A Toxoplasma gondii-derived ROP38 protein engages Ser15/Thr18-linked p53 activation and DNA damage-associated apoptotic signaling in human cancer cells

Abstract

Background

The tumor suppressor p53 integrates diverse cellular stresses to regulate DNA damage responses, cell-cycle progression, and apoptosis-associated signaling. Toxoplasma gondii secretes rhoptry effectors that remodel host signaling networks, but the host targets and signaling functions of the rhoptry kinase ROP38 remain poorly defined.

Methods

GFP-tagged T. gondii ME49 ROP38 full-length protein (ROP38 Full) and its kinase-domain fragment (ROP38 KiD) were stably expressed in HCT116 and HepG2 human cancer cells. Kinase-associated activity was assessed in anti-GFP immunoprecipitates, and a candidate co-immunoprecipitation screen was used to identify ROP38-associated host proteins. ROP38–p53 association and phospho-p53 species were further examined by co-immunoprecipitation. Purified recombinant ROP38 and human p53 were used for direct interaction and in vitro phosphorylation assays. Whole-cell p53 signaling, DNA damage/apoptosis-associated markers, and cell-cycle distribution were also analyzed. Statistical analyses used one- or two-way ANOVA, as appropriate, followed by Tukey’s multiple comparisons test.

Results

ROP38 Full- and ROP38 KiD-containing immunoprecipitates showed increased kinase-associated activity. Among the candidate proteins examined, p53 was the only ROP38-associated host factor detected in both cell lines. ROP38 Full and ROP38 KiD co-immunoprecipitated with total p53 and preferentially enriched p53 phosphorylated at Ser15 and Thr18. Purified recombinant ROP38 directly associated with recombinant p53 and promoted Ser15/Thr18 phosphorylation in the presence of ATP. At the whole-cell level, ROP38 expression increased p53 abundance and multisite phosphorylation, with prominent Ser15/Thr18 induction, and was accompanied by increased γH2AX, PUMA, cleaved caspase-3, and p21 expression. ROP38-expressing cells also showed decreased G₁-phase populations with increased S- and G₂/M-phase populations.

Conclusions

These data identify a candidate ROP38-associated p53 signaling module in human cancer cells and provide biochemical evidence that ROP38 can directly associate with p53 and promote Ser15/Thr18 phosphorylation in vitro. The requirement for intrinsic ROP38 catalytic activity and the physiological relevance of this relationship during T. gondii infection remain to be established.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s12964-026-03257-5.

Background

The tumor suppressor p53 functions as a central signaling node that integrates diverse cell stresses, including DNA damage, oncogenic signaling, and metabolic perturbations, to orchestrate cell-cycle arrest, DNA damage-based responses, and apoptosis-associated signaling [1–3]. In cancer cells, p53 activity is tightly regulated at multiple levels, particularly through protein stability control and phosphorylation-dependent activation [4, 5]. Phosphorylation events within the N-terminal transactivation domain (TAD), including Ser15 and Thr18, are widely associated with early activation steps that facilitate p53 maintenance and downstream transcriptional responses [5–7].

Intracellular pathogens commonly secrete effectors to remodel host signaling pathways and cell-states in ways that favor pathogen survival, persistence, and host adaptation [8]. These can target phosphorylation-dependent networks, reshaping stress responses, survival programs, immune signaling, and cell-fate decisions [9, 10]. Elucidating the host factors engaged by such effectors provides a powerful route to uncover concealed connections between infection-associated molecules and core host-regulatory circuits [8, 11].

Toxoplasma gondii is an obligate intracellular parasite that can establish persistent infection and modulate diverse host-cell responses [12–14]. Several of its secreted rhoptry proteins (ROPs) harbor kinase-like domains and are implicated in modulating host signaling [15]. ROP16 and ROP18 are among the most-characterized rhoptry kinases: ROP16 phosphorylates host STAT3 and STAT6 to modulate immune signaling [16], whereas ROP18 functions as a serine-threonine kinase that phosphorylates host immunity-related GTPases (IRGs) and contributes to parasite survival [17, 18]. These effectors illustrate that T. gondii rhoptry kinases can engage specific host signaling nodes through kinase-dependent mechanisms. In contrast, ROP38 is an annotated rhoptry kinase that has been implicated in the modulation of host transcriptional responses and MAPK-associated genes; however, its defined host protein partners and downstream mechanistic signaling pathways remain incompletely characterized [19, 20].

Previous studies have established that T. gondii infection can modulate host cell-cycle fate, DNA damage responses, and genome integrity. Infection has been reported to induce γH2AX-associated DNA double-strand breaks and ATM/CHK2 pathway activation [21]. Recent studies further showed that T. gondii-induced host DNA damage can be strain-dependent and associated with chromosomal instability and activation of the ATM-dependent homologous recombination pathway [22]. In addition, early infection has been linked to host cell-cycle arrest, DNA damage foci, binucleation, and micronuclei formation, supporting a connection between T. gondii infection and host genome instability [23]. ROP16 has also been shown to interact with p53, promote p53 phosphorylation at Ser15 and Ser37, and induce p53-dependent apoptosis and G₁ cell-cycle arrest [24]. Together, these studies indicate that p53-, γH2AX-, and genome-integrity-associated host responses have previously been linked to T. gondii infection and effector biology. Therefore, the present study focuses specifically on whether the poorly characterized rhoptry kinase ROP38 can engage p53-associated signaling under isolated expression conditions in human cancer cells.

T. gondii effectors influence host stress- and cancer-associated signaling outputs when expressed in mammalian cells, including pathways linked to AKT/MAPK, NF-κB, PTEN, and tumor suppressor-associated programs [25–29]. Such observations raise the possibility that lesser-characterized rhoptry kinases may engage host stress-signaling networks beyond canonical immune-regulatory pathways. However, whether ROP38 interacts with specific signaling-related host proteins and how such associations are linked to downstream cell stress responses remains unclear.

Human cancer cell models provide a useful experimental context for interrogating effector-driven host signaling because stress-response pathways, including p53-dependent transcriptional and apoptosis-associated signaling programs, can be assessed in defined genetic backgrounds [25–29]. In particular, p53-proficient cancer cell lines allow the assessment of whether a parasite-derived effector can engage tumor suppressor-associated signaling outputs, such as p53 accumulation, phosphorylation, DNA damage-associated markers, and downstream transcription- or apoptosis-related readouts [26, 28, 29]. Thus, the stable expression of poorly characterized T. gondii effectors in human cancer cells can serve as a tractable discovery platform for identifying parasite–host signaling interfaces. However, such tumor-derived models should not be interpreted as models of normal host-cell DNA damage responses, primary-cell infection responses, tissue-level responses, or in vivo T. gondii infection. Rather, they provide a tractable cancer-cell signaling context in which ROP38-associated host signaling outputs can be examined under isolated expression conditions.

From a signaling perspective, activation-linked p53 phosphorylation provides informative data for stress-associated regulatory events. Ser15 phosphorylation stimulates p53-dependent transactivation, which can subsequently facilitate Thr18 phosphorylation [6, 7, 30]. Therefore, analyzing p53 phospho-state regulation, particularly at Ser15 and Thr18, can provide mechanistic insights into whether an upstream signaling input engages an early p53 activation signature.

