c‐Abl Kinase Targets Tight Junction Protein ZO‐2 in Regulation of Cell Migration and Morphology

ABSTRACT

c‐Abl is a non‐receptor tyrosine kinase involved in the regulation of cell migration and morphogenesis, but the underlying mechanism remains unclear. Here, we report the identification of tight junction protein ZO‐2 as a bona fide substrate of c‐Abl. We show that c‐Abl directly binds to and phosphorylates the C‐terminus of ZO‐2. In addition, c‐Abl stimulates the activity of JAK1, which subsequently phosphorylates the N‐terminus of ZO‐2. Using the RNAi‐mediated knockdown/rescue strategy, we demonstrate that c‐Abl regulates cellular morphology and migration through targeting ZO‐2 for phosphorylation. c‐Abl activity is also associated with decreased traction forces exerted on the cell substrate, thus corroborating c‐Abl kinase activity‐mediated inhibition of cell migration. Collectively, our data uncover ZO‐2 as a novel mediator for c‐Abl‐dependent regulation of cell migration.

c‐Abl binds and phosphorylates ZO‐2 directly at its C‐terminus and indirectly at the N‐terminus via JAK1. These phosphorylation events weaken cytoskeletal contractility, lower traction forces, and reduce cell migration, establishing ZO‐2 as a central mediator linking c‐Abl activity to changes in cell morphology and motility.

1. Introduction

Cell migration is a critical process in development, immune response, and homeostasis. Although the Abelson tyrosine kinase (c‐Abl) is best known for its roles in oncogenic transformation and nuclear signaling, accumulating evidence implicates its cytoplasmic functions in cytoskeletal regulation and, consequently, in migration. c‐Abl participates in diverse cellular processes including morphogenesis, proliferation, survival [1] and central nervous system development [2]. In the cytoplasm, c‐Abl interacts with F‐actin and G‐actin to regulate cell migration and morphological changes through actin cytoskeletal reorganization [3]. During the early stages of cell spreading, integrins stimulate c‐Abl kinase activity [4], which subsequently interacts with focal adhesion proteins [4, 5] and phosphorylates cytoskeletal regulatory proteins [6, 7, 8]. Through these phosphorylation events, c‐Abl can also indirectly influence both single‐cell motility and collective migration [9, 10, 11]. Together, these observations suggest that c‐Abl can modulate cellular dynamics relevant to cell motility, although the specific downstream effectors that couple c‐Abl to cytoskeletal remodeling remain incompletely defined.

In normal cells, the catalytic activity of c‐Abl tyrosine kinases is tightly regulated as a critical component of diverse signaling responses [12, 13]. c‐Abl can shuttle between the nucleus and the cytoplasm, facilitated by three nuclear localizing signals and a single nuclear exporting signal [14, 15]. c‐Abl performs distinct functions dependent on its subcellular distribution: nuclear c‐Abl is associated with G1 cell cycle arrest [16], DNA damage response [17], apoptosis [18], DNA repair and RNA polymerase II activation [19]. In contrast, less is known about the cytoplasmic functions of c‐Abl. Deregulated c‐Abl kinase activity is implicated in abnormal motility associated with diverse pathological consequences including chronic myeloid leukemia (CML) [20], acute lymphocytic leukemia (ALL) [21] and neurodegenerative diseases such as Alzheimer's [22] and Parkinson's diseases [23]. c‐Abl knockout in mice results in embryonic or postnatal lethality, morphological abnormalities, T and B cell lymphopenia [24, 25] and cardiac abnormalities [26]. In addition to these functions, c‐Abl is also reported to influence cell shape changes and morphogenetic programs in both development and disease contexts [8, 9, 27]. Together, these findings emphasize the broad physiological and pathological significance of c‐Abl, while also pointing to unresolved questions about its cytoplasmic functions in cytoskeletal regulation and cell migration.

Tight junction proteins ZO (Zonula Occludens, TJP)‐1, ZO‐2, and ZO‐3 are phosphoproteins and belong to the MAGUK family. ZO proteins are multidomain scaffolds that recruit multiple and diverse molecules at the tight junction (TJ) region. Deregulation of ZO‐2 function is associated with conditions such as familial hypercholanemia [28], nonsyndromic progressive hearing loss [29] and various epithelial cancers including breast cancer [30], testicular carcinoma [31] and pancreatic duct adenocarcinomas [32]. The loss or mutation of ZO proteins can result in the disruption of cell polarity [33] and tissue architecture [34]. It has been reported that ZO proteins contain multiple interaction domains (PDZ, SH3, and a guanylate kinase‐like domain) that organize junctional complexes and link them to the actin cytoskeleton. Among them, ZO‐2 (encoded by TJP2) has been implicated not only in the maintenance of paracellular barrier function and epithelial polarity but also in nuclear signaling and regulation of cytoskeletal architecture through direct and indirect interactions with F‐actin and junctional components [35, 36]. Moreover, ZO‐2 contributes to tissue fluidity and coordinated migration; post‐translational modification of ZO proteins, including tyrosine phosphorylation, alters junctional interactions and mechanical coupling between cells and the extracellular matrix [37, 38, 39].

Recent studies indicate that changes in ZO‐2 expression or phosphorylation status modulate collective migration and junctional plasticity, providing a potential mechanistic link between tight junction remodeling and tissue morphogenesis. These findings suggest that ZO‐2 is not merely a structural component of junctions but also an active regulator of cytoskeletal dynamics and multicellular coordination. Importantly, such roles parallel the cytoskeletal effects attributed to c‐Abl, raising the possibility of functional interplay between these two molecules [9, 10, 11, 37, 38].

As such, c‐Abl and ZO‐2 have both been individually implicated in cytoskeletal dynamics and cell junction regulation; however, the possibility of a direct regulatory relationship between them has not previously been explored. In this study, we have identified ZO‐2 as a novel substrate of c‐Abl, which directly binds to and phosphorylates the ZO‐2 protein. Multiple lines of evidence are provided to demonstrate that c‐Abl‐mediated ZO‐2 phosphorylation plays a critical role in regulating cellular morphology and migration. We believe that these findings will enhance our understanding of the cytoplasmic functions of c‐Abl and its key role in cell motility and morphogenesis, providing new insights into the molecular mechanisms underlying these processes. Furthermore, by combining inducible c‐Abl activation, biochemical mapping of ZO‐2 phosphorylation, and quantitative analyses of traction forces and monolayer tension, we demonstrate that ZO‐2 serves as a critical mediator linking c‐Abl activity to cytoskeletal organization and collective cell mechanics, thereby integrating junctional signaling with tissue‐level behaviors such as migration and morphogenesis.

