A dual-pronged host-directed therapeutic targeting cyclophilin A and pathogenic interferon response abrogates virus-triggered pregnancy pathologies

Abstract

Pathogenic viruses threaten fetal development by achieving vertical transmission and instigating placental immunopathology, while the pathobiological mechanisms and effective therapeutics remain critical gaps. Here, we reveal cyclophilin A (CypA) as a crucial host factor necessary for Zika virus (ZIKV) replication in human placental trophoblasts, acting independently of its canonical functions. ZIKV infection recruits CypA into the viral replication organelle and reconfigures its interactome, thereby subverting host RNA decay machinery and stress granule-mediated antiviral surveillance. Both genetic ablation of CypA and its pharmacological inhibition with clinically approved drug ciclosporin A (CsA) restrict ZIKV transplacental transmission and corresponding placental and fetal pathologies. Beyond its anti-ZIKV potency, CsA concurrently counteracts pathological type I interferon signaling by targeting the JAK1-STAT1/2 pathway, broadly ameliorating pregnancy-specific interferonopathies driven by viral infection or endogenous double-stranded RNA stress. Our findings elucidate CypA-governed ZIKV pathogenesis and license CsA as a promising dual-action therapeutic to counteract congenital viral infections.

Vertical transmission of Zika virus (ZIKV) poses significant risks to fetal development. Here the authors show that cyclophilin A (CypA) is essential for ZIKV replication in placental cells, and demonstrate that ciclosporin A is protective by inhibiting CypA-driven ZIKV infection and the pathogenic interferon response in trophoblasts.

Introduction

Infections during pregnancy remain a major public health concern and pose a serious threat to maternal and fetal health, frequently culminating in miscarriage, intrauterine growth restriction (IUGR), and other pathogen-specific complications. While the placenta functions as a formidable antimicrobial barrier against vertical transmission, TORCH pathogens (Toxoplasma, other, rubella, cytomegalovirus (CMV) and herpesviruses), as well as certain newly emerging viruses, have evolved sophisticated strategies to breach placental defenses and ultimately infect the fetus^1–3. Beyond direct viral invasion, perturbation of the finely tuned immune equilibrium at the maternal-fetal interface is increasingly appreciated as a key driver of viral pathogenesis by compromising the normal placental development and barrier functions^2,4–7. Of note, although the activation of the interferon (IFN) pathway is a cornerstone of innate immunity and plays a critical role in limiting viral infection, recent studies in mice have demonstrated that a spectrum of congenital abnormalities arises primarily from undue IFN responses, rather than from viral burden within the fetoplacental unit^5,8. Thus, these findings imply that antiviral therapy alone is insufficient to reverse the distinctive sequelae of viral infections in the context of pregnancy.

Zika virus (ZIKV) preferentially targets placental tissues and has deleterious effects on fetal development, both through direct transplacental infection and aberrant activation of placental innate immunity^5,9. Despite this, the specific host factors co-opted for ZIKV infection and pathogenesis within the placenta are largely unknown. To address this, we recently performed an unbiased genome-wide CRISPR screening in placental trophoblasts and identified a machinery specific for ZIKV entry¹⁰. In addition, our screen also highlighted cyclophilin A (CypA), encoded by the peptidylprolyl isomerase A (PPIA) gene, as a leading candidate devoid of any established role in ZIKV biology. Elucidating the precise function and mechanistic underpinnings of CypA in ZIKV vertical dissemination is therefore crucial for advancing our understanding of viral pathogenesis during pregnancy.

ZIKV possesses a positive-sense, single-stranded RNA genome that is translated by host ribosomes into a single polyprotein and subsequently cleaved into three structural (C, prM, E) and seven non-structural (NS) proteins. The NS proteins orchestrate extensive remodeling of endoplasmic reticulum (ER) membranes to scaffold specialized, protective compartments known as replication factories (RFs)¹¹. In response, the host deploys multiple intrinsic antiviral defenses to restrict infection, including sequestration of viral RNA into stress granules (SGs), induction of interferon-stimulated genes (ISGs), and activation of intracellular RNA decay pathways^8,12–14. Although ZIKV is known to counteract these antiviral barriers, the precise molecular mechanisms underlying this evasion remain poorly understood.

Here, we uncover that CypA serves as a viral RNA-binding protein (RBP) that facilitates ZIKV replication and vertical transmission by shielding the virus from the host RNA decay machinery and undermining SG assembly. Importantly, we identify the clinically approved drug cyclosporine A (CsA) as a potential therapeutic antiviral that addresses ZIKV-driven pathologies by limiting viral transplacental transmission and mitigating virus-instigated interferonopathies. Our findings reveal a novel CypA-directed cellular process underlying ZIKV infection and highlight a viable therapeutic to prevent congenital viral transmission and IFN-associated pregnancy complications.

Results

CypA is required for ZIKV replication in human trophoblasts

To systemically investigate the function of CypA on ZIKV infection in trophoblasts, we generated CypA knockout (KO) JEG-3 cells using a CRISPR-associated protein 9 (CRISPR-Cas9) system (Supplementary Fig. 1a, b). Multiple assays consistently demonstrated that depletion of CypA significantly restricted ZIKV infection, as evidenced by a decrease in ZIKV-positive cells and a markedly diminished production of infectious viral progeny (Fig. 1a, and Supplementary Fig. 1c–e). Restoration experiments confirmed that reintroduction of CypA via gene overexpression or exogenous recombinant CypA treatment could fully reinstate ZIKV susceptibility in CypA^-/- cells (Fig. 1b, c). To delineate the specific stages of the ZIKV life cycle influenced by CypA, we assessed viral binding, internalization, and subsequent replication. Real-time quantitative PCR (RT-qPCR) analyses revealed comparable levels of ZIKV attachment and internalization between Ctrl and CypA-KO cells, suggesting that CypA is unlikely to participate in the ZIKV entry process (Fig. 1d). Instead, infection kinetics showed a pronounced decrease in ZIKV infection at 24- and 48-hour post-infection (hpi) (Fig. 1a and Supplementary Fig. 1f), implying that CypA exerts a pro-ZIKV effect predominantly by targeting the replication phase. To substantiate this, we utilized an in vitro-transcribed ZIKV replicon system, wherein luciferase activity serves as a proxy for vRNA replication and host factor requirements independent of viral entry^10,15. Silencing of CypA resulted in an over 50% reduction of luciferase signal (Fig. 1e), further underscoring the critical role of CypA in promoting ZIKV RNA synthesis. Additionally, we also generated CypA-knockdown (KD) human trophoblast stem cells (hTSCs) via shRNA-mediated lentiviral transduction. CypA depletion had no overt effects on cell viability or apoptosis, and the expression of key stemness markers (Supplementary Fig. 1g–j). Consistent with the observation in JEG-3 cells, CypA KD markedly disrupted ZIKV infection at both 24 hpi and 48 hpi hTSCs (Fig. 1f). Collectively, our data highlight CypA as a host factor supporting ZIKV replication in trophoblasts.

Fig. 1. CypA mediates ZIKV replication in human trophoblasts.

a Flow cytometry analysis showing the ZIKV-positive cells. MOI = 0.1. n = 5. b Representative immunofluorescence of ZIKV-E (bottom) and western blot of the reconstituted cyclophilin A (CypA) expression. 48-hour post-infection (hpi), MOI = 0.1. c RT-qPCR of ZIKV mRNA in CypA-KO cells following CypA overexpression (OE) or recombinant CypA (rCypA) treatment at 24 hpi. MOI = 0.1. n = 3. d RT-qPCR assessment of ZIKV binding and internalization. MOI = 10. n = 3. e Luciferase activity of the ZIKV replicon at 48 h. n = 6. f RT-qPCR time course of ZIKV mRNA. MOI = 0.5. n = 3. g Co-immunofluorescence of ZIKV-E and TP63 (an hTSC marker) at 48 hpi. CsA2, 2 μM cyclosporine A (CsA); CsA4, 4 μM CsA. h RT-qPCR of ZIKV mRNA at 48 hpi. MOI = 0.5. n = 3. i Experimental design using human placental organoids (hPO). j Representative ZIKV-E immunofluorescence in hPO. k Quantification of ZIKV-E+ cells in hPOs. Mock, n = 3; DMSO, CsA2, and CsA4, n = 6/group. l RT-qPCR of ZIKV mRNA in hPO supernatants. n = 3. m Representative light microscopy images of hPO across six days. n, o Quantitative analysis of hPO growth. Surface area measurements (n) and size increase on day 6 vs. day 0 (o). n = 4. p Diagram of inhibitor targets. mPTP, mitochondrial permeability transition pore. q Summary table of IC₅₀, CC₅₀, and SI values. r RT-qPCR analysis of ZIKV mRNA following 48 h of inhibitor treatment. n = 3. Data are presented as mean values ± SEM. n represents biologically independent samples. Statistical significance was performed by One-Way (c, e, k, o, r) or Two-way (a, d, f, l, n) ANOVA followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

Pharmacological inhibition of CypA protects trophoblasts against ZIKV infection

We then assessed whether an FDA-approved CypA inhibitor, CsA, could effectively repress ZIKV infection in trophoblast models. In JEG-3 cells, CsA treatment profoundly decreased intracellular ZIKV titers at 24 and 48 hpi with a half-maximal inhibitory concentration (IC₅₀) of 2.233 μM and half-maximal cytotoxic concentration value (CC₅₀) of 11.2 μM (Supplementary Fig. 1k, l). In hTSCs, CsA treatment resulted in a dose-dependent reduction in ZIKV-E+ cells, intracellular ZIKV RNA levels and viral titers in supernatants, without affecting cell stemness and viability (Fig. 1g, h and Supplementary Fig. 1m–p). Furthermore, CsA administration in hTSCs phenocopied the genetic loss of CypA, potently suppressing ZIKV replicon activity (Supplementary Fig. 1q). Subsequently, we experimentally determined if CsA exerts an antiviral effect through CypA blockade in CypA-silenced trophoblasts. The finding that CsA provided no additional antiviral effect following CypA depletion suggests that its mechanism of action largely depends on targeting CypA (Supplementary Fig. 1r, s). To corroborate the antiviral activity of CsA with a more physiologically relevant system, we employed the three-dimensional human placenta organoids (hPOs), which recapitulate the functional complexity of the in vivo human placenta (Fig. 1i). CsA conferred substantial resistance to ZIKV infection in hPOs, as manifested by a marked reduction in ZIKV-E positive cells and progeny virions release (Fig. 1j–l). Correspondingly, ZIKV-driven growth retardation of hPOs was robustly mitigated by 4 μM CsA treatment (Fig. 1m–o). Above all, these findings demonstrate that CsA exerts a potent antiviral effect against ZIKV in human placentas.