Given the central role of phosphorylation networks in determining stress and survival outcomes, we hypothesized that ROP38 may engage host signaling nodes and promote measurable stress-associated signaling responses in human cells. This study was not designed to recapitulate the physiological delivery, spatiotemporal regulation, or multi-effector context of natural T. gondii infection. To test this hypothesis, we established stable human cancer cell models expressing T. gondii ME49 ROP38 full-length protein (ROP38 Full) or its kinase-domain fragment (ROP38 KiD), assessed kinase-associated activity within immunopurified ROP38-containing complexes, and performed a hypothesis-driven candidate co-immunoprecipitation (Co-IP) screen to identify host factors associated with ROP38. We then examined ROP38–p53 association and p53 phosphorylation in cancer cells and used purified recombinant proteins to assess direct ROP38–p53 association and Ser15/Thr18 phosphorylation in vitro. Downstream cellular responses were further evaluated by examining p53 abundance and phosphorylation, γH2AX, PUMA, cleaved caspase-3, p21 expression, and cell-cycle distribution. Here, we identify a candidate ROP38-associated p53 signaling module in p53-proficient human cancer cells and provide biochemical evidence linking ROP38 to p53-associated signaling under isolated expression conditions.

Methods

Reagents and antibodies

Pierce™ Classic Magnetic IP/Co-IP Kit (Thermo Fisher Scientific, Waltham, MA, USA; 88804) was used for immunoprecipitation. Kinase-associated activity in immunoprecipitated protein complexes was measured using a Universal Kinase Assay Kit (Fluorometric) (Abcam, Cambridge, UK; ab138879). Total RNA was isolated with a HiGene™ Total RNA Prep Kit (BIOFACT, Daejeon, Republic of Korea; RP101-050), and cDNA was synthesized employing a Reverse-Transcription Premix (Elpis Biotech, Daejeon, Republic of Korea; EBT-1515). Proteins were quantified using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific; 23225). Western blot signals were developed with a Pierce™ ECL Western Blotting Substrate (Thermo Fisher Scientific; 32106). Restriction enzymes and ligases utilized included PspXI (NEB, Ipswich, MA, USA; R0656S), BamHI (Enzynomics, Daejeon, Republic of Korea; R003S), and T4 DNA ligase (Enzynomics; M001S). Polybrene (Santa Cruz Biotechnology, Dallas, TX, USA; sc-134220) was used for lentiviral transduction. Protein A/G PLUS-Agarose beads (Santa Cruz Biotechnology; sc-2003) were used for recombinant protein-based interaction assays. Kinase Buffer (Cell Signaling Technology, Danvers, MA, USA; 9802) and ATP (Cell Signaling Technology; 9804) were used for in vitro kinase assays. Propidium iodide and RNase A (Sigma-Aldrich, St. Louis, MO, USA) were used for cell-cycle analysis. All primary and secondary antibodies utilized for immunoprecipitation, immunoblotting, and immunofluorescence staining; phospho-specific detection, including antibodies targeting GFP, total p53, phospho-p53, ERK–MAPK, and PI3K–AKT–mTOR pathway candidate proteins; and loading controls are detailed in Additional file 1: Table S1.

Plasmid construction

The coding sequences encoding a T. gondii ME49 ROP38 full-length protein (1,626 nt; 541 aa) and its kinase domain (927 nt; 309 aa) were amplified from cDNA synthesized using total RNA isolated from ME49 tachyzoites (50611, ATCC, Manassas, VA, USA) propagated in HFF-1 cells. Amplicons and the pWPXL backbone (Addgene, Watertown, MA, USA; 12257) were digested with PspXI (New England Biolabs, Ipswich, MA, USA; R0656S) and BamHI (Enzynomics; R003S) and fused using T4 DNA ligase (Enzynomics; M001S). Competent E. coli DH5α (Enzynomics; CP011) were transformed with the ligations, and positive clones were identified via screening and plasmid purification. Plasmid construction and sequence validation were performed at Cosmogenetech Co., Ltd. (Seoul, Republic of Korea). Final constructs—pWPXL–ROP38 Full and pWPXL–ROP38 KiD were verified by Sanger sequencing and used in subsequent experiments. The ROP38 Full construct contained the complete 541 aa coding sequence, including the endogenous N-terminal signal peptide (1–29 aa), whereas a distinct pro-domain has not been annotated or experimentally defined for ROP38. Because the protein was expressed directly in mammalian cells, its native parasite-specific rhoptry trafficking and invasion-coupled secretion were not expected to be reproduced and were not examined in this study. Here, ROP38 KiD denotes the kinase-domain fragment and does not indicate a kinase-dead mutant or knockdown construct.

Cell culture

HCT116 cells (KCLB, Seoul, Republic of Korea; 10247) were cultured in RPMI1640 medium (Welgene, Gyeongsan, Republic of Korea; LM011-01). HepG2 (KCLB; 88065), Lenti-X 293T (Takara Bio, Shiga, Japan; 632180), and HFF-1 (ATCC; SCRC-1041) cells were cultured in DMEM (Welgene; LM001-05). Culture media were supplemented with 10% fetal bovine serum (Thermo Fisher Scientific; A5256701) and 1% antibiotic–antimycotic (Cellpick, Seoul, Republic of Korea; CAA001); cells were maintained at 37℃ in a humidified incubator with 5% CO₂ unless otherwise indicated. T. gondii ME49 tachyzoites (ATCC 50611) were propagated in HFF-1 cells as a source for RNA used for cloning.

Lentivirus production and stable cell line generation

Recombinant lentiviruses were produced in Lenti-X 293T cells (Takara Bio; 632180) by co-transfection of pWPXL-based expression plasmids (Addgene; 12257) with the packaging psPAX2 (Addgene; 12260) and envelope pMD2.G (Addgene; 12259) plasmids using standard protocols. Viral supernatants were collected 48 h post-transfection, clarified by centrifugation, and used to transduce HCT116 and HepG2 cells with polybrene (Santa Cruz Biotechnology; sc-134220) at 8 µg/mL. GFP-positive cells were enriched by fluorescence-activated cell sorting (FACS; SH-800 S; Sony, Tokyo, Japan) to generate stable populations expressing GFP-only vector, GFP-ROP38 Full, or GFP-ROP38 KiD. Parental cells without lentiviral transduction served as controls.

RT-PCR verification of ROP38 transgene expression

To verify the expression of ROP38 Full and ROP38 KiD, reverse transcription PCR (RT-PCR) was performed using cDNA synthesized from the total RNA from each stable cancer cell population with the HiGene™ Total RNA Prep Kit (BIOFACT, Daejeon, Republic of Korea; RP101-050), and a RT premix containing oligo d(T) primers (Elpis Biotech, Daejeon, Republic of Korea; EBT-1515). The cDNA obtained served as a PCR template, and primer sets specific for EF-1α, ROP38 Full, and ROP38 KiD were employed; primer sequences are provided in Additional file 2: Table S2. PCR was performed on a SureCycler 8800 Thermal Cycler (Agilent Technologies, Santa Clara, CA, USA). The program employed was- initial denaturation at 95℃ for 2 min; 30 cycles of 95℃ for 20 s, 58℃ for 40 s, and 72℃ for 90 s; and a final extension at 72℃ for 5 min. Amplicons were visualized after 1% agarose gel electrophoresis on the Gel Doc XR + Gel Documentation System (Bio-Rad Laboratories, Hercules, CA, USA). The product sizes were 758 bp (EF-1α), 1,626 bp (ROP38 Full), and 927 bp (ROP38 KiD).

GFP fluorescence microscopy

To verify GFP expression at the cell level, HCT116 and HepG2 cells expressing GFP-tagged ROP38 proteins were imaged with a DMI6000B inverted fluorescence microscope (Leica, Wetzlar, Germany) in brightfield and GFP channels. Representative images were acquired under identical exposure conditions. Images were acquired and processed uniformly across experimental groups.

Immunoprecipitation for IP–Western blotting

Immunoprecipitation with the Pierce™ Classic Magnetic IP/Co-IP Kit (Thermo Fisher Scientific; 88804) was performed to selectively recover GFP-tagged ROP38-containing protein complexes. After cell culture, the media were removed, cells were washed once with PBS, and broken in ice-cold IP Lysis/Wash Buffer for 5 min on ice. Lysates were clarified by centrifugation at 13,000× g for 10 min, and protein concentrations were determined using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific; 23225).