2. Materials and Methods

2.1. Cell Culture, Transfection, and c‐Abl Induction

Human 293 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% heat‐inactivated fetal bovine serum (FBS), 100 μg/mL streptomycin and 100 units/mL penicillin and incubated at 37°C in 5% CO₂. Tet system approved FBS (Clontech) was used for 293FT‐TREX inducible cell lines (active c‐Abl KA and inactive c‐Abl KR). KA cells were engineered to express FLAG‐tagged wild‐type c‐Abl and KR cells were engineered to express a kinase‐inactive mutant [40]. For transfection assays, cells were seeded at least 12 h before transfection. Cells were transfected with Lipofectamine 2000 reagent (Life Technologies). Lipofectamine 2000 reagent and plasmids were diluted in Opti‐MEM medium (Sigma). For tetracycline induction, 293FT‐TREX KA and KR inducible cell lines were treated with 1 μg/mL of tetracycline for indicated time points to induce c‐Abl expression.

2.2. Generation and Validation of ZO‐2 Knockout (KO) Cell Lines Using CRISPR/Cas9

Target‐specific single guide RNA (sgRNA) sequences were designed using the Bioneer AccuTool platform (Bioneer, Daejeon, Korea), based on the presence of a 5′‐NGG‐3′ PAM motif. The selected sgRNA sequence was 5′‐AAAGATGGCAACCTTCACGA‐3′ (20 nt, excluding PAM). The sgRNA was synthesized and provided as a plasmid (1–2 μg; Bioneer, Cat. no. ATC‐0050). For control experiments, CCR5‐targeting sgRNA plasmids (Bioneer, Cat. no. ATS‐0055) were used. For Cas9 delivery, pRGEN Cas9 expression plasmids were obtained from Bioneer AccuTool. Depending on the experimental purpose, CMV/T7‐Puro‐RFP vectors (Bioneer, Cat. no. ATS‐0062) were used. Plasmids were prepared at 5 μg, according to the manufacturer's specifications. For transfection, cells were seeded at a density of 2 × 10⁶ cells in a 25T flask 24 h prior to plasmid delivery. Two micrograms of Cas9 plasmid and 2 μg sgRNA plasmid were introduced using AccuFect Transfection Reagent (Bioneer, Cat. no. K‐7920), following the manufacturer's protocol. The medium was replaced with fresh medium 6 h post‐transfection. At 24 h post‐transfection, cells expressing fluorescent reporters were sorted by fluorescence‐activated cell sorting (FACS). Single fluorescent‐positive cells were deposited into individual wells of 96‐well plates containing complete culture medium supplemented with Antibiotic–Antimycotic to prevent contamination. Only viable single‐cell–derived colonies were expanded and maintained for subsequent analyses. Genomic DNA was extracted from transfected cells using a genomic DNA preparation kit (EZ Genomic DNA Prep Kit, Cat. no. EP401‐50N), according to the manufacturer's protocol. The isolated gDNA samples were submitted to Macrogen Inc. (Seoul, Korea) for PCR amplification and Sanger sequencing of the target locus. The primers used for PCR were forward 5′‐GCATTGCATTTGGTTTTTATGTA‐3′ and reverse 5′‐ACTGTTACGACCCTCAATGTCTG‐3′. Sequencing results were analyzed using BioEdit software to quantify indel frequency and genome editing efficiency.

Then, we validated the KO efficiency through western blot (WB). For immunoblotting, cell lysates were prepared, and equal amounts of protein (40 μg per lane) were resolved by SDS–PAGE. Samples were separated on 8% polyacrylamide gels for 2 h and then were transferred onto PVDF membrane for 60 min. Membranes were blocked with 5% (w/v) skim milk for 1 h at room temperature and then incubated overnight at 4°C with primary antibody anti‐ZO‐2 (1:500; Abcam, Cat. no. ab224314, RRID: AB_95570) diluted in 3% skim milk. After washing with TBST, membranes were incubated with HRP‐conjugated secondary antibodies (1:1000, diluted in 5% skim milk) for 1 h at room temperature. Protein bands were visualized using enhanced chemiluminescence (ECL) reagents and imaged with a chemiluminescence detection system. β‐actin was used as a loading control (Sigma‐Aldrich, Cat. no. A5441, RRID: AB_476744).

2.3. Immunoprecipitation and Western Blot Assay

Harvested cells were rinsed with PBS and lysed in lysis buffer (50 mM Tris pH 8.0, 150 mM NaCl, 5 mM EDTA, 0.5% NP‐40, 2 mM PMSF, 20 mg/mL aprotinin, 25 mM NaF and 0.2 mM sodium orthovanadate). After centrifugation, the supernatant was incubated with Anti‐flag M2 magnetic beads (Sigma) for 4 h at 4°C. The samples were washed three times, and the supernatant was removed. Twice added SDS loading buffer was added and boiled. Prepared samples were loaded on a 12% SDS‐PAGE gel. Gels were transferred to a nitrocellulose membrane and incubated with blocking buffer, primary antibody, and secondary antibody (washed with TBS‐T in between each incubation time). The blots were visualized by ECL. Antibodies used in Western blot analysis were c‐Abl monoclonal antibody (K2, Santa Cruz), flag monoclonal antibody (Sigma‐Aldrich, Cat. no. F3165, RRID: AB_259529), ZO‐2 polyclonal antibody (Santa Cruz, sc‐11448 [H‐110], RRID: AB_2203583), phospho‐tyrosine monoclonal antibody (4G10, Cell Signaling Technology, Cat. no. 96215, RRID: AB_3096295), β‐actin monoclonal antibody (Sigma‐Aldrich, Cat. no. A5441, RRID: AB_476744), Jak1 polyclonal antibody (Cell Signaling Technology, Cat. no. 3332, RRID: AB_2128499), and phospho‐Jak1 (Tyr1022/1023) polyclonal antibody (Cell Signaling Technology, Cat. no. 3331, RRID: AB_2265057).

2.4. Binding Assay and In Vivo Kinase Assay

Either c‐Abl KA or KR was co‐transfected with the ZO‐2 into 293 cells. ZO‐2 was isolated by immunoprecipitation (IP) using Anti‐flag M2 magnetic beads (Sigma). Western blot analysis was performed with anti‐c‐Abl antibody (K2, Santa Cruz), anti‐flag antibody (Flag M2, Sigma‐Aldrich, Cat. no. F3165, RRID: AB_259529), anti‐phosphotyrosine antibody (4G10, Cell Signaling Technology, Cat. no. 96215, RRID: AB_3096295) and anti‐β‐actin antibody (Sigma‐Aldrich, Cat. no. A5441, RRID: AB_476744).

