The stimuli-responsive CRISPR-Cas12a system for modulating the selective aggregation of cell membrane receptors

Abstract

While CRISPR systems exhibit remarkable programmability in the field of nucleic acid editing, their extension to protein engineering faces a fundamental challenge, namely the traditional CRISPR tools lack the design to efficiently convert stimulus signals into the selective clustering of membrane receptors. This study develops a stimulus-responsive membrane-confined CRISPR-Cas12a platform that enhances selective clustering of membrane receptors for functional regulation. Specifically, a membrane-anchored DNA tetrahedral framework (TD-apt) was designed, which leverages vascular endothelial growth factor (VEGF) to activate Cas12a. Compared with unconfined CRISPR-Cas12a, membrane-confined CRISPR-Cas12a exhibits stronger cleavage activity, the interaction between the cellular-mesenchymal epithelial transition factor (c-Met) receptor and transferrin receptor (TfR) on A549 cells was efficiently modulated by nucleic acid assembly. This manipulation selectively inhibited c-Met function through spatial steric hindrance of TfR, modulating cellular behavior. Notably, the system’s generality was validated by engineering of c-Met homodimerization for activation. This cascading regulatory paradigm of environmental sensing (VEGF response)-nucleic acid computation (CRISPR-based nucleic acid molecular computation)-protein assembly (receptor topological remodeling) effectively extends CRISPR’s application boundaries to the field of non-genetic regulation protein-protein interaction (PPI) and establishes a versatile toolkit for dynamic and precise functional regulation.

Graphical abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s12951-026-04405-2.

Introduction

Cell surface receptors act as the gatekeepers of the cell and can activate intracellular downstream cascades by sensing extracellular signaling molecules, thereby precisely regulating cellular behaviors [1–3]. Among these, the receptor tyrosine kinase (RTK) family, as a class of ubiquitous membrane receptors, is frequently overexpressed in epithelial-derived solid tumors (e.g., lung and breast cancers), where it drives proliferation, migration, invasion, and angiogenesis [4, 5]. Consequently, inhibiting RTK function has become an essential strategy in targeted cancer therapy [6, 7]. Beyond oncology, RTK activation also plays critical roles in embryonic development and wound healing [8, 9]. Notably, RTKs typically function not as monomers but through dimerization-induced structural approximation, which activates their intracellular kinase domains and trigger downstream cellular events [10, 11]. This mechanistic paradigm renders the artificial control of RTK dimerization a compelling approach for user-customizable programming of cellular activities.

Traditional genetic engineering has served as the primary strategy for modulating receptor oligomerization. However, it’s time-consuming nature and unpredictable outcomes have spurred urgent demand for non-genetic alternatives [12, 13]. Among these, DNA architectures-mediated receptor oligomerization has become one of the most attractive of the non-genetic engineering strategies. It has evolved from the early use of bivalent DNA aptamer assemblies as receptor dimerization agonists [14, 15] or steric hindrance-based inhibitors (protein pairing) [16], to dynamically adjustable oligomerization systems that can respond to exogenous stimuli such as light and chemicals [17–21], and recently to smart-responsive strategies leveraging pathological microenvironmental cues for autonomous receptor clustering [22–24]. This adaptable and versatile stimuli-responsive paradigm aligns with the growing demand for precision regulation, positioning DNA nanotechnology as a reliable platform for developing intelligent receptor modulation strategies. For example, in the tumor microenvironment, vascular endothelial growth factor (VEGF), secreted by tumor cells, is typically highly expressed as a prototypical pathological microenvironmental cue and serves as a major driver of tumor angiogenesis [23, 25, 26]. While endogenous stimulus responses ensure high specificity and effectively reduce side effects, their weaker signal strength compared with exogenous stimuli poses a challenge for reliable receptor modulation.

In recent years, RNA-guided clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated proteins (Cas) systems have emerged as revolutionary tools in genome editing and molecular diagnostics due to their powerful nucleic acid-editing capabilities [27–30]. Beyond these applications, their stringent molecular recognition and robust trans-cleavage amplification have been harnessed to develop ultrasensitive biosensors by many researchers, serving as signal-amplifying elements for detecting diverse analytes [31–34]. However, current research on CRISPR system remain underexplored for spatial regulation of PPI and membrane receptor arrangement due to the traditional CRISPR tools lack the design to efficiently convert stimulus signals into the selective clustering of membrane receptors. Therefore, there is an urgent need to design new stimuli-responsive strategies based on CRISPR/Cas12a that couples environmental stimulus signals (VEGF) to Cas12a-mediated signal amplification, enabling efficient, precise and selective aggregation of cell membrane receptors, thereby extending the functional range of the CRISPR system and establishing new paradigms for stimulus-responsive regulation of protein interactions.