For antigen–antibody complex formation, 1.0 mg of total protein was mixed with 5 µg of a GFP-specific antibody and incubated at 4℃ overnight (16 h). Then, 25 µL of Pierce Protein A/G Magnetic Beads (Thermo Fisher Scientific; 88802) were added and incubated for 1 h at room temperature with mixing. Beads were washed twice with IP Lysis/Wash Buffer, and immune complexes were eluted using 100 µL of Elution Buffer at 96℃ for 10 min. Eluates were collected for Western blotting to assess protein complex formation. Antibodies used for IP and Western blotting are listed in Additional file 1: Table S1.

Recombinant ROP38 Full and human p53 proteins

Recombinant ROP38 Full protein was produced by BIONICS Co., Ltd. (Seoul, Republic of Korea). The recombinant construct encoded an N-terminal BR2 cell-penetrating peptide, a caspase-3-cleavable linker, the complete T. gondii ME49 ROP38 Full sequence, and a C-terminal 6×His tag and was expressed from the pET21a vector in Escherichia coli BL21(DE3) cells. The recombinant protein was recovered from the insoluble fraction, refolded, and purified by Ni-NTA affinity chromatography. The calculated molecular weight was 64 kDa, and the final purity was greater than 90%, as assessed by SDS-PAGE and Coomassie blue staining. The purified protein was supplied in buffer containing 3 M urea, 20 mM Tris-HCl (pH 8.0), 150 mM NaCl, and 10% glycerol. Recombinant human p53 protein was purchased from Active Motif (Carlsbad, CA, USA; 81091).

Recombinant protein-based direct interaction assay

To examine the direct association between recombinant ROP38 Full and human p53 in the absence of cellular lysates, a recombinant protein-based co-immunoprecipitation assay was performed using Protein A/G PLUS-Agarose beads (Santa Cruz Biotechnology; sc-2003). The binding buffer contained 25 mM HEPES (pH 7.5), 150 mM NaCl, 0.05% NP-40, and 5% glycerol, whereas the washing buffer contained 25 mM HEPES (pH 7.5), 250 mM NaCl, 0.1% NP-40, and 5% glycerol.

Protein A/G PLUS-Agarose beads (20 µL) were washed with binding buffer and incubated with 5 µL of an anti-His tag antibody (Cell Signaling Technology; 2365) for 1 h at 4 °C with rotation. The antibody-coupled beads were blocked for 30 min at 4 °C using binding buffer supplemented with 0.5% BSA. Recombinant human p53 (0.6 µg) was separately pre-cleared using Protein A/G PLUS-Agarose beads and anti-His tag antibody for 1 h at 4 °C, and the resulting supernatant was used in subsequent binding reactions. The antibody-coupled beads were incubated with 2 µg of recombinant His-tagged ROP38 Full or recombinant GFP protein (Thermo Fisher Scientific; A42613) for 1 h at 4 °C. Recombinant GFP was used as an unrelated recombinant protein control. After washing, pre-cleared recombinant p53 was added to the indicated groups, and the samples were incubated for 2 h at 4 °C with rotation. Control reactions lacking the indicated recombinant protein or antibody components were processed in parallel. The beads were washed five times with washing buffer, and bound proteins were eluted in 1× sample buffer by heating at 95 °C for 5 min. After centrifugation, the supernatants were subjected to SDS-PAGE and Western blotting. Recombinant p53 was detected using an anti-p53 antibody (Santa Cruz Biotechnology; sc-126) and an HRP-conjugated mouse IgG κ-binding protein (Santa Cruz Biotechnology; sc-516102). Signals were developed using Pierce™ ECL Western Blotting Substrate (Thermo Fisher Scientific; 32106).

In vitro kinase assay using recombinant ROP38 and p53

To examine whether recombinant ROP38 Full promotes phosphorylation of recombinant human p53, an in vitro kinase assay was performed using recombinant ROP38 Full and recombinant human p53. Each reaction contained 0.5 µg of recombinant p53 and either 0.5–1 µg of recombinant ROP38 Full in a final volume of 15 µL. Reactions were prepared in 1× Kinase Buffer (Cell Signaling Technology; 9802) in the presence or absence of 200 µM ATP (Cell Signaling Technology; 9804). The reaction groups consisted of recombinant p53 with ATP but without ROP38 Full; recombinant p53 with 1 µg of ROP38 Full but without ATP; recombinant p53 with 0.5 µg of ROP38 Full and ATP; recombinant p53 with 1 µg of ROP38 Full and ATP; and recombinant p53 with 1 µg of heat-inactivated ROP38 Full and ATP. For the heat-inactivated control, recombinant ROP38 Full was heated at 95 °C for 5 min and immediately cooled on ice before use.

Kinase reactions were initiated by adding ATP to the designated groups, whereas an equivalent volume of nuclease-free water was added to the ATP-free control, followed by incubation at 30 °C for 30 min. Reactions were terminated by adding 4 µL of 5× SDS sample buffer, followed by heating at 95 °C for 5 min. Samples were subjected to SDS-PAGE and Western blotting. Phosphorylation of recombinant p53 was assessed using antibodies specific for p53 phosphorylated at Ser15 and Thr18. Total p53 and ROP38 Full were detected to verify the presence of the substrate and recombinant ROP38 protein, respectively.

Immunofluorescence staining

To examine p53 abundance and phospho-p53 signals at the cell level, stable HCT116 and HepG2 populations were stained for immunofluorescence. Cells were fixed with 4% paraformaldehyde for 10 min at 25℃ and washed with PBS (pH 7.5). The cells were then permeabilized with PBS containing 0.2% Triton X-100 for 15 min at room temperature and blocked with PBS containing 0.1% Tween-20 (PBS-T) supplemented with 1% bovine serum albumin (BSA) and 22.52 mg/mL glycine for 1 h at room temperature. After washing with PBS-T, cells were incubated with primary antibodies against total p53 or phospho-p53 species, including pSer15, pThr18, pSer20, pSer33, pSer46, and pThr81, for 16 h at 4 °C. After washing with PBS-T, cells were incubated with Alexa Fluor 594-conjugated goat anti-rabbit IgG (H + L) or donkey anti-mouse IgG (H + L) secondary antibodies for 1 h at room temperature in the dark. All primary and secondary antibodies used are listed in Additional file 1: Table S1. Nuclei were counterstained with 5 µg/mL Hoechst 33,342 (Sigma-Aldrich, St. Louis, MO, USA) diluted in PBS for 5 min at room temperature in the dark. After a final wash with PBS-T, bright-field and fluorescence images were acquired with a DMI6000B inverted fluorescence microscope (Leica, Wetzlar, Germany). Representative images were acquired under identical exposure settings across experimental groups. Fluorescence intensity was quantified using ImageJ software (National Institutes of Health, Bethesda, MD, USA), and mean fluorescence intensity values were used for quantitative comparison among experimental groups.

Western blotting (whole-cell lysates)

Whole-cell lysates were prepared using M-PER™ Mammalian Protein Extraction Reagent (Thermo Fisher Scientific; 78501), and proteins were quantified with the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific; 23225). For each condition, 30 µg of total protein was separated by 12% SDS-PAGE and transferred to PVDF membranes utilizing a Mini Trans-Blot Electrophoretic Transfer Cell (Bio-Rad Laboratories, Hercules, CA, USA) at 100 V for 1 h at 4℃. Membranes were blocked in Tris-buffered saline (pH 7.6, containing 0.1% Tween-20) supplemented with 5% BSA for 1 h at room temperature, followed by incubation with primary antibodies at 4℃ for 16 h. After washing, membranes were incubated with HRP-conjugated antimouse or antirabbit secondary antibodies for 1 h at room temperature. Protein signals were detected employing Pierce™ ECL Western Blotting Substrate (Thermo Fisher Scientific; 32106). Densitometric analysis of immunoblot bands was performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Target protein signals were normalized to β-actin, and relative expression levels were calculated against the Control group. The primary and secondary antibodies are detailed in Additional file 1: Table S1.