The canonical functions of CypA are dispensable for ZIKV infection in trophoblasts

CypA is a key immunophilin that exhibits peptidyl-prolyl isomerase (PPIase) activity, facilitating protein folding and cellular signaling. Upon binding to CsA, the CypA-CsA complex can suppress calcineurin phosphatase activity, thereby blocking nuclear factor of activated T cells (NFAT) translocation and exerting immunosuppressive effects¹⁶. To dissect the molecular mechanisms underlying CypA-dependent ZIKV replication in trophoblasts (Fig. 1p), we first interrogated the requirement for its PPIase activity. Treatment with TMN355, a CypA inhibitor ~27‑fold more potent than CsA in blocking isomerase activity, failed to impair ZIKV infection (Supplementary Fig. 2a), hinting that the enzymatic function of CypA is not essential¹⁷. Consistently, CypA^-/- JEG-3 cells reconstituted with either WT CypA or the catalytically inactive R55A mutant harbored equivalent viral titers (Supplementary Fig. 2b, c). As PPIase activity of CypA is known to support hepatitis C virus (HCV) RNA synthesis, these findings suggest virus-specific functional adaptations of CypA¹⁸. Next, we assessed the involvement of CypA downstream target calcineurin signaling in ZIKV infection. FK506, which represses the calcineurin phosphatase activity independently of CypA, had no effect on ZIKV replication (Supplementary Fig. 2d)¹⁷. Likewise, pharmacological blockade of NFAT, the transcriptional effector of calcineurin, did not restrict ZIKV infection (Supplementary Fig. 2e)¹⁹. By contrast, treatment of trophoblasts with NIM811, a CsA derivative unable to target calcineurin, profoundly suppressed viral replication, dissociating the proviral role of CypA from the calcineurin/NFAT signaling axis (Fig. 1q and Supplementary Fig. 2f). Parallel experiments in hTSCs yielded consistent results, with only NIM811 displaying antiviral potency (Fig. 1r and Supplementary Fig. 2g). Moreover, we revealed that CsA and NIM811 exhibited broad-spectrum virucidal function across the tested ZIKV strains, including PRVABC59 and Paraiba 2015, whereas other inhibitors remained ineffective (Supplementary Fig. 2h). Given that CsA and NIM811 also inhibit mitochondrial permeability transition pore (mPTP) formation, we investigated whether mPTP contributes to ZIKV infection^20,21. RT-qPCR analysis showed that inhibiting pore opening and associated Ca²⁺ overload using mPTP-IN-1 and BAPTA‑AM did not affect ZIKV infection in trophoblasts (Fig. 1q and Supplementary Fig. 2i, j). Together, our data reveal that the CypA regulates ZIKV replication in trophoblasts through mechanisms that are non-enzymatic and independent of canonical signaling pathways.

CypA redistributes to the ZIKV replication factory for enhancing viral RNA stability

Intriguingly, we observed that ZIKV infection provoked a dramatic relocalization of CypA from widespread distribution into perinuclear puncta that colocalize with viral dsRNA, a hallmark of the RFs (Fig. 2a–c). Given that RFs contain multiple essential viral proteins required for vRNA synthesis, especially for NS3 and NS5, the recruitment of CypA into RFs raised the hypothesis that CypA directly engages the viral replicases^22,23. To this end, we performed co-immunoprecipitation (Co-IP) assays using HEK293T cells co-transduced with HA-tagged CypA and individual ZIKV NS proteins, and found that CypA was selectively associated with ZIKV-NS1, NS3, NS2A, and NS4A (Fig. 2d). In the context of an authentic viral infection, co-IP results also revealed robust interactions between CypA and these identified viral NS proteins (e.g., NS3, NS1) in hTSCs (Fig. 2e). We selected NS3 as a representative NS for further immunofluorescence analysis, owing its essential catalytic role in vRNA replication within the RFs^11,24. Confocal imaging clearly visualized the co‑localization of endogenous CypA aggregates with NS3 in ZIKV-infected hTSCs (Fig. 2f, g). Overall, these data reveal that ZIKV infection triggers the recruitment of CypA to the RF, where it participates in the viral replication complex via physical interaction with core replication factors.

Fig. 2. CypA redistributes into the ZIKV replication factory and protects the virus against host RNA decay.

a Representative confocal images revealing the distribution of CypA and dsRNA. Arrow, the region for colocalization analysis. b, c Fluorescence intensity profile illustrating the distribution of CypA (b) and its spatial overlap with dsRNA (c). d, e Co-Immunoprecipitation assays showing the interaction between ectopically expressed CypA and ZIKV nonstructural proteins in HEK293T cells (d) or ZIKV-infected hTSC (e). f, g Representative confocal images (f) and quantitative colocalization analysis (g) depicting the colocalization of NS3 with CypA in ZIKV-infected cells. Arrow, the region for colocalization analysis. h Schematic pipeline of the chromatin isolation by RNA purification (ChIRP). i ChIRP-WB analysis showing the interplay between viral RNA (vRNA) and CypA. j hTSCs expressing CypA-Flag or vector were infected with ZIKV (MOI = 0.1) for 48 h. The enriched vRNA was immunoprecipitated using the anti-Flag antibody and subsequently quantified by RT-qPCR. n = 4 biological replicates. k Diagram of the potential mechanisms by which recruitment of CypA into the replication factory (RF) suppresses host IFN-mediated antiviral immunity or RNA degradation system to facilitate ZIKV replication. RIG-I Retinoic acid-inducible gene I; MDA Melanoma differentiation-associated protein 5; TBK1 TANK-binding kinase 1; IRF3 Interferon regulatory factor 3. l Schematic of the experimental outline. m RT-qPCR quantification of ZIKV RNA decay over time upon genetic or pharmacological blocking of CypA. n = 3 biological replicates. Data are presented as mean values ± SEM. Statistical significance was performed by unpaired two-tailed Student’s t test (j) or Two-way ANOVA followed by Tukey’s multiple comparisons test (m). Source data are provided as a Source Data file.

Considering that CypA is associated with the ZIKV replication compartment, we asked whether it acts as a non-classical RNA-binding protein engaging ZIKV vRNA. To capture ZIKV-interacting proteins in infected hTSCs systematically, we conducted chromatin isolation by RNA purification (ChIRP) using a panel of biotinylated antisense oligos targeting the ZIKV genome (Fig. 2h). Western blot (WB) analysis of the ChIRP eluates demonstrated a substantial co-enrichment of CypA specifically with the vRNA (Fig. 2i). Additionally, immunoprecipitation of CypA-Flag robustly pulled down ZIKV vRNA, providing complementary evidence of a direct interplay (Fig. 2j). Therefore, these orthogonal approaches conclusively unveil the formation of ZIKV vRNA-CypA complexes in the RFs.

The RF provides an “immune-privileged” niche that shields replicating RNA from host innate immune sensing and degradation²³. Noting the abundant CypA within the RF, we speculated that CypA may play a role in masking ZIKV from host antiviral surveillance (Fig. 2k). To this end, we first evaluated the IFN responses following CypA depletion. WB data revealed that the phosphorylation of TANK-binding kinase 1 (TBK1) and interferon regulatory factor 3 (IRF3), the key signaling mediators linking pathogen recognition to interferon induction, was not further activated upon CypA depletion (Supplementary Fig. 3a, b). Consistently, qRT‑PCR analysis of IFNB1, IFNL1, and the representative ISGs (MX2 and IFIT2) at 12 hpi showed no significant differences between control and CypA‑silenced trophoblasts. Thus, these data indicate that CypA deficiency did not affect initial viral sensing and IFN signaling (Supplementary Fig. 3c, e). The observed decline in ISG expression at 24–48 h coincided with reduced viral replication, suggesting it is likely a secondary effect of lower viral titer rather than direct modulation of IFN signaling (Supplementary Fig. 3d, f). Next, we investigated whether CypA regulates ZIKV vRNA stability by treating infected cells with a flavivirus replication inhibitor NITD008 (Fig. 2l). Genetic ablation of CypA or pharmacological inhibition by CsA markedly accelerated vRNA decay (Fig. 2m and Supplementary Fig. 3g). Taken together, our data imply that ZIKV hijacks CypA within the RFs to protect its genome from host RNA-degradation defenses, thus ensuring efficient replication.

