage over UV- and blue light–based Cas9 induction systems. Moreover, FAST may offer neuroscientists an alternative to the presently popular optical fiber implantation–based approaches for optogenetic-based gene editing research.

In summary, we have developed a FAST system that is apparently safe (negligible phototoxicity to mammalian cells, high tissue permeability, and noninvasiveness). With FRL as its fundamental basis, the FAST system offers excellent tunability (robust induction of gene editing and almost negligible background activity) and precise controllability (illumination intensity dependent, exposure time dependent, and strong spatiotemporal specificity), making it suitable and practical for the many biological and biomedical applications that require gene editing in vivo, especially for processes that occur within animal tissues.

MATERIALS AND METHODS

Design and construction of the FAST system

The FAST system consists of the following main components: the FRL sensors (BphS and p65-VP64-BldD) (31), interacting proteins (cohesion Coh2 and dockerin DocS from C. thermocellum) (32), and the N- and C-terminal fragments of Streptococcus pyogenes Cas9 [Cas9(N) (residues 2 to 713) and Cas9(C) (residues 714 to 1368)] (13). Complementary DNAs (cDNAs) encoding BphS and p65-VP64-BldD were prepared, as previously described (31). cDNAs encoding Coh2 and DocS were chemically synthesized by the company Genewiz Inc. cDNAs encoding the N- and C-terminal fragments of Cas9 fused with a nuclear localization signal from SV40 T antigen were amplified from the Addgene plasmid 42230. The inducible Cas9 was constructed on the basis of the Cas9(N) and Cas9(C) fragments fused with Coh2 and DocS, respectively, which were cloned through Gibson assembly according to the manufacturer’s instructions [Seamless Assembly Cloning Kit; catalog no. BACR(C) 20144001; OBiO Technology Inc.]. All genetic components have been validated by sequencing (Genewiz Inc.). Plasmids constructed and used in this study are provided in table S1.

sgRNA constructions

The sgRNAs targeting CCR5, EMX1, CXCR4, VEGFA, BMP1, tdTomato stop cassette, and PLK1 were generated by annealed oligos and cloned into the BbsI site of a constitutive mammalian PU6-driven sgRNA expression vector (pYH49). The PU6-sgRNA fragment was PCR amplified from the Addgene plasmid 58767 and then cloned into the corresponding sites (MluI/XbaI) of pcDNA3.1(+) to obtain the pYH49 expression vector. The target sequences and oligonucleotides used for sgRNA construction are listed in table S2.

Cell culture and transfection

All cell types {HEK-293 [CRL-1573; American Type Culture Collection (ATCC)], HeLa (CCL-2; ATCC), telomerase-immortalized human mesenchymal stem cells (43), and HEK-293–derived Hana3A cells engineered for constitutive expression of RTP1, RTP2, REEP1, and Gαoλϕ} were cultured at 37°C in a humidified atmosphere, containing 5% CO2 in Dulbecco’s modified Eagle’s medium (DMEM; catalog no. C11995500BT; Gibco) supplemented with 10% fetal bovine serum (FBS; catalog no. 16000-044; Gibco) and 1% (v/v) penicillin/streptomycin solution (catalog no. ST488-1/ST488-2; Beyotime Inc.). All cell lines were regularly tested for the absence of mycoplasma and bacterial contamination. Cells were transfected with an optimized polyethyleneimine (PEI)–based protocol (44). Briefly, cells were seeded in a 24-well cell culture plate (6 × 104 cells per well) 18 hours before transfection and were subsequently cotransfected with corresponding plasmid mixtures for 6 hours with 50 μl of PEI and DNA mixture [PEI and DNA at a ratio of 3:1 or 5:1 (w/w)] (PEI molecular weight, 40,000; stock solution of 1 mg/ml in ddH2O; catalog no. 24765; Polysciences Inc.). At 12 hours after transfection, the culture plate was placed below a custom-designed 4 × 6 LED array (1 mW/cm2; 730 nm) for illumination.