Real-time qPCR

Total RNA was isolated from each experimental group with the HiGene™ Total RNA Prep Kit (BIOFACT; RP101-050) and reverse-transcribed into cDNA using a Reverse-Transcription Premix (Elpis Biotech; EBT-1515). Real-time qPCR was performed on a CFX96 Touch Real-Time PCR Detection System (Bio-Rad Laboratories, Hercules, CA, USA) utilizing TOPreal™ qPCR 2X PreMIX (SYBR Green with low ROX) (Enzynomics; RT500S). PCR program was- initial denaturation at 95℃ for 15 min, followed by 40 cycles of 95℃ for 20 s, 59℃ for 20 s, and 72℃ for 30 s. Fluorescence signals were collected and analyzed using iQ™5 Optical System Software (Bio-Rad Laboratories, Hercules, CA, USA). Relative expression levels were normalized to GAPDH C_t values and compared to the control group. Sequences for p21 and GAPDH-specific primers obtained from OriGene™ Technologies (Rockville, MD, USA) are provided in Additional file 2: Table S2.

Cell-cycle analysis by flow cytometry

Cell-cycle distribution was analyzed in stable Control, Vector, ROP38 Full, and ROP38 KiD HCT116 and HepG2 cell populations. Cells were harvested and fixed in ice-cold 70% ethanol for 1 h at − 20 °C. After fixation, the cells were washed with PBS and stained in the dark at room temperature for 20 min with 400 µL of propidium iodide solution containing 50 µg/mL propidium iodide, 3.8 mM sodium citrate, and 100 µg/mL RNase A (Sigma-Aldrich, St. Louis, MO, USA). DNA content was measured using a BD FACSCanto™ II flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA). At least 1 × 10⁴ cells were acquired per sample, and the proportions of cells in the G₁, S, and G₂/M phases were analyzed using FlowJo software version 7.6 (Tree Star, Ashland, OR, USA).

Immunoprecipitation-based kinase activity assay

To evaluate kinase-associated activity in ROP38 Full- or ROP38 KiD-containing complexes expressed in human cancer cells, an immunoprecipitation-coupled kinase activity (IP-based kinase activity) was assayed. Proteins were extracted from stable Control, Vector, ROP38 Full, and ROP38 KiD cell populations, and immunoprecipitated using the Pierce™ Classic Magnetic IP/Co-IP Kit (Thermo Fisher Scientific; 88804). A GFP-specific antibody was utilized to selectively recover GFP-tagged ROP38 Full or ROP38 KiD complexes. Immunoprecipitated protein samples were quantified, and 1 µg of them was employed for downstream kinase activity measurements.

Kinase-associated activity was assayed based on ADP generation and using the Universal Kinase Assay Kit (Fluorometric) (Abcam; ab138879) following the manufacturer’s instructions. Briefly, immunoprecipitated samples were transferred to black microplates and combined with the kit components provided to initiate kinase reactions. After incubation at room temperature, ADP-dependent fluorescence was measured with a Spark fluorescence plate reader (Tecan, Männedorf, Switzerland) under the recommended settings and presented as relative fluorescence units (RFU, a.u.). As this assay used anti-GFP immunoprecipitates, the ADP-dependent signal measured represents kinase-associated activity within GFP-ROP38-containing complexes rather than the intrinsic catalytic activity of purified recombinant ROP38 alone.

Statistical analysis

HCT116 and HepG2 cells were separately analyzed. Quantitative data are presented as the mean ± SD. Statistical analyses were performed in GraphPad Prism 8 (https://www.graphpad.com/), and p < 0.05 was considered statistically significant. For comparisons of multiple groups, including immunofluorescence intensity, densitometric protein quantification, p21 mRNA expression, and kinase-associated activity, ordinary one-way analysis of variance (ANOVA) with Tukey’s multiple comparisons test was used. Cell-cycle distribution was analyzed using two-way ANOVA followed by Tukey’s multiple comparisons test. Statistical significance was indicated using the symbols- *indicates ROP38 Full vs. Control; †indicates ROP38 Full vs. Vector; §indicates ROP38 KiD vs. Control; and ¶indicates ROP38 KiD vs. Vector.

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Results

Construction and validation of GFP-tagged ROP38 Full- and kinase-domain-expressing human cancer cell populations

To enable functional interrogation of the secreted T. gondii rhoptry kinase ROP38 in mammalian cells, we generated GFP-tagged expression constructs encoding ME49 ROP38 full-length (ROP38 Full; 1,626 nt/541 aa) and its kinase-domain fragment (ROP38 KiD; 927 nt/309 aa) (Fig. 1A, B). Based on sequence annotation, the kinase domain within the full-length ROP38 sequence was defined and selected for domain-specific expression (Fig. 1A). The corresponding inserts were cloned into the multiple cloning site of the pWPXL backbone to produce the pWPXL–ROP38 Full and pWPXL–ROP38 KiD (Fig. 1B). Final plasmids were sequence-verified to match the ME49 ROP38 reference, providing validated tools for subsequent expression and signaling analyses in human cancer cell lines.

Fig. 1.

Generation and validation of stable human cancer cell populations expressing GFP-tagged ROP38 Full or KiD. A DNA and protein sequence annotation of T. gondii ME49 ROP38 showing the location of the annotated kinase domain within the full-length sequence. ROP38 Full comprises 1,626 nt/541 aa, and the kinase-domain fragment (ROP38 KiD) comprises 927 nt/309 aa. B Schematic representation of the GFP-tagged lentiviral expression constructs. ROP38 Full or ROP38 KiD was cloned into the multiple cloning site of the pWPXL backbone to generate pWPXL–ROP38 Full and pWPXL–ROP38 KiD. C, D Flow cytometry histograms showing enrichment of GFP-positive populations following lentiviral transduction in (C) HCT116 and (D) HepG2 cells. Black traces indicate GFP-negative control cells, and red traces indicate GFP fluorescence distributions in ROP38 Full- or ROP38 KiD-transduced cells. The dashed line indicates the fluorescence threshold used for GFP-positive cell enrichment. The experiments were independently performed three times (n = 3), and representative histograms are shown. E, F RT-PCR validation of transgene expression in (E) HCT116 and (F) HepG2 stable cell populations. Primer sets specific for EF-1α, ROP38 Full, or ROP38 KiD were used. Expected amplicon sizes are indicated: EF-1α, 758 bp; ROP38 Full, 1,626 bp; and ROP38 KiD, 927 bp. G, H Representative bright-field and GFP fluorescence images of (G) HCT116 and (H) HepG2 stable cell populations. GFP fluorescence was detected in ROP38 Full- and ROP38 KiD-expressing cells, whereas control cells showed no detectable GFP signal. The experiments were independently performed three times (n = 3), and representative images are shown. M, DNA marker. Scale bars, 75 μm

To generate uniform populations for downstream analyses, HCT116 and HepG2 cells were transduced with lentiviruses encoding GFP-tagged ROP38 Full or ROP38 KiD and enriched by FACS based on GFP intensity. Compared with GFP-negative control cells (black peaks), ROP38 Full- and ROP38 KiD-transduced populations exhibited a clear rightward shift of the FITC-A signal (red peaks), and GFP-high cells were collected utilizing the indicated sorting gate (dashed line) to obtain enriched GFP-positive populations (Fig. 1C, D). ROP38 transgene expression was confirmed by RT-PCR using cDNA synthesized from total RNA isolated from each stable cell population. PCR-based amplification using primer sets specific for EF-1α, ROP38 Full, or ROP38 KiD yielded products of the expected sizes only in the relevant cell populations (Fig. 1E, F). Consistent with the stable expression of GFP-tagged proteins, fluorescence microscopy demonstrated robust GFP signals in ROP38 Full- and ROP38 KiD-expressing HCT116 and HepG2 cells, whereas controls lacked detectable GFP fluorescence (Fig. 1G, H). Together, these results validate the establishment of stable GFP-positive cell populations expressing ROP38 Full or ROP38 KiD for subsequent biochemical and signaling studies.