ZIKV infection modulates Cyclophilin A proteo-interactome

Prompted by the striking re-distribution of CypA in ZIKV-positive cells, we hypothesized that infection may reshape the CypA interactome. To address this, we performed proteomics profiling of CypA-interacting proteins by IP-mass spectrometry (Fig. 3a). Principal-component analysis revealed pronounced intergroup segregation, indicative of a profound rewiring of the CypA-centred interaction network (Fig. 3b). Remarkably, over 21% of the CypA-bound proteins were significantly influenced upon ZIKV infection (Fold change >1.5, p < 0.05), with 208 increased and 417 decreased in abundance (Supplementary Fig. 4a). Gene ontology (GO) analysis of the upregulated partners revealed a strong enrichment in ribosome biogenesis, RNP complex biogenesis, rRNA metabolic process, and protein targeting to ER processes, which echoes the altered subcellular localization of CypA into ZIKV RFs (Fig. 3c and Supplementary Fig. 4b). In contrast, the interplay between CypA and proteins implicated in mRNA transport, splicing and RNA stability were significantly decreased, suggesting ZIKV-driven subversion of host RNA processing (Fig. 3c). Notably, proteins associated with the regulation of innate immune response, response to virus, regulation of canonical NF-κB signal transduction, stress granule assembly, and regulation of pattern recognition receptor signaling pathway were diminished in response to ZIKV infection (Fig. 3c and Supplementary Fig. 4c), indicating that ZIKV erodes CypA-mediated host immune defense^25,26. Additionally, we noted that over 70% of modulated CypA partners (e.g., HNRNPA1, PABPC1, G3BP1) contain intrinsically disordered regions (IDRs) and participate in phase separation and membrane-less organelle formation, such as SGs (Fig. 3d), raising the possibility that remodelled protein condensate dynamics may facilitate viral replication²⁷. Independent co-immunoprecipitation assays validated our proteomics data, revealing that ZIKV infection induces the dissociation of CypA from representative host factors, including G3BP1 and adenosine deaminase acting on RNA 1 (ADAR1) (Fig. 3e). These data collectively uncover an extensive reprogramming of the CypA binding network by ZIKV.

Fig. 3. ZIKV disrupts the host stress granule assembly by reshaping the CypA interactome.

a Proteomic profiling of CypA interactome in mock- and ZIKV-infected hTSCs. b PCA of global proteomic datasets. The percentage of total variance explained by each principal component is indicated in the axis labels. n = 3 biological replicates. c Bubble plots of GO-defined biological processes for upregulated (left) and downregulated (right) hits after ZIKV infection. d Heatmap for representative proteins with intrinsically disordered regions (IDRs). e Co-Immunoprecipitation validation of the interaction between CypA and indicated proteins. ADAR1, Adenosine deaminase acting on RNA 1; G3BP1, Ras GTPase-activating protein-binding protein 1. f Schematic of the potential interaction between stress granule (SG) dynamics and ZIKV infection. g SG formation upon ZIKV infection. Representative confocal images of G3BP1 and ZIKV-E 24 hpi, followed by 1 h sodium arsenite treatment (0.5 mM). h Quantification of SG number and size. i Representative images of G3BP1 and ZIKV-NS3 in hTSCs transfected with individual ZIKV-NS3 for 24 h, followed by 1 h sodium arsenite treatment. j Quantification of SG number and size. Data are presented as mean values ± SEM. n = 15 independent fields. Statistical significance was performed by One-Way ANOVA followed by Tukey’s multiple comparisons test (h, j). Source data are provided as a Source Data file.

ZIKV dampens SG formation largely dependent on CypA

SGs function as an essential antiviral hub to antagonize viral replication and RF formation by condensing and sequestering vRNA. In response, ZIKV disassembles these granules to subvert the antiviral state and enhance its survival by an undefined mechanism^13,28. The substantial perturbations in CypA-associated SG components observed in our proteomic profile during ZIKV infection prompt the hypothesis that CypA may be integral to the viral disruption of SGs (Fig. 3f). To confirm this, we assessed SG dynamics by quantifying the number and size of G3BP1-positive signals in trophoblasts following the treatment with SG inducer (sodium arsenite). In uninfected cells, either CypA depletion or CsA treatment produced discernible effects on G3BP1 expression and SG formation (Supplementary Fig. 5a–d). Conversely, ZIKV infection robustly impeded SG assembly, consistent with a prior report²⁸. Strikingly, this suppression was reversed in CypA-deficient and CsA-treated trophoblasts (Fig. 3g, h, and Supplementary Fig. 5b–g). To dissect these findings and exclude the indirect effects from reduced viral load, we expressed the FLAG-tagged ZIKV NS3 protein (a known pathogenic factor of ZIKV in SG inhibition) in hTSCs. As anticipated, compared to controls, cells expressing NS3 exhibited a ∼50% and ∼60% reduction in the average number and size of SGs, respectively, which was significantly attenuated by either CypA ablation or CsA administration (Fig. 3i, j and Supplementary Fig. 5h–j). Together, these results demonstrate that ZIKV co-opts CypA to inactivate SG-driven antiviral defenses, thereby improving vRNA stability.

CypA inhibition restricts ZIKV maternal-fetal transmission and adverse pregnancy outcomes

To interrogate the physiological functions of CypA in vivo, we set up a mouse model in which Ppia^+/- sires were crossed with Ppia^+/- female mice. This breeding strategy results in placentas and fetuses with all three genotypes within the same litter, thus enabling exclusion of potential maternal systemic influences¹⁰. One day after administration of an anti-IFNAR-1 antibody, dams were subcutaneously inoculated with 10³ PFU ZIKV at either embryonic day (E) 8.5 or E12.5 (Fig. 4a). The virologic data demonstrated that Ppia-deficient placentas and the corresponding amniotic fluid and fetal brain harboured >10-fold lower ZIKV levels relative to their Ppia^+/+ or Ppia^+/- counterparts, both at early (E8.5-E12.5) and late gestation (E12.5-E16.5) stages (Fig. 4b, c). Additionally, no significant sex-specific differences in viral titers were detected in placentas or fetuses (Supplementary Fig. 6a). Therefore, these findings support CypA as a pivotal placental factor facilitating ZIKV transplacental transmission and a candidate therapeutic target to tackle congenital ZIKV infection.

Fig. 4. CypA loss-of-function counteracts ZIKV vertical transmission and pathogenicity in mice.

a Schematic depiction of ZIKV infection experiments in Ppia^+/- mice. b, c RT-qPCR of ZIKV mRNA in placenta, fetal brain, and amniotic fluid at E12.5 (b) and E16.5 (c). For placenta and fetal brain, n = 15 (Ppia^+/+), 25 (Ppia^+/-), 14 (Ppia^-/-); amniotic fluid, n = 12, 21, 12, respectively (b). For placenta, n = 11 (Ppia^+/+), 21 (Ppia^+/-), 14 (Ppia^-/-); fetal brain, n = 10, 17, 13, respectively (c). d Timeline of drug treatment in pregnant mice. e RT-qPCR measurement of viral mRNA in placentas (n = 16/group), amniotic fluid (n = 8 for DMSO group, n = 9 for CsA group), and fetal brain (n = 17 for DMSO group, n = 16 for CsA group). f Representative immunofluorescence of placental sections co-stained for ZIKV-E and pan-CK (a trophoblast marker). g, h Quantification of ZIKV RNA levels in maternal serum (g) and representative tissues (h). n = 6/group. i Representative appearance of the E12.5 placenta and fetus. j The number and rate of fetal resorption. k Scatterplot of differential gene expression between the indicated placentas. l GO analysis of pathways significantly altered by CsA in ZIKV-infected placentas. m GSEA analysis of IFN-response pathways. n Heatmap displays z-score normalized FPKM expression values for selected ISGs from placental tissue. Circle size, log₂(fold change). Circle color, adjusted p value (Bonferroni method). Data are presented as mean values ± SEM. Statistical significance was performed by unpaired two-tailed Student’s t test (e, g) or One-way ANOVA followed by multiple comparisons test (b, c, h). Source data are provided as a Source Data file.

Next, to evaluate the antiviral efficacy of CsA in vivo, we utilized our established mouse model of gestational ZIKV infection^10,29. Ifnar1⁻/⁻ dams are mated with WT sires to generate Ifnar1⁺/⁻ placentas, thereby rendering the dams susceptible to infection while largely preserving intact placental interferon signaling. Dams were treated with CsA following a dosing regimen (10 mg/kg intraperitoneally every 48 h) adapted from prior studies^30–32, which we confirmed herein was not associated with maternal hepatotoxicity (Fig. 4d and Supplementary Fig. 6b). CsA administration resulted in an approximately 5 ~ 10-fold reduction of viral RNA levels in placentas, amniotic fluid, and fetal brains compared with controls (Fig. 4e). Immunostaining for ZIKV-E protein consistently demonstrated significantly fewer ZIKV-positive cells in the CsA-treated placentas (Fig. 4f). Interestingly, while CsA treatment resulted in a modest reduction in maternal blood viral burden, viral titers in several maternal tissues remained unchanged (Fig. 4g, h). We interpret this mild reduction in maternal viremia as potentially resulting from decreased ZIKV spread from the placental reservoir.

In accordance with virological outcomes, CsA-treated mice showed a reduced rate of fetal resorption and improved embryonic and placental development (Fig. 4i, j). Considering that ZIKV infection-associated adverse pregnancy outcomes are largely linked to placental dysfunctions, we focused on placental tissues to further corroborate the protective effects of CsA treatment³³. ZIKV challenge caused severe placental insufficiency, with more than 50% reductions in total placental area and labyrinth layer, which was profoundly mitigated in the CsA-treated group (Supplementary Fig. 6c–e). To gain a comprehensive understanding of the gene signature driven by CsA administration, we performed bulk RNA-seq on mouse placentas and noted that the majority of ZIKV‑perturbed transcripts were reversed upon CsA treatment (Fig. 4k). GO analysis uncovered significant downregulation of biological processes in the CsA group related to viral infections, including response to virus, regulation of viral process, and viral genome replication, confirming the in vivo antiviral activity of CsA (Fig. 4l). Gene Set Enrichment Analysis (GSEA) revealed that CsA treatment could promote trophoblast differentiation, facilitate blood vessel formation, inhibit trophoblast apoptosis, and improve placental nutrient transportation, collectively indicating a beneficial effect on placenta development (Supplementary Fig. 6f–h). Corroborating these findings, immunostaining revealed a marked improvement in the structure and integrity of the placental labyrinth, the core maternal-fetal interface (Supplementary Fig. 6i). Above all, our compelling results underscore CsA as an efficacious medicine to avert ZIKV pathogenicity during pregnancy.