For HDR-mediated genome editing experiments, 6 × 105 HEK-293 cells were nucleofected with the FAST system plasmids (pXY137, 200 ng; pYH20, 100 ng; and pYH102, 200 ng), sgRNA expression vector (pYH227, 100 ng; targeting EMX1), and 10 μM single-stranded oligonucleotide donor using the SF Cell Line 4D-Nucleofector X Kit L (catalog no. V4XC-2024; Lonza) and the CM-130 program (4D-Nucleofector System; Lonza). At 24 hours after nucleofection, cells were illuminated by FRL (1 mW/cm2; 730 nm) for 4 hours once a day for 2 days, and then cells were collected at 48 hours after the first illumination for analysis. Genomic DNA was isolated using a TIANamp Genomic DNA Extraction Kit (catalog no. DP304; TIANGEN Biotech Inc.) according to the manufacturer’s instructions.

Mismatch-sensitive T7E1 assay

Genomic DNA was extracted from cells or tissues using the TIANamp Genomic DNA Extraction Kit (catalog no. DP304; TIANGEN Biotech Inc.) according to the manufacturer’s instructions. The genomic region containing the target sites was PCR amplified using the 2× Taq Plus Master Mix II (Dye Plus) DNA polymerase (catalog no. P213; Vazyme Inc.). The primers used for PCR amplification are listed in table S3. The PCR amplicons were purified using HiPure Gel Pure Micro Kits (catalog no. D2111-03; Magen Inc.) according to the manufacturer’s protocol. Purified PCR products (300 ng) were mixed with 1.5 μl of 10× M buffer for restriction enzyme (catalog no.1093A; Takara Bio) and ultrapure water to a final volume of 15 μl and reannealed (95°C, 5 min; 94°C, 2 s, −0.1°C per cycle, 200 times; 75°C, 1 s, −0.1°C per cycle, 600 times; and 16°C, 5 min) to form heteroduplex DNA. After reannealing, the heteroduplexed DNA was treated with 5 U of T7E1 (catalog no. M0302; New England BioLabs) for 1 hour at 37°C and then analyzed by 1.5% agarose gel electrophoresis. Gels were stained with GelRed (catalog no. 41003; Biotium) and imaged with Tanon 3500 gel imaging system (Tanon Science & Technology Inc.). Relative band intensities were calculated by ImageJ software. Indel percentage was determined by the formula 100% × [1 − (1 − (b + c)/(a + b + c))1/2], in which a is the integrated intensity of the undigested PCR product, and b and c are the integrated intensities of each cleavage product.

DNA sequence analysis

Sequence of the gene region containing the target sequence was amplified by PCR. Purified PCR amplicons from the nuclease target site were cloned into the T-vector pMD19 (catalog no. 3271; Takara Bio). Thirty clones were randomly selected and sequenced using each gene’s PCR forward primers by the Sanger method (45). Primers used for PCR amplification are listed in table S3.

TIDE analysis

Target regions were amplified by PCR. Purified PCR samples were analyzed by Sanger sequencing. The sequencing data files (.ab1 format) were imported into the TIDE Web tool (https://tide.nki.nl/) (46) to quantify nature and frequency of generated indels.

Detection of the HDR-mediated modification in endogenous genes

The genomic PCR and purification were performed, as described above. Purified PCR products were mixed with 15 U of HindIII (catalog no. 1060B; Takara Bio), 2 μl of 10× M buffer for restriction enzyme, and ultrapure water to a final volume of 20 μl and then incubated at 37°C for 3 hours. The digested products were analyzed by agarose gel electrophoresis. Gel staining and imaging were performed, as described above. Quantification was calculated on the basis of relative band intensities. The HDR percentage was determined by the formula 100% × (b + c)/(a + b + c), in which a is the intensity of the undigested PCR product, and b and c are the intensities of each HindIII-digested product.

Flow cytometry

HEK-293 cells (6 × 104) were cotransfected with the FAST system (pXY137, 100 ng; pYH20, 50 ng; and pYH102, 100 ng), the sgRNA targeting d2EYFP (pYH410, 50 ng), and the d2EYFP reporter plasmid (pYW110, 200 ng). At 12 hours after transfection, cells were illuminated (1 mW/cm2; 730 nm) for 4 hours once a day for 2 days and were harvested after trypsinization and washed in phosphate-buffered saline (PBS) for three times. About 10,000 events were collected per sample and analyzed with a BD LSRFortessa cell analyzer (BD Biosciences) equipped for d2EYFP [488-nm laser, 513-nm longpass filter, and 520/30 nm emission filter (passband centered on 530 nm; passband width of 30 nm)] detection. Data were analyzed using the FlowJo V10 software.

