https://orcid.org/0000-0001-8262-0876
Figures
Abstract
Trichinella spiralis (T. spiralis), a zoonotic nematode that causes severe myositis and systemic morbidity, sustains chronic muscle parasitism through evolutionary adaptations; however, this globally prevalent disease lacks targeted therapies to disrupt chronic infection. Although the heme transport protein HRG-1 has been characterized as an intervention target in free-living species (e.g., Caenorhabditis elegans) and hematophagous parasites (e.g., Haemonchus contortus), the molecular machinery governing heme acquisition in the nonhematophagous parasite T. spiralis remains uncharacterized, and no drugs targeting HRG-1 have been reported until now. Herein, we demonstrate that T. spiralis, a parasite that lacks the ability to synthesize heme autonomously, has evolved a sophisticated mechanism to scavenge and utilize heme from its host. By employing an aspartic protease to degrade host hemoglobin and myoglobin in the parasitic niche, T. spiralis is able to liberate heme for its own growth and survival. The structurally and functionally conserved Ts-HRG-1 protein plays a key role in transporting heme to the entire worm, particularly to functional organs, such as the cuticle and stichosome. More importantly, we discovered that the interaction between Ts-HRG-1 and Ts-ATP6V0C results in the formation of a functional complex that is essential for the parasite’s heme acquisition. The intervention effect achieved by Ts-ATP6V0C RNAi or inhibiting the activity of Ts-ATP6V0C with bafilomycin A1 (BafA1) was consistent with Ts-HRG-1 RNAi, resulting in impaired heme uptake, developmental arrest and a reduced larval burden in mouse hosts. These findings enhance our understanding of the parasite’s heme acquisition mechanism and identify the development of drugs that target proteins that interact with HRG-1 as a new direction in anthelminthic drug research.
Author summary
Chronic Trichinella spiralis (T. spiralis) infections currently lack effective anthelmintic drugs, particularly for the persistent muscle-dwelling phase. Exploring the physiological activities of T. spiralis during muscle parasitism as a breakthrough for drug development, we discovered that the T. spiralis aspartic protease degrades myoglobin to release heme, just as it degrades hemoglobin. As an organism incapable of synthesizing heme de novo, protein molecules that regulate heme uptake are undoubtedly potential intervention targets. Based on previous studies, we functionally confirmed that structurally conserved HRG-1 is a key protein involved in heme uptake in T. spiralis. Although reducing HRG-1 expression via RNAi affects larval growth and development, notably, no drugs blocking HRG-1 have been developed to date. In this work, Ts-ATP6V0C was identified as an HRG-1-interacting protein through coimmunoprecipitation, and most importantly, the intervention effect achieved by pharmacological inhibition of Ts-ATP6V0c was identical to that of RNAi. These findings suggest that the development of inhibitors targeting HRG-1-interacting proteins represents a new therapeutic direction for T. spiralis infections and even helminth infections.
Citation: He Y, Wang Y, He X, Lv Q, Vallee I, Boireau P, et al. (2026) Bafilomycin A1 is a promising therapeutic agent against T. spiralis infection by inhibiting the heme-transporting ATP6V0C/HRG-1 complex. PLoS Pathog 22(3): e1014042. https://doi.org/10.1371/journal.ppat.1014042
Editor: Raffi V. Aroian, University of Massachusetts Chan Medical School, UNITED STATES OF AMERICA
Received: September 22, 2025; Accepted: February 26, 2026; Published: March 16, 2026
Copyright: © 2026 He et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data generated or analyzed during this study are included in this published article, and its supplementary information files. The mass spectrometry raw data that support the findings of this study could be accessed in iProX (https://www.iprox.org/) with Project ID IPX0015439000 (https://www.iprox.cn/page/project.html?id=IPX0015439000).
Funding: This study was supported by The National Key Research and Development Program of China (https://service.most.gov.cn/) (2023YFE0107300 to J. D.), National Natural Science Foundation of China (https://www.nsfc.gov.cn/) (32302960 to J.D.; 32373032 to X.L.), and Natural Science Foundation of Jilin Province of China (https://jlky.yuntu.cn/dashboard/analysis) (No. 20220508052RC to J.D.). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Trichinella spiralis (T. spiralis) is a zoonotic parasite, and the majority of human infection cases are associated with the consumption of raw or undercooked meat containing infective larvae [1]. Despite the global prevalence (with an estimated 10,000 human cases annually worldwide, as reported by the WHO) and significant clinical impact of T. spiralis infections [2], targeted therapeutic strategies remain elusive, particularly for its encysted parasitic stage in muscle tissue. Elucidating critical metabolic pathways governing parasite growth and developmental transitions provides a foundational framework for advancing stage-specific interventions.
Heme, a metalloporphyrin with an iron protoporphyrin IX core, is an essential cofactor for diverse biological processes of living organisms, including oxygen transport, electron transfer, and xenobiotic detoxification [3,4]. Although most metazoans synthesize heme through a conserved biosynthetic pathway, parasitic nematodes have evolutionarily lost key enzymes in this pathway, rendering them obligate heme auxotrophs [5–7]. The obligatory reliance of these parasites on the acquisition of heme from host sources establishes their associated transport machinery as high-value chemotherapeutic targets.
Recent advances in heme trafficking mechanisms have revealed a conserved and multifaceted regulatory network with significant pathophysiological implications. The free-living nematode Caenorhabditis elegans (C. elegans) exerts this complexity, with at least seven identified HRG transporters (HRG-1 to HRG-4, HRG-7, HRG-9, and HRG-10) orchestrating heme homeostasis through specialized roles [8–13]. Among these, Ce-HRG-1 facilitates intracellular heme availability through endosomal- and/or lysosomal-related compartments, whereas its paralog Ce-HRG-4 mediates heme uptake across the plasma membrane [8]. Ce-HRG-3 was identified as a secreted protein that mediates intertissue heme transport from maternal intestinal stores to developing embryos, ensuring developmental homeostasis [11]. Furthermore, Ce-HRG-7 orchestrates systemic heme balance through a gut–neuron signaling axis involving the DBL-1/BMP pathway, highlighting the critical role of noncell-autonomous signaling in heme regulation [12]. In the same organism, Ce-HRG-9 and its human homolog TANGO2 were demonstrated to be essential heme chaperones in lysosome-related organelles and mitochondria, respectively, with functional defects leading to mitochondrial heme accumulation and metabolic disturbances, providing mechanistic insights into human TANGO2-associated encephalopathy and cardiac arrhythmias [13]. Similarly, in hematophagous parasites such as Haemonchus contortus, HRG-1 was shown to be indispensable for heme uptake and intracellular trafficking, with Hc-HRG-1 RNAi blocking host infection [14]. Together, these studies delineate an evolutionarily conserved framework for heme trafficking with broad implications for antiparasitic strategies. Critically, although heme acquisition represents a druggable vulnerability, no pharmacologic agents targeting HRG family transporters currently exist—a key barrier to anthelminthic drug development.
Unlike free-living nematodes (e.g., Caenorhabditis elegans) and hematophagous parasites (e.g., Haemonchus contortus), which acquire heme through environmental uptake or blood-feeding, respectively, the molecular basis of heme acquisition in the nonhematophagous parasite T. spiralis remains uncharacterized. T. spiralis establishes chronic infection by invading host striated muscle cells during its muscle larvae (ML) stage [15]. This distinct survival strategy suggests that T. spiralis may utilize myoglobin-derived heme as an alternative pathway alongside the canonical hemoglobin utilization systems observed in hematophagous species to sustain long-term parasitism [16]. Although aspartic protease (Ts-Asp2) in T. spiralis was previously shown to degrade hemoglobin in vitro [17,18], its capacity to hydrolyze myoglobin—the dominant heme reservoir in skeletal muscle—remains uncharacterized. Furthermore, the molecular mechanisms governing heme acquisition in T. spiralis remain poorly characterized.