CsA abrogates ZIKV pathogenesis during pregnancy by dampening detrimental IFN responses

We observed an intense downregulation of pathways associated with the regulation of innate immune response and response to type I IFN in ZIKV-infected placentas treated with CsA (Fig. 4m). Notably, the expression of IFN-related genes (e.g., Mx2, Myd88, Ifitm3, Isg15) and placental developmental parameters were completely restored to baseline levels comparable to those in uninfected tissues (Fig. 4n and Supplementary Fig. 6d, e), representing a more pronounced recovery than the reduction in ZIKV titers. Therefore, we speculated a direct role for CsA in mitigating the pathogenic IFN‑I signaling known to drive ZIKV‑associated placental disorders⁵. To address this, we employed a mouse model in which poly(I:C), a synthetic analogue of dsRNA, was administered to mimic viral infections and exclude possible pleiotropic effects resulting from decreased ZIKV replication. Pregnant mice received poly(I:C) (200 μg) at E10.5, while CsA was administered every other day from E9.5 to E12.5 (Fig. 5a). Consistent with our hypothesis, CsA treatment substantially impeded poly(I:C)-induced IFN responses in placentas, as shown by the downregulated expression of multiple ISGs (Oas1a, Isg15, Ifitm3, Irf7) (Fig. 5b). As excessive IFN signaling and pathogenic ISG induction (such as IFITMs and GBP5) are detrimental to the developing fetuses by inhibiting the formation of the placental syncytiotrophoblast (STB) barrier, we next assessed the embryonic and placental development following CsA administration^34,35. CsA intervention markedly raised embryonic survival rate from 24.3 % to 93.9%, and significantly mitigated growth retardation phenotypes relative to poly(I:C)-treated fetus (Fig. 5c, d and Supplementary Fig. 7a). Histological analysis further revealed that the thickness of the placental labyrinth and junctional layer (JZ) in poly(I:C)-treated dams was restored upon CsA treatment (Supplementary Fig. 7b–e). Furthermore, immunostaining for markers of the labyrinth zone cells demonstrated restoration of multinucleated STBs (MCT1+ SynTI and MCT4+ SynTII) and reconstruction of Laminin+ endothelial cells lining the fetal capillaries in the CsA group (Fig. 5e-h). Therefore, we ascertain that, beyond direct antiviral effects, blunted overactivation of IFN-I responses also underlies the preventive effects of CsA against ZIKV pathogenesis during pregnancy.

Fig. 5. CsA sustains placental barrier integrity and fetal development by limiting hyperactivation of the IFN response.

a Experimental design of the mouse model. b RT-qPCR of placental ISG expression at E12.5. PBS + DMSO (n = 3), PBS + CsA (n = 3), poly(I:C) + DMSO (n = 6), and poly(I:C) + CsA (n = 6). c Gross morphology of placenta and fetus harvested at E12.5. d The number and rate of fetal resorption. e, f Representative immunostaining of laminin (e) and its quantification (f) in E12.5 placentas. n = 6 placentas from 3 dams. g, h Representative images for MCT1/4 (g) and quantification of MCT1/4 positive area (h) in placentas. n = 6 placentas from 3 dams. i Workflow for evaluating CsA effects in hTSCs. j, k Western blot (j) and RT-qPCR (k) of ISG induction in hTSCs. l Schematic for assessing CsA effects on syncytiotrophoblast (STB) differentiation. m, n RT-qPCR showing hTSC and STB markers upon poly(I:C) (m) and ZIKV challenge (n) with or without CsA. n = 3. Data are presented as mean values ± SEM. Statistical significance was performed by Two-way ANOVA followed by multiple comparisons test (b, f, h). Source data are provided as a Source Data file.

To circumvent potentially confounding influences from immune cells at the maternal-fetal interface in vivo and specifically delineate autonomous IFN responses to CsA treatment in human trophoblasts, we treated hTSCs with CsA in the presence of poly(I:C) and observed markedly impaired ISG augmentation triggered by poly(I:C), compared to the DMSO controls (Fig. 5i–k and Supplementary Fig. 7f). Of note, in the absence of poly(I:C) stimulation, ISG levels in CsA-treated mouse placentas and human trophoblasts were comparable to those in DMSO-treated controls (Fig. 5b, k). These data suggest that CsA effectively restrains aberrant IFN responses driven by viruses while sustaining placenta-intrinsic antiviral defenses. To further substantiate the protective role of CsA in placental development observed in mice, we evaluated the differentiation potential of hTSCs upon CsA treatment under virus challenges (Fig. 5l). Remarkably, CsA-treated hTSCs were resistant to poly(I:C)- and ZIKV-driven inhibition of trophoblast fusion, as displayed by robust increased expression of STB markers (ERVW-1 and CGB) in parallel with decreased levels of hTSC markers (CDH1 and TP63) (Fig. 5m, n and Supplementary Fig. 7g, h). Collectively, our findings indicate that CsA administration is beneficial for preserving trophoblast function and placental barrier integrity by modulating toxic IFN responses under infectious conditions.

CsA represses virus-instigated IFN-I signaling hyperactivation via targeting the JAK1-STAT1/2 pathway

To determine whether CsA restricts IFN-I response via its canonical receptor CypA, we stimulated CypA-depleted trophoblasts with poly(I:C) and found that ablation of CypA had no discernible impact on the magnitude of ISG induction compared with controls. On the contrary, CsA retained full potency in suppressing dsRNA‑elicited ISG expression in CypA-KO trophoblasts (Fig. 6a–c and Supplementary Fig. 8a, b). To confirm the CypA independence, we treated cells with two alternative CypA inhibitors (TMN355 and NIM811). Both compounds failed to impair IFN responses triggered by dsRNA (Fig. 6d, e and Supplementary Fig. 8c), suggesting that the immunosuppressive action we observe is exclusive to CsA and does not arise from CypA blockade per se.

Fig. 6. CsA attenuates IFN-I signaling overactivation by inhibiting the JAK1-STAT1/2 pathway.

a, b Representative immunoblots (a) and quantification (b) of interferon-stimulated genes (ISG) expression in hTSCs. n = 3. c RT-qPCR showing ISG expression in hTSCs under the indicated conditions. n = 3. d, e Immunoblots (d) and quantification (e) of ISG expression in hTSCs after poly(I:C) treatment with or without NIM811 and TMN355. n = 3. f Schematic of IFN-I responses during viral infection. g, h Time-course immunoblots (g) of TBK1/IRF3 phosphorylation in hTSCs and corresponding protein quantification (h). n = 3. i–k Western blot (i, j) and RT-qPCR (k) of ISG induction in hTSCs following poly(dA:dT) transfection. n = 3. l, m Time-resolved WB showing the JAK1 and STAT1/2 phosphorylation (l) and quantification (m). n = 3. n, o Immunofluorescence imaging of STAT2 subcellular localization 1 h post IFN-β treatment and quantification of nuclear STAT2 intensity (o, 15 fields). p,q Western blot (p) and RT-qPCR (q) analysis of ISG levels upon indicated treatment. Data are presented as mean values ± SEM. n represents biologically independent samples. Statistical significance was performed by unpaired two-tailed Student’s t test (m) or Two-way ANOVA followed by multiple comparisons test (b, c, e, h, j, o). Source data are provided as a Source Data file.

Poly(I:C) triggers ISG expression through two sequential stages: an initial TBK1/IRF3 and nuclear factor κB (NF-κB)-dependent wave leading to IFN-I and cytokine production, followed by a second phase where IFN-I drives further ISG expression through the JAK-STAT1/2 pathway (Fig. 6f)^36,37. To pinpoint the potential targets of CsA, we first monitored the early events upon dsRNA recognition. WB and IF assays showed that CsA treatment neither abrogated phosphorylation of TBK1/IRF3 nor counteracted nuclear translocation of NF-κB invoked by dsRNA (Fig. 6g, h and Supplementary Fig. 8d–f). Concordantly, the transcriptional upregulation of IFN-I (IFNA1 and IFNB1) and pro-inflammatory cytokines (IL1B and IL6) was equivalent between DMSO- and CsA-treated hTSCs in response to dsRNA (Supplementary Fig. 8g, h). Therefore, our data suggest that CsA desensitized virus-mediated ISG induction independently of initial pathogen sensing and subsequent IFN-I production. In support of this, CsA also potently curtailed the expression of ISGs elicited by dsDNA mimic poly(dA:dT) without altering TBK1/IRF3 and NF-κB pathway transduction (Fig. 6i–k and Supplementary Fig. 8i–k).

We proceeded to investigate whether CsA impeded ISG expression by interfering with the downstream of IFN-I. Robust activations of JAK1 and STAT1/ STAT2 occurred upon IFN-β treatment for 2 and 4 hours, whereas CsA substantially weakened JAK1 and STAT1/2 phosphorylation, as well as STAT2 nuclear localization (Fig. 6l-o and Supplementary Fig. 8l). This disruption led to a dose-dependent reduction of key ISG expression (e.g., total STAT1/2, IFITM2/3, OAS1) in both JEG-3 cells and hTSCs (Fig. 6p, q and Supplementary Fig. 8m, n). To assess whether the well-known immunosuppressive target of CsA, NFAT signaling, also contributes to the inhibition of IFN-β responses, we treated cells with a selective NFAT inhibitor during IFN-β administration. The pharmacological blockade of NFAT had no impact on IFN-β‑induced pathogenic ISG augmentation (IFITMs and GBP5) in either trophoblast type (Fig. 6p, q and Supplementary Fig. 8m, n), demonstrating that the suppression of IFN-I signaling operates independently of the canonical NFAT horizon. Collectively, these results reveal that CsA tempers virus-triggered IFN-I signaling, at least in part by targeting the JAK1-STAT1/2 signaling.