SEAP reporter assay

The production of human placental SEAP in cell culture medium was quantified using a p-nitrophenylphosphate–based light absorbance time course assay, as previously reported (31). Briefly, 120 μl of substrate solution [100 μl of 2× SEAP buffer containing 20 mM homoarginine, 1 mM MgCl2, and 21% (v/v) diethanolamine (pH 9.8) and 20 μl of substrate solution containing 120 mM p-nitrophenylphosphate] were added to 80 μl of heat-inactivated (65°C, 30 min) cell culture supernatant. The time course of absorbance at 405 nm was measured by using a Synergy H1 hybrid multimode microplate reader (BioTek Instruments Inc.) installed with the Gen5 software (version 2.04).

MTT assay

Cell viability was assayed using an MTT [3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide] cytotoxicity assay kit (catalog no. E606334-0250; Sangon Biotech Inc.) according to the manufacturer’s instructions. Briefly, 10 μl of MTT reagent (5 mg/ml) was added to each well of 96-well plates. The samples were mixed gently and incubated for 4 hours in a CO2 incubator. Formazan solubilization solution (100 μl) was added into each well. The plate was put on a shaker to mix gently for 10 min to dissolve the formazan crystals, and then the plate was read with a Synergy H1 microplate reader (BioTek Instruments Inc.) at 570 nm.

Off-target analysis

The off-target sites of the BMP1 gene were examined according to the previously reported procedure (33). Genomic DNA was extracted, as described above, and the region of genome containing the possible nuclease off-target sites was PCR amplified using appropriate primers (table S3). The following procedures were similar to those of on-target examination by T7E1 assay, as described above.

Minicircle DNA vector production

Minicircles are episomal DNA vectors that allow sustained transgene expression in quiescent cells and tissues. Minicircle DNA vectors were prepared, as previously described (36). Minicircle-producing system contains the Escherichia coli strain ZYCY10P3S2T (a genetically modified minicircle-producing bacterial strain) and the empty minicircle-producing plasmid pMC.BESPX (gene of interest would be cloned into this plasmid). Briefly, ZYCY10P3S2T competent cells prepared with standard protocol, as previously described (36), were transformed with the minicircle-producing plasmid pMC.BESPX carrying the gene of interest. The transformed cells were cultured and induced by 0.01% l-arabinose to produce minicircle DNA vectors that were devoid of the bacterial plasmid DNA backbone and contain only genes of interest.

Synthesis of the in vivo transfection reagent APC

The in vivo DNA delivery reagent APC is a cationic polymer-coated nanoparticle composed of biocompatible polystyrene sulfonate and β-cyclodextrin–PEI (Mw, 25 kDa) and prepared, as previously reported (41). First, the seed solution was prepared by adding freshly prepared 600 μl of NaBH4 (10 mM) into 5-ml mixture of HAuCl4·3H2O (0.5 mM) and cetyltrimethylammonium bromide (CTAB; 0.1 M) and incubated at 30°C for 30 min. Ten milliliters of HAuCl4·3H2O (1 mM), 10 ml of CTAB (0.2 M), 120 μl of AgNO3 (0.1 M), and 600 μl of hydroquinone (0.1 M) were mixed together as growth solution. When the color of the growth solution turned from yellow to colorless, 320 μl of seed solution was added. The desired longitudinal surface plasmon resonance peak was obtained after keeping the reaction mixture undisturbed in dark at 30°C for 12 hours. The products were then gathered by centrifugation at 7000 RCF (relative centrifugal force) for 10 min at 30°C. The supernatant was removed, and the precipitate was resuspended in 2 ml of 30°C ultrapure water. Furthermore, 1 ml of the products from last step [Au (0.2 mg/ml)] was added to 10 ml of polysodium 4-styrenesulfonate (2 mg/ml) dissolved in NaCl (1 mM) solution and stirred for 1 hour at 30°C. The solution was centrifuged at 7000 RCF for 10 min, and the residue was resuspended to obtain 2 ml of biocompatible polystyrene sulfonate–coated nanoparticle solution. Last, 1 ml of biocompatible polystyrene sulfonate–coated nanoparticles was added to 10 ml of β-cyclodextrin–PEI (2 mg/ml) dispersed in NaCl (1 mM) solution and stirred for 1 hour at 30°C to obtain APC.