In this study, in addition to validating the heme transport function of the conserved HRG-1 in T. spiralis via yeast rescue assays, we further defined the molecular role of a V-type proton ATPase proteolipid subunit (Ts-ATP6V0C) as its critical interacting partner. Pharmacological targeting of Ts-ATP6V0C with bafilomycin A1 (BafA1), a specific inhibitor of vacuolar ATPase activity [19], recapitulated the hallmark pathological effects of Ts-HRG-1 RNAi, including impaired heme uptake, developmental arrest, and diminished larval burdens in vivo. Elucidating this heme metabolic network is critical not only for understanding T. spiralis’ adaptation to myoglobin-rich niches but also for establishing the direct inhibition of Ts-ATP6V0C as a mechanistically innovative and therapeutically promising strategy against trichinellosis.
Results
The heme auxotroph T. spiralis utilizes aspartic protease to degrade myoglobin and release heme
To confirm whether T. spiralis is a heme auxotroph, we measured the activity of ALAD (δ-aminolevulinic acid dehydratase) in the parasite [20–22]. Using Escherichia coli, which is capable of heme synthesis, as a positive control, homogenates of E. coli and T. spiralis were incubated with 5-aminolevulinic acid (ALA) for 4 hours, followed by quantification of the downstream product PBG (porphobilinogen). The results revealed that PBG synthesis in E. coli decreased with lower ALA concentration, whereas no PBG was detected in T. spiralis homogenates, confirming that T. spiralis cannot synthesize heme de novo and is a heme auxotroph (Fig 1A and 1B).
(A) Colorimetric detection of ALAD activity. Reaction mixtures containing δ-aminolevulinic acid (ALA, 2.5–50 μg/μL) were incubated with T. spiralis (Ts) lysates, E. coli lysates (positive control), or PBS (negative control). Porphobilinogen (PBG) was detected with Ehrlich’s reagent. A pink-to-red colorimetric signal (indicative of PBG formation) was observed in E. coli lysates in an ALA concentration-dependent manner. No visible color change was detected in Ts lysates or the PBS control. (B) Absorbance quantification at 555 nm. E. coli lysates exhibited dose-dependent PBG production (p < 0.0001 vs. Ts/PBS). Ts lysates and PBS showed negligible absorbance across all ALA concentrations, with no significant difference between Ts and PBS. Statistical analysis was performed using two-way ANOVA (n = 3). (C) TMB-based proteolytic assay. Mb (1 μg/μL) was incubated with rTs-Asp2 (0.5 μg/μL) ± Pepstatin A (10 μM, aspartic protease inhibitor) at 37°C for 4 h. Mb alone exhibited the strongest blue coloration (oxidized TMB), followed by rTs-Asp2 + Mb + PepA, whereas rTs-Asp2 + Mb showed minimal coloration. No signal was detected in the rTs-Asp2-alone group. (D) Absorbance spectra of the TMB oxidation products. Mb alone had the highest peaks at 370 and 652 nm (TMB oxidation markers). rTs-Asp2 + Mb exhibited significantly reduced signals, whereas rTs-Asp2 + Mb + PepA displayed intermediate peaks. No peaks were detected for rTs-Asp2 alone. Statistical analysis was performed using two-way ANOVA (n = 3). (E) SDS‒PAGE analysis of Mb integrity. Mb alone and rTs-Asp2 + Mb + PepA had intact 15 kDa bands, whereas rTs-Asp2 + Mb had a significantly fainter band. (F) Hemin-dependent molting rate. Larvae cultured with hemin (0–100 μM) for 24 h exhibited dose-dependent molting (0 μM vs. 100 μM, p < 0.0001; n = 3 biological replicates, one-way ANOVA). (G) Microscopic visualization of molting. Shed cuticles (translucent sheaths, arrows) were rare at 0 μM but increased with increasing hemin concentration. Scale bars: 200 μm (main panels), 100 μm (insets). (H) ZnMP fluorescence imaging. Larvae cultured in PBS, 25 μM hemin, 6.25 μM hemoglobin (Hb), or 25 μM myoglobin (Mb) presented the strongest anterior-localized granular fluorescence, with diminished signals in the Hemin/Hb/Mb groups (excitation/emission: 561/595 nm). (I) ZnMP fluorescence intensity in larvae (one-way ANOVA, n = 3).
Previous studies have reported that T. spiralis degrades host hemoglobin via a proteolytic cascade system to release heme for metabolic needs, with aspartic proteases serving as key enzymes [17]. Given that T. spiralis parasitizes inside skeletal muscle cells, which are rich in myoglobin but lack hemoglobin, during the ML stage [15,16], we hypothesized that an aspartic protease, TsASP2, may also exhibit myoglobin-degrading activity. To test this hypothesis, recombinant TsASP2 (rTsASP2) was obtained through prokaryotic expression and assayed for enzymatic activity using myoglobin as a substrate. Compared with the myoglobin-only (Mb) group, the rTsASP2 + Mb group presented significantly reduced TMB chromogenic reactivity and absorbance at 370 nm and 652 nm, indicating myoglobin degradation by rTsASP2. Furthermore, the addition of the aspartic protease inhibitor Pepstatin A (PepA) resulted in partial chromogenic reactivity and absorbance levels in the rTsASP2 + Mb + PepA group, confirming that rTsASP2 activity is suppressed by PepA (Fig 1C and 1D) [23]. SDS‒PAGE analysis revealed a prominent myoglobin band in the Mb group, whereas this band was markedly diminished in the rTsASP2 + Mb group (Fig 1E). These results establish that TsASP2 degrades not only hemoglobin but also myoglobin, demonstrating a mechanism enabling free heme acquisition in T. spiralis.
Hemin promotes larval molting and competitively inhibits heme analog uptake
Given that T. spiralis is a heme auxotroph that requires external heme acquisition, we supplemented hemin in culture medium to assess its effects on larval growth and development. The rate of molting increased in a dose-dependent manner with increasing exogenous hemin supplementation in serum-free, hemin-free basal medium. After a 24-hour incubation period (as determined by preliminary experiments) with 50 and 100 µM hemin, the molting rate reached approximately 60%, suggesting that heme promotes parasite growth and development (Fig 1F and 1G) [24]. In standard RPMI-1640 medium, ML exhibited significantly greater uptake of Zn (II) Mesoporphyrin IX (ZnMP, a fluorescent heme analog) than did the groups supplemented with 25 µM hemin, 6.25 µM hemoglobin (Hb), or 25 µM myoglobin (Mb) (Fig 1H and 1I). The present findings demonstrate the essentiality of exogenous heme for the development and molting of T. spiralis larvae, and also reveal that heme uptake and accumulation are strictly regulated, as has been previously observed in the nematode C. elegans [8].
Ts-HRG-1 demonstrates structural conservation and the capacity to bind heme in vitro
Given the critical role of heme in the growth and development of T. spiralis, we focused on the evolutionarily conserved heme transporter HRG-1 in nematodes. Ts-HRG-1 is transcribed at different developmental stages, with the lowest level in the ML stage (S1A Fig). To explore the evolutionary conservation and structural features of Ts-HRG-1, we conducted phylogenetic analysis of its homologous proteins. These homologs are categorized into four groups based on their biological classification and parasitic traits: animal parasitic nematodes, plant-parasitic nematodes, non-parasite nematodes, and non-nematodes. Phylogenetic tree analysis revealed that Ts-HRG-1 forms a sister branch with HRG-1 from C. elegans (Ce-Hrg-1), indicating a close evolutionary relationship within the phylum Nematoda (Fig 2A).
(A) A maximum-likelihood phylogenetic tree of HRG-1 homologs (MEGA) grouped T. spiralis (Ts) HRG-1 within the animal parasitic nematode clade, which is distinct from that of plant parasitic nematodes, non-parasitic nematodes, and non-nematodes. (B) Sequence alignment (MEGA) of Ts-HRG-1 with homologs from Hc, Ce, mouse, and zebrafish revealed four conserved transmembrane domains (TMD1-4, purple boxes) and an intact heme-binding Y-x-R-x-R motif (yellow box). (C) Homology models (ChimeraX) depicting HRG-1 structures across species, color-coded to show extracellular loops (pink), transmembrane helices (green), and the surrounding phospholipid bilayer (red: outer leaflet; blue: inner leaflet). TMDs are spatially conserved, which is consistent with a shared transmembrane architecture.