Herein, this study proposes a stimuli-responsive membrane-confined CRISPR-Cas12a system for modulating membrane receptor aggregation by leveraging the unique sensing and amplification features of Cas12a. An intelligent TD-apt was designed to couples VEGF stimulation with Cas12a activation, wherein Cas12a-mediated cleavage of stem-loop hairpins expands the VEGF-triggered receptor assembly from a 1:1 linear response to a 1:n amplified response. This design ultimately enables nucleic acid-directed selective clustering of membrane receptors, thereby regulating intracellular downstream signaling and cellular behavior (Fig. 1). Specifically, TD was anchored to the cell membrane by cholesterol, with an extended targeting strand (TS) that complements the VEGF aptamer, forming TD-apt. In addition, probes H1 and H3 were meticulously designed to target c-Met and TfR, incorporating DNA hairpin and RNA linker structure based on the c-Met and TfR aptamer sequence [16, 35]. Notably, Cas12a’s trans-cleavage activity has no effect on RNA sequences, so the RNA linker is preserved, providing a sequence basis for subsequent hybridization. When VEGF is present, it binds to the VEGF aptamer, exposing the TS to activate the CRISPR-Cas12a. The responsiveness of VEGF can better ensure the tumor specificity of the stimulus-responsive regulatory strategy. The trans-cleavage activity of CRISPR-Cas12a disrupts the hairpin structure of H1 and H3, exposing their complementary sequences, which further promotes the dimerization of the c-Met and TfR mediated by nucleic acid assembly, inhibiting c-Met function via the steric hindrance effect of TfR. To demonstrate the versatility of this strategy, when the TfR-targeting probe H3 is replaced with another c-Met-targeting probe H2, it further induces c-Met homodimerization and activates c-Met function. In summary, this environmental sensing-nucleic acid computation-protein assembly strategy achieves selective receptor aggregation, offering a novel and customizable approach for artificially manipulating receptor functions and cellular behaviors.

Fig. 1.

Schematic illustration of the stimuli-responsive CRISPR-Cas12a system for modulating the selective aggregation of cell membrane receptors.

Experimental

Flow cytometry assay

Flow cytometry was employed to analyze changes in cellular fluorescence intensity under different experimental conditions. For instance, to quantitatively assess the dynamic stability of TD-apt on the cell membrane surface, flow cytometry was used to monitor the signal changes of fluorescently labeled probes. A549 cells in the logarithmic growth phase were digested and centrifuged, and the cell density was adjusted to 2 × 10⁵ cell/mL. The cells were incubated with FAM-TD and FAM-TD-apt-BHQ1, respectively. After incubation, flow cytometric analysis was performed at 0, 0.5, 1, 1.5, and 2 h to detect the changes in their FAM fluorescence intensity.

Fluorescence measurement

The FAM-TD-apt-BHQ1 (FAM is quenched state due to neighboring BHQ1) complex was prepared by complementary hybridization of FAM-TD with apt-BHQ1. Different concentrations of VEGF (0.5 µg/ml-4 µg/ml) were co-incubated with FAM-TD-apt-BHQ1 (200 nmol/L) in a thermostatic metal shaker at 37 ℃ with 500 r/min for 30 min. After that, 200 µL of the mixture of VEGF and FAM-TD-apt-BHQ1 was added into a micro cuvette, and the excitation wavelength was set at the optimal excitation wavelength of 496 nm for FAM, and the scanning range of the emission wavelength was 510–600 nm, and the width of the slit was 1.5 nm. The VEGF-responsive performance of TD-apt was determined by analyzing the changes in FAM fluorescence intensity.

Confocal laser scanning microscopy (CLSM) imaging

A549 cells in the logarithmic growth phase were digested, centrifuged, and adjusted to a density of 1 × 10⁵ cells/mL. The cells were then seeded into confocal dishes and cultured for 24 h. The cells were incubated with TD-apt and 200 nmol/L Cy3-H1-BHQ2/ Cy5-H3-BHQ2 aptamer probes for 15 min, followed by the addition of VEGF for further incubation for 30 min. The unbound probes were removed by washing with PBS, and the Cas12a reaction system was added for co-incubation at 37 °C for 30 min. Finally, live-cell real-time imaging was performed using a confocal microscope. The H1 probes were excited with a 552 nm laser, and fluorescence signals between 560 and 650 nm were collected and pseudocolored green.