To determine whether ROP38-containing complexes expressed in human cancer cells exhibit kinase-associated activity, we performed an IP-based assay using anti-GFP immunoprecipitates from HCT116 and HepG2 cells. Compared with the Control and Vector immunoprecipitates, ROP38 Full and ROP38 KiD samples showed significantly higher ADP-dependent fluorescence in the immunoprecipitation-based kinase activity assay (Additional file 3: Figure S1A, B).

A candidate co-IP screen identifies p53 as a ROP38-associated host factor

To explore potential host signaling nodes associated with ROP38, we performed a hypothesis-driven candidate co-IP screen in HCT116 and HepG2 cells. Candidate proteins were selected to represent ROP38-associated MAPK-related signaling reported previously, together with interconnected phosphatase, growth, and stress-regulatory pathways. We first probed anti-GFP immunoprecipitates for core ERK–MAPK pathway components and MAPK phosphatases—MEK, ERK, DUSP3/6/7/9, and PTPRR—but under these conditions, these factors were not detectably co-enriched with ROP38 compared with controls (Fig. 2A, F). We next examined PP2A-related regulators—PP2A, PP2Cα, PP2A-B55, and PP2A-B56—which were not detectably co-precipitated with ROP38 (Fig. 2B, G). Components of the parallel PI3K–AKT–mTOR signaling axis—PI3K, PDK1, AKT, mTOR, and PTEN—were also not detectably co-immunoprecipitated (Fig. 2C, H), and additional regulators linked to MAPK/stress signaling—USP7, PEA15, and BIK—were not detectably co-enriched (Fig. 2D, I). In contrast, among the p53/GSK3β regulatory candidates, p53 was detected in ROP38 immunoprecipitates from both HCT116 and HepG2 cells, whereas GSK3β was not detectably co-enriched (Fig. 2E, J). Collectively, among the candidate proteins examined, p53 was the only ROP38-associated host factor detected in both cell lines, providing a rationale for subsequent p53-focused interaction and phosphorylation analyses.

Fig. 2.

Candidate co-immunoprecipitation screen identifies p53 as a ROP38-associated host factor. Anti-GFP immunoprecipitation (IP: GFP) was performed using lysates from HCT116 and HepG2 cells expressing GFP-only Vector or GFP-tagged ROP38 Full. Inputs and immunoprecipitates were analyzed by immunoblotting for the indicated candidate proteins. A–E Candidate co-IP screen in HCT116 cells. F–J Candidate co-IP screen in HepG2 cells. A, F ERK–MAPK/DUSP signaling candidates (MEK, ERK, DUSP3, DUSP6, DUSP7, DUSP9, and PTPRR). B, G PP2A-related signaling candidates (PP2A, PP2Cα, PP2A-B55, and PP2A-B56). C, H PI3K–AKT–mTOR signaling candidates (PI3K, PDK1, AKT, mTOR, and PTEN). D, I MAPK/stress-associated candidates (USP7, PEA15, and BIK). E, J p53/GSK3β regulatory candidates. Red boxes and arrows indicate p53 detected in GFP-ROP38 Full immunoprecipitates from both HCT116 and HepG2 cells. Among the candidate proteins examined, p53 was the only protein detected in GFP-ROP38 Full immunoprecipitates from both cell lines. The panels shown represent one exploratory candidate co-immunoprecipitation screening experiment. M, protein marker

ROP38 Full and KiD co-immunoprecipitate with p53 and preferentially enrich p53 pSer15/pThr18 species

Building on the candidate co-IP screen (Fig. 2), we next validated the ROP38–p53 association and tested whether such an interaction is retained by the ROP38 KiD and whether specific p53 phospho-species are preferentially enriched in the ROP38 complexes. Anti-GFP immunoprecipitation of HCT116 and HepG2 cells, including Control, Vector, ROP38 Full, and ROP38 KiD, consistently recovered total p53 in the latter two immunoprecipitates (Fig. 3A, B), suggesting that the kinase-domain fragment is sufficient to maintain detectable p53 association under such conditions. Notably, p53 phosphorylated at Ser15 and Thr18 was prominently detected in ROP38 immunoprecipitates, whereas other phospho-sites tested (Ser20, Ser33, Ser46, and Thr81) were not detectably enriched under these conditions (Fig. 3A, B). This preferential enrichment pattern suggested that ROP38-containing complexes engage a p53 activation-linked phospho-state, prompting us to further examine whether ROP38 can directly associate with p53 and promote Ser15/Thr18 phosphorylation using purified recombinant proteins.

Fig. 3.

ROP38 Full and KiD associate with p53 and preferentially enrich p53 phosphorylated at Ser15 and Thr18. A, B Anti-GFP immunoprecipitation (IP: GFP) was performed using lysates from (A) HCT116 and (B) HepG2 cells, including Control (parental, GFP-negative), Vector (GFP-only), ROP38 Full, and ROP38 KiD cell populations. Input lysates and immunoprecipitates were analyzed by immunoblotting for total p53 and p53 phosphorylated at Ser15, Thr18, Ser20, Ser33, Ser46, and Thr81. Total p53 was detected in both ROP38 Full and ROP38 KiD immunoprecipitates. Among the phospho-p53 species examined, p53 phosphorylated at Ser15 and Thr18 was prominently detected in ROP38 Full and ROP38 KiD immunoprecipitates, whereas p53 phosphorylated at Ser20, Ser33, Ser46, and Thr81 was not detected under the same conditions. β-Actin was examined in input lysates, and GFP-tagged constructs were detected in input and IP fractions to verify expression and immunoprecipitation. The experiments were independently performed three times (n = 3), and representative blots are shown. M, protein marker

Recombinant ROP38 directly associates with p53 and promotes Ser15/Thr18 phosphorylation in vitro

To determine whether the ROP38–p53 association observed in cancer cells could occur independently of other cellular proteins, we performed a recombinant protein-based co-immunoprecipitation assay using purified recombinant full-length ROP38 (recombinant ROP38) and recombinant human p53 (Additional file 4: Figure S2). Recombinant p53 co-precipitated with His-tagged recombinant ROP38, whereas no p53 signal was detected in the negative-control reactions, including the recombinant GFP control (Fig. 4A). These results support a direct association between ROP38 and p53 in the absence of cellular lysates. We next examined whether recombinant ROP38 could promote phosphorylation of recombinant p53 at Ser15 and Thr18 in vitro. Phosphorylation at both sites was detectable in reactions containing recombinant ROP38 and ATP and was more prominent with 1 µg than with 0.5 µg of recombinant ROP38. In contrast, phosphorylation at Ser15 and Thr18 was not detected in reactions without recombinant ROP38 or ATP or in the heat-inactivated recombinant ROP38 control (Fig. 4B). Together, these results support that recombinant ROP38 can directly associate with p53 and promote Ser15/Thr18 phosphorylation in a purified recombinant protein system.

Fig. 4.