Topological modeling and amino acid sequence alignment revealed that Ts-HRG-1 shares sequence similarity with homologs from mouse, zebrafish, C. elegans, and Haemonchus contortus, all harboring four transmembrane domains and a conserved structural region. Notably, Ts-HRG-1 contains key heme-binding residues (H90 in TMD2) and a Y-x-R-x-R motif at the C-terminal tail, suggesting functional conservation in heme transport (Fig 2B and 2C) [25–27]. Molecular docking predicted the heme-binding capacity of Ts-HRG-1, identifying a hydrophobic pocket as the binding site. The lowest-energy conformation (–7.59 kcal/mol) revealed the stabilization of the hemin porphyrin ring through hydrogen bonds with ARG33, LYS145, and TYR153, supplemented by Pi-cation and Pi-sigma interactions with LYS145 (Fig 3A and 3B). We subsequently validated the interaction between recombinant Ts-HRG-1 (rTs-HRG-1) and hemin in vitro using TMB staining. A distinct band was observed in the reaction mixture containing rTs-HRG-1 and hemin. The absorbance measurements revealed that the characteristic peak of free hemin at 390 nm shifted to 370 nm in the presence of rTs-HRG-1 (Fig 3C and 3D). These results suggest that Ts-HRG-1 exhibits structural conservation and is capable of binding to heme in vitro.
(A) Molecular docking (AutoDock) identified Arg33, Lys145, and Tyr153 as key residues that form hydrogen bonds and hydrophobic interactions with hemin. Electrostatic surface models highlight the hemin-binding pocket. (B) Binding affinity analysis (Discovery Studio) confirmed the hemin-Ts-HRG-1 interaction, with schematic diagrams depicting binding poses (see in-fig legends). (C) Nondenaturing PAGE of rTs-HRG-1 incubated with hemin (37°C, 2 h). TMB staining revealed a distinct band in the rTs-HRG-1 + hemin lane (hemin-bound complex), which was absent in the rTs-HRG-1 alone lane. Coomassie blue staining confirmed equal protein loading. Hemoglobin (positive control) showed intrinsic heme-dependent TMB reactivity. (D) Absorbance spectra (340–750 nm). Free hemin exhibited a peak at 390 nm (Soret band), which shifted to 370 nm in the rTs-HRG-1 + hemin group, indicating hemin‒protein interactions. No peak was observed for rTs-HRG-1 alone.
Ts-HRG-1 possesses a conserved heme transport function identical to that of Ce-HRG-4 in a heme-auxotrophic yeast model
We evaluated the function of Ts-HRG-1 in Saccharomyces cerevisiae engineered with a heme-auxotrophic phenotype (Δhem1) [21,28]. CRISPR-Cas9-mediated disruption of HEM1 resulted in a strain incapable of endogenous heme synthesis without 5-aminolevulinic acid (ALA, a heme biosynthesis precursor) and unable to acquire exogenous heme, as confirmed by PCR amplification and sequencing of the disrupted locus (Figs 4A, S1B and S1C). Phenotypic validation confirmed that the Δhem1 strain failed to grow on YPD medium lacking ALA, whereas growth was restored upon ALA supplementation, verifying its heme auxotrophy (Fig 4B). To investigate whether Ts-HRG-1 shares functional similarities with the heme transporter Ce-HRG-4 from C. elegans, we cloned Ts-HRG-1 and Ce-HRG-4 into yeast expression vectors and transformed them into the Δhem1 strain to assess their rescue capabilities under heme-deficient conditions. After confirming successful transformation by Western blotting (Fig 4C), we assessed the growth of the control vector, Ts-HRG-1, and Ce-hrg-4 transformants in a spot assay on SD medium with or without ALA. All the transformants showed no growth in the absence of ALA but exhibited comparable colony density in the ALA-supplemented medium, indicating that the introduction of exogenous genes did not alter the auxotrophic phenotype of the yeast and validating the experimental approach (Fig 4D). To further evaluate the function of Ts-HRG-1, we assessed growth recovery using spot assays with three concentrations of exogenous hemin (0.25, 2.5, and 10 μM). The results demonstrated that the Δhem1 strains expressing Ts-HRG-1 or Ce-HRG-4 presented significant growth restoration across all hemin concentrations, whereas the empty vector control did not (Fig 4E). Growth curve analysis at 2.5 μM hemin further confirmed robust recovery in Ts-HRG-1/Ce-HRG-4 transformants compared with the vector control (Fig 4F). These findings indicate that Ts-HRG-1 possesses a heme transport function similar to that of Ce-HRG-4.
(A) Schematic diagram of HEM1 knockout and complementation in S. cerevisiae BY4741. The CRISPR/Cas9 system (pCAS plasmid) with HEM1-targeting sgRNA was used to generate the Δhem1 strain, which was subsequently transformed with empty vector (pYES2-CT), Ce-HRG-4, or Ts-HRG-1 constructs. (B) Validation of Δhem1 knockout: parental and Δhem1 strains were cultured on YPD ± 250 μM δ-aminolevulinic acid (ALA). Δhem1 failed to grow without ALA, confirming auxotrophy. (C) Western blotting (with an anti-His tag) confirmed the expression of Ce-HRG-4 and Ts-HRG-1 in the transformed strains. (D) Growth validation of transformed Δhem1 strains: Empty vector-, Ce-HRG-4-, and Ts-HRG-1-expressing strains grew similarly on SD ± ALA, confirming plasmid neutrality. (E) Hemin concentration-dependent growth rescue. Δhem1 strains expressing Ts-HRG-1, Ce-HRG-4, or the empty vector were cultured in SD medium supplemented with hemin (0.25, 2.5, or 10 μM). Ts-HRG-1 restored growth at 2.5 μM hemin, with further enhancement at 10 μM, mimicking that of Ce-HRG-4. No rescue occurred at 0.25 μM or with the empty vector. (F) Growth curves of Δhem1 yeast strains expressing the empty vector (control), Ts-HRG-1, or Ce-HRG-4 in SD medium containing 2.5 μM hemin. The indicated P value (p < 0.0001) represents the comparison of OD600 values at the 22-hour time point, analyzed by one-way ANOVA (n = 3).
The subcellular localization of Ts-HRG-1 supports its role in heme uptake and transport in T. spiralis
Immunofluorescence experiments using Ts-HRG-1-specific antiserum revealed that Ts-HRG-1 is predominantly localized to the cuticle and stichosome in ML (Fig 5A and 5B), with identical localization observed in the stichosome in adult worms 6 days after infection (AD6) (S1D Fig). Its presence in the cuticle, the outermost protective layer of the parasite, suggests a potential role in heme acquisition from the host environment. Additionally, localization in the stichosome, a specialized secretory organ, implies involvement in heme storage or transport.
(A) Immunofluorescence localization of Ts-HRG-1 in ML sections using a Ts-HRG-1 polyclonal antibody (red; Alexa Fluor 555-conjugated secondary antibody and Alexa Fluor 647-conjugated secondary antibody) and DAPI (blue, nuclei). The negative control serum (preimmune serum with identical secondary antibodies) showed no specific signal. Ts-HRG-1 localized to the stichosome (arrows). Scale bars: 20–50 μm. (B) Immunofluorescence localization of Ts-HRG-1 in intact ML using a Ts-HRG-1 polyclonal antibody (red; Alexa Fluor 555-conjugated secondary antibody and Alexa Fluor 647-conjugated secondary antibody) and DAPI (blue, nuclei). The negative control serum (preimmune serum with identical secondary antibodies) showed no specific signal. Ts-HRG-1 localized to the cuticle (arrows). Scale bars: 20–50 μm. (C) Colocalization of Ts-HRG-1-GFP (green) with organelle markers: Golgi, lysosomes, and plasma membrane. Strong colocalization (yellow) was observed with the plasma membrane (arrowheads), but minimal overlap with the Golgi or lysosomes was detected. (D) Colocalization analysis. Pearson’s correlation coefficient (Rr): plasma membrane = 0.782, Golgi = 0.274, and lysosomes = 0.239. The intensity profiles show high overlap between the Ts-HRG-1-GFP and mCherry signals.