CsA counteracts interferonopathies driven by the undue accumulation of immunogenic dsRNA in placentas

Unrestricted or prolonged IFN-I responses driven by viral infection or genetic mutations underpin a spectrum of special autoinflammatory diseases with high morbidity and mortality termed interferonopathies^38,39. In the setting of pregnancy, aberrant IFN-I signaling, as seen in conditions such as Aicardi-Goutieres Syndrome (AGS) and systemic lupus erythematosus, frequently leads to adverse fetal outcomes that closely resemble congenital infections^40,41. Indeed, our recent work demonstrated that placenta-specific deletion of Adar1 (hereafter Adar1^PKO), an essential A-to-I RNA editing enzyme implicated in AGS, confers the accumulation of self-derived dsRNA and anomalous IFN activation, culminating in pregnancy-context interferonopathies such as placental insufficiency, fetal growth restriction, and even embryonic lethality in mice⁴². The observation that CsA suppressed IFN-I signaling hyperactivation prompted us to investigate whether CsA could also alleviate endogenous dsRNA-mediated IFN responses and resultant pregnant complications. To this end, we administered daily injections of DMSO or CsA to pregnant Adar1^WT and Adar1^PKO dams from E10.5 to E14.5 (Fig. 7a). As anticipated, CsA treatment significantly suppressed the expression of representative ISGs (Ifitm3, Isg15, and Oas1a) in Adar1^PKO placentas, reinforcing its potent immunosuppressive role in trophoblast-derived innate immune responses (Fig. 7b). In parallel, CsA administration effectively improved fetal weight, gross appearance, placental efficiency, and yolk sac development compared to DMSO-treated Adar1^PKO controls (Fig. 7c–e). Additionally, key placental features sensitive to Adar1 depletion, including junctional zone thickness, Tpbpa expression or positive area, glycogen storage (PAS staining), and the abundance of placenta-specific glycoproteins (Psg17, Psg19, and Ceacam3), were prominently rescued in the Adar1^PKO placenta by CsA treatment (Fig. 7f-j). Therefore, our findings underscore CsA treatment as a promising therapeutic strategy to ameliorate pregnancy complications linked to interferonopathies.

Fig. 7. CsA alleviates pregnancy-associated adverse outcomes induced by placental endogenous dsRNA overload.

a Schematic of CsA treatment in placenta-specific Adar1 knockout (Adar1^PKO) mice. b RT-qPCR of placental ISG expression at E14.5. Adar1^WT + DMSO (n = 3), Adar1^WT + CsA (n = 3), Adar1^PKO + DMSO (n = 6), and Adar1^PKO + CsA (n = 6). c Representative morphology of the yolk sac, placenta, and fetus at E14.5. d, e Assessments of fetal weight (d) and embryo/placenta weight ratio (e). Adar1^WT + DMSO (n = 14), Adar1^WT + CsA (n = 15), Adar1^PKO + DMSO (n = 6), and Adar1^PKO + CsA (n = 6). f Representative H&E staining, in situ hybridization of junctional zone marker Tpbpa, and PAS staining of glycogen‑rich trophoblast layers of E14.5 placentas. g, h JZ thickness (g) and JZ/total placental area ratio (h) at E14.5. n = 3 from 3 dams. i Quantification of Tpbpa positive area. n = 6 placentas from 3 dams. j RT-qPCR of pregnancy-specific secreted glycoprotein genes in E14.5 placentas. Adar1^WT + DMSO (n = 3), Adar1^WT + CsA (n = 3), Adar1^PKO + DMSO (n = 5), and Adar1^PKO + CsA (n = 5). Data are presented as mean values ± SEM. Statistical significance was performed by Two-way ANOVA followed by multiple comparisons test (b, d, e, g–j). Source data are provided as a Source Data file.

Discussion

The ability of ZIKV to destroy and traverse the placental barrier is central to its teratogenic potential, yet the host determinants and therapeutics governing these pregnancy-specific pathologies remain incompletely identified. Our study demonstrates CypA as an essential factor that orchestrates viral replication within trophoblasts and is indispensable for efficient in utero dissemination. Mechanistic investigations reveal ZIKV infection provokes the relocalization of CypA into viral replication compartments and reconfigures its interaction network, disrupting host RNA decay and SG formation. Therapeutically, the pharmacological inhibitor CsA confers dual protective effects against ZIKV-driven pathological phenotypes during pregnancy by suppressing viral replication and simultaneously antagonizing maladaptive placental IFN-I responses.

Cell-intrinsic antiviral mechanisms are fundamental obstacles for invading viruses, yet ZIKV has evolved a multifaceted arsenal to dismantle these defences^1,14. For instance, ZIKV nonstructural proteins (such as NS2B3, NS4A, NS4B, and NS5) interfere with the phosphorylation and stability of human STAT1/STAT2, thereby attenuating IFN signaling and diminishing antiviral ISG expression^43,44. Concurrently, ZIKV hijacks autophagy, a pathway normally dedicated to pathogen degradation, to impede inflammatory antiviral responses, promote endosomal viral RNA release, and furnish necessary resources for viral replication^45–48. Our findings further reveal that CypA is actively recruited to ZIKV replication sites, where it interacts with multiple viral nonstructural proteins and viral RNA. This relocalization likely disrupts its association with innate immune effectors. Given the important role of CypA in modulating RIG-I–mediated antiviral signaling, the reprogrammed CypA interactome may represent an additional layer of immune evasion employed by ZIKV²⁶. In parallel, although growing evidence indicates that ZIKV is refractory to host-mediated viral RNA degradation by promoting subgenomic vRNA formation and SG disassembly, the molecular underpinnings of these phenomena are largely unknown^28,49. Our study suggests that CypA plays a critical role in ZIKV-driven RNA decay resistance and SG disintegration, which may foster a less hostile intracellular milieu for ZIKV replication and propagation. Above all, we speculate that CypA emerges as a dual-function host factor in ZIKV biology, both acting as a structural scaffold within the replication complex and manipulating host antiviral mechanisms.

Here, we observed that CypA inhibition restricted ZIKV infection in placental trophoblasts, whereas no reduction in viral titers was detected in several maternal tissues examined. This tissue-restricted phenotype underscores the context-dependency of host factor involvement in viral infections. Indeed, recent high-resolution CRISPR screens in ZIKV, CMV, and SARS-CoV-2 infection have shown that host factor requirements can vary markedly across cell types or infection stages^10,50,51. This likely arises from cell-type-specific expression patterns, differential availability of essential cofactors, or distinct local immune environments. Further investigation into the role of CypA in ZIKV infection across distinct tissue contexts will be essential to inform the tissue-selective application of host-directed strategies while minimizing potential off-target effects.

Despite expanding mechanistic insights into ZIKV pathogenesis, safe and effective therapeutic interventions for pregnant individuals remain extremely scarce. Experimental antiviral strategies, including nucleoside analogues (e.g., sofosbuvir), monoclonal antibodies, vaccines, and phytochemicals, have shown encouraging efficacy in preclinical models^29,52–55. However, concerns regarding fetal teratogenicity and undefined placental transfer profiles have hindered their clinical translation during pregnancy. By contrast, repurposing drugs with established clinical use offers a pragmatic pathway to accelerate clinical application by leveraging known pharmacokinetics, dosing regimens, and pre‑existing safety databases for pregnant populations. Here, we report that CsA, an FDA‑approved medication widely deployed to prevent graft rejection, potently abrogates ZIKV placental replication and vertical transmission. Notably, CsA has already been applied in pregnancy management under certain circumstances, improving live-birth rates in women with unexplained recurrent miscarriage or autoimmune disorders^56–60. Pharmacokinetic data further indicate that CsA levels in umbilical-cord blood at delivery reach only ~6% of maternal blood concentrations, suggesting limited fetal exposure⁶¹. Importantly, the CsA dose employed in this study (10 mg/kg in mice, equivalent to 50 mg for a 60 kg individual) is substantially lower than the standard clinical dose (150-200 mg/day), underscoring its favourable safety margin⁶². It is worth mentioning that previous studies have also demonstrated CsA’s capacity to impair the infectivity of other positive-sense RNA viruses, including HCV and human immunodeficiency virus type 1, across diverse models^63–65. Together, these observations highlight CsA as a promising candidate for repurposing against congenital viral infections. Nonetheless, CsA is a potent inhibitor of multidrug transporters, particularly P-glycoprotein, and high-dose regimens have been associated with adverse outcomes such as impaired implantation, increased prematurity, and long-term nephron deficits^66–69. These risks underscore the need for rigorous dose optimization and vigilant monitoring during any potential therapeutic application of CsA in pregnancy, particularly to avoid adverse drug-drug interaction.

In this study, we reveal that, apart from the direct antiviral effects via CypA inhibition, CsA simultaneously preserves placental barrier integrity by dampening hyperactivated IFN-I response and ensuing toxic ISG induction triggered by viral dsRNA, thereby mitigating immune-mediated placental damage and pregnancy complications. Mounting evidence frames the placental IFN response as a physiological double-edged sword. While essential for viral restriction, their dysregulation drives adverse pregnancy outcomes, such as IUGR, miscarriage and preterm birth^{1,5,6,14,34,70,71}. Thus, the immunomodulatory property delineated here uniquely qualifies CsA as a promising candidate for the management of pregnancy-associated viral infections. Moreover, related arboviruses, such as Chikungunya virus and West Nile virus, could also contribute to fetal pathologies by eliciting potent placental innate immune activation in mice despite inefficient transplacental transmission^72–74. Accordingly, by demonstrating that CsA efficiently counteracted poly(I:C)-induced placental insufficiency and fetal demise, our viral mimicry experiments highlight its potential as a broad-spectrum strategy to combat IFN-driven gestational complications across diverse infectious contexts.