Recombinant ROP38 directly associates with p53 and promotes Ser15/Thr18 phosphorylation in vitro. A Recombinant protein-based co-immunoprecipitation assay examining the association between recombinant His-tagged ROP38 Full and recombinant human p53 in the absence of cellular lysates. Protein A/G PLUS-Agarose beads coupled to an anti-His tag antibody were incubated with the indicated combinations of recombinant ROP38 Full, recombinant p53, or recombinant GFP, which served as an unrelated recombinant protein control. Bound p53 was detected by immunoblotting. Recombinant p53 was recovered in the presence of ROP38 Full but not in the corresponding negative-control reactions, supporting a direct association between recombinant ROP38 Full and p53. B In vitro kinase assay using recombinant ROP38 Full and recombinant human p53. Each reaction contained 0.5 µg of recombinant p53 and the indicated amount of recombinant ROP38 Full (0.5–1 µg) in the presence or absence of 200 µM ATP. Heat-inactivated ROP38 Full (1 µg; 95 °C for 5 min) was included as an additional control. Phosphorylation of p53 at Ser15 and Thr18 was assessed by immunoblotting, together with total p53 and ROP38 Full. Phosphorylation signals at Ser15 and Thr18 were detectable with 0.5 µg of recombinant ROP38 Full and were more prominent with 1 µg of ROP38 Full in the presence of ATP. The experiments in (A) and (B) were each independently performed three times (n = 3), and representative blots are shown. M, protein marker

ROP38 enhances p53 abundance and multisite p53 phosphorylation with prominent Ser15/Thr18 induction

To determine whether the preferential enrichment of p53 pSer15/pThr18 species in ROP38 complexes (Fig. 3) is accompanied by broader whole-cell changes, we examined p53 abundance and multisite p53 phosphorylation in HCT116 and HepG2 cells expressing ROP38 Full or ROP38 KiD. Immunofluorescence exhibited increased total p53 and phospho-p53 signals in ROP38 Full- and ROP38 KiD-expressing cells compared with Control and Vector-containing cells of both lines, with quantitative fluorescence intensity analysis supporting these increases (Fig. 5A, B). Signals for p53 phosphorylated at Ser15 and Thr18 were prominently elevated, and additional phospho-p53 signals, including Ser20, Ser33, Ser46, and Thr81, were also identified in ROP38-expressing cells. Consistent with the immunofluorescence results, immunoblotting of whole-cell lysates indicated that ROP38 Full and ROP38 KiD increased total p53 abundance and enhanced phosphorylation across multiple sites in HCT116 and HepG2 cells (Fig. 5C, D). Among the phospho-sites examined, Ser15 and Thr18 were prominently induced, consistent with phospho-state selectivity observed in ROP38 immunoprecipitates (Fig. 3). Overall, these data indicate that isolated ROP38 expression is associated with increased p53 abundance and multisite p53 phosphorylation, with particularly prominent Ser15/Thr18 induction.

Fig. 5.

ROP38 increases p53 abundance and multisite p53 phosphorylation with prominent Ser15/Thr18 induction. A, B Immunofluorescence analysis of total p53 and phosphorylated p53 species in (A) HCT116 and (B) HepG2 cells. Control, Vector, ROP38 Full, and ROP38 KiD cells were stained for total p53 or phospho-p53 species, including Ser15, Thr18, Ser20, Ser33, Ser46, and Thr81, together with Hoechst 33,342 nuclear counterstaining. Mean fluorescence intensities of the respective p53 signals were quantified and are presented in arbitrary units (a.u.). Scale bars, 75 μm. C, D Whole-cell lysates from (C) HCT116 and (D) HepG2 cells were analyzed for total p53 and phosphorylated p53 species at Ser15, Thr18, Ser20, Ser33, Ser46, and Thr81 by immunoblotting, with densitometric quantification shown. Fold changes are expressed relative to Control, set to 1. β-actin served as a loading control. Immunofluorescence staining and immunoblotting analyses were performed using three independent experiments (n = 3). Representative images and blots are shown. Quantitative data are presented as the mean ± SD. Statistical analyses, including ordinary one-way ANOVA with Tukey’s multiple comparisons test, were performed in GraphPad Prism 8; p < 0.05 was considered statistically significant. *indicates ROP38 Full vs. Control; †indicates ROP38 Full vs. Vector; §indicates ROP38 KiD vs. Control; ¶indicates ROP38 KiD vs. Vector

ROP38 expression is associated with DNA damage/apoptosis-related molecular outputs and altered cell-cycle distribution

Because ROP38 expression increased p53 abundance and phosphorylation, we next examined the downstream signaling outputs associated with DNA damage and apoptosis-related responses. In HCT116 cells, ROP38 Full and ROP38 KiD increased γH2AX and PUMA proteins and enhanced cleaved caspase-3 signals compared with the Control and Vector cells (Fig. 6A). RT-qPCR further showed increased p21 mRNA expression in ROP38 Full- and ROP38 KiD-expressing HCT116 cells (Fig. 6A). A similar pattern was observed in HepG2 cells, in which ROP38 Full and ROP38 KiD increased γH2AX and PUMA protein contents, elevated cleaved caspase-3 signals, and enhanced p21 mRNA levels compared with the Control and Vector cells (Fig. 6B). We next examined whether these molecular changes were accompanied by alterations in cell-cycle distribution. In both HCT116 and HepG2 cells, ROP38 Full and ROP38 KiD expression markedly decreased the proportion of cells in the G₁ phase and increased the proportions of cells in the S and G₂/M phases compared with the Control and Vector groups (Fig. 6C, D). These findings indicate a redistribution of the cell-cycle profile from G₁ toward S and G₂/M phases in ROP38-expressing cells. Together, these results indicate that isolated ROP38 expression is associated with increased p53 phosphorylation-linked DNA damage/apoptosis-related molecular markers and altered cell-cycle distribution in the two human cancer cell lines.

Fig. 6.

ROP38 expression is associated with DNA damage/apoptosis-related signaling and altered cell-cycle distribution. A, B Whole-cell lysates from (A) HCT116 and (B) HepG2 cells, including Control, Vector, ROP38 Full, and ROP38 KiD populations, were analyzed by immunoblotting for γH2AX, PUMA, and pro-/cleaved caspase-3. β-Actin served as a loading control. Densitometric quantification is shown as relative protein expression normalized to Control. Relative p21 mRNA expression was measured by RT-qPCR, normalized to GAPDH, and presented relative to the Control group. C, D Cell-cycle distribution of (C) HCT116 and (D) HepG2 cell populations was analyzed by propidium iodide staining and flow cytometry. Representative DNA-content histograms and quantitative distributions of cells in the G₁, S, and G₂/M phases are shown. ROP38 Full- and ROP38 KiD-expressing cells showed decreased G₁-phase populations accompanied by increased S- and G₂/M-phase populations compared with Control and Vector cells. Immunoblotting and cell-cycle analyses were each independently performed three times (n = 3). RT-qPCR analysis of p21 expression was performed in one experiment using five independent biological samples (n = 5). Representative blots and flow-cytometry histograms are shown, and quantitative data are presented as the mean ± SD. For immunoblotting and RT-qPCR data, statistical analyses were performed using ordinary one-way ANOVA followed by Tukey’s multiple comparisons test. Cell-cycle distribution was analyzed using two-way ANOVA followed by Tukey’s multiple comparisons test. p < 0.05 was considered statistically significant. *indicates ROP38 Full vs. Control; †indicates ROP38 Full vs. Vector; §indicates ROP38 KiD vs. Control; ¶indicates ROP38 KiD vs. Vector

Discussion

This study identifies a ROP38-associated p53 signaling module in human cancer cells. Under isolated ROP38 expression conditions, T. gondii ME49 ROP38 Full and its kinase-domain fragment associated with p53 and preferentially enriched p53 phosphorylated at Ser15 and Thr18 in ROP38-containing complexes. Purified recombinant ROP38 also directly associated with recombinant p53 and promoted Ser15/Thr18 phosphorylation in vitro. At the whole-cell level, ROP38 expression was accompanied by increased p53 abundance and multisite phosphorylation, elevated γH2AX, PUMA, cleaved caspase-3, and p21 expression, together with redistribution of the cell-cycle profile from G₁ toward S and G₂/M phases. Collectively, these findings support a ROP38-associated p53 signaling phenotype under the experimental conditions used here, while its relevance to physiological T. gondii infection remains to be established.