2.5. In Vitro Kinase Assay

ZO‐2 was enriched with IP and incubated with either active c‐Abl KA or inactive c‐Abl KR in kinase buffer (25 mM Tris pH 7.4, 10 mM MgCl2, 1 mM MnCl2, 0.5 mM DDT and 10 μM ATP) at 30°C for 15 min. The reaction was stopped by boiling in SDS sample buffer, and separated by SDS‐PAGE. A Western blot assay was performed using anti‐phosphotyrosine antibody (4G10, Cell Signaling Technology, Cat. no. 96215, RRID: AB_3096295), anti‐c‐Abl antibody (K2, Santa Cruz), anti‐flag antibody (Flag M2, Sigma‐Aldrich, Cat. no. F3165, RRID: AB_259529), and anti‐β‐actin antibody (Sigma‐Aldrich, Cat. no. A5441, RRID: AB_476744).

2.6. Site‐Specific Mutation

To generate phospho‐resistant mutant plasmids of ZO‐2 by substitution, PCR was performed with synthetic oligonucleotide primers using the QuikChange site‐directed mutagenesis kit (Agilent, Stratagene). To digest the parental DNA template and to select for mutation‐containing synthesized DNA, the mutant plasmids were treated with Dpn I endonuclease (target sequence: 5′‐Gm6ATC‐3′), which is specific for methylated and hemimethylated DNA. The nicked plasmids incorporating the desired mutations were then transformed into Escherichia coli . The mutated plasmids were then purified.

2.7. Quantitative Real‐Time PCR

Total RNA was isolated from KA cells using NucleoZOL (Macherey‐Nagel), and RNA quality and concentration were assessed with a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific). Complementary DNA (cDNA) was synthesized from 1 μg RNA using RevertAid Reverse Transcriptase with Oligo(dT)20 primers and dNTP Mix (Thermo Fisher Scientific), supplemented with RNase inhibitor (Takara Bio). Quantitative PCR was performed with SYBR Green (Toyobo) on a CFX Connect Real‐Time PCR Detection System (Bio‐Rad). GAPDH served as the internal control, and primers were purchased from BIONICS.

2.8. Immunofluorescence Imaging and Analysis

Cells were fixed in 3.7% formaldehyde solution in PBS, permeabilized with TX‐100 (0.01%), and incubated with blocking buffer (1% bovine serum albumin) and washed with PBS in between each incubation time. The samples were incubated with the indicated primary antibodies overnight at 4°C and then, with all secondary antibodies including DAPI stain to visualize nuclei, mouse Alexa fluor 488 and rabbit Alexa fluor 647 conjugated antibodies. Cells were mounted and visualized using fluorescence microscopy.

To quantitatively analyze actin stress fiber thickness, we used ImageJ software (NIH). A perpendicular line was drawn across each actin stress fiber in the cross‐sectional immunofluorescence images, and the corresponding fluorescence intensity profile was extracted (Figure S1). The full width at half maximum (FWHM) obtained by Gaussian curve fitting of the intensity profile was taken as the actin stress fiber thickness. To quantify paxillin signal intensity, ImageJ software (NIH) was also used. Cross‐sectional confocal images of paxillin near the basal plane were obtained from z‐stack images, and the average intensity within the cellular area of each cross‐section was measured (Figure S1).

2.9. siRNA Transfection and Quantitative‐Real‐Time‐PCR (q‐RT‐PCR) Analysis

Cells were transfected with siRNA using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions. The culture medium was replaced after 24 h, and total RNA was extracted using TRIzol reagent (Invitrogen) followed by chloroform extraction. cDNA was synthesized from 1 μg of total RNA using SuperScript III Reverse Transcriptase (Invitrogen). Quantitative PCR was performed using the QuantiTect SYBR Green PCR Kit (Qiagen) on a Monitor thermocycler.

2.10. Mass‐Spectrometry Analysis

Flag‐ZO‐2 was co‐transfected with either c‐Abl KA or KR. Cells were lysed in lysis buffer (50 mM Tris pH 8.0, 150 mM NaCl, 5 mM EDTA, 0.5% NP‐40, 2 mM PMSF, 20 mg/mL aprotinin, 25 mM NaF and 0.2 mM sodium orthovanadate). ZO‐2 was pulled down with flag tag‐conjugated beads and washed three times.

2.11. Optical Microscopy

Time lapse images of the cells were captured every 5 min using phase contrast on a DMI6000B microscope stand with a 5× NA 0.12 objective and a DFC345FX CCD camera (Leica). The imaging environment was maintained at 37°C/5% CO₂ in a heated enclosure. For the EGTA assay, images were captured every 2.5 s.

2.12. Wound Healing Assay

For more accurate quantification of the wound closure rate, circular wounds were introduced in 293FT KA/KR cell monolayers grown in a 96‐well plate. To make a circular wound, we placed a polydimethylsiloxane (PDMS) pillar (diameter = 0.5 mm) in each well before seeding the cells. The PDMS pillars were made by mixing the silicone elastomer and the cross‐linker (Sylgard 184, Dow Corning) at a 10:1 ratio and cured overnight on a hot plate at 80°C. Two c‐Abl inducible lines (KA and KR) were plated and transfected with either control siRNA or ZO‐2 siRNA. The cells were counted and plated in the prepared 96‐well plates with each well having a PDMS pillar (Figure 3). The next day, either wild type or each plasmid of ZO‐2 phospho‐resistant mutants was transfected into ZO‐2‐depleted KA cells. Cells were incubated for 24–30 h. For active c‐Abl induction, 1 μg/mL of tetracycline was treated to the samples. Six hours later, the pillar was removed leaving a cell‐free area. Immediately after removing the PDMS pillar, phase contrast images of every wound in each well of the 96‐well plate were captured every 10 min for 14 h. We obtained images for various cells in parallel using the microscope's motorized stage.

FIGURE 3.

Cell morphology changes by c‐Abl‐mediated ZO‐2 phosphorylation. (A) Identification of tyrosine residues phosphorylated by c‐Abl using Mass‐Spectrometry analysis. (ZO2A: ZO‐2 and KA were co‐transfected into HEK 293 cells, ZO2R: ZO‐2 and KR were co‐transfected into HEK 293 cells). (B) Schematic of the ZO‐2 structure and location of tyrosine phosphorylation sites. (C) List of the phospho‐resistant mutants of ZO‐2 and its abbreviations. (D) c‐Abl‐mediated cell morphology change was examined as a rescue experiment. As a control, either KA or KR cells were transfected with ZO‐2 siRNA and treated with tetracycline. After siRNA transfection, either wild type (WT) or each of phospho‐resistant mutants of ZO‐2 was reintroduced into KA cells. Then, tetracycline (Tet) was added to induce active c‐Abl.