Previous studies have primarily characterized CsA as an immunosuppressant that acts through inhibition of NFAT signaling and inflammatory responses in immune cells - the mechanisms that underpin its clinical use in organ transplantation^75,76. However, emerging evidence reveals that CsA exerts a much broader spectrum of biological activities. For example, CsA has been shown to modulate Wnt signaling in human hair follicles and inhibit the CypA-independent mPTP, underscoring its pleiotropic molecular targets and pharmacological diversity^21,77. Here, we broadened the immunomodulatory repertoire of CsA: suppression of the JAK1-STAT1/2 signaling cascade in response to viral nucleic acid (both RNA and DNA) and IFN-β. Inspired by this action, we showed that CsA ameliorated placental and fetal malformation driven by overzealous activation of IFN-I pathways in models of both endogenous and exogenous dsRNA stress. As poly(I:C) stimulation or Adar1 deficiency also recapitulate features of certain non-infectious pregnancy complications, CsA-mediated blockage of IFN-I responses may hold translational potential well beyond infectious settings^35,41,78. Future studies should elucidate how CsA intersects with kinase activation and STAT phosphorylation, as well as determine whether additional signaling pathways or co-regulatory factors contribute to ISG repression, to refine targeted anti-IFN strategies during pregnancy.

Altogether, our study comprehensively defines the subversion of host CypA as the pivotal mechanism facilitating ZIKV invasion across the maternal-fetal barrier, and demonstrates that repurposed CsA grants both robust antiviral efficacy and placental immune modulation to safeguard pregnancies from virus-associated pathologies. These insights lay a mechanistic and therapeutic foundation for the rational design of pregnancy-tailored antiviral strategies against emerging viral threats.

Methods

Cells

Vero (ATCC, CRL-1586) and HEK293T (ATCC, CRL-11268) cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, #C11965500BT). JEG-3 cells (ATCC, HTB-36) were cultured in DMEM: Nutrient Mixture F-12 (Gibco, #C11330500BT). The culture media were supplemented with 10% (v/v) heat-inactivated fetal bovine serum (FBS) (Gibco, #2036224 C) and 1% Penicillin-Streptomycin (PS, Thermo Fisher Scientific, #15140122). hTSCs were derived from a male first-trimester placenta and cultured according to methods detailed in our prior studies^10,79,80. Briefly, villous tissues were digested sequentially with 0.25% trypsin (30 min) followed by 10 U/ml DNase I (15 min). The resulting single-cell suspension was then isolated and plated in 5 μg/mL collagen Type IV (RD, #3410-010-01)-coated plates and maintained in a defined trophoblast stem cell culture medium composed of Advanced DMEM/F12 (Gibco, #12634-010), 0.2% FBS, 0.5% PS, 0.1 mM β-mercaptoethanol (Thermo Fisher Scientific, #21985023), 0.3% BSA (Sigma-Aldrich, #A9418), 1.5 μg/mL L-ascorbic acid (Wako, #013-12061), 1% ITS-X supplement (Thermo Fisher Scientific, #41400045), 50 ng/mL EGF (MedChemExpress, #HY-P7109), 0.5 μM A83-01 (MedChemExpress, #HY-10432), 2 μM CHIR99021 (Selleck, #S2924), 1 μM SB431542 (Wako, #031-24291), 0.8 mM VPA (Wako, #227-01071) and 5 μM Y27632 (Selleck, #S1049) following our previous protocol^10,79. All cells were cultured in a 5% CO₂ incubator at 37 °C. The medium was replaced every 48 hours. Passaging was performed using TrypLE Express when cells reached ~80% confluency. All procedures involving the isolation and utilization of hTSCs were approved by the Ethics Committees of Women and Children’s Hospital at the School of Medicine of Xiamen University (IRB: KY-2022-076-h01). Written informed consent was obtained from each donor before sample collection and cell isolation.

For human placental organoids (hPOs), hTSCs in the exponential growth phase were harvested and resuspended in Matrigel at a density of 5000 cells/30 μL. The Matrigel drops were plated into pre-warmed culture plates and solidified at 37 °C. hTO medium was prepared as previously described^10,79,81: Advanced DMEM/F12, N2 supplement (Gibco, #17502048), B27 supplement (Gibco, #12587010), 1% PS, 1.25 mM N-acetyl-L cysteine (Sigma-Aldrich, #616-91-1), 2 mM L-glutamine (Sigma-Aldrich, 25030024), 50 ng/mL recombinant human EGF, 1.5 μM CHIR99021, 80 ng/mL recombinant human R-spondin-1 (MedChemExpress, #HY-P7114), 100 ng/mL recombinant human FGF2 (Peprotech, #100-18 C), 50 ng/mL recombinant human HGF (Peprotech, #100-39-10), 500 nM A83-01, 2.5 μM prostaglandin E2 (MedChemExpress, #HY-101952), and 2 μM Y-27632. Organoids were cultured at 37 °C with 5% CO₂, and the medium was refreshed every 2-3 days. This method yields organoids with cytotrophoblasts on the surface and interior STBs⁸⁰. The size of hTOs with indicated treatment was monitored under brightfield microscopy using Leica v4.12 Cell3 Imaging System. Organoid growth rates were quantified by dividing the measured size at 4 days post-infection (dpi) by the initial size at 0 dpi. hTOs were fixed with 4% paraformaldehyde (PFA) for histomorphological assay.

STBs differentiation

hTSCs were seeded on 2.5 mg/mL Collagen IV-coated dishes and exposed to poly(I:C) (20 ng) (InvivoGen, #tlrl-pic) or ZIKV (MOI = 0.1) in the presence or absence of 2 µM CsA (MedChemExpress, #HY-B0579) for 24 h in maintenance medium. The medium was switched to the STB differentiation medium, consisting of Advanced DMEM/F12, 0.5% PS, 0.1 mM 2-mercaptoethanol, 0.3% BSA, 1% ITS-X supplement, 2.5 μM Y27632, 2 μM Forskolin, and 4% KSR and proceeded for 48 h.

Viruses

ZIKV strains (IbH30656 and PRVABC59) were purchased from ATCC (Manassas, VA, USA). The virus stocks were generated in Vero cells and titrated via plaque assay on Vero cells. All virus manipulations were performed in a BSL2 facility according to the corresponding biosafety protocols.

Mice

The mouse experiments were approved by the Xiamen University Ethics Committee (XMULAC20200093) and conducted following the guidelines of the Xiamen University Laboratory Animal Centre. All mice utilized in this study were on a C57BL/6 J background. Specifically, Ifnar1^-/- mice were generously provided by Dr. Jiahuai Han at Xiamen University. Ppia^-/- (Strain number: T017425) mice were purchased from GemPharmatech. Female mice, aged 10-12 weeks, were randomly assigned to experimental groups and housed in pathogen-free conditions at the Xiamen University Laboratory Animal Center (23 °C, a 12/12 h light/dark cycle with free access to food and water). For timed mating, females were paired with males overnight, and the embryonic day 0.5 (E0.5) was designated upon detection of a vaginal plug the following morning. For drug administration, pregnant Ifnar1^−/− mice mated with WT sires received intraperitoneal injections of either CsA (10 mg/kg) or vehicle control on E6.5. Two days after treatment, dams were anaesthetized and inoculated via footpad injection with 10³ Plaque Forming Units (PFU) of ZIKV or PBS as a mock control. To suppress type I IFN response in Ppia^+/- mice, dams were subjected to an intraperitoneal injection of 2 mg monoclonal anti-mouse IFNAR-1 blocking antibody (MAR1-5A3, Leinco, #I-401-100) one day before infection. To mimic viral infection, poly(I:C) (200 μg in 100 μL of sterile PBS) was delivered intraperitoneally at E10.5. In the placenta-specific Adar1 KO mouse model, Adar1^PKO or Adar1^WT dams received daily intraperitoneal injections of either DMSO or CsA from E10.5 through E14.5. All maternal and fetal tissues were harvested at the indicated time points for subsequent analysis.

Plaque Assay

Vero cells were seeded in 24-well plates at a density of 10⁵ cells/well. Once confluent, the cells were infected with serial dilutions of supernatants collected from infected cultures and incubated for 2 h at 37 °C. Subsequently, the viral inoculum was removed, and a 1 % methyl cellulose overlay (Sigma-Aldrich, #M0512) was applied to the cells. After 5 days of culture, cells were fixed and stained with crystal violet solution (2 % crystal violet dissolved in 40 % formaldehyde and 60 % anhydrous ethanol). The virus titer, expressed as PFU per milliliter (PFU/mL), was calculated by counting the number of plaques and multiplying by the corresponding dilution factor.

Molecular cloning and overexpression

Expression plasmids encoding human CypA and CypA-R55A were generated by RT-PCR amplification from hTSCs cDNA. Plasmids expressing individual ZIKV NS proteins were PCR-amplified from the ZIKV strain IbH30656. All indicated proteins were cloned into the pLVX plasmid and verified by Sanger sequencing. Reconstitution of CypA or CypA-R55A into CypA KO cells was performed via lentivirus-mediated transduction. JEG-3 cells or hTSCs were infected with lentiviral supernatant for 48 h and selected with puromycin for at least 7 days. The overexpression efficiency was assessed by Western blotting. Primer sequences used for cloning are listed in Supplementary Table 1. To generate PPIA shRNA plasmids, oligonucleotides targeting PPIA were cloned into the pLKO.1 plasmid (Addgene, #15257) and subsequently packaged into lentiviruses in HEK293T cells. Primers for shRNA targeting sequence: PPIA#1, CACAACTTCGTGGAGGTCTTA; PPIA#2, GCCAAGTGGTACATGCAGTTT.

Generation of CypA-depleted cell lines

CypA KO JEG-3 cell lines were generated using the CRISPR-Cas9 genome editing system. Oligonucleotides encoding the sgRNA sequences were synthesized, annealed, and cloned into the lentiCRISPRv2 plasmid (Addgene, #52961) following our previously published protocol¹⁰. For lentivirus production, HEK293T cells were co-transfected with the sgRNA-containing lentiCRISPRv2 plasmid, psPAX2 (Addgene, #12260), and pMD2.G (Addgene, #12259) using Lipofectamine 3000. Viral supernatants were collected 48 h post-transfection and filtered through a 0.45 µm filter. JEG-3 cells were exposed to viral supernatants supplemented with 8 µg/mL polybrene. After 48 hours, transduced cells were selected with puromycin (10 µM) for 7 days. Surviving cells were dissociated into single cells and seeded at low density in 96-well plates to allow the growth of single-cell-derived clones. Two independent CypA KO lines were confirmed by Sanger sequencing and western blot analysis.