Annexin V–FITC apoptosis detection

Apoptosis analysis at the cellular level was assessed using the Annexin V–fluorescein isothiocyanate (FITC)/propidium iodide (PI) Apoptosis Detection Kit (catalog no. E606336; Sangon Biotech Inc.). Briefly, A549 cells (3 × 104) cotransfected with the minicircle iterations of the FAST system and the sgRNA targeting PLK1 {pYH412 (PhCMV-p65-VP64-BldD-pA::PhCMV-BphS-P2A-YhjH-pA, 135 ng), pYH414 [PFRL-NLS-Cas9(N)-Linker-Coh2-pA, 77 ng], and pYH420 [PU6-sgRNA (PLK1)::PhCMV-DocS-Linker-Cas9(C)-NES-pA, 288 ng]} were illuminated by FRL (1 mW/cm2; 730 nm) for 4 hours once a day for 2 days and were then collected at 48 hours after the first illumination for analysis. The subsequent procedures were performed according to the manufacturer’s instructions and analyzed by flow cytometry (BD LSRFortessa cell analyzer; BD Biosciences). The LSRFortessa was equipped with green fluorescence channel (488-nm laser, 530/30 nm emission filter, 505 nm longpass dichroic mirror) and red fluorescence channel (561-nm laser, 610/20 nm emission filter, 595 nm longpass dichroic mirror). A gate was applied on forward scatter and side scatter to remove debris from cell populations. Data were analyzed using the FlowJo V10 software.

qRT-PCR analysis

Total RNA of cells or tissues was extracted using the RNAiso Plus kit (catalog no. 9109; Takara Bio). A total of 500 ng of RNA was reverse transcribed into cDNA using a PrimeScript RT Reagent Kit with the genomic DNA Eraser (catalog no. RR047; Takara Bio). Quantitative PCR (qPCR) reactions were performed on the LightCycler 96 real-time PCR instrument (Roche Life Science) using the SYBR Premix Ex Taq (catalog no. RR420; Takara Bio). Program for qPCR amplifications were as follows: 95°C for 10 min, followed by 40 cycles at 95°C for 10 s, 60°C for 15 s, and 72°C for 10 s, and then 95°C for 10 s, 60°C for 60 s, 97°C for 1 s, and last, 37°C for 30 s. The qPCR primers used in this study are listed in table S4. Samples were normalized to the housekeeping gene glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as the endogenous control. Standard ΔΔCt method was used to obtain relative mRNA expression level.

Hollow fiber implantation

Wild-type mice [8 week old, male, C57BL/6J, East China Normal University (ECNU) Laboratory Animal Center] were randomly divided into two groups. The semipermeable KrosFlo polyvinylidene fluoride hollow fiber membrane (Spectrum Laboratories Inc.; notably, the light-absorption properties of this material to lights of 300 to 1000 nm are almost the same) implants containing optogenetically engineered HEK-293 cells (pairs of 2.5-cm hollow fibers containing a total of 5 × 106 engineered cells) were subcutaneously implanted beneath the dorsal skin of the mice under anesthesia (two 2.5-cm hollow fibers in each mouse). At 1 hour after implantation, the mice were illuminated by FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off, alternating, to avoid the thermal discomfort in mice caused by continuous illumination) for 4 hours once a day for 2 days. The control mice were kept in dark. Cells were then collected from the implanted hollow fibers at 48 hours after the first illumination, and the genomic DNA was extracted for mismatch-sensitive T7E1 assay to quantify the indel mutations of the endogenous gene CCR5.

Gene editing mediated by FAST in tdTomato reporter mouse model

The transgenetic Ai14 tdTomato reporter mice [6 week old, female, Gt(ROSA)26Sortm14(CAG-tdTomato)Hze, from the Jackson laboratory; Ai14 is a Cre reporter allele designed to have a loxP-flanked stop cassette, preventing the transcription of a CAG promoter–driven red fluorescent tdTomato, all inserted into the Gt(ROSA)26Sor locus] were randomly divided into three groups (vehicle, FAST without illumination, and FAST with FRL). The minicircle DNA vectors encoding the FAST system {pYH412 (PhCMV-p65-VP64-BldD-pA::PhCMV-BphS-P2A-YhjH-pA, 81 μg), pYH413 [PU6-sgRNA (tdtomato stop cassette)::PhCMV-DocS-Linker-Cas9(C)-NES-pA, 173 μg], and pYH414 [PFRL-NLS-Cas9(N)-Linker-Coh2-pA, 46 μg]} were dissolved in Ringer’s solution [NaCl (8.6 g/liter), KCl (0.3 g/liter), and CaCl2 (0.28 g/liter)] and injected into mice’s tail vein by hydrodynamic injection. The injection volume of the DNA mixture solution was 100 μl per mouse weight (gram). Twenty-four hours after injection, mice were illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off, alternating, to avoid the thermal discomfort in mice caused by continuous illumination) for 4 hours per day for 3 days (according to the time schedule in Fig. 5A). A second-round injection of the minicircle-based FAST system was performed on the fifth day, followed by 4 hours of daily illumination for three additional days. On the 15th day after the first hydrodynamic injection, mice were euthanized, and the livers were isolated for fluorescence imaging or histological analysis. The tdTomato signal from isolated liver was detected using IVIS Lumina II in vivo imaging system (PerkinElmer, USA) and frozen tissue section histological analysis.