The ROP38–p53 link emerged from a hypothesis-driven candidate co-immunoprecipitation survey in which multiple signaling nodes were tested in parallel. Under our experimental conditions, representative ERK–MAPK- and PP2A-related regulators and PI3K–AKT–mTOR pathway components were not detectably co-enriched with ROP38, whereas p53 was detected in ROP38 immunoprecipitates from both HCT116 and HepG2 cells. Among the candidate proteins examined, p53 was the only ROP38-associated host factor detected in both cell lines, providing a basis for subsequent p53-focused biochemical analyses. Notably, this pattern distinguishes ROP38 from other well-characterized rhoptry kinases. ROP16 engages STAT3 and STAT6 to modulate host immune signaling [16], whereas ROP18—an active serine-threonine kinase whose catalytic domain has been studied—directly phosphorylates host IRGs to protect the parasite from immune clearance [17, 18]. Given the structural and evolutionary diversity of the ROPK family [17], our identification of p53 as a ROP38-associated host factor suggests that individual ROPKs may engage host signaling pathways beyond canonical immune-regulatory circuits, including p53-centered stress signaling.

A key biochemical finding is the phospho-state selectivity observed with ROP38-associated complexes. Phospho-specific analyses revealed a bias toward p53 pSer15/pThr18, whereas the other phospho-sites examined—Ser20, Ser33, Ser46, and Thr81—did not exhibit enrichment comparable to pSer15/pThr18 under the same conditions, indicating site-selective association rather than a nonspecific increase in p53 phosphorylation. Ser15 phosphorylation is widely associated with DDR-linked ATM/ATR signaling and broader DNA damage-related signaling dynamics [31–33]. Seminal biochemical work established that Ser15 phosphorylation stimulates p53-dependent transactivation through enhanced interaction with the p300 coactivator, an effect that operates independently of direct modulation of MDM2 binding [7]. Sequential phosphorylation models further indicate that Ser15 phosphorylation can facilitate subsequent Thr18 phosphorylation by CK1δ, thereby contributing to p53 stabilization [6]. Such selectivity is mechanistically notable because Ser15 and Thr18 reside within the p53 N-terminal TAD, where early phosphorylation events are closely linked to p53 stabilization and functional activation [5, 6, 34]. Thr18 phosphorylation weakens p53–MDM2 binding, thereby favoring p53 accumulation and downstream transcriptional competence [35–38]. Whether ROP38-induced pThr18 also functionally alters MDM2–p53 binding in cells remains unresolved. Importantly, the preferential Ser15/Thr18 pattern observed in ROP38 immunocomplexes was further supported by experiments using purified recombinant proteins, in which recombinant ROP38 directly associated with recombinant p53 and promoted phosphorylation of Ser15 and Thr18 in the presence of ATP. These findings strengthen the biochemical link between ROP38 and an early p53 activation-associated phospho-state, although the requirement for ROP38 catalytic activity cannot yet be established in the absence of a kinase-dead ROP38 mutant. For illustrative purposes, AlphaFold-predicted structures of ROP38 and p53 obtained from the AlphaFold Protein Structure Database [39, 40] were schematically arranged to indicate the ROP38 kinase-domain region and the p53 N-terminal region containing Ser15 and Thr18 (Additional file 5: Figure S3).

Previous studies have linked T. gondii effector biology to host p53-associated cell-cycle and apoptosis-related pathways. Chang et al. reported that ROP16 interacts with p53, promotes p53 phosphorylation at Ser15 and Ser37, and induces p53-dependent apoptosis and G₁ cell-cycle arrest [24]. Thus, the present study does not claim that p53-dependent host cell-fate regulation is newly described in T. gondii biology. Rather, our findings identify ROP38 as an additional rhoptry kinase associated with p53 signaling. The convergence of ROP16 and ROP38 on p53 raises the possibility that distinct rhoptry kinases may engage a shared host stress-regulatory node in a stage- or context-dependent manner. Because ROP16 expression has been reported to be highest in intracellular tachyzoites, whereas ROP38 is induced during bradyzoite differentiation [41], these effectors may exert temporally distinct or complementary effects on p53-associated host responses during different phases of parasite development. However, this possibility remains speculative, as the present study did not directly compare ROP16 and ROP38 or examine their functions in stage-specific parasite infection models.

The present findings should also be considered in relation to Peixoto et al., who reported that ROP38 overexpression in a type I parasite background dampened host transcriptional responses during infection, including genes associated with MAPK signaling, apoptosis, and cell proliferation [19]. These findings implicated ROP38 in MAPK-associated host transcriptional regulation, whereas the present study identifies p53 as a ROP38-associated host factor under isolated expression conditions. The apparent difference from our findings may reflect the distinct experimental contexts of intact parasite infection and isolated ectopic ROP38 expression. During infection, ROP38 is present within a broader parasite effector network, whereas the present study examined ROP38-associated signaling independently of other parasite-derived effectors. Thus, these findings may represent context-dependent rather than directly contradictory host responses. Importantly, our data do not establish that ROP38 induces a p53-dependent apoptotic program during physiological T. gondii infection.

Consistent with this complex-level enrichment pattern, whole-cell analyses showed that ROP38 expression enhances total p53 abundance and phosphorylation across multiple p53 sites, with prominent induction at Ser15 and Thr18. These biochemical changes were accompanied by increased γH2AX and PUMA protein levels, enhanced cleaved caspase-3, and elevated p21 transcription, indicating that isolated ROP38 expression is associated with DNA damage/apoptosis-related molecular outputs in p53-proficient human cancer cells. Cell-cycle analysis further showed a marked reduction in the G₁-phase population accompanied by increased S- and G₂/M-phase populations in both HCT116 and HepG2 cells, indicating redistribution of cell-cycle progression rather than a simple G₁-arrest phenotype. The increase in γH2AX observed in the present ectopic ROP38 expression system should also be interpreted in light of previous infection-focused studies showing that T. gondii infection can induce host DNA double-strand breaks, γH2AX accumulation, ATM/CHK2 or ATM-dependent DNA damage response signaling, and genome-instability-associated phenotypes, including binucleation, micronuclei formation, and chromosomal instability [21–23]. However, unlike these infection-based studies, our system examines isolated ROP38 expression in human cancer cells and therefore does not establish that ROP38 is responsible for infection-induced DNA damage or genome instability during natural T. gondii infection. The convergence of selective pSer15/pThr18 enrichment within ROP38 immunocomplexes with downstream p53-associated molecular readouts provides a framework for a candidate ROP38-associated p53 signaling module under ectopic expression conditions. A broader multisite p53 phosphorylation observed at the whole-cell level—in contrast to the selective pSer15/pThr18 enrichment in the ROP38 complex—may reflect secondary propagation through host kinase networks following ROP38–p53 engagement. Host kinases, such as ATM, ATR, or CK1δ may therefore still contribute to additional phosphorylation events on p53 in cells, although the current data do not define the responsible cellular kinase cascade.

More broadly, T. gondii secretes effectors that remodel host kinase-dependent signaling programs [42, 43]. Canonical examples include ROP16-mediated modulation of STAT signaling [16] and GRA24-mediated engagement of host p38 MAPK signaling [25], illustrating that parasite-derived effectors can interface with defined host signaling nodes through kinase-dependent or kinase pathway-modulating mechanisms. In parallel, infection-focused studies support that T. gondii can modulate apoptotic pathways and broader host-response programs, providing a biological context for effector-associated regulation of stress and cell-fate signaling [43, 44]. Within this broader framework, the present study shows that isolated ROP38 expression in human cancer cells is associated with a p53-centered signaling phenotype characterized by Ser15/Thr18-linked phosphorylation and DNA damage/apoptosis-associated molecular outputs. The physiological relevance of this phenotype during T. gondii infection, however, remains to be established.