Stress granule induction

SGs were induced in the indicated trophoblasts by treatment with sodium arsenite (Ars). Briefly, Cells were incubated with 0.5 mM (Sigma-Aldrich, #S7400) Ars for 30 min and then cultured in fresh medium without Ars for an additional 30 min. Following stress induction, cells were fixed with 4% PFA for 15 min at room temperature, then permeabilized using 0.1% Triton X-100 in PBS. SGs were visualized by immunostaining with primary antibodies against G3BP1 (Proteintech, #13057-2-AP, 1:500). Imaging was performed using a confocal microscope (Zeiss LSM 880) with a 63× oil immersion objective. The number and size of SGs in cells were calculated using ImageJ software with manual threshold adjustments.

ZIKV infection assays

To assess ZIKV binding, trophoblasts were washed twice with cold PBS and incubated with ZIKV (MOI = 10) at 4 °C for 1 h. The unbound virus was removed by washing three times with cold PBS. For viral entry analysis, cells were challenged with ZIKV (MOI = 10) at 37 °C for 1 h. Following incubation, uninternalized virus was removed by treating cells with trypsin (Sigma-Aldrich, #T9935) for 2 min at 37 °C. To evaluate ZIKV replication kinetics, trophoblasts were infected with ZIKV (MOI = 0.1) at 37 °C for 2 h, followed by three washes with PBS. The fresh medium was added to the cells, and supernatants and cell lysates were collected at the indicated time points post-infection. Bound and intracellular viral RNA was extracted using the RNAprep Pure Cell Kit (Qiagen, #74134), and viral RNA in supernatants were isolated using the QIAamp Viral RNA Mini Kit (Qiagen, #52906) following the manufacturer’s instructions.

Infectivity inhibition assay

To test the anti-ZIKV effect of individual drugs, JEG-3 cells were seeded at a density of 10⁵ cells/well in 12-well plates. Cells were infected with the IbH30656 strain of ZIKV at an MOI of 0.1 in combination with the indicated compounds at various concentrations. Following 48 h of co-treatment, cell lysates were collected and subjected to one-step RT-qPCR. The half-maximal inhibitory concentration values (IC₅₀) were calculated by using the log (inhibitor) versus normalized control (DMSO)-variable slope analysis.

Cytotoxicity assay

JEG-3 cells or hTSCs were seeded in 96-well plates at a density of 4 × 10³ cells/well. After 24 h, cells were incubated with indicated concentrations of individual drugs (CsA, NFAT inhibitor (MedChemExpress, #HY-P1026), FK506 (MedChemExpress, #HY-13756), TMN355 (MedChemExpress, # HY-107635), NIM811 (MedChemExpress, #HY-P0025)), BAPTA‑AM (MedChemExpress, #HY-100545), and mPTP-IN-1 (MedChemExpress, #HY-172959) for 48 h. The half-maximal cytotoxic concentration value (CC₅₀) was determined using the crystal violet assay, expressed as a percentage of the control.

ZIKV replicon assay

Trophoblasts were seeded in a 12-well plate at a density of 10⁵ cells per well. ZIKV replicon was transduced into cells using Lipofectamine 3000 according to the manufacturer’s instructions. At 24, 48, or 72 hours post-transfection, cells were washed with PBS and lysed in 200 µL of luciferase-compatible lysis buffer (1% Triton X-100, 25 mM glycyl-Glycine (pH 7.8), 15 mM MgSO₄, 4 mM EGTA, 1 mM DTT). The luciferase substrate was added to lysates for luminescence measurement. Data were normalized to total protein concentration and expressed as fold changes relative to controls.

Flow cytometry analysis

JEG-3 cells were exposed to ZIKV for 12, 24, and 48 h. Following infection, cells were detached using trypsin and fixed with BD CytoFix/CytoPerm solution for 30 min at 4 °C. The cells were then washed twice with PBS and blocked in PBS containing 1% BSA and 0.5% Triton X-100 for 1 h at room temperature. Intracellular viral antigen was detected by incubation with the anti-ZIKV E antibody (GeneTex, #GTX133314, 1:2000) for 1 h at 4 °C, followed by the secondary antibody staining for 1 h at 4 °C in the dark. Flow cytometry was carried out using a CytoFLEX S Flow Cytometer (Beckman Coulter), and data analysis was performed using FlowJo software.

Real-time Quantitative PCR (RT-qPCR)

Total RNA was extracted from cultured cells or tissue samples using TRIzol (Thermo Fisher Scientific, #15596018) or RNAprep Pure Cell Kit (Qiagen, #74134) according to the manufacturer’s instructions. Reverse transcription was performed using PrimeScript™ RT reagent Kit (Takara, #RR047A) following the manufacturer’s protocol. qPCR was carried out using SYBR Green Master Mix (Vazyme, #Q712-03) in a final reaction volume of 10 μL. Each sample was analyzed in duplicate using a QuantStudio5 thermocycler. The following cycling conditions were used: initial denaturation at 95 °C for 10 min, followed by 40 cycles of 95 °C for 15 seconds, and 60 °C for 1 min. Primer sequences for all target genes were listed in Supplementary Table 2. Relative mRNA expression was calculated using the ΔΔCt method and normalized to the housekeeping gene (β-actin).

Viral RNA was extracted from amniotic fluid or cell culture supernatants using the QIAamp Viral RNA Mini Kit (Qiagen, #52906) and determined using the Luna® Universal Probe One-Step RT-qPCR Kit (New England Biolabs, #E3006L) following the manufacturer’s instructions. A total of 10 μL reaction volume was prepared, consisting of 5 μL of 2×reaction mix, 0.8 μL forward and reverse primers (10 μM), 0.2 μL ZIKV probe (10 μM) and 1 μL of RNA template. ZIKV-specific primers and probes: Forward, 5′-CCGCTGCCCAACACAAG-3′; Reverse, 5′-CCACTAACGTTCTTTTGCAGACAT-3′; Probes, 5′-/FAM/TGACAAGCAATCAGACACTCAA/TAM-3′. Viral RNA levels were expressed as copies per unit (g for tissues, μg RNA for cells, mL for supernatants or amniotic fluid).

RNA-seq analysis

Total RNA was extracted from mouse placentas using RNAprep Pure Cell Kit according to the manufacturer’s instruction. High-depth transcriptomic sequencing was conducted on the BGISEQ-500 platform (China, BGI). The raw sequencing reads are aligned to the mouse reference genome (GRCm39) using HISAT2 alignment tools. Gene expression levels are quantified with featureCounts. Differential gene expression analysis was performed using DESeq2 (v1.32.0), with a significance threshold set at pvalue < 0.05 and |fold change | > 1.5. Bioinformatic analyses and data visualization, including volcano plots, GO enrichment, heatmaps, and GSEA enrichment, were performed using the OmicStudio tools at https://www.omicstudio.cn/tool.

RNA immunoprecipitation (RIP)-qPCR

hTSCs overexpressing CypA were infected with ZIKV (MOI = 1) for 24 h. The cells were resuspended in lysis buffer containing protease and RNase inhibitors. The lysates were incubated on ice for 30 min and centrifuged at 12,000 g for 10 min to remove cellular debris. The resulting supernatant was incubated with Anti-FLAG® Magnetic Beads (Sigma-Aldrich, #M8823) at 4 °C overnight with gentle rotation. The bead-bound complexes were washed three times with cold lysis buffer to eliminate non-specific binding. RNA was extracted from the immunoprecipitated protein-RNA complexes using TRIzol reagent. Viral RNA enrichment was quantified by one-step RT-qPCR as described above.

Western blotting

Cells were lysed in RIPA buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, and 0.1% SDS) supplemented with protease and phosphatase inhibitors. Lysates were incubated on ice for 30 min and clarified by centrifugation at 12,000 g for 30 min at 4 °C. Protein concentrations were determined using the BCA Protein Assay Kit (Thermo Fisher, #23225). Equal amounts of protein (20 μg) were denatured in 1×Laemmli sample buffer and resolved by SDS-PAGE on 10 % polyacrylamide gels. Proteins were then transferred to a PVDF membrane (Bio-Rad, #1620177) by semi-dry transfer at 15 V for 30 min or wet transfer at 100 V for 90 min at 4 °C. Membranes were blocked with 5% non-fat dry milk (BD Bioscience, #232100) in TBST, followed by overnight incubation with the indicated primary antibodies (anti-β-ACTIN (ABclonal, # AC026, 1:20000), anti-CypA (Proteintech, #10720-1-AP, 1:3000; Aladdin, #Ab098507, 1:1000), anti-G3BP1 (Proteintech, #13057-2-AP, 1:1000), anti-NS3 (GeneTex, #GTX133309, 1:1000), anti-NS5 (GeneTex, #GTX133312, 1:1000), anti-ZIKV-E (GeneTex, #GTX133314, 1:2000), anti-Flavivirus (GeneTex, #GTX57154, 1:500), anti-Flag (Sigma-Aldrich, #F1804, 1:1000), anti-HA (Cell Signaling Technology, #3724 T, 1:1000), anti-E-Cadherin/CDH1 (Cell Signaling Technology, #3195 T, 1:1000), anti-Syndecan-1 (Abcam, #ab34164, 1:1000), anti-CGB/HCG (Abcepta, # AP13036B, 1:1000), anti-OAS1 (Abcam, #284864, 1:2000), anti-IFITM2 (Thermo Fisher Scientific, #MA5-27503, 1:1000), anti-IFITM3 (ZENBIO, #382077, 1:1000), anti-TBK1 (Cell Signaling Technology, #38066S, 1:1000), anti-p-TBK1 (Cell Signaling Technology, #5483 T, 1:1000), anti-IRF3 (ABclonal, #A11118, 1:1000), anti-p-IRF3 (Cell Signaling Technology, #29047S, 1:1000), anti-JAK1 (Cell Signaling Technology, #3344 T, 1:1000), anti-p-JAK1 (Cell Signaling Technology, #74129 T, 1:1000), anti-Stat1 (Cell Signaling Technology, #14994 T, 1:1000), anti-p-Stat1 (Cell Signaling Technology, #7649 T, 1:1000), anti-Stat2 (Cell Signaling Technology, #72604 T, 1:1000), anti-p-Stat2 (Cell Signaling Technology, #4441 T, 1:1000), anti-GAPDH (Cell Signaling Technology, #2118S, 1:20000), anti-ADAR1 (Cell Signaling Technology, #81284 T, 1:1000) at 4 °C. After washing, membranes were incubated with HRP-conjugated secondary antibodies (1: 1000) for 1 h at room temperature. Protein bands were detected using enhanced chemiluminescence (ECL, Bio-Rad, #1705060), and images were captured using an e-Blot Touch Imager. Band intensities were quantified using ImageJ software (National Institutes of Health, Bethesda, MD, USA), and the signal for each target protein was normalized to its corresponding loading control. Uncropped and unprocessed scans of all blots are provided in the Source Data.