Frozen tissue section

First, dissected liver tissue blocks were soaked in 4% paraformaldehyde for 2 hours. Subsequently, the tissue blocks were stepwise dehydrated with 15% sucrose solution overnight and then soaked in 30% sucrose solution for another 3 hours. After being washed three times with PBS, freshly dissected tissue blocks (<5 mm thick) were placed on to a prelabeled tissue base mold and embedded in Tissue-Tek optimal cutting temperature (O.C.T.) compound (catalog no. 4583; Sakura). These tissue blocks were stored at −80°C for freezing until ready for sectioning. The tissues were sliced into frozen sections with 5-μm thickness using Cryostat Microtome (Clinical Cryostat; CM1950; Leica) for further processing or stored at −80°C ultralow-temperature freezer.

Gene editing mediated by the FAST system in tumor mouse model

A total of 5 × 106 of A549 cells were suspended in 0.2 ml of sterile PBS and subcutaneously injected onto the back of the 6-week-old female BALB/c nude mice (ECNU Laboratory Animal Center). When the tumor size reached about 80 to 100 mm3, APC/FAST complex containing 20 μl of APC and the minicircle iteration of the FAST system {pYH412 (PhCMV-p65-VP64-BldD-pA::PhCMV-BphS-P2A-YhjH-pA, 2.7 μg), pYH414 [PFRL-NLS-Cas9(N)-Linker-Coh2-pA, 1.5 μg], and pYH420 [PU6-sgRNA (PLK1)::PhCMV-DocS-Linker-Cas9(C)-NES-pA, 5.8 μg]} were injected intratumorally. These injected mice were randomly divided into two groups (dark and illumination). Injections were conducted under anesthesia once every 2 days for five times. Twenty-four hours after every injection, mice were illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off, alternating, to avoid the thermal discomfort in mice caused by continuous illumination) according to the time schedule in Fig. 6A or kept in dark. Mice of the vehicle control group were intratumorally injected with 20 μl of APC and 50 μl of PBS and were then illuminated with FRL (10 mW/cm2; 730 nm; 2-min on, 2-min off), as indicated in Fig. 6A. The tumor sizes and the body weights of mice were measured every 2 days. On the 15th day after the first intratumor injection, all mice were sacrificed and tumor weights were recorded. The tumor volumes were measured using a digital caliper and calculated by the following formula: tumor volume = [length of tumor × (width of tumor)2]/2. Then, tumors were isolated for indel mutation analysis and tumor apoptosis detection by hematoxylin and eosin (H&E) staining, TUNEL, and caspase-3–labeling assays.

H&E staining of frozen tumor tissue sections

Glass slides that hold the frozen tissue sections were washed with PBS three times for 5 min each time, transferred to 0.5% Triton X-100 (dissolved in PBS; Sigma-Aldrich) for 10 min, and washed with PBS twice for 5 min each time. The slides were rinsed in running tap water at room temperature for 1 min. The samples were then stained in hematoxylin staining solution (catalog no. E607317; Sangon Biotech Inc.) for 8 min and washed in running tap water for 10 min. Next, the samples were differentiated in 1% acid alcohol for 10 s, washed in running tap water for 30 min, and were then counterstained in eosin staining solution (catalog no. E607321; Sangon Biotech Inc.) for 30 s to 1 min and washed in running tap water for 10 min. Last, the tissue sections were sealed by a drop of mounting medium over the tissue and then covered by a coverslip. The prepared slides were then observed by a microscope (DMI8; Leica) equipped with an Olympus digital camera (Olympus DP71; Olympus).