Preparation of cell lysates

A549 cells were seeded in 6-well plates and cultured until reaching 80% confluency. The culture medium was then replaced with serum-free medium (FBS-free) for serum starvation treatment over 24 h. Subsequently, TD-apt and H1/H3 probes were incubated with the cells for 15 min, followed by VEGF stimulation for 30 min. Unbound probes were removed by washing with PBS, and the Cas12a reaction system was added for further incubation at 37 °C for 30 min. After washing off the Cas12a system with PBS, cells were treated with 30 ng/mL HGF for 30 min. Control groups were treated equivalently with PBS instead of the TD-apt/Cas12a system. All cells were rinsed three times with PBS, and 150 µL of ice-cold lysis buffer (supplemented with protease and phosphatase inhibitors) was added to each well. The cells were gently pipetted on ice for 10 min, and the lysates were collected and centrifuged at 120,000 ×g for 10 min. The supernatants were aliquoted and stored at -80 °C for subsequent analysis.

Western blot assay

Protein concentrations of the cell lysates from each group were quantified using a BCA assay kit. Equal amounts of protein (20 µg) were taken from each group and separated by electrophoresis on 7.5% SDS-PAGE, then electrophoretically transferred onto PVDF membrane. After transfer, the PVDF membranes were blocked with a blocking buffer at room temperature for 2 h. Next, it was incubated with the primary antibody corresponding to the protein overnight at 4 °C, after which the membrane was washed three times with TBST. The corresponding secondary antibody was added and incubated at room temperature for 1 h, and again washed three times again with TBST. Finally, a pre-made chemiluminescent substrate was added to the PVDF membrane and imaged by a chemiluminescence imaging system.

Cell proliferation assay

A549 cells in the logarithmic growth phase were digested, centrifuged, and adjusted to a density of 5 × 10⁴ cells/mL. The cell suspension was seeded into 96-well plates (100 µL/well) and cultured for 18 h until adherence. The medium was replaced with serum-free medium for 24 h to synchronize the cell cycle. Cells were then incubated with TD-apt and varying concentrations of H1/H3 probes for 15 min, followed by VEGF stimulation for 30 min. After removing unbound probes, the Cas12a reaction system was added and co-incubated at 37 °C for 30 min. Finally, 30 ng/mL HGF was added to the cells and incubated for 24 h. Following the manufacturer’s instructions, CCK-8 reagent was added to each well and incubated in the dark for 1 h. Absorbance at 450 nm was measured using a multifunctional microplate reader.

Wound healing assay

The well-grown A549 cells were digested, centrifuged, and resuspended to an appropriate density. The cell suspension was seeded into 12-well plates and cultured until reaching 80% confluency. Serum-free medium was then used for 24 h starvation treatment. A uniform scratch was created by vertically pressing a 200 µL sterile pipette tip against the bottom of the wells and dragging steadily. After washing away detached cells with PBS, images of the scratch area were captured under a microscope and recorded as the 0 h time point. For the treatment group, pre-prepared TD-apt, H1/H3 probes, VEGF, and the Cas12a reaction system were sequentially added and incubated at 37 °C, followed by HGF stimulation. The control group underwent identical procedures with PBS replacing the TD-apt/Cas12a system. Pre-marked fields were re-imaged at 4 h, 8 h, 12 h, and 24 h to monitor wound closure dynamics.

Transwell invasion assay

50 µL of diluted Matrigel was spread evenly in the upper chamber of the Transwell, and left at 37 ℃ for 4 h to solidify. After digestion and centrifugation of the well-grown cells, 200 µL of cell suspension was seeded into the upper chamber, and the lower chamber was added with medium containing FBS. After 24 h incubation, cells were serum-starved in a serum-free medium for 24 h to eliminate basal migration activity. For the treatment group, pre-prepared TD-apt, H1/H3 probes, VEGF, and the Cas12a reaction system were sequentially added to the upper chamber and incubated at 37 °C, followed by 24 h stimulation with HGF. The control group received PBS instead of the TD-apt/Cas12a system while maintaining other procedures.

Finally, Transwell inserts were removed, gently washed with PBS, fixed with 4% paraformaldehyde for 30 min, and stained with 1% crystal violet for 20 min. Non-migrated cells on the upper membrane were removed using a cotton swab. Migrated cells were imaged under an inverted microscope and quantified using ImageJ software by counting cells in five randomly selected fields per insert.