2.13. Traction Force Microscopy and Monolayer Stress Microscopy

Polyacrylamide gels were prepared as a substrate for computing cell tractions. Gels with Young's modulus of 1.2 kPa were made by preparing a solution of 3% acrylamide (Bio‐Rad), 0.11% bisacrylamide (Bio‐Rad), 2 mg/mL acrylic acid N‐hydroxysuccinimide ester (NHS; Sigma), 0.014% 0.5 μm fluorescent particles (Life Technologies) and 0.033% ammonium persulfate (Bio‐Rad). 0.05% N,N,N′,N′‐tetramethylethylenediamine (TEMED; Bio‐Rad) was used to catalyze the reaction. Twenty‐four microliters of the polyacrylamide solution was placed onto a glass‐bottomed dish and covered with an 18‐mm diameter coverslip. During polymerization, the gels were centrifuged upside down at 115× g for 8 min to move the fluorescent particles to the top surface of each gel. After centrifuging, the gels swelled in water overnight at 4°C. To grow cells on the gel, the gel surface was functionalized by adding 200 μL of 2 mM sulfosuccinimidyl 6 (4′‐azido‐2′‐nitrophenyl‐amino) hexanoate (sulfo‐SANPAH; ProteoChem) to the surface of the gels and photoactivating with a UV lamp for 10 min. The gels were then rinsed and coated with 250 μL of 100 μg/mL collagen type I (Advanced BioMatrix) for about 8 h at 4°C. After the surface treatment, gels were rinsed 3 times with phosphate buffered saline and warmed up to room temperature. Then, cells were trypsinized from the culture flask and seeded on the gel. A small amount (4 μL) of dense cell suspension (8 × 10⁶ cells/mL) was plated on each gel covered with pre‐warmed culture medium, and the gels were kept in the incubator for about 48 h to allow cells to form a monolayer. To induce c‐Abl, half of the monolayers were incubated with tetracycline for 12 h (KA + Tet) and the other half was kept in tetracycline‐free medium (KA).

Time‐lapse images of cells (phase contrast) and particles beneath the polyacrylamide gel surface (fluorescence) were captured every 5 min for 6 h using the Leica microscope described above. After the time lapse, the cell medium was removed, the cells were rinsed with phosphate buffered saline, and trypsin was added. After waiting 10 min, the cells were gently aspirated to remove them from the polyacrylamide substrate, and a final image of the fluorescent particles was captured in the stress‐free state. Polyacrylamide substrate displacements were computed using image correlation. Tractions applied by the cells to the substrate were computed from the displacements with Fourier transform traction microscopy [41], using an implementation for finite substrate thickness [42, 43]. Stresses between the cells were computed by applying a force balance to the traction data using monolayer stress microscopy [44, 45]. The in‐plane stress tensor computed with monolayer stress microscopy has three unique components. We report here the largest principal stress (which is defined as the largest eigenvalue of the stress tensor), which is an indicator of the maximum component of stress transmitted within the monolayer.

2.14. Statistical Analysis

Data are presented as mean values with standard deviations. Differences between two groups were assessed using the Student's t‐test or one‐way ANOVA with Tukey's correction for multiple comparisons.

3. Results

3.1. c‐Abl Kinase Activity Induces Cell Morphology Changes

To investigate the mechanism underlying c‐Abl‐mediated alterations in cell morphology and cytoskeletal organization, we used tetracycline‐inducible 293 cell lines that express either FLAG‐tagged wild‐type c‐Abl (KA) or a kinase‐inactive mutant (KR) as previously described by Stuart et al. [40]. Western blot analysis confirmed that the expression of both c‐Abl (KA) and c‐Abl (KR) was markedly induced at 24 and 48 h after the addition of tetracycline (Tet) (Figure 1A). As expected, tyrosine kinase activity was detected only in c‐Abl (KA) cells, as evidenced by the anti‐Tyr immunoblot (Figure 1A) and was absent in c‐Abl (KR) cells. Interestingly, there was a drastic change in cell morphology specifically associated with c‐Abl kinase activity. When comparing c‐Abl kinase‐activated cells (KA + Tet) to the control cells (KA) without Tet, a significant alteration of the cell monolayer was observed, resulting in island‐like cell aggregates and a loss of typical single‐layer epithelial cell morphology (Figure 1B). In contrast, c‐Abl kinase‐inactive cells (KR + Tet) showed no change in the morphology.

FIGURE 1.

c‐Abl induced cell morphology change and potential substrates of c‐Abl. (A) c‐Abl‐inducible cells, FLAG‐tagged wild‐type c‐Abl (KA) and a kinase‐inactive mutant (KR), were incubated for 24 h and 48 h after tetracycline treatment (1 μg/mL). c‐Abl (theoretical 120–124 kDa) was observed at ~135 kDa. (B) c‐Abl‐inducible cell lines were induced using tetracycline to express KA and KR for 48 h. (C) Eight putative candidates of c‐Abl were selected by the consensus sequence of c‐Abl. (D) Each of the 8 candidates was depleted by siRNA‐mediated knockdown in KA cells with tetracycline (Ct siRNA: Control siRNA, KA: Kinase active c‐Abl, KR: Kinase inactive c‐Abl, +Tet: Tetracycline was added).

Given that the observed morphological changes were dependent on c‐Abl kinase activity, we hypothesized that c‐Abl induces these changes by phosphorylating cellular proteins. We tested the hypothesis by identifying potential substrates of c‐Abl with a focus on cytoskeletal proteins. For this purpose, Flag‐tagged c‐Abl complexes were affinity‐purified with anti‐Flag beads and analyzed by LC–MS/MS after proteolytic digestion, allowing unbiased identification of associated candidate proteins. In a proteomic analysis (Pull‐down/MS), we detected tyrosine phosphorylation of 26 cytoskeletal proteins exclusively in cells expressing the kinase‐active (KA) form (Table S1). From these, we selected eight candidates that contained the c‐Abl phosphorylation consensus sequence (I/V/L‐Y‐X‐X‐P/F) [46, 47] (Figure 1C).

We reasoned that the protein has to be expressed in order to mediate the effects of c‐Abl. Using c‐Abl activity‐induced morphology change as a functional readout, we assessed each of the eight candidates via siRNA‐mediated knockdown. Knockdown efficiency of each siRNA was validated by qRT‐PCR (Figure S1, Table S1), with additional Western blot confirmation for ZO‐2 (Figure S1). The results revealed that reduced expression of ZO‐2, PTPN14 (Tyrosine‐protein phosphatase non‐receptor type 14), NUMA1 (Nuclear Mitotic Apparatus Protein 1), and PRC1 (Protein Regulator of Cytokinesis 1) significantly attenuated c‐Abl activity‐induced morphology changes (Figure 1D). In contrast, knockdown of EPHA7 (Ephrin Type‐A Receptor 7), GEF2 (Rho/Rac Guanine Nucleotide Exchange Factor 2), SEPT2 (Septin‐2) and MEMO1 (Mediator of Cell Motility 1) did not induce a similar attenuation and kept the aggregated morphology (Figure 1D). KR‐expressing cells were included as controls to demonstrate that knockdown of each candidate gene alone did not have a significant impact on cell morphology. Among the four candidates showing attenuation, NUMA1 knockdown produced only a modest effect and was excluded from further analysis. We then prioritized ZO‐2 over PTPN14 and PRC1 because (i) as a tight‐junction scaffold, ZO‐2 is structurally and functionally linked to epithelial morphology, whereas PTPN14 and PRC1 are not directly implicated in maintaining cell morphology or migration in our system; and (ii) subsequent immunoprecipitation assays confirmed direct binding between c‐Abl and ZO‐2, providing a strong rationale for in‐depth analysis.