To further validate its function, colocalization analysis in HeLa cells expressing Ts-HRG-1-GFP revealed a strong plasma membrane association (Pr = 0.782) but negligible colocalization with the Golgi (Rr = 0.274) or lysosomes (Rr = 0.239) (Fig 5C and 5D). This quantitative compartmental specificity (>0.5 threshold) supports the primary role of Ts-HRG-1 in mediating extracellular heme uptake at the plasma membrane in T. spiralis.
Ts-HRG-1-mediated heme transport governs ML development and reproductive capacity in T. spiralis
To assess the impact of Ts-HRG-1 on larval development and growth in T. spiralis, we employed RNA interference (RNAi) targeting Ts-HRG-1 in ML. RNAi efficacy was confirmed by significant reductions in both the mRNA and protein levels of Ts- HRG-1 (S2 Fig).The transcriptional level of HRG-1 in untreated ML exhibited a gradual increase under low concentrations of heme (0–25 μM), but decreased at high heme concentrations (100 μM) (Fig 6A). This phenomenon could be interpreted as an adaptive response resulting from the toxicity associated with elevated heme concentrations [29]. Consequently, in subsequent experiments, an exogenous heme concentration of 25 μM was selected to verify the effect of HRG-1 RNAi on heme uptake. The results demonstrated that the transcriptional and protein levels of HRG-1 in the RNAi worms increased following the supplementation with exogenous heme (Fig 6B–6D). The molting rate in the RNAi worms exhibited a consistent decrease, which was followed by an increase after the administration of exogenous heme (Fig 6E and 6F).
(A) ML were cultured with 0–100 μM hemin for 24 h, and the expression of Ts-HRG-1 was analyzed by qPCR. Statistical significance was determined by one-way ANOVA (n = 3). (B) qPCR analysis of Ts-HRG-1 expression in ML treated with either NC (nontargeting control) or Ts-HRG-1 RNAi and cultured with 0 or 25 μM Hemin for 24 h. Statistical significance was determined by one-way ANOVA (n = 3). (C) Western blot (anti-Ts-HRG-1, with α-tubulin as a loading control). (D) Grayscale quantification of the WB bands. Statistical analysis was performed using one-way ANOVA (n = 3). (E) Microscopy images of ML molting. (F) Molting rate quantification. Statistical analysis was performed using one-way ANOVA (n = 3). (G) Schematic of the mouse infection assay. PBS-, NC-, or Ts-HRG-1 RNAi-treated ML (200 larvae/mouse) were orally administered to the mice, with the larvae collected on Day 6 (AD6) and Day 35 (ML stage). (H) Larval burden analysis. AD6: No difference across groups. Statistical analysis was performed using one-way ANOVA (n = 6). (I) NBL: The RNAi group presented reduced larval counts (p < 0.05 vs. the NC group). Statistical analysis was performed using one-way ANOVA (n = 12). (J) The Day 35 ML: RNAi group presented a significantly lower ML burden (p < 0.05 vs. the NC group). Statistical analysis was performed using one-way ANOVA (n = 6).
When RNAi-treated ML was orally administered to the mice, the number of Ad6 in the Ts-HRG-1 knockdown group was not significantly different from those in the PBS and negative control (NC) groups. However, parasite reproductive capacity was markedly reduced. At 35 days post infection, the number of recovered ML was significantly lower, demonstrating that Ts-HRG-1 knockdown impaired the reproductive ability of T. spiralis and diminished the worm burden in the host (Fig 6G–6J). These results collectively establish Ts-HRG-1 as a heme transporter essential for larval development and reproductive fitness in T. spiralis.
Ts-ATP 6V0C and Ts-HRG-1 regulate heme transport to sustain development across T. spiralis life stages
Given the absence of targeted inhibitors for Ts-HRG-1, we sought to identify its interaction partners as potential therapeutic candidates. An ATP-associated protein (Ts-V-ATPase_V1_A) as a potential interaction partner of Ts-HRG-1 was identified by coimmunoprecipitation (Co-IP) of polyclonal antibodies with T. spiralis crude extract (S1 Table). However, subsequent colocalization analysis, Co-IP, and yeast two-hybrid assays failed to validate this interaction (S3A–S3D Fig). Based on prior reports of HRG-1 interaction with the V-type proton ATPase proteolipid subunit in mammalian cells [30], homology analysis identified Ts-ATP6V0C as its ortholog in T. spiralis. Yeast two-hybrid assays demonstrated a direct interaction between Ts-HRG-1 and Ts-ATP6V0C (Fig 7A). Subsequent Co-IP in 293T cells coexpressing Ts-HRG-1-GFP and Ts-ATP6V0C-mCherry confirmed their specific binding in cellular lysates (Fig 7B), with immunofluorescence analysis in the same transfected 293T cells demonstrating robust plasma membrane colocalization (Rr = 0.773) (Fig 7C and 7D). These results establish both direct physical interactions and functional colocalization of Ts-HRG-1 and Ts-ATP6V0C, implicating their partnership in heme transport mechanisms.
(A) Yeast two-hybrid assay. AD-Ts-ATP6V0C and BD-Ts-HRG-1 were cotransformed into yeast. Growth on SD-TLHA + 10 mM 3-AT confirmed the interaction, with no autoactivation in the controls (AD/BD empty vectors). (B) Coimmunoprecipitation (Co-IP). Lysates from HeLa cells coexpressing Ts-HRG-1-GFP and Ts-ATP6V0C-mCherry were immunoprecipitated with anti-GFP. Ts-ATP6V0C-mCherry was detected in the IP fraction (anti-mCherry blot), confirming the physical interaction. No signal was observed in the GFP-empty controls. (C) Colocalization analysis. Pearson’s correlation coefficient (Rr = 0.773) and distance-dependent intensity profiles confirmed the spatial proximity between Ts-HRG-1 and Ts-ATP6V0C. (D) Colocalization in HeLa cells. Ts-HRG-1-GFP (green) and Ts-ATP6V0C-mCherry (red) strongly colocalized (yellow) in the cell. Scale bar: 5 μm.
To further explore the functional role of Ts-ATP6V0C, we investigated the effects of RNAi and its specific inhibitor BafA1 on the viability of T. spiralis at different developmental stages. The results demonstrated that the effective knockdown of Ts-ATP6V0C (S3E–S3G Fig) or its activity inhibition by the drug resulted in a reduction in ZnMP uptake and a decline in the molting rate (Figs 8A–8C and S4A). By applying various drug concentrations at different developmental stages of T. spiralis, it was observed that as the drug concentration increased, the mortality rates of NBL and AD6 exhibited a gradual rise, while the molting rate of ML demonstrated a gradual decrease (Fig 8D–8F and S4B–S4D). Subsequent analysis of the transcriptional level of Ts-ATP6V0C after the drug acted on ML revealed that BafA1 did not cause significant changes in the transcriptional level of Ts-ATP6V0C (Fig 8G). The hypothesis posits that this phenomenon is attributable to the unique mechanism of BafA1, which exerts its influence on the activity of Ts-ATP6V0C rather than on its expression. However, it is noteworthy that BafA1 treatment of ML or RNAi of Ts-ATP6V0C in ML resulted in an increase in the transcriptional level of Ts-HRG-1 (Fig 8H). In order to corroborate this unanticipated phenomenon, an analysis was conducted on the transcriptional levels of Ts-HRG-1 in T. spralis treated with varying concentrations of BafA1 at different developmental stages in vitro. The results demonstrated that the transcription of Ts-HRG-1 exhibited a fluctuating pattern, with a trend of increasing transcription at low drug concentrations and decreasing transcription at high drug concentrations (Fig 8I–8K). Although the transcription of Ts-HRG-1 in T. spiralis showed an increasing trend after treatment with low concentration of BafA1, the loss of the auxiliary transport function due to the inhibition of Ts-ATP6V0C activity still led to the inhibition of the worm’s vitality. The dose-dependent transcriptional dysregulation of Ts-HRG-1 caused by BafA1 supports the functional coupling between Ts-ATP6V0C and Ts-HRG-1, as well as the compensatory regulatory mechanism of interacting proteins. This is because pharmacological disruption of the transport complex can directly regulate the expression of the heme sensor.