Results

Assembly and characterization of TD-apt complex

TD-apt was composed of five DNA single strands: TH1, TH2, TH3, TS and VEGF aptamer. For assembly, the four single strands of TH1, TH2, TH3 and TS were firstly heated at 95 °C for 5 min, then gradually cooled to room temperature. Next, the mixture was reacted at 25 °C to generate TD, followed by the addition of VEGF aptamer to continue the reaction for 30 min, which ultimately resulted in the generation of the TD-apt complex. Next, polyacrylamide gel electrophoresis (PAGE) analysis was used to verify the assembly of TD-apt. As shown in Fig. 2a, lane 1 is the DNA Marker, lanes 2–6 are TH1, TH2, TH3, TS and VEGF aptamer single-stranded, respectively; lane 7 (TH1 + TH2) showed a new band with a lower migration rate than single-stranded, indicating the formation of double-stranded DNA; lanes 8 (TH1 + TH2+TH3) and 9 (TD) sequentially showed the characteristic bands at higher molecular weight positions, confirming the successful assembly of TD step by step. Notably, when VEGF aptamer was added, the band in lane 10 (TD-apt) shifted upward compared with that in lane 9 (TD), indicating that VEGF aptamer was successfully anchored to TD to form a higher molecular weight complex (TD-apt). The atomic force microscope image also confirmed the successful preparation of TD (Fig. 2b). Additionally, the hydrodynamic diameter of TD-apt analyzed by dynamic light scattering was 17.66 nm (Fig. S1), which was larger than that of TD (11.05 nm); Zeta potential measurements showed that the Zeta potential of TD-apt was − 7.19 mV (Fig. S2), which was also more negative than that of TD (-6.45 mV). All these results indicated that TD-apt was successfully prepared on the basis of TD.

Fig. 2.

Construction and characterization of TD-apt. (a) Gel electrophoresis characterization of the assembly of TD (lane 9) and TD-apt (lane 10). (b) The atomic force microscope image of TD. (c) Analysis of the stability of TD-apt anchored on cell membranes by cholesterol modification. (d) Normalized fluorescence spectra analysis of the response performance of TD-apt to VEGF. λex = 496 nm. (e) The CLSM images of A549 cells incubated with FAM-TD (i), FAM-TD-apt-BHQ1 (ii) and FAM-TD-apt-BHQ1 + VEGF (iii), respectively. λex = 488 nm. Scale bar: 50 μm

Following the successful assembly of TD-apt, this study further evaluated its cholesterol-mediated membrane anchoring capability and stability. First, FAM-labeled TS chains as well as BHQ1 quenching motif-modified VEGF aptamers were designed to construct FAM-TD (fluorescence-on state) and FAM-TD-apt-BHQ1 (fluorescence-quenched state) probes, respectively. Therein, the non-small cell lung cancer cell line (A549), which exhibits high c-Met expression, was selected as the model for analysis. Flow cytometry assay (Fig. 2c) showed that the fluorescence intensity of the cells was significantly increased after incubation with FAM-TD, while the FAM-TD-apt-BHQ1 group exhibited fluorescence quenching, and the fluorescence quenching signals on the cell membrane remained stable for 120 min. These results demonstrate that cholesterol modification enables effective and sustained anchoring of TD-apt on cell surfaces.

The stimulatory response of the TD-apt to VEGF

On the above basis, the intelligent responsiveness of TD-apt to VEGF was further validated. The specific principle is that after TD-apt recognizes VEGF, the interaction between VEGF and its aptamer exposes the TS in the TD that was originally occupied by VEGF aptamer. As shown in Fig. 2d, fluorescence spectroscopy analysis showed that when FAM-TD binds to VEGF aptamer-BHQ1 to form FAM-TD-apt-BHQ1, the fluorescence intensity of FAM was significantly reduced; upon VEGF addition, the fluorescence recovery was observed. Concentration-dependent experiments showed a positive correlation between fluorescence recovery intensity and VEGF concentrations (0–4 µg/mL) (Figs. S3a and S3b), further confirming target-activated fluorescence switching.

Encouraged by the above results, this study continued to verify the performance of TD-apt anchored to the cell membrane in response to VEGF using flow cytometry and confocal laser scanning microscopy (CLSM). The CLSM images showed that when A549 cells were incubated with FAM-TD, the FAM fluorescence signal was visible on the cell membrane (Fig. 2e-i). When incubated with FAM-TD-apt-BHQ1, the fluorescence signal on the cell membrane disappears because FAM fluorescence was quenched by BHQ1 (Fig. 2e-ii), and the situation changed markedly when VEGF was added, due to the interaction of VEGF with the VEGF aptamer-BHQ1, resulting in the restoration of FAM fluorescence signal on the cell membrane (Fig. 2e-iii). The quantitative results of cell fluorescence intensity in Fig. 2e are shown in Fig. S4. The results of the flow cytometry analysis (Fig. S5) were consistent with the CLSM imaging results, and the FAM fluorescence signal of the cells was effectively restored after the addition of VEGF. Therefore, TD-apt is expected to act as a VEGF responder to further activate CRISPR-Cas12a and promote selective aggregation of cell membrane receptor.

Stimuli-responsive TD-apt activates CRISPR-Cas12a system-mediated selective aggregation of cell membrane receptors

CRISPR-Cas12a, a nuclease derived from the Type V CRISPR system, exhibits trans-cleavage activities (activation-induced nonspecific single-stranded DNA cleavage), making it an ideal signal amplification module. Previous studies have demonstrated that stem-loop hairpin probes exhibit superior binding affinity to the catalytic pocket of Cas proteins compared to conventional linear probes [36]. Leveraging this principle, we designed a TS within the TD as the CRISPR-Cas12a activator and integrated TD-apt with CRISPR-Cas12a forming a VEGF-responsive signal-processing hub. This system converts VEGF stimulation into CRISPR-Cas12a activation, thereby harnessing its trans-cleavage activity to achieve precise cleavage of loops in hairpin probes H1 and H3 (Fig. 3a).