3.2. Tight Junction Protein ZO‐2 Is a Novel Substrate of c‐Abl

To determine whether ZO‐2 binds to c‐Abl, we performed immunoprecipitation (IP). Lysates prepared from cells co‐expressing Flag‐ZO‐2 and either c‐Abl (KA) or c‐Abl (KR) were subjected to anti‐Flag IP. The result of immunoblot showed that ZO‐2 binds to both c‐Abl (KA) and c‐Abl (KR), indicating that this binding is independent of kinase activity. Consistent with the proteomic data, anti‐p‐Tyr immunoblot detected tyrosine phosphorylation of ZO‐2 specifically in c‐Abl (KA) expressing cells, but not in c‐Abl (KR) expressing cells (Figure 2A).

FIGURE 2.

Tight junction protein ZO‐2 as a new substrate of c‐Abl. (A) Immunoprecipitation of ZO‐2 and either KA (active) or KR (inactive) c‐Abl. (B) Three truncated mutants (FL: full length, M: middle part of ZO‐2, ΔC: C‐terminal deletion mutant, ΔN: N‐terminal deletion mutant). (C) Either KA or KR was co‐transfected with one of the four ZO‐2 constructs (FL, ΔN, ΔC, and M) into HEK 293 cells. Then, ZO‐2 was pulled down with flag antibody (flag‐ZO‐2). (D) In vivo kinase assay with phospho‐resistant mutant (Y1118F) and wild type of ZO‐2. (E) In vitro kinase assay with FL and three deletion mutants of ZO‐2. (F) Endogenous JAK1 and phosphorylated JAK1 were detected after induction of either KA or KR (7 and 15 μg—amount of sample loaded). (G) The tyrosine phosphorylation levels of ΔC mutant with JAK1 inhibitor and JAK1 siRNA. c‐Abl was detected at ~135 kDa. Full‐length ZO‐2 (134 kDa, 1190 aa) migrated at ~150 kDa, likely due to post‐translational modifications. ZO‐2 truncations (ΔN: ~87 kDa, ΔC: ~98 kDa, M: ~54 kDa) appeared at ~110, 100, and 60 kDa. JAK1 and phospho‐JAK1 were observed at ~130 kDa.

To further characterize the interaction between c‐Abl and ZO‐2, we generated ZO‐2 deletion mutants by dividing it into three fragments: N‐terminal deleted (ΔN), only middle (M), and C‐terminal deleted (ΔC) peptides (Figure 2B). IP‐Western analysis revealed that c‐Abl binds to both full length (FL) and ΔN fragments of ZO‐2, but not to ΔC or the middle fragments (Figure 2C), indicating that c‐Abl binds to the C‐terminus of ZO‐2.

Consistent with the location of the c‐Abl consensus phosphorylation site (Y1118) at the C‐terminus, anti‐p‐Tyr detected strong phosphorylation of ΔN as well as FL ZO‐2. Unexpectedly, the ΔC‐terminus, which had no detectable binding to c‐Abl, was also found intensely phosphorylated by c‐Abl (Figure 2C), suggesting the presence of an intermediate tyrosine kinase mediating the phosphorylation of the ZO‐2 N‐terminus. In agreement with this possibility, the phospho‐resistant (Y1118F) mutant of ZO‐2 remained albeit slightly less phosphorylated (Figure 2D).

To ascertain the indirect phosphorylation of the ZO‐2 N‐terminus by c‐Abl, we carried out an in vitro kinase assay. Purified full‐length and deletion mutants of ZO‐2 were incubated with either c‐Abl (KA) or c‐Abl (KR). Only the full‐length and ΔN mutant of ZO‐2, but not the ΔC mutant, was phosphorylated by c‐Abl (KA), consistent with the notion that active c‐Abl directly phosphorylates the C‐terminal of ZO‐2 and indirectly the N‐terminus (Figure 2E).

Non‐receptor tyrosine kinase JAK1 was reported to phosphorylate the N‐terminus of ZO‐2 [48], and JAK1 is known to be activated by v‐Abl [49], raising the possibility that JAK1 might be the intermediate kinase. To test this possibility, we first examined whether c‐Abl could stimulate JAK1 activity using a specific phospho‐JAK1 (Tyr1022/1023) antibody as a surrogate marker of JAK1 activation [50]. Immunoblot analysis with this phospho‐specific antibody detected JAK1 phosphorylation specifically in c‐Abl (KA) but not c‐Abl (KR) expressing cells, consistent with previous findings that c‐Abl can activate JAK1 (Figure 2F) [51]. We next investigated whether JAK1 is responsible for the phosphorylation of the N‐terminus of ZO‐2 using a combination of a JAK1 specific inhibitor and JAK1 siRNA. Phospho‐tyrosine analysis indicated that inhibition of JAK1 was indeed associated with diminished phosphorylation of ZO‐2 N‐terminal peptides (Figure 2G). Collectively, these data support the conclusion that JAK1 acts as an intermediate kinase downstream of c‐Abl to phosphorylate ZO‐2 (Figure 2G).

3.3. C‐Abl‐Mediated ZO‐2 Phosphorylation Plays Significant Role in Cellular Morphology

The results shown in Figure 2C–E prompted us to identify additional phosphorylation sites on ZO‐2 beyond Y1118. To this end, we performed phospho‐proteomic analysis. Immuno‐purified ZO‐2 from either c‐Abl (KA) or c‐Abl (KR) expressing cells was subjected to MS/MS analysis of tyrosine phosphorylation. Indeed, apart from Y1118, we identified several additional sites of tyrosine phosphorylation in ZO‐2 isolated from c‐Abl (KA) but not c‐Abl (KR) cells (Figure 3A).