(A) ZnMP fluorescence in treated larvae. ML pretreated with Ts-HRG-1 RNAi, Ts-ATP6V0C RNAi or 2.5μg/mL BafA1 (for 24 h) and fed ZnMP displayed markedly reduced intracellular fluorescence, suggesting impaired heme uptake. (B) Quantification of ZnMP fluorescence intensity. The intensity was significantly reduced in the Ts-HRG-1 RNAi, Ts-ATP6V0C RNAi and BafA1 groups. Statistical analysis was performed using one-way ANOVA (n = 3). (C) ML molting rate analysis of the CON, Ts-HRG-1 RNAi, Ts-ATP6V0C RNAi and BafA1 (2.5 μg/mL) groups; data are presented as the means ± SEMs (one-way ANOVA, n = 3). (D) Mortality of NBL: Concentration-dependent mortality following 24-h BafA1 exposure. One-way ANOVA vs. control, n = 3. (E) Mortality of AD6: Dose‒dependent mortality after 24 h of BafA1 treatment (0–10 μg/mL). One-way ANOVA vs. control, n = 3. (F) Molting rate of ML: BafA1-induced suppression of molting after 24 h of treatment. One-way ANOVA vs. control, n = 3. (G) ML: Ts-ATP6V0C mRNA measurement following 24 h of BafA1 treatment. Data = mean ± SEM; one-way ANOVA vs. control. 18S rRNA internal control (n = 3). (H) Ts-HRG-1 qPCR analysis: CON, Ts-HRG-1 RNAi, Ts-ATP6V0C RNAi and BafA1 (2.5 μg/mL) groups with means ± SEMs; one-way ANOVA (18s rRNA reference). (I) NBL: Ts-HRG-1 mRNA quantification after 24 h of BafA1 treatment. Data = mean ± SEM; one-way ANOVA vs. control. 18S rRNA internal control, n = 3. (J) AD6: Ts-HRG-1 mRNA levels after 24 h of BafA1 treatment. Data = mean ± SEM; one-way ANOVA vs. control. 18S rRNA internal control, n = 3. (K) ML: Ts-HRG-1 mRNA measurement following 24 h of BafA1 treatment. Data = mean ± SEM; one-way ANOVA vs. control. 18S rRNA internal control, n = 3.
Bafilomycin A1 exhibits therapeutic potential against T. spiralis infection in mice
To evaluate the therapeutic potential of BafA1 for treating trichinellosis, we conducted in vivo experiments. Mice orally infected with ML were randomly divided into seven groups and administered three concentrations of BafA1 (0.1, 0.5, or 1 mg/kg) via oral gavage during either the intestinal phase (Days 0–7 post infection) or the encystment phase (Days 21–29 post infection). At 35 days postinfection, ML were collected and quantified. Compared with the control groups, all the BafA1-treated groups presented significantly lower larval burdens (Fig 9A and 9B). Histopathological evaluation of diaphragm sections further revealed a marked decrease in intramuscular cyst counts in BafA1-treated mice (Fig 9C), confirming the efficacy of BafA1 in disrupting both larval viability and cyst formation.
(A) Experimental design. Mice (n = 6/group) were administered BafA1 (0.1, 0.5, or 1 mg/kg) via oral gavage during the intestinal phase (0–7 dpi; Int. ph) or encystment phase (21–29 dpi; Encyst. ph). ML were recovered at 35 dpi for quantification. (B) Larval burden analysis. BafA1 treatment during the intestinal phase (0–7 dpi; Int. ph) significantly reduced the ML count (p < 0.05), and treatment during the encystment phase (21–29 dpi; Encyst. ph) reduced the ML count (p < 0.05). The data were analyzed with GraphPad Prism (mean ± SEM). Statistical analysis was performed using one-way ANOVA (n = 6). (C) H&E staining of representative diaphragm sections. Treatment groups: Nondosing group (CON), BafA1-administered at 0.1, 0.5, or 1 mg/kg during the intestinal phase (Int. ph) and encystment phase (Encyst. ph). Scale bar: 200 μm.
Fig 10 presents a schematic of a proteolysis–transport axis for extracellular heme acquisition in T. spiralis, wherein secreted TsASP2 liberates hemin from host myoglobin or hemoglobin for subsequent internalization via the Ts-ATP6V0C/Ts-HRG-1 transporter complex to sustain parasitic development. Critically, pharmacologic inhibition of Ts-ATP6V0C by BafA1 disrupts this axis, arresting ML development and reducing infectivity in mice. These findings highlight the potent efficacy of BafA1 against the persistent encysted larval stage of T. spiralis, positioning it as a promising therapeutic candidate for chronic trichinellosis.
Key findings are integrated: Ts-Asp2 degrades host myoglobin (Mb) or hemoglobin (Hb) to release hemin, which is transported by Ts-HRG-1 and its interacting protein Ts-ATP6V0C.
Discussion
The persistent evolutionary interplay between parasites and their hosts has fostered the development of intricate mechanisms for nutrient acquisition in parasitic nematodes [31]. Our study confirms the evolutionary conservation of HRG-1-mediated heme transport in T. spiralis while revealing two groundbreaking advances: the first demonstration of myoglobin-derived heme acquisition through the aspartic protease TsASP2 and the novel therapeutic inhibition of the Ts-ATP6V0C to affect heme transport using bafilomycin A1, a macrolide antibiotic that reduces larval burden in murine infection models. Collectively, these findings establish a mechanistically grounded strategy to combat trichinellosis by targeting parasitic heme homeostasis through the Ts-ATP6V0C/Ts-HRG-1 complex.
Like other parasitic nematodes, T. spiralis exhibits auxotrophy for heme biosynthesis, as evidenced by the absence of δ-aminolevulinic acid dehydratase (ALAD) activity, thereby necessitating reliance on exogenous heme. Notably, T. spiralis parasitizes within striated muscle cells—an intracellular niche rich in myoglobin but devoid of hemoglobin—where NBL undergo >10-fold volumetric expansion within 20 days postinvasion to establish chronic infection, a process demanding substantial heme acquisition to sustain massive biomass production. Our study revealed that the parasite secretes an aspartic protease (TsASP2) capable of releasing heme from host myoglobin, a mechanism that may expand its repertoire of heme acquisition pathways. This adaptation highlights the parasite’s metabolic plasticity in exploiting tissue-specific resources. Such convergent evolution underscores the selective pressure to optimize heme acquisition in diverse parasitic niches, where nutrient availability varies drastically. Importantly, genomic analyses confirm that Ts-HRG-1 is the sole annotated HRG homolog in T. spiralis, underscoring its nonredundant function in heme acquisition. The structural conservation of HRG-1 across nematodes, particularly in transmembrane domains (TMD2 H90) and the C-terminal Y-x-R-x-R motif, strongly supports its capacity for heme binding, a function pivotal to maintaining heme homeostasis [25–27]. This is further corroborated by molecular docking analyses showing that these motifs form a hydrophobic pocket stabilized by residues critical for heme coordination (e.g., ARG33 and TYR153). In T. spiralis, HRG-1-mediated transport underpins a conserved heme acquisition strategy, a mechanism maintained across nematodes for heme homeostasis. This conservation underscores HRG-1’s indispensable role in nematode physiology—irrespective of the heme source exploited—reinforcing an evolutionary blueprint for heme transporter function.
Immunofluorescence analysis revealed that Ts-HRG-1 localized to the cuticle and stichosome, whereas ZnMP—a fluorescent heme analog—accumulated predominantly in the midgut lumen. Colocalization studies further demonstrated the presence of Ts-HRG-1 at the plasma membrane, suggesting its role in transporting heme from the midgut to peripheral tissues. This spatial separation suggests that orally ingested hemin may first be absorbed through midgut epithelial cells and subsequently transported to peripheral tissues via Ts-HRG-1 localized at the plasma membrane. The stichosome—a glandular organelle composed of longitudinally aligned stichocytes [32] —shows specific localization of Ts-HRG-1, suggesting a potential role in heme metabolism. The detection of Ts-HRG-1 at the cuticle—a structural barrier permeable to small molecules based on studies in other nematodes (i.e., C. elegans)—suggests its potential association with small-molecule transport, although T. spiralis cuticle permeability remains to be experimentally validated. Critically, the compartmentalized distribution of Ts-HRG-1 underscores its indispensable role in systemic heme trafficking, indicating that this transporter is a high-value therapeutic target. Despite its mechanistic promise, there are currently no pharmacologic agents targeting HRG-1—an urgent gap in anthelminthic development.