Fig. 3.

Activated CRISPR-Cas12a exhibits trans-cleavage activity. (a) Schematic diagram of CRISPR-Cas12a being activated and exerting trans-cleavage activity. (b) Normalized fluorescence spectra of Cy3-H1-BHQ2 probes with/without TD-activated CRISPR-Cas12a treatment on A549 cells. λex = 550 nm. (c) Normalized fluorescence spectra of Cy5-H3-BHQ2 probes with/without TD-activated CRISPR-Cas12a treatment on A549 cells. λex = 638 nm. (d) The CLSM images of A549 cells incubated with Cy3-H1-BHQ2 (i), Cy3-H1-BHQ2 + TD-activated CRISPR-Cas12a (ii) and Cy3-aptamer (iii), respectively. λex = 552 nm. (e) The CLSM images of A549 cells incubated with Cy5-H3-BHQ2 (i), Cy5-H3-BHQ2 + TD-activated CRISPR-Cas12a (ii) and Cy5-aptamer (iii), respectively. λex = 638 nm. Scale bar: 50 μm

To evaluate the cleavage activity of CRISPR-Cas12a, fluorescence-based validation of hairpin probe cleavage efficiency was first performed. The H1 probe (targeting c-Met) was labeled with Cy3 at its midsection and BHQ2 at the 3ʹ-end (Cy3-H1-BHQ2), while the H3 probe (targeting TfR) was labeled with Cy5 at its midsection and BHQ2 at the 3ʹ-end (Cy5-H3-BHQ2). In the intact hairpin configuration, the spatial proximity of the fluorophores (Cy3/Cy5) to the quencher (BHQ2) suppressed fluorescence emission. As shown in Fig. 3b, the Cy3-H1-BHQ2 probe alone exhibited minimal fluorescence, but its intensity significantly increased upon treatment with TD-activated CRISPR-Cas12a. A similar fluorescence recovery was observed for the Cy5-H3-BHQ2 probe (Fig. 3c), confirming that TD-activated CRISPR-Cas12a efficiently cleaved the hairpin structures of both probes, thereby separating the fluorophores from the quenchers and restoring fluorescence.

To verify the receptor-targeting capability of H1 and H3 probes and the cleavage activity of membrane-anchored TD-activated Cas12a, further validation was performed using CLSM imaging. When cells were incubated with Cy3-H1-BHQ2 probes alone, no Cy3 fluorescence was detected on the membrane (Fig. 3d-i). However, distinct Cy3 signals emerged on the cell surface after co-treatment with TD and CRISPR-Cas12a (Fig. 3d-ii). As a positive control, when the cells were incubated with the Cy3-aptamer alone, a significant fluorescence signal was also observed on the cell surface (Fig. 3d-iii). Similarly, activated CRISPR-Cas12a-mediated cleavage of the Cy5-H3-BHQ2 probe generated clear Cy5 fluorescence signals (Fig. 3e). The quantitative results of cell fluorescence intensity in Fig. 3d and e are shown in Fig S6. Compared with the aptamer labeled with Cy3/Cy5, the cleavage efficiency of the activated CRISPR-Cas12a-mediated probes for H1 and H3 is above 80%. These results demonstrate that H1 and H3 probes have good receptor-targeting capability and the TS within TD effectively activates the CRISPR-Cas12a system, enabling its trans-cleavage activity to disrupt the hairpin structures of H1 and H3 probes. This foundational validation underscores TD-activated CRISPR-Cas12a-mediated molecular processing, providing critical support for subsequent investigations.

Subsequently, we investigated whether this approach could effectively cleave the hairpin structures of H1 and H3 probes to expose their complementary sequences, thereby mediating c-Met and TfR aggregation via DNA hybridization. First, fluorescence resonance energy transfer (FRET) between H1-Cy3 and H3-Cy5 was employed for validation. Fluorescence spectra(Fig. S7) revealed that upon 550 nm excitation, A549 cells treated with Cy3-H1-BHQ2, Cy5-H3-BHQ2 and activated CRISPR-Cas12a exhibited a 7-fold increase in Cy5 fluorescence intensity at 650–700 nm compared to the control group (without CRISPR-Cas12a). These results demonstrate that CRISPR-Cas12a cleavage disrupts the hairpin constraints, liberating complementary sequences to drive Cy3-H1-H3-Cy5 hybridization and generate FRET signals, indicating spatial proximity between receptors. Next, CLSM imaging further confirmed the feasibility of this strategy (Fig. 4a). Upon VEGF activating the stimuli-responsive CRISPR-Cas12a, distinct Cy5 fluorescence signals (excited at 552 nm) emerged on the cell membrane (Fig. 4a-iii). In contrast, control groups lacking CRISPR-Cas12a (Fig. 4a-i) or containing non-activated Cas12a (Fig. 4a-ii) showed no detectable Cy5 signals (excited at 552 nm). These above findings indicate that VEGF unlocks CRISPR-Cas12a enzymatic activity to cleave Cy3-H1-BHQ2 and Cy5-H3-BHQ2 probes, enabling exposed complementary sequences to form a “molecular zipper” through hybridization. This forces spatial rearrangement of c-Met and TfR within the membrane plane.