To determine the functional consequence of ZO‐2 phosphorylation by c‐Abl, we substituted each tyrosine (Y) residue with phenylalanine (F) individually or in combination (Figure 3B,C) and assessed each mutant for its ability to affect c‐Abl activity‐induced cell morphology changes. Specifically, we employed a depletion/rescue strategy to replace endogenous ZO‐2 with each phospho‐resistant mutant. To ensure the expression of exogenous ZO‐2, we used ZO‐2 siRNA targeting the 3′UTR of ZO‐2 so that only the endogenous ZO‐2 protein was knocked down. The results showed that re‐expression of wild‐type ZO‐2 in ZO‐2‐depleted cells almost completely restored c‐Abl‐induced morphology changes, indicating that exogenously expressed ZO‐2 was able to functionally replace endogenous ZO‐2 (Figure 3D). The Y1118F mutant of ZO‐2 exhibited markedly reduced ability to rescue when compared with wild‐type ZO‐2, indicating the functional importance of Y1118 phosphorylation. Expression of other phospho‐resistant mutants of ZO‐2, with mutation sites in addition to Y1118F, in ZO‐2‐depleted cells resulted in even more compromised rescue, suggesting that these additional phosphorylation sites are also functionally important (Figure 3D).

3.4. C‐Abl‐Mediated ZO‐2 Phosphorylation Plays a Significant Role in Cell Migration

To examine how c‐Abl‐induced ZO‐2 phosphorylation affects cell migration, we employed a wound‐healing assay that allows quantitative analysis. The control cells nearly filled the wound or gap within 14 h (Figure 4, and Videos S1, S2, S3). In sharp contrast, the activation of c‐Abl resulted in an almost complete halt of cell migration, leaving the gap largely unfilled (Figure 4A KA + Tet, and Video S4). This effect appears to be kinase activity‐dependent, as the expression of c‐Abl (KR) under the same condition did not impair wound healing.

FIGURE 4.

Inhibition of cell migration by c‐Abl kinase activity and ZO‐2 phosphorylation. (A) Wound healing assay. Tetracycline inducible KR cells and KA cells were used. Either wild type or each of phospho‐resistant mutants was reintroduced into ZO‐2 depleted KA cells and then c‐Abl was induced using tetracycline. (Time‐lapse imaging‐10 min intervals, 0: Starting point). (B) Quantification of wound closure (% of initial area). One‐way ANOVA revealed significant group differences (p < 0.0001), followed by Tukey's HSD post hoc test. Significance is indicated as ***p < 0.001, ****p < 0.0001. Data are based on 6–9 independent experiments.

We next investigated whether ZO‐2 mediates c‐Abl‐induced inhibition of cell migration by employing a gene depletion/rescue strategy. Interestingly, c‐Abl‐induced inhibition of cell migration was almost completely lost upon siRNA‐mediated depletion of ZO‐2, as evidenced by the mostly filled gap (Figure 4A KA + ZO‐2 siRNA + Tet, and Video S5). Remarkably, the re‐expression of wild‐type ZO‐2 in ZO‐2‐depleted cells fully restored the c‐Abl‐dependent inhibition of migration, demonstrating that ZO‐2 is largely responsible for mediating this inhibitory effect on migration.

We then assessed the significance of ZO‐2 phosphorylation in c‐Abl‐induced inhibition of cell migration by replacing wild‐type ZO‐2 Figure 4A KA + ZO‐2 siRNA + WT ZO‐2 + Tet and Video S6 with various phospho‐resistant mutants of ZO‐2 (Figure 4A). The results confirmed the critical role of c‐Abl‐mediated ZO‐2 phosphorylation in the regulation of cell migration. Among the identified phosphorylation sites, Y1118 seemed to play a major role, while phosphorylation at additional sites also contributed partially to the effect. This was demonstrated by comparing the ratio of the wound area at the start of the assay to the area of the same wound upon assay completion (Figure 4B).

3.5. Active c‐Abl Induces Cell Mechanical Force Changes Across the Cellular Monolayer Through ZO‐2

As shown, active c‐Abl induces marked alterations in cell morphology, most notably the formation of cellular aggregates that appear to be mediated by ZO‐2 (Figure 3D). In addition, the wound‐healing assay (Figure 4) further supported the significant role of c‐Abl‐mediated ZO‐2 phosphorylation in cell migration. Since cellular shape and migratory behavior are often governed by a balance of mechanical forces transmitted across the cell body, cell–matrix/substrate adhesions, and cell–cell junctions [44, 52, 53, 54, 55], the observed phenotypic changes could be related to modifications in these underlying force distributions. Therefore, we next examined whether c‐Abl activation might also alter such mechanical outputs. To address this, we applied traction force microscopy (TFM) and monolayer stress microscopy (MSM) to quantify cell–substrate tractions and intercellular stresses, respectively [41, 43, 44, 45]. Through these approaches, the mechanical interactions between cells and the underlying matrix/substrate and the mechanical stress across the monolayer and cell–cell junctions can be directly and quantitatively assessed.

As depicted in Figure 5A, traction refers to the force per area exerted by cells onto the substrate. Traction was analyzed in cellular monolayers under four conditions: KA, KA + Tet, KA‐ZO2‐KO, and KA‐ZO2‐KO + Tet. To first determine whether c‐Abl activation alters mechanical outputs associated with the observed changes in cellular morphology and migration, KA and KA + Tet cells were compared. The KA‐ZO2‐KO is a ZO‐2 knockout derivative of the active c‐Abl–inducible KA cell line, which is used to test whether these phenotypic changes were mediated through ZO‐2. Representative traction maps are shown in Figure S1. For quantitative comparison between c‐Abl‐non‐activated and c‐Abl–activated states, the root‐mean‐square of traction was computed over space, excluding data near the edges of the image as shown in Figure 5B. This process was repeated for many different cell monolayers under all conditions. As shown in Figure 5C, tractions exerted by c‐Abl‐activated cells (KA + Tet) were significantly lower than those of KA cells without Tet, implying that active c‐Abl perturbs and decreases the cell‐substrate traction forces. However, the traction force did not decrease significantly by active c‐Abl when ZO‐2 was depleted (KA‐ZO2‐KO + Tet) (Figure 5C), consistent with the notion that the traction reduction included by active c‐Abl was through ZO‐2.

FIGURE 5.

Reduced traction by active c‐Abl acting on ZO‐2 and cytoskeletal changes within c‐Abl‐induced cells. (A) Schematic representation of traction and tension within a group of cells. Black arrows indicate traction exerted on the substrate, and gray arrows represent tension across the cell monolayer. (B) Image of a representative cell monolayer with yellow shading indicating region used for analyzing traction and stress. (C, D) Traction and stress for the KA and KA‐ZO2‐KO cells with and without Tet. Dots indicate the root‐mean‐square traction and mean stress values for each monolayer; horizontal lines indicate means across all experiments in each group. In panel C, for KA cells, Tet− and Tet+ are statistically different (p = 0.0017, one‐way ANOVA with Tukey). In panel D, there are no statistical differences (p = 0.13, one‐way ANOVA). (E) Immunofluorescence image of KA cells with and without tetracycline. Paxillin (red), actin (green), and nucleus (blue) were stained. Scale bar = 20 μm. (F) Quantification of paxillin signal intensity in KA and KA + Tet cells (n = 20). (G) Quantification of actin stress fiber thickness in KA and KA + Tet cells (n = 10). Signification is indicated as ** p < 0.01, ***p < 0.001.