From the parasite’s perspective, the relationship between T. gondii infection and host-cell apoptosis is complex and context-dependent. Both apoptosis-inhibitory and apoptosis-associated responses have been reported in different experimental systems, and these divergent outcomes may reflect differences in host-cell type, parasite strain, infection stage, and the combined activity of multiple parasite effectors [24, 45, 46]. Accordingly, the apoptosis-associated molecular outputs observed following isolated ROP38 expression in human cancer cells are not necessarily inconsistent with previous infection-based findings. In the present reductionist system, ROP38 was expressed independently of other parasite effectors, whereas during natural infection the net host-cell response reflects the coordinated activity of multiple parasite-derived factors. Therefore, the current findings should not be interpreted as evidence that ROP38 induces apoptosis during physiological T. gondii infection. Rather, they indicate that isolated ROP38 expression can engage apoptosis-associated signaling under the experimental conditions used here, a possibility that will require validation using physiologically relevant infection models, including wild-type, ROP38-deficient, and complemented parasite strains.

From a translational perspective, whether ROP38-mediated p53 activation can influence cancer-cell survival or therapeutic sensitivity remains an open question that will require dedicated functional and pharmacological studies.

Several limitations of this study should be explicitly acknowledged. First, the use of a heterologous overexpression system—stable lentiviral expression of GFP-tagged ROP38 in human cancer cell lines—does not fully recapitulate the physiological context of T. gondii infection. Second, HCT116 and HepG2 are tumor-derived cancer cell lines and do not represent non-tumoral host cells, primary cells, tissues, or in vivo infection settings. Cancer cells may harbor rewired DNA damage response, p53/checkpoint signaling, apoptosis regulation, and altered DNA repair pathway usage, including ATM/ATR- and BRCA-associated repair programs [47]. In addition, tumor-derived cells may exhibit chromosomal instability, aneuploidy, and structural rearrangements [48]. Moreover, p53-linked stress signaling in tumor-derived cells may result in cell-fate outcomes distinct from those in normal cells, including senescence or mitotic catastrophe rather than canonical apoptosis [49]. Therefore, the increases in γH2AX, PUMA, cleaved caspase-3, and p21 observed here should be interpreted as DNA damage/apoptosis-associated molecular outputs in p53-proficient human cancer cells, not as evidence of a normal host-cell DNA damage response or a physiological apoptotic response during T. gondii infection. Validation in non-tumoral or primary host cells, including fibroblasts, immune cells, and macrophages, will be required to determine the broader relevance of the ROP38–p53 relationship. Third, although purified recombinant ROP38 directly associated with recombinant p53 and promoted Ser15/Thr18 phosphorylation in the presence of ATP, the present study did not include a catalytically inactive ROP38 mutant. Therefore, the requirement for ROP38 catalytic activity in this phosphorylation reaction remains unresolved. In addition, the broader multisite p53 phosphorylation observed in ROP38-expressing cells may involve secondary host kinase signaling that was not defined in the present study. Fourth, the structural representation shown in Additional file 5: Figure S3 is an illustrative arrangement of individually predicted ROP38 and p53 structures and should not be interpreted as a predicted protein complex, docking model, or experimentally defined interaction interface.

Thus, several future directions remain important for resolving the mechanism and physiological relevance of the ROP38–p53 relationship. These include: (i) generation and characterization of kinase-dead ROP38 mutant constructs to determine the contribution of catalytic activity to the p53 phenotypes observed; (ii) experimental T. gondii infections with ROP38-deficient and complemented strains to assess the infection-context relevance of the ROP38-associated p53 phenotype; (iii) validation in non-tumoral or primary host cells; (iv) genome-wide transcriptome and chromatin profiling to define the broader scope of ROP38-associated p53 target gene regulation; and (v) structural studies to further define the molecular basis of the ROP38–p53 association.

Conclusions

In conclusion, this study identifies a candidate ROP38-associated p53 signaling module in human cancer cells. Under isolated ROP38 expression conditions, ROP38 Full and its kinase-domain fragment associated with p53 and preferentially enriched p53 pSer15/pThr18 species. Purified recombinant ROP38 also directly associated with recombinant p53 and promoted Ser15/Thr18 phosphorylation in vitro. At the cellular level, ROP38 expression was accompanied by increased p53 abundance and multisite phosphorylation, elevated DNA damage/apoptosis-associated molecular markers, and redistribution of the cell-cycle profile from G₁ toward S and G₂/M phases. Together, these findings provide biochemical evidence that ROP38 can engage p53-associated signaling under the experimental conditions used here. However, the requirement for intrinsic ROP38 catalytic activity and the physiological relevance of this relationship during T. gondii infection or in non-tumoral host-cell contexts remain to be established using kinase-dead ROP38 mutants, primary/non-tumoral cells, and ROP38-modified parasite infection models.

Abbreviations

ADP

Adenosine diphosphate

AKT

Protein kinase B

ATM

Ataxia-telangiectasia mutated

ATR

ATM and Rad3-related

BIK

BCL2-interacting killer

CDKN1A (p21)

Cyclin-dependent kinase inhibitor 1 A

Co-IP

Co-immunoprecipitation

DDR

DNA damage response

DUSP

Dual-specificity phosphatase

EF-1α

Elongation factor-1 alpha

ERK

Extracellular signal-regulated kinase

FACS

Fluorescence-activated cell sorting

GAPDH

Glyceraldehyde-3-phosphate dehydrogenase

GFP

Green fluorescent protein

HRP

Horseradish peroxidase

IP

Immunoprecipitation

IRG

Immunity-related GTPase

KiD

Kinase domain

MAPK

Mitogen-activated protein kinase

MCS

Multiple cloning site

MDM2

Mmouse double minute 2 homolog

mTOR

Mechanistic target of rapamycin

NF-κB

Nuclear factor kappa B

p21 (CDKN1A)

Cyclin-dependent kinase inhibitor 1 A

PDK1

3-phosphoinositide-dependent protein kinase-1

PEA15

Phosphoprotein enriched in astrocytes 15

PI3K

Phosphoinositide 3-kinase

PP2A

Protein phosphatase 2 A

PP2Cα

Protein phosphatase 2 C alpha

PP2A-B55

PP2A regulatory subunit B55 (B family)

PP2A-B56

PP2A regulatory subunit B56 (B′ family)

PTEN

Phosphatase and tensin homologs

PTPRR

Protein tyrosine phosphatase receptor type R

PUMA

P53 upregulated modulator of apoptosis

RFU

Relative fluorescence units

ROPK

Rhoptry protein kinase

ROP38

Rhoptry kinase family protein 38

TAD

Transactivation domain

T. gondii

Toxoplasma gondii

USP7

Ubiquitin-specific protease 7

γH2AX

Phosphorylated histone H2AX (Ser139)

Authors’ contributions

SHS, JEL, and EHS contributed to the conceptualization of the study. SHS and JEL contributed to the study design and methodology. SHS, JEL, and EJC performed the experiments and conducted the investigation. SHS and JEL acquired and curated the data. SHS and EHS analyzed and interpreted the data. SHS drafted the original manuscript. SHS and EHS substantively reviewed and edited the manuscript. EHS and SHS acquired funding for the study. All authors read and approved the final submitted manuscript and agree to be accountable for their own contributions and for ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

Funding

This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea government (MSIT) (IRIS No. RS-2025-00516274 and RS-2025-00559259). The funder had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Data availability

All data supporting the findings are included in this article and its additional files. Additional raw data supporting the conclusions are available from the corresponding author and can be provided upon reasonable request.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.