Remarkably, although no direct HRG-1-targeting agents exist, targeting its functional interactor (Ts-ATP6V0C) showed potent anthelmintic efficacy in our study. Furthermore, Ts-ATP6V0C RNAi and inhibition of Ts-ATP6V0C (by BafA1) triggered compensatory Ts-HRG-1 upregulation, revealing a feedback loop counteracting heme deficiency. This functional crosstalk establishes the Ts-ATP6V0C/Ts-HRG-1 complex as an essential heme trafficking axis for parasite development. Most importantly, pharmacologic inhibition of Ts-ATP6V0C recapitulated the phenotypic outcomes of Ts-HRG-1 or Ts-ATP6V0C RNAi silencing—impaired ML molting and reduced larval burden—demonstrating that disrupting HRG-1 interactors achieves equivalent disruption of heme homeostasis.
Bafilomycin A1 (BafA1) is a macrolide antibiotic that acts as a highly specific inhibitor of vacuolar (V-type) H ⁺ -ATPase (V-ATPase) by directly binding to its c-subunit (ATP6V0C) [19,33]. Beyond its applications in oncology and autophagy inhibition, BafA1 exhibits antiparasitic activity, including potent effects against Plasmodium falciparum and synergistic action with dihydroartemisinin (DHA) against this parasite [34]. BafA1 also reduces the intracellular burden of Toxoplasma gondii [35]. In this study, we report for the first time BafA1 significant anthelmintic activity against the nematode, and mechanistically link this effect to the disruption of the parasite’s heme acquisition pathway via Ts-ATP6V0C. It should be noted that as a V-ATPase inhibitor, BafA1 may also affect other ATPase targets in T. spiralis beyond Ts-ATP6V0C. While our data demonstrate that inhibition of Ts-ATP6V0C phenocopies Ts-HRG-1 knockdown, future studies employing genetic ablation or more specific inhibitors are warranted to definitively confirm the on-target specificity of BafA1 in this context. What’s more, although our in vivo data indicate a favorable tolerability at efficacious doses (0.1-1 mg/kg), the therapeutic potential of BafA1 must be balanced against its potential toxicity as a inhibitor of V-ATPase that is ubiquitously expressed in host cells.
Numerous studies indicate that BafA1 is not the sole anticancer drug that exhibits antiparasitic activity. For example, methotrexate and imatinib have demonstrated efficacy against Toxoplasma and Trypanosoma cruzi infections, respectively [36,37]. Conversely, classical antiparasitic drugs also show anticancer potential—albendazole targets lung cancer and melanoma [38], praziquantel synergistically enhances paclitaxel-induced tumor suppression [39], and artemisinin derivatives exhibit activity against breast cancer, lung cancer, and leukemia [40–42]. These bidirectional repurposing cases indicate fundamental biological similarities between parasite development and tumor pathogenesis, highlighting drug repositioning as a promising strategy for novel therapeutic development. Notably, the antiparasitic mechanism of BafA1 is distinct from that of conventional anthelmintics. Principally, the effectiveness of conventional anthelmintics like albendazole is strongly dependent upon early administration and high-dose regimens [43], as their efficacy diminishes after the larvae have encysted within muscles—a stage at which most patients are diagnosed. In striking contrast, our findings demonstrate that BafA1 administration during the encystment phase (days 21–29 post-infection) significantly reduces larval burden, positioning it as a promising therapeutic candidate for treating chronic trichinellosis.
It is particularly important to emphasize here that the mechanism by which helminth regulates heme excretion is also a field worthy of in-depth study. We observed that expression of the key transporter Ts-HRG-1 was downregulated at high hemin concentrations, hinting at a feedback regulatory mechanism to manage intracellular heme levels and potentially mitigate toxicity. This pattern is not without precedent; similarly, in C. elegans, the expression of HRG-10—which mediates heme export from enriched compartments—also shows non-monotonic changes under hemin treatment as high as 400 μM [13]. Mammals or protozoa degrade heme via heme oxygenases (iron-dependent enzymes that convert heme into biliverdin, CO, and free iron) [44], but heme oxygenases in T. spiralis are currently unreported. T. spiralis may instead rely on heme-sequestering proteins such as glutathione transferases (GSTs), as observed in hookworms [45]—a hypothesis requiring experimental validation. In addition to these established mechanisms, future studies employing heme-affinity proteomics or genome-wide RNAi screening should prioritize the identification of new heme transporters and their interacting proteins, given the parasite’s exceptional heme demand during volumetric expansion in muscle cells. The discovery of more molecules and their auxiliary components regulating heme transport will contribute to the mapping of the heme metabolism blueprint of parasites, revealing new targets for combinatorial disruption of heme trafficking.
In conclusion, this work delineates a coordinated “degrade-transport-regulate” axis for heme acquisition in T. spiralis, bridging evolutionary conservation with parasite-specific adaptations. By exploiting metabolic dependencies unique to parasitic nematodes, these insights pave the way for novel anthelmintics that target heme homeostasis—a conserved therapeutic target across helminth pathogens. Future studies should prioritize targeting HRG-1 interaction partners, develop inhibitors against these complexes, evaluate their cross-reactivity in diverse nematode species, and explore synergistic interactions with current therapeutic regimens. Given the increasing resistance to conventional anthelmintic agents, targeting evolutionarily constrained metabolic dependencies—such as heme transport machinery—represents a strategically viable approach to alleviate the global health burden imposed by multidrug-resistant nematode infections.
Materials and methods
Ethics statement
All procedures strictly followed protocols approved by the Animal Welfare and Research Ethics Committee and Institutional Animal Care and Use Committee of Jilin University (Approval No. KT202202140).
Parasites and animals
The Trichinella spiralis T1 strain (maintained at the State Key Laboratory of Zoonotic Diseases, Jilin University) was propagated through serial passages in female ICR mice (6–8 weeks old; Liaoning Changsheng Biotechnology Co.). ML were isolated from infected skeletal muscle via digestion in artificial gastric juice (1% pepsin/1% HCl, 37°C, 2 h), filtered sequentially through 200 and 100 μm sieves, and washed with PBS [46]. AD6 were collected from small intestines at 6 days postoral inoculation with 300–350 ML. NBL were obtained by culturing Ad6 in RPMI-1640 medium (Bio-Channel) under 5% CO₂ at 37°C for 16–24 h, followed by 100 μm filtration and centrifugation [47]. All parasite stages were maintained in serum- and hemin-free RPMI-1640 medium unless otherwise specified in the experimental procedures.
For infection experiments, female BALB/c mice (6–8 weeks; Yisi Laboratory Animal Technology Co.) and New Zealand rabbits (Yisi Laboratory Animal Technology Co.) were housed under 12-h light/dark cycles with ad libitum access to feed/water.
Gene cloning and plasmid construction
The Ts-Asp2 (GenBank accession number: XP_003380300.1), Ts-HRG-1 (GenBank accession number: KRY41175.1), and Ts-ATP6V0C (GenBank accession number: XP_003381793.1)genes were amplified by PCR. Ts-Asp2 was cloned and inserted into the pCold I vector (Takara Biomedical Technology) using Gibson Assembly-based seamless cloning (Biomed, Beijing, China). Ts-HRG-1 was subcloned and inserted into the following vectors: pET30a (Sangon Biotech, Shanghai, China) for prokaryotic expression, pEGFP-C1 (Sangon Biotech) for eukaryotic localization, pYES2-CT (Miaolingbio, Wuhan, China) for yeast expression and pBT3STE (Sangon Biotech, Shanghai, China) for yeast two-hybrid analysis. Ts-ATP6V0C was cloned and inserted into pET-28a, pPR3N (Sangon Biotech, Shanghai, China) and pcDNA3.1-mCherry, and Ce-HRG-4 (GenBank accession number: NP_001129857.1) was inserted into pYES2-CT (Miaolingbio, Wuhan, China). All the constructs were verified by bidirectional Sanger sequencing (Sangon Biotech, Shanghai, China) to confirm sequence integrity. S2 Table lists the oligonucleotide primers designed for PCR-based cloning procedures.