Fig. 4.

VEGF activates the CRISPR-Cas12a system to edit PPI through nucleic acid assembly. (a) The CLSM images of A549 cells incubated with Cy3-H1-BHQ2 and Cy5-H3-BHQ2 (i), Cy3-H1-BHQ2 and Cy5-H3-BHQ2 + non-activated CRISPR-Cas12a (ii) and Cy3-H1-BHQ2 and Cy5-H3-BHQ2 + activated CRISPR-Cas12a using VEGF (iii), respectively. λex = 552 nm. (b) The CLSM images of FRET fluorescence intensity between Cy3-H1-BHQ2 and Cy5-H3-BHQ2 mediated by TS-apt (i) Td-apt (ii) and TD-apt (iii) combined with VEGF and CRISPR-Cas12a on A549 cells. λex = 552 nm. Scale bar: 50 μm. (c) Schematic diagram of modulating the interaction between c-Met and TfR proteins using H1 and H3 probes treated with activated CRISPR-Cas12a. (d) Western blot analysis of the inhibitory effects of H1 and H3 probes on the c-Met in A549 cells under different conditions

To investigate the central role of TD-apt in VEGF signal response and CRISPR-Cas12a activation, we designed two unconfined CRISPR-Cas12a groups (TS-apt without a tetrahedral scaffold and Td-apt without cholesterol modification) for comparative analysis. As shown in Fig. S8, compared to unconfined CRISPR-Cas12a (TS-apt and Td-apt), membrane-confined CRISPR-Cas12a (cholesterol-modified TD-apt) enhanced the efficiency of VEGF-triggered CRISPR-Cas12a activation, thereby achieving Cy3-H1-H3-Cy5 complementary hybridization, with the highest FRET fluorescence efficiency in these group. Further, CLSM imaging results showed that the FRET fluorescence intensity on cells in the TD-apt group (Fig. 4b-iii) was significantly higher than in the two unconfined CRISPR-Cas12a groups (Fig. 4b-i and b-ii). This is attributed to the cholesterol-modified TD-apt, which enables TD-CRISPR-Cas12a to anchor to the cell membrane surface and form the membrane-confined CRISPR-Cas12a. The spatial confinement effect (increasing its local concentration) and the natural mobility of the cell membrane effectively enhance the cleavage activity of CRISPR-Cas12a, thereby improving the efficiency of receptor clustering. To better utilize the precise regulation of VEGF-activated CRISPR-Cas12a trans-cleavage activity, the key reaction parameters of the CRISPR-Cas12a system were optimized in this study using a one-factor-at-a-time approach. Based on the FRET efficiency between H1-Cy3 and H3-Cy5 probes, the Cas12a concentration of 25 nmol/L, the ratio of crRNA/Cas12a of 1:1, the buffer of NEBuffer r2.1, the reaction time of 30 min, and the reaction temperature of 37 °C were finally selected as the optimal experimental conditions (Figs. S9-S13), which laid the foundation for the subsequent modulation of receptor function.

Stimuli-responsive CRISPR-Cas12a system for functional regulation of the c-Met by modulating PPI

The function of the c-Met relies on hepatocyte growth factor (HGF)-induced dimerization and autophosphorylation of its intracellular kinase domain, which triggers downstream PI3K/Akt and MAPK/Erk signaling cascades (Fig. 4c). To evaluate the functional impact of stimuli-responsive CRISPR-Cas12a-mediated c-Met-TfR aggregation on c-Met signaling, phosphorylation levels of c-Met and key pathway molecules were assessed by Western blot (WB) in A549 cells. As shown in Fig. 4d, cells exhibited growth arrest with minimal phosphorylation of c-Met (p-Met) and downstream effectors (p-Akt and p-Erk1/2) in the Serum starvation group. While the treatment with HGF significantly induced c-Met dimerization and pathway activation, markedly elevating p-Met, p-Akt, and p-Erk1/2 levels; on which the H1 and H3 probes intervention showed negligible effects on c-Met at this time. Based on H1 and H3 probes, only the VEGF-activated CRISPR-Cas12a intervention effectively blocked HGF-driven signaling, substantially reducing the expression of p-Met, p-Akt, and p-Erk1/2. These data demonstrate that the stimuli-responsive CRISPR-Cas12a system inhibits c-Met function by forcing c-Met and TfR co-aggregation, thereby disrupting the homodimerization of c-Met.