Then, to quantify intercellular forces within the monolayer, we performed MSM [44, 45]. MSM computes stresses (force per unit area) within the cell monolayer from the traction data by applying the principle of force equilibrium within the monolayer. From these data, we calculated the first principal stress within, which is transmitted within and between cells in the monolayer (Figure 5A). As shown in Figure 1B, induction of active c‐Abl in KA cells with tetracycline led to aggregate formation, which can create patches of cell‐free areas. To ensure reliable analysis under conditions of preserved monolayer integrity, MSM measurements were restricted to within 24 h of tetracycline treatment and to regions where the monolayer remained as a continuous cell sheet. As shown in Figure S1, measurements were performed only in areas where monolayer continuity was maintained. Under KA + Tet conditions, the monolayer edge appeared slightly shrunken and retracted but overall continuity was preserved. In contrast to traction, however, the stress across the cell monolayer did not show statistically significant differences among the experimental conditions (Figure 5D). Hence, the balance of traction and tension was altered in the c‐Abl activated cells, with a reduction in the magnitude of traction. Given that tractions pull the cells into free space [56], the altered tractions are consistent with the reduced rate of wound closure in the KA + Tet conditions.

3.6. Molecular and Protein Level Changes in Active c‐Abl‐Induced Cells Related to the Cellular Contractility

Next, we sought to identify molecular mechanisms underlying the observed traction changes, focusing on the cytoskeleton and adhesion complexes, which are known to play key roles in force generation and transmission. Quantitative PCR analysis revealed that myosin II mRNA, encoding a motor protein essential for actomyosin contractility, was markedly reduced when c‐Abl was activated (KA + Tet) compared with KA controls (Figure S1). Whereas this reduction was not observed in ZO‐2–knockout cells upon c‐Abl activation (KA‐ZO2‐KO + Tet versus KA‐ZO2‐KO; Figure S1). Alongside myosin II, we also assessed the expression of RhoA and Rac1, two upstream regulators of cytoskeletal organization, as well as actin, a major structural component of the cytoskeleton. In contrast to myosin II, the mRNA levels of RhoA, Rac1, and actin remained largely unchanged upon c‐Abl activation, both in KA cells (KA vs. KA + Tet) and in ZO‐2–knockout cells (KA‐ZO2‐KO vs. KA‐ZO2‐KO + Tet). We also examined focal adhesion components, including paxillin and focal adhesion kinase (FAK). Paxillin mRNA levels did not differ significantly between experimental groups, whereas FAK expression was markedly decreased upon c‐Abl activation (KA + Tet versus KA; Figure S1). Similar to myosin II, this reduction in FAK was not observed in ZO‐2–knockout cells (KA‐ZO2‐KO + Tet versus KA‐ZO2‐KO; Figure S1).

Lastly, we labeled and imaged the major machineries responsible for contractile and adhesive forces in cells, such as actin and focal adhesion proteins. The cell body appeared somewhat shrunken following aggregation in c‐Able activated cells (KA + Tet) compared with the control cells (KA) (Figure 5E). For paxillin, one of the major components of focal adhesions, quantitative analysis for paxillin dots was not feasible because the paxillin signal did not form clearly discernible focal‐adhesion puncta (Figure 5 and Figure S1). Signal intensity analysis from the cross‐sectional image of paxillin also did not exhibit a significant difference between KA and KA + Tet (Figure 5F). In contrast, actin stress fibers became visibly less prominent and thinner in KA + Tet cells compared with KA controls (Figure 5E). Quantitative analysis of immunostaining confirmed this observation, demonstrating a significant 37% decrease in actin stress fiber thickness in KA + Tet cells compared with KA controls (Figure 5G).

4. Discussion

c‐Abl, a member of the non‐receptor Src tyrosine kinase family, is known to contribute to signaling events that regulate diverse cellular functions. Depending on the localization of its substrates, c‐Abl can influence both nuclear and cytoplasmic cellular processes. In this study, we focused on the cytoplasmic activity of c‐Abl given that many cytoskeletal proteins were found to be tyrosine phosphorylated in response to c‐Abl activation. Using cellular morphology as a functional readout to screen for c‐Abl putative substrates, we identified ZO‐2 as the protein chiefly responsible for c‐Abl kinase‐dependent regulation of cell morphology and migration (Figures 3 and 4).

Binding studies demonstrated a direct association of c‐Abl to the C‐terminus of ZO‐2 in a kinase‐independent manner. This binding allows c‐Abl to target ZO‐2 for phosphorylation (Figure 2A,C). Specifically, c‐Abl phosphorylates Y1118, a residue within the consensus sequence at the ZO‐2 C‐terminus (Figure 2E). Apart from this direct phosphorylation site, c‐Abl also induces phosphorylation at additional sites indirectly through other tyrosine kinases, including JAK1, as revealed by siRNA‐mediated knockdown and the use of a specific inhibitor (Figure 2G). In agreement with the presence of additional phosphorylation sites, phospho‐proteomic analysis (Pull‐down/MS) uncovered eight new phosphorylation sites in the ZO‐2 protein (Figure 3A). Collectively, these results indicate that c‐Abl activity induces ZO‐2 phosphorylation at multiple sites via a combination of direct and indirect modes of action.

The critical role of ZO‐2 in mediating c‐Abl activity‐induced effects on cell morphology was investigated by depletion/rescue experiments. We first established the essential role of ZO‐2 by showing that knockdown of ZO‐2 abrogated, whereas the re‐expression of ZO‐2 restored c‐Abl‐dependent regulation of cell morphology and migration (Figures 3D and 4). In line with this kinase activity‐dependent regulation, we showed that substituting wild‐type ZO‐2 with phospho‐resistant mutants resulted in compromised rescue. Quantitative analysis revealed a major role of phosphorylation at Y1118, as well as other sites albeit to a lesser extent, in c‐Abl activity‐dependent regulation of cell migration.

The wound‐healing assay further supported the inhibitory role of c‐Abl kinase activity in cell migration. In line with our results, Kain and Klemke [8] have shown that c‐Abl reduces the rate of cell migration. Specifically, embryonic fibroblasts derived from abl ^−/− arg ^−/− mice fill in a wounded area more rapidly than normal counterparts, whereas reintroduction of wild‐type c‐Abl into abl ^−/− arg ^−/− cells with inhibits cell migration by preventing the formation of the Crk‐p130^CAS (CAS) complex. Additionally, cell movement is enhanced by treatment with the Abl inhibitor STI571 and inhibited by overexpression of active c‐Abl, reinforcing the notion that c‐Abl plays a negative role in cell migration [57].