δ-Aminolevulinic acid (ALA) assay and porphobilinogen (PBG) detection
ML or E. coli (positive control) were lysed in RIPA buffer containing 1 mM PMSF and incubated on ice for 30 min. Lysates were clarified by centrifugation at 8,000 × g for 10 min, and the supernatants were collected. ML lysates or E. coli (positive control) supernatants were then incubated with 2.5–50 μg/mL δ-aminolevulinic acid (ALA; Solarbio, Beijing, China) in PBS at 37°C for 4 h. The resulting mixtures were directly mixed with Ehrlich’s reagent (1:1 v/v) [22,48], and the absorbance at 555 nm was measured using an Epoch microplate reader (BioTek, Vermont, USA). Color development (blue) was visually assessed to confirm PBG formation.
Recombinant protein expression and purification
Recombinant Ts-Asp2 (expressed from the pCold I vector) and Ts-HRG-1 (expressed from the pET-30a(+) vector) were produced in E. coli BL21 (DE3) cells. Protein expression was induced with 0.1 mM IPTG at 16°C for 20 h when cultures reached an OD600 of 0.6–0.8. The cells were harvested by centrifugation (5,000 × g, 15 min, 4°C) and lysed in buffer (20 mM Tris-HCl, 300 mM NaCl, 10 mM imidazole, pH 8.0) containing protease inhibitors. After sonication and centrifugation (12,000 × g, 30 min, 4°C), the supernatant was purified using a HisTrap HP column (Cytiva, USA) following the manufacturer’s protocol. Proteins were eluted with an imidazole gradient, dialyzed into 20 mM Tris-HCl (pH 8.0), 150 mM NaCl, and 10% glycerol, and concentrated. Protein purity was verified by SDS‒PAGE, and protein concentrations were determined spectrophotometrically. The purified proteins were stored at −80°C.
Proteolytic activity assay of rTs-Asp2
Recombinant Ts-Asp2 (20 μg) was incubated with 20 μg of myoglobin (Sigma‒Aldrich, USA) in PBS (pH 7.4) at 37°C for 8 h. Experimental groups included rTs-Asp2 with myoglobin, myoglobin alone, rTs-Asp2 alone, and rTs-Asp2 with myoglobin plus 10 μM Pepstatin A (PepA) [23]. For colorimetric analysis, an equal volume of 3,3’,5,5’-tetramethylbenzidine (TMB, Aladdin, T117927) was added to the reaction mixtures, and absorbance spectra (320–750 nm) were recorded immediately using an Epoch microplate reader (BioTek, Vermont, USA). For SDS‒PAGE validation, the reactions were terminated by the addition of 5 × SDS loading buffer (Beyotime Biotechnology, Shanghai, China), followed by boiling (10 min) and resolution on 15% polyacrylamide gels. Myoglobin degradation and rTs-Asp2 stability were directly assessed by Coomassie Brilliant Blue R-250 staining.
Hemin effects on molting
T. spiralis ML were cultured in RPMI-1640 medium (a hemin-free formulation) supplemented with Hemin (MCE, USA), which was dissolved in DMSO (Thermo Fisher, Massachusetts, USA) according to the manufacturer’s instructions, at concentrations ranging from 0–100 μM for 24 h at 37°C under 5% CO₂ [24]. After incubation, the larvae were washed with PBS, and the molting rates were quantified by light microscopy (Nikon ECLIPSE C1, 40 × objective) by counting sheathed versus unsheathed individuals across three biological replicates
ZnMP uptake under heme protein exposure
T. spiralis ML were cultured in RPMI-1640 medium containing 25 μM hemin, 6.25 μM hemoglobin (Hb; Sigma‒Aldrich, St. Louis, USA), 25 μM myoglobin (Mb; Sigma‒Aldrich, St. Louis, USA), or no additives (control) for 24 h at 37°C under 5% CO₂. After initial incubation, 50 μM ZnMP (Frontier Scientific) was added to all groups for an additional 24 h [8,14]. Larvae were then washed three times with PBS, fixed in 4% paraformaldehyde for 15 min, and mounted on glass slides. ZnMP fluorescence was quantified using an OLYMPUS FV3000 confocal microscope (excitation/emission: 561/595 nm). AD6 and NBL were assayed under identical conditions to compare stage-specific uptake efficiency. Fluorescence intensity analysis was performed using ImageJ.
Bioinformatic identification and phylogenetic analysis of HRG-1 homologs
To identify Ts-HRG-1 homologs across species, BLASTp searches were conducted against the NCBI nonredundant (nr) protein database [49]. Verified sequences from parasitic nematodes, free-living nematodes, plant-parasitic nematodes, and nonnematode parasites were compiled for phylogenetic reconstruction. Multiple sequence alignment was performed using MUSCLE, followed by Maximum Likelihood tree construction in MEGA X (Jones‒Taylor‒Thornton model, 1,000 bootstrap replicates) [50]. Additionally, pairwise sequence identities between Ts-HRG-1 and homologs from Homo sapiens, Mus musculus, Danio rerio, and C. elegans were calculated in MEGA X. Structural models of orthologous proteins were visualized in UCSF ChimeraX v1.9 with explicit annotation of transmembrane helices to assess topological conservation [51].
Molecular docking and interaction analyses
The three-dimensional structure of T. spiralis HRG-1 was obtained from the UniProt database (accession: A0A0V1BW58) [52]. Molecular docking simulations between Ts-HRG-1 and Hemin were conducted in AutoDock 4.2 with Lamarckian genetic algorithm parameters (50 independent runs, population size 150, energy evaluations 2.5 × 106) [53]. The dominant binding conformation exhibiting the lowest Gibbs free energy was selected for interaction analysis. Polar contacts (hydrogen bonds: donor‒acceptor distance ≤3.5 Å, angle ≥120°) and hydrophobic interfaces were characterized using the Discovery Studio 2021 Molecular Simulation Package. Critical binding residues were mapped through spatial occupancy analysis and visualized using the PyMOL Molecular Graphics System (v3.1.3.1) with electrostatic surface rendering [54].
Hemin binding assays
Purified HRG-1 (20 μg) was incubated with 10 μM hemin chloride in phosphate-buffered saline (PBS, pH 7.4) at 37°C for 2 h. Absorption spectra (330–750 nm) were recorded using an Epoch microplate reader (BioTek, Vermont, USA). For native PAGE analysis, samples were mixed with nonreducing loading buffer (Beyotime Biotechnology, Shanghai, China) and resolved on 8% nondenaturing polyacrylamide gels at 100 V for 90 min in Tris‒glycine running buffer (pH 8.3, without SDS) [55]. The gels were stained with Coomassie Brilliant Blue R-250 to visualize protein bands or with 3,3’,5,5’-tetramethylbenzidine (TMB) to detect heme-associated peroxidase activity [56]. Control experiments included free hemin and hemoglobin (rabbit) under identical incubation conditions.
Yeast complementation assay
The heme transport function of Ts-HRG-1 was assessed by heterologous complementation in a heme-auxotrophic yeast strain, with C. elegans HRG-4 (Ce-HRG-4) serving as a positive control. A Saccharomyces cerevisiae BY4741 strain with a HEM1 gene knockout (Δhem1), generated via CRISPR-Cas9 (pCAS plasmid) as previously described [57], was used. This strain cannot synthesize heme endogenously and requires supplementation with either δ-aminolevulinic acid (ALA, a heme precursor) or exogenous heme for growth.
The Δhem1 strain was transformed with the following plasmids: the experimental construct pYES2-CT-TsHRG1, the positive control construct pYES2-CT-CeHRG4, and the empty vector pYES2-CT as a negative control. Transformants were selected on SD medium lacking uracil (SD/-Ura) [58]. S2 Table lists the sgRNA sequences along with the oligonucleotide primers designed for PCR-based cloning procedures. For the functional assay, overnight cultures of each transformant were adjusted to an OD600 of 1.0, and 10-fold serial dilutions were prepared. A 5 µL aliquot of each dilution was spotted onto SD/-Ura solid medium plates containing a range of hemin concentrations (0, 0.25, 2.5, 10, and 100 µM) but lacking ALA. Growth was assessed after incubation at 30°C for 48–72 hours. The ability of Ts-HRG-1 to restore growth in the absence of ALA, comparable to the positive control Ce-HRG-4, was taken as direct evidence of its heme uptake activity. Protein expression from all constructs was confirmed by Western blot analysis of parallel liquid cultures using an anti-His tag antibody.