To evaluate the tumor-targeting specificity of stimuli-responsive CRISPR-Cas12a system combing with the probes, this study also examined its regulatory effects on human normal bronchial epithelial cells (BEAS-2B cell line). As shown in Fig. S14a, WB analysis revealed that c-Met expression in A549 cells was approximately 2-fold higher than in BEAS-2B cells. Subsequently, CLSM imaging further confirmed that the fluorescence intensity of A549 cells was significantly higher than that of BEAS-2B cells when incubated with Cy3-c-Met aptamer (Fig. S14b), and the results were consistent with WB. When BEAS-2B cells were subjected to identical treatments (Fig. S15), the stimuli-responsive CRISPR-Cas12a system failed to significantly suppress HGF-induced signaling, and phosphorylation levels of p-Met, p-Akt, and p-Erk1/2 showed little change in the experimental group versus the HGF-only control. These results collectively demonstrate that the VEGF-mediated TD-apt-activated CRISPR-Cas12a system selectively disrupts c-Met signaling in tumor cells. This cellular specificity likely stems from insufficient c-Met expression in BEAS-2B cells to support receptor clustering and subsequent pathway blockade.

Universal validation of CRISPR-Cas12a system-mediated selective aggregation of cell membrane receptors

After validating that the CRISPR-Cas12a system inhibits c-Met function in A549 cells by mediating c-Met and TfR co-aggregation, we further assessed the generality of this receptor-clustering strategy. To explore whether CRISPR-Cas12a could induce c-Met homodimerization (rather than hetero-aggregation), we redesigned the H3 probe into a structurally analogous H2 probe that also targets c-Met. This provides a new tool for further research into CRISPR-Cas12a-mediated H1 and H2 hybridization to explore c-Met homodimerization and activation (Fig. 5c). First, fluorescence spectroscopy(Fig. S16) confirmed that TD-activated CRISPR-Cas12a efficiently cleaved the hairpin structure of the Cy5-H2-BHQ2 probe, restoring Cy5 fluorescence. CLSM imaging validation as shown in Fig. 5a, no fluorescent signal of Cy5 was observed when cells were incubated with Cy5-H2-BHQ2 probes alone (control groups), and distinct membrane-localized Cy5 fluorescence emerged after treatment with activated CRISPR-Cas12a.

Fig. 5.

Universal validation of CRISPR-Cas12a system-mediated PPI through nucleic acid assembly. (a) The CLSM images of A549 cells incubated with Cy5-H2-BHQ2/Cy5-H2-BHQ2 + TD-activated CRISPR-Cas12a. λex = 638 nm. (b) The CLSM images of A549 cells incubated with Cy3-H1-BHQ2 and Cy5-H2-BHQ2 (i), Cy3-H1-BHQ2 and Cy5-H2-BHQ2 + non-activated CRISPR-Cas12a (ii), and Cy3-H1-BHQ2 and Cy5-H2-BHQ2 + activated CRISPR-Cas12a (iii), respectively. λex = 552 nm. Scale bar: 50 μm. (c) Schematic diagram of modulating the interaction between c-Met and c-Met using H1 and H2 probes treated with activated CRISPR-Cas12a. (d) Western blot analysis of the activation effects of H1 and H2 probes on the c-Met receptor in A549 cells under different conditions

Having confirmed the effective cleavage of the H2 probe by activated CRISPR-Cas12a, we further investigated the hybridization between H1 and H2 probes via FRET. Fluorescence spectra revealed a 10-fold increase in Cy5 intensity (650–700 nm) under 550 nm excitation in cells co-treated with activated CRISPR-Cas12a, Cy3-H1-BHQ2 and Cy5-H2-BHQ2 probes compared with the control group (only Cy3-H1-BHQ2 and Cy5-H2-BHQ2 probes) (Fig. S17). Similarly, CLSM imaging demonstrated distinct Cy5 signals (552 nm excitation) on cell membranes after CRISPR-Cas12a activation (Fig. 5b-iii), whereas control groups showed negligible fluorescence (Fig. 5b-i and b-ii). These results indicate that VEGF unlocks CRISPR-Cas12a activity to trigger H1 and H2 hybridization, thereby inducing c-Met homodimer formation. Critically, WB analysis (Fig. 5d) confirmed that stimulus-activated CRISPR-Cas12a mediated functional c-Met dimerization and activation, its autophosphorylation levels reached a level equivalent to that after HGF treatment. This demonstrates the system’s capacity to mimic HGF ligand function and may position it as a viable alternative to HGF. Collectively, these findings establish that the CRISPR-Cas12a system can bidirectionally switch receptor modulation modes (inhibition or activation) through the rational design of probes, highlighting its exceptional programmability and versatility.