Given the aggregated morphology and reduced migration observed in c‐Abl–activated cells, alterations in the distribution of mechanical forces were expected. The MSM results revealed that c‐Abl activation does not alter monolayer stress in either KA or KA‐ZO2‐KO cells (Figure 5D). Since monolayer stress arises primarily from forces transmitted through cell–cell junctions, particularly cadherin‐based adherens junctions [44, 58], this finding suggests that the overall capacity of the monolayer to transmit intercellular stresses is preserved. However, MSM measures mechanical stress transmission rather than the molecular strength of adhesive complexes [59, 60], and thus more subtle alterations in junctional composition or bonding affinity cannot be excluded.

Whereas monolayer stress did not change following c‐Abl activation, traction force measurements revealed a significant reduction in c‐Abl–activated KA cells (Figure 5C). Importantly, no such reduction was observed in ZO‐2–knockout cells upon c‐Abl activation (Figure 5C). In ZO‐2–knockout cells, baseline traction forces were slightly lower than in KA controls. Prior studies have shown that the depletion of other tight junction proteins can alter traction in an isoform‐dependent manner; for instance, ZO‐1 knockdown increased traction, whereas ZO‐3 knockdown decreased it [58]. Therefore, the removal of ZO protein can variably influence baseline traction, and ZO‐2 depletion itself also appeared to affect cellular baseline traction in our system. The central point for our study is that once ZO‐2 was knocked out, c‐Abl activation no longer led to a further reduction in traction. This indicates that ZO‐2 mediates the c‐Abl–induced decrease in traction forces.

Since traction forces represent the contractile force transmitted to the substrate through actin stress fibers and focal adhesions, these findings indicate that c‐Abl activation decreases cellular contractility. Supporting this interpretation, molecular analyses showed that myosin II—the key effectors of actomyosin contractility—was significantly reduced at the gene level in KA cells (Figure S1). Consistently, these molecular changes were not observed in KA‐ZO2‐KO cells, further supporting the role of ZO‐2 as an essential mediator (Figure S1). In line with this, actin stress fibers became thinner in c‐Abl–activated KA cells (Figure 5E,G). However, weakening of mechanical force (traction or cellular contractility) alone is unlikely to be sufficient to drive aggregation. KA‐ZO2‐KO cells display baseline traction comparable to KA + Tet yet do not exhibit robust aggregation, indicating that additional processes beyond changes in traction—such as junctional organization or cell–cell adhesion dynamics—are required. Accordingly, mechanical weakening is an important contributing factor, but not the sole driver, of aggregation, and targeted analyses of junctional remodeling and adhesion signaling will be needed to further delineate detailed downstream mechanisms.

A reduction in cellular contractility, reflected by decreased traction force, could in turn weaken cell–substrate adhesions and thereby contribute to the aggregation phenotype. This interpretation is supported by the mRNA level of FAK (Figure S1) and is consistent with previous studies showing that reduced cell‐substrate anchorage predisposes cells to cluster or form aggregates [61, 62, 63]. For instance, when cultured on a non‐adhesive surface, cells lost their attachment to the ECM/substrate and typical morphology, forming various aggregation patterns [62]. Moreover, because contractility and adhesion are well‐established prerequisites for cell migration [64, 65], their reduction provides a mechanistic link between c‐Abl activation, and the impaired migratory capacity observed in KA + Tet cells.

In addition to these adhesion‐related mechanisms, ZO‐2 has also been reported to directly interact with junctional adhesion molecule‐A (JAM‐A or JAM‐1), which regulates adhesion and migration through dimerization by indirectly modulating β1 integrin levels [63]. We therefore speculate that c‐Abl–dependent phosphorylation of ZO‐2 may alter its association with JAM‐A, providing a complementary mechanism through which adhesion and migration are regulated [64, 65]. Previous studies have also reported functional impacts of ZO‐2 phosphorylation in other contexts. For example, Saito et al. showed that ZO‐2 can be phosphorylated by the tyrosine kinase c‐Src and interacts with its negative regulator, C‐terminal Src kinase (Csk) [66]. In addition, tyrosine phosphorylation of ZO‐1 and ZO‐2 by v‐Src has been shown to weaken junctional sealing [67]. In contrast, our findings indicate that c‐Abl activation and subsequent ZO‐2 phosphorylation did not alter monolayer stress, suggesting that phosphorylation by c‐Abl may have distinct consequences compared to Src‐family kinases. Clarifying these differences will require more detailed investigation to fully understand the broader impact of ZO‐2 phosphorylation.

In conclusion, our data establish ZO‐2 as a pivotal mediator of c‐Abl–induced alterations in cell morphology and migration. Direct and indirect phosphorylation of ZO‐2 by c‐Abl reduces traction forces and cellular contractility, leading to marked morphological changes such as aggregate formation and ultimately contributing to impaired migration (Figure 6A,B). This phosphorylation‐mediated regulation of ZO‐2 may have important implications in various physiological and pathological contexts involving cell–cell interactions, cell–matrix adhesion, cell migration, and epithelial organization.

FIGURE 6.

Schematic diagram of the changes within the cells by active c‐Abl. (A) Illustrates the morphological and architectural changes in cells due to active c‐Abl, and (B) depicts the signaling pathway activated by c‐Abl. The activation of c‐Abl leads to changes in cell contractility, cell‐matrix transferred force (traction), and cell‐matrix adhesion, and overall cell morphology and migration.

Author Contributions

D.E.C.: conceptualization, methodology, validation, formal analysis, investigation, data curation, writing – original draft, writing – review and editing, project administration; B.G.: conceptualization, methodology, investigation, writing – original draft, writing – review and editing, funding acquisition; J.N.: software, validation, data curation, writing – review and editing; H.K.: investigation, writing – original draft; X.‐S.L.: investigation, methodology; J.J.F.: resources, writing – review and editing.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Data S1: fsb271214‐sup‐0001‐DatasetS1.pdf.

Video S1: fsb271214‐sup‐0002‐VideoS1.avi.

Video S2: fsb271214‐sup‐0003‐VideoS2.avi.

Video S3: fsb271214‐sup‐0004‐VideoS3.avi.

Video S4: fsb271214‐sup‐0005‐VideoS4.avi.

Video S5: fsb271214‐sup‐0006‐VideoS5.avi.

Video S6: fsb271214‐sup‐0007‐VideoS6.avi.

Choi D. E., Gweon B., Notbohm J., Kim H., Liu X.‐S., and Fredberg J. J., “ c‐Abl Kinase Targets Tight Junction Protein ZO‐2 in Regulation of Cell Migration and Morphology,” The FASEB Journal 39, no. 22 (2025): e71214, 10.1096/fj.202402159R.

Data Availability Statement

The data that support the findings of this study are available in the Materials and Methods, Results, and/or [Link], [Link] of this article.