ChIRP-WB

JEG-3 cells were infected with either ZIKV (Nigerian strain, MOI = 0.1) or mock-treated with PBS. At 48 hpi, cells were detached with trypsin, collected by centrifugation at 1400 g for 5 min, washed twice with PBS, and resuspended in 3% (v/v) formaldehyde. Crosslinking was carried out for 30 min at 25 °C with gentle rocking, then quenched by adding glycine to a final concentration of 125 mM for 5 min at 25 °C. Crosslinked cells were pelleted at 2000 g for 5 min and frozen at −80 °C for storage. For lysate preparation, ~100 mg of cell pellet was thawed on ice and suspended in 1 mL lysis buffer (50 mM Tris‑HCl, pH 7.0, 10 mM EDTA, 1 % SDS). Samples were sonicated on a focused‑ultrasonicator (DIAGENODE) until the average RNA fragment size was ~500 nt as verified by agarose gel electrophoresis. An aliquot corresponding to 1% of each lysate was reserved as “input.” The remaining lysate was processed for ChIRP according to Ooi et al., including pre‑clearing, hybridization with biotinylated probes, capture on streptavidin beads, and sequential high‑stringency washes⁸². The enriched proteins on the beads were eluted using RIPA buffer (25 mM Tris-HCl, pH 7.4, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) for further WB analysis.

Immunofluorescence staining of cells

Cells were fixed with 4% PFA at room temperature for 10 min, followed by blocking with a solution containing 10% BSA and 0.2% Triton X-100 for 1 h. Cells were then incubated with the indicated primary antibody (anti-NF-κB, Abmart, #T55034F, 1:500; anti-STAT2, Cell Signaling Technology, #72604 T, 1:500) overnight at 4 °C. After three washes with PBST, cells were incubated at room temperature for 1 h with AlexaFluor-conjugated secondary antibodies and counterstained with 4′,6-diamidino-2-phenylindole (DAPI, Thermo Fisher Scientific, #D1306, 1:1000). Fluorescence imaging was captured using a Zeiss LSM 880 inverted Confocal Laser Scanning Microscope (ZEISS). Image processing and analysis were performed using NIS-Elements Viewer 5.21 and ImageJ software (v2.0.0).

Histopathology, Immunofluorescence, and Immunohistochemistry assays

Mouse placentas were sectioned at a thickness of 6 µm, and sections were mounted on pre-coated glass slides. For hematoxylin and eosin (H&E) staining, sections were deparaffinized in xylene and rehydrated through a descending ethanol series. Slides were stained with hematoxylin for 3–5 min, differentiated in acid alcohol and counterstained using an H&E staining kit (Solarbio, #G1120) according to the manufacturer’s protocol. For periodic acid-schiff (PAS) staining, deparaffinized sections were incubated in 0.5% periodic acid (Solarbio, # G1280) for 10 min at room temperature. After rinsing in running tap water for 5 min, slides were transferred to Schiff’s reagent for 15 min in the dark. For immunofluorescence analysis, the deparaffinized sections underwent antigen retrieval through microwaving in 10 mM sodium citrate buffer (pH 6.0) for 15 min and permeabilized with 0.3% Triton X-100 in PBS for 15 min at room temperature. Non-specific binding was blocked using 10 % BSA for 1 h at room temperature. Primary antibodies specific to the target proteins (anti-MCT1, Sigma-Aldrich, #AB1286-I, 1:500; anti-MCT4, Sigma-Aldrich, #AB3314P, 1:500, and anti-ZIKV-E, GeneTex, #GTX133314, 1:500) were diluted in a blocking buffer and incubated overnight at 4 °C. After three PBS washes, sections were incubated with fluorescently labeled secondary antibodies for 1 h at room temperature in the dark. Nuclei were counterstained with DAPI for 5 min. Slides were mounted using an anti-fade mounting medium and imaged using a confocal microscope (ZEISS). For immunohistochemical staining, retrieved slides were incubated with 3% H₂O₂ in methanol for 10 min to quench endogenous peroxidase activity, followed by blocking with 5% BSA in PBS containing 0.1% Triton X-100 for 2 h. Sections were incubated with polyclonal anti-Laminin (Sigma-Aldrich, #L9393, 1:500) overnight at 4 °C. After three PBS washes, HRP-conjugated secondary antibody (1:400) was applied for 1 hour at room temperature. Detection was performed using the DAB substrate kit (Abcam, #ab64238). Slides were counterstained with hematoxylin, dehydrated, cleared, and mounted. Images were captured using a bright-field microscope. The MCT1/4, laminin, and Tpbpa-positive areas were quantified using ImageJ/Fiji software and normalized to the number of DAPI-positive nuclei per field.

Co-immunoprecipitation

hTSCs and HEK293T cells were transduced with the indicated proteins for 48 h using the lentiviral-mediated method or Lipofectamine 3000, respectively. The cells were then infected with or without ZIKV infection for 24 h and lysed in ice-cold lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP-40, 0.1% SDS, and 1× protease and phosphatase inhibitor cocktail) for 30 min on ice. The lysates were clarified by centrifugation at 14,000 g for 15 min at 4 °C and incubated with Anti-FLAG®M2 Magnetic Beads overnight at 4 °C with gentle rotation. The immunoprecipitated protein complexes were eluted by boiling the beads in 1× Laemmli buffer for 5 min at 95 °C. Eluted proteins were subjected to WB analysis.

Mass spectrometry

CypA-overexpressed hTSCs were exposed to PBS or ZIKV (MOI = 1, n = 3 biological replicates/group) for 48 h. The protein complexes bound to CypA were enriched with Anti-FLAG® M2 Magnetic Beads and then eluted by incubation with 50 µL of elution buffer (0.1 M glycine, pH 2.5, or 8 M urea) for 5 min. Proteins were separated by 10% SDS-PAGE gels and stained with Coomassie Blue according to the manufacturer’s protocol (Thermo Fisher Scientific, #LC6065). Gel bands were sliced and digested with sequencing-grade trypsin (1:50 enzyme-to-substrate ratio) at 37 °C overnight. Samples were reduced, alkylated, and digested with trypsin in solution. The resulting peptides were dried and analyzed on an EASY-nLC 1200 (Thermo Scientific) coupled to an Orbitrap Fusion Lumos (Thermo Scientific) equipped with an EASY-IC ion source. Peptides were dissolved in 10 μL 0.1% formic acid and were auto-sampled directly onto a homemade C18 column (35 cm × 75 μm i.d., 2.5μm 100 Å). Samples were then eluted for 120 min with linear gradients of 3–35% acetonitrile in 0.1% formic acid at a flow rate of 300 nL/min. The raw files were analyzed by Proteome Discoverer 2.5 software against the UniProtKB human database. Statistical analysis was carried out using Perseus (version 2.1.0.0) and R (version 4.3.1). Differential abundance was assessed by a two‑tailed t‑test with Benjamini‑Hochberg FDR correction.

Statistical analysis

Statistical analysis and data visualization were conducted using GraphPad version 9 (La Jolla, CA, USA). Data from at least three independent experiments are presented as the mean ± standard error of the mean (SEM), with individual data points displayed as dots. Normality and homogeneity of variance were assessed using the Shapiro-Wilk and Brown-Forsythe, respectively. Parametric comparisons were performed using unpaired two-tailed Student’s t-test (two groups) or one-way ANOVA with Tukey’s post hoc test (multiple groups). Non-parametric comparisons were performed using the Mann-Whitney U test (two groups) or the Kruskal-Wallis test with Dunn’s correction (multiple groups). A pvalue < 0.05 was considered statistically significant.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

W.Y. and B.C. designed research; W.Y., H.C., Z.D., J.C., Z.S., and B.Z. performed research; W.Y., H.C., D.C., X.H., and B.C. analyzed and supervised data; W.Y., X.H., and B.C. wrote and reviewed the paper.

Peer review

Peer review information

Nature Communications thanks Jiahui Ding, who co-reviewed with Anthony Maxwell and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Funding

This work was supported by grants from the National Natural Sciences Foundation in China (82130047 to B.C., 82401973 to W.Y., and 82571928 to X.H.), the National Key Research and Development Program of China (2022YFC2702400 to B.C. and 2022YFC2704702 to X.H.), Natural Science Foundation of Xiamen, China (3502Z202372001 to X.H.), Open Funding of State Key Laboratory of Reproductive Medicine and offspring health (SKLRM-K202401 to B.C.), Fundamental Research Funds for the Central Universities (20720240113 to X.H.).

Data availability

The bulk RNA-seq data generated in this study have been deposited in the NCBI Gene Expression Omnibus (GEO) under accession code GSE308795. The IP-MS proteomics data have been deposited to the ProteomeXchange Consortium via the iProX partner repository under the dataset identifier PXD071302 (https://www.iprox.cn//page/project.html?id=IPX0014425000)⁸³. All other data and materials are available from the corresponding author (B.C.). Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-73497-z.