TUNEL staining of frozen tumor tissue sections

A TUNEL Apoptosis Assay Kit (catalog no. 30063; Beyotime Biotechnology Inc.) was used to evaluate tumor tissue apoptosis according to the manufacturer’s instructions. After washing three times with PBS, the slides were incubated with 4′,6-diamidino-2-phenylindole (DAPI) solutions (5 μg/ml; catalog no. C1002; Beyotime Inc.) for 2 to 5 min at room temperature. The slides were further washed three times with PBS and mounted with the antifade mounting media. Last, the slides were sealed and observed by a fluorescence microscope (DMI8; Leica) equipped with an Olympus digital camera (Olympus DP71; Olympus). TUNEL-positive nuclei were stained green, and all other nuclei were stained blue.

Caspase-3 labeling of frozen tumor tissue sections

Isolated tumor frozen tissue sections were thawed at room temperature for 15 min and rehydrated in PBS for 10 min. The tissue samples were surrounded with a hydrophobic barrier using a barrier pen after draining the excess PBS. Then, the slides were soaked in 0.5% Triton X-100 (dissolved in PBS; catalog no. 9002-93-1; Sigma-Aldrich) for 20 min. Nonspecific staining between the primary antibodies and the tissue samples was blocked by incubating sections in the block buffer (1% FBS in PBS) for 1 hour at room temperature. After incubating with the anti–caspase-3 antibody (1:100; catalog no. ab32351; Abcam) overnight at 4°C, the slides were washed three times for 15 min each time in PBS and then incubated with the Alexa Fluor 555 goat anti-rabbit immunoglobulin G antibody (1:500; catalog no. ab150078; Abcam) for 1 hour at room temperature. After washing three times with PBS, the slides were incubated with DAPI solutions (5 μg/ml; catalog no. C1002; Beyotime Inc.) for 2 to 5 min at room temperature. The slides were further washed three times with PBS and mounted with the antifade mounting media. Last, the slides were sealed and observed by a fluorescence microscope (DMI8; Leica) equipped with an Olympus digital camera (Olympus DP71; Olympus). Caspase-3–positive cytoplasm was stained red, and all nuclei were stained blue.

Ethics

All experiments involving animals were conducted in strict adherence to the guidelines of the ECNU Animal Care and Use Committee and in direct accordance with the Ministry of Science and Technology of the People’s Republic of China on Animal Care. The protocols were approved by the ECNU Animal Care and Use Committee (protocol IDs, m20180105 and m20190607). All mice were euthanized after the termination of the experiments.

Statistical analysis

All in vitro data represent means ± SD and are described separately in the figure legends. For the animal experiments, each treatment group consisted of randomly selected mice (n = 3 to 5). Comparisons between groups were performed using Student’s t test, and the results are expressed as means ± SEM. GraphPad Prism software (version 6) was used for statistical analysis.

SUPPLEMENTARY MATERIALS

Supplementary material for this article is available at http://advances.sciencemag.org/cgi/content/full/6/28/eabb1777/DC1

https://creativecommons.org/licenses/by-nc/4.0/

This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

REFERENCES AND NOTES

Acknowledgments: We are grateful to all the laboratory members for cooperation in this study, especially J. Jiang, S. Zhu, and X. Yang. Funding: This work was financially supported by the grants from the National Key R&D Program of China, Synthetic Biology Research (no. 2019YFA0904500), the National Natural Science Foundation of China (NSFC; no. 31971346 and no. 31861143016), the Science and Technology Commission of Shanghai Municipality (no. 18JC1411000), the Thousand Youth Talents Plan of China, and the Fundamental Research Funds for the Central Universities to H.Y. This work was also partially supported by NSFC no. 31901023 to N.G. We also thank the ECNU Multifunctional Platform for Innovation (011) for supporting the mouse experiments and the Instruments Sharing Platform of School of Life Sciences, ECNU. Author contributions: H.Y. conceived the project. H.Y. and Y.Y. designed the experiment, analyzed the results, and wrote the manuscript. Y.Y., X.W., J.S., H.L., and Y.C. performed the experimental work. Y.P., D.L., and N.G. analyzed the results and revised the manuscript. All authors edited and approved the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors. All genetic components related to this paper are available with a material transfer agreement and can be requested from H.Y. (hfye{at}bio.ecnu.edu.cn).
View Abstract

Stay Connected to Science Advances

Navigate This Article