Stimuli-responsive CRISPR-Cas12a system regulates the behavior of A549 cells through modulating c-Met-TfR aggregation

Given the pivotal role of HGF-mediated c-Met signaling in tumor progression, such as cell migration and proliferation are closely linked to cancer metastasis. Therefore, after investigating that the stimuli-responsive CRISPR-Cas12a system could functionally regulate the c-Met receptor, its effect on cell malignant phenotypes by inhibiting c-Met receptor function was further explored. First, cell counting kit-8 (CCK-8) assays assessed the effect of H1 and H3 probes (with stimuli-responsive CRISPR-Cas12a) on A549 cell proliferation. The results showed that the proliferation inhibition rate increased significantly with increasing concentrations of H1 and H3 probes in a dose-dependent manner, in which the proliferation inhibition in the 100 nmol/L treatment group achieved 30% at 24 h (Fig. 6a). Furthermore, scratch wound healing assays evaluated migration capacity under different treatments (Fig. S18). Only the CRISPR-Cas12a-activated H1 and H3 system (100 nmol/L) significantly blocked HGF-induced migration, and cell migration rates were reduced compared to the control group (HGF group) at 4 h, 8 h, 12 h, and 24 h time points (Fig. 6b). Similarly, Transwell invasion assays were employed to determine the cell invasion ability (Fig. 6c). The results revealed that activated CRISPR-Cas12a combined with H1 and H3 probes (100 nmol/L) reduced invasive capacity to 40% of control levels (Fig. 6d). Collectively, these findings establish that CRISPR-Cas12a-directed blockade of c-Met signaling via H1 and H3 probes effectively suppresses malignant behaviors in tumor cells.

Fig. 6.

Activated CRISPR-Cas12a combined with H1 and H3 probes influence cell behaviors. (a) The CCK-8 assay to analyze the effect of activated CRISPR-Cas12a combined with H1 and H3 probes on the proliferation capacity of A549 cells. (b) Quantification analysis of the effects of H1 and H3 probes without/with activated CRISPR-Cas12a on the migration capacity of A549 cells. (c) Transwell invasion assay of A549 cells after treatment with different conditions. (d) Quantification analysis the effects of H1 and H3 probes without/with activated CRISPR-Cas12a on the invasion capacity of A549 cells

Conclusions

This study has successfully developed a stimuli-responsive membrane-confined CRISPR‑Cas12a platform. By coupling VEGF signals with the amplification effect of Cas12a, the platform achieves non‑genetical, efficient, selective clustering of cell membrane receptors, thereby realizing receptor functional regulation. Compared with unconfined CRISPR-Cas12a, membrane-confined CRISPR-Cas12a exhibits stronger cleavage activity towards H1 and H3 probes, thereby effectively improving the regulatory efficiency of the interaction between c-Met and TfR proteins. WB results and cell behavior experiments confirmed that this significantly suppresses c-Met receptor signaling and downstream transduction pathways in A549 cells, thereby reducing tumor cell proliferation and invasive capabilities. Meanwhile, compared with normal cells, this strategy showed good selectivity for tumor cells that highly express c-Met. Further studies indicate that by replacing aptamer probes, this strategy can be extended to c-Met homodimerization modulating, activating receptors and downstream pathways. This study transforms CRISPR-Cas12a from a nucleic acid manipulator into a non-genetic platform for regulating PPI via TD-apt, creating an “environment sensing-nucleic acid computing-protein assembly” cascade interaction platform. In a nutshell, this strategy achieves artificially controllable programming of selective aggregation of cell membrane receptors by the membrane-confined CRISPR-Cas system, establishing a new paradigm for dynamically precise interventions. However, this study has not yet conducted in vivo experiments in mice for validation, which is one of the key limitations of the current study. In future studies, we will focus on designing, verifying, and ensuring the in vivo translational application of this stimulus-responsive CRISPR-Cas12a strategy.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Ya Wang: Methodology, Validation, Formal analysis, Writing - original draft, Visualization. Yujing Guo: Validation, Methodology, Investigation. Yamin Xiong: Conceptualization, Resources, Supervision. Xinyi Ren: Investigation, Validation. Yunli Zhao: Resources, Formal analysis. Lulu Song: Formal analysis, Validation. Leiliang He: Conceptualization, Supervision, Writing - review & editing, Funding acquisition.

Data availability

All data needed to support the conclusions in the paper are presented in the manuscript and the Electronic Supplementary Material. Additional data related to this paper may be requested from the corresponding author upon request.

Declarations

Competing interests

The authors declare no competing interests.