Polyclonal antibody production and immunofluorescence analysis
Recombinant T. spiralis HRG-1 (rTs-HRG-1) and ATP6V0C (rTs-ATP6V0C) was emulsified with Freund’s adjuvant and used to immunize New Zealand White rabbits. Antisera were collected after four boosts and the specific antibodies were affinity-purified using a HiTrap Protein A HP (Cytiva, USA) column. Specificity was confirmed by Western blotting. For immunofluorescence, T. spiralis ML and AD6 were fixed in 4% paraformaldehyde (15 min), permeabilized in 1% Triton X-100 for 24 h, and blocked with 5% BSA. The sections were incubated with an anti-Ts-HRG-1 rabbit polyclonal antibody (1:200) or preimmune serum (control) overnight at 4°C, followed by an Alexa Fluor 555-conjugated goat anti-rabbit IgG or Alexa Fluor 647-conjugated goat anti-rabbit IgG (1:5000, Abcam, Cambridge, UK) for 1 h at RT. Nuclei were stained with DAPI (1 μg/mL, MCE, USA). Images were acquired using an OLYMPUS FV3000 confocal microscope (AF555: Ex/Em = 561/595 nm; AF647: Ex/Em = 651/667 nm; DAPI: 405/450 nm). The fluorescence intensity of Ts-HRG-1 in ML sections was analyzed in ImageJ by outlining individual larvae and measuring the integrated density (ID) in the AF555 or 647 channels.
Subcellular localization of Ts-HRG-1-GFP in HeLa cells
The preconstructed Ts-HRG-1-GFP plasmid (pEGFP-C1 vector) was used for transfection. HeLa cells were transiently transfected with the Ts-HRG-1-GFP plasmid (2 μg) using Lipofectamine 2000 (Thermo Fisher, Massachusetts, USA) according to the manufacturer’s protocol. To assess subcellular localization, transfected cells were costained with organelle-specific markers: Golgi-Tracker Red (Beyotime Biotechnology, Shanghai, China), LysoTracker Red (Beyotime Biotechnology, Shanghai, China), and a Cell Plasma Membrane Staining Kit with DiI (Beyotime Biotechnology, Shanghai, China) for labeling the Golgi apparatus, lysosomes, and plasma membrane, respectively. The staining procedures followed the manufacturer’s instructions. Nuclei were counterstained with DAPI (Beyotime Biotechnology, Shanghai, China) for 5 min at room temperature.
Confocal images were acquired using an OLYMPUS FV3000 confocal microscope with sequential scanning to avoid spectral overlap. The fluorescence signals were captured using the manufacturer-recommended filter settings for each probe. Colocalization analysis was performed in ImageJ using the Colocalization Finder plugin [59], with Pearson’s correlation coefficient (Pr value) calculated to quantify spatial overlap between Ts-HRG-1-GFP and organelle markers.
RNA interference and functional analysis
To investigate the role of T. spiralis HRG-1 and ATP6V0C, ML were electroporated (250 V, 10 ms pulse) with 5 μM Ts-HRG-1- or Ts-ATP6V0C-specific siRNA or scrambled siRNA (control) and cultured in RPMI-1640 medium containing hemin for 24 h (37°C, 5% CO2). The molting rates were quantified by light microscopy [60].
To validate the gene silencing efficiency, hemin-treated ML were homogenized in TRIzol for RNA extraction. cDNA was synthesized using PrimeScript RT Master Mix, and qPCR was performed with SYBR Green Supermix. The primers used included Ts-HRG-1- or Ts-ATP6V0C-specific primers and 18S rRNA-specific primers. S2 Table lists the oligonucleotide sequences of the primers used in the qPCR analysis. Relative mRNA levels were calculated using the 2−ΔΔCt method. Western blotting was performed with rabbit anti-Ts-HRG-1 polyclonal serum (1:500), rabbit anti-Ts-ATP6V0C polyclonal serum (1:500) and a mouse anti-α-tubulin antibody (1:1,000, Santa Cruz Biotechnology, California, USA). Band intensities were analyzed in ImageJ and normalized to those of α-tubulin.
For in vivo infection, BALB/c mice (n = 6 per group) were orally inoculated with 200 Ts-HRG-1 RNAi-treated ML. AD6 were collected from the small intestine at 6 days post infection (dpi) and cultured in vitro for 24 h to quantify NBL production. Muscle larvae were recovered by pepsin-HCl digestion at 35 dpi, and total worm counts were recorded. Statistical analysis was conducted using GraphPad Prism 9.0.
Yeast two-hybrid interaction assay
The interaction between Ts-HRG-1 and Ts-ATP6V0C was analyzed using the membrane yeast two-hybrid (MYTH) system [61]. The bait plasmid pBT3STE-Ts-HRG-1 and prey plasmid pPR3N-Ts-ATP6V0C were cotransformed into Saccharomyces cerevisiae strain NMY51 via lithium acetate-mediated transformation. The transformants were plated on synthetic dropout (SD) medium lacking tryptophan and leucine (SD-TL) for selection. Self-activation and functional expression were assessed on SD-TLHA (lacking Trp, Leu, His, or Ade) supplemented with 5 mM or 10 mM 3-aminotriazole (3AT). Protein interactions were evaluated by growth on SD-TLHA + 3AT plates, as reconstitution of the transcription factor due to bait-prey binding activates the HIS3 and ADE2 reporter genes, enabling growth on this selective medium. Positive (pNubG-Fe65/pTSU2-APP) and negative (pPR3N/pTSU2-APP) controls were included. Yeast cultures were incubated at 30°C for 4 days, and colony growth was documented.
Protein interaction and colocalization analysis
HEK293T cells were cotransfected with pEGFP-C1-TsHRG1 and pcDNA3.1-mCherry-TsATP6V0C using Lipofectamine 3000 (Thermo Fisher, Massachusetts, USA) at a 1:1 plasmid ratio. After 24 h, the cells were lysed in RIPA buffer (Beyotime, P0013B) supplemented with 1 × protease inhibitor cocktail (Beyotime Biotechnology, Shanghai, China) on ice for 30 min. The lysates were subsequently centrifuged (12,000 × g, 15 min, 4°C) to remove debris, and the supernatants were incubated with anti-GFP agarose beads (AlpaLifeBio, Shenzhen, China) for 4 h at 4°C. The beads were washed four times with TBS-T (20 mM Tris, 150 mM NaCl, 0.1% Tween-20), and the bound proteins were directly mixed with 1 × SDS loading buffer and boiled at 95°C for 10 min. The immunoprecipitates and whole-cell lysates (WCLs) were resolved by SDS‒PAGE, transferred to PVDF membranes, and probed with anti-GFP (1:5,000, Affinity, USA) and anti-mCherry (1:1,000, Affinity, USA) antibodies.
For colocalization studies, transfected cells on poly-L-lysine-coated coverslips were fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton X-100, and mounted with ProLong Diamond Antifade Mountant containing DAPI. Confocal images were acquired using a Zeiss LSM 980 microscope (GFP: 488/550 nm; mCherry: 594/660 nm; DAPI: 405/450 nm). Colocalization was analyzed in ImageJ using the JACoP plugin to calculate the Pearson correlation coefficient (Pr value).
In vitro bafilomycin A1 treatment and HRG-1 mRNA expression in T. spiralis
T. spiralis AD6, NBL, and ML were cultured in DMEM supplemented with various concentrations of BafA1 for 24 h at 37°C under 5% CO2. After treatment, the parasites were homogenized in TRIzol reagent, and total RNA was extracted for cDNA synthesis. Ts-HRG-1 mRNA levels were quantified by qPCR using SYBR Green Supermix and normalized to 18S rRNA via the 2−ΔΔCt method. For functional validation, BafA1-treated or Ts-HRG-1-RNAi larvae were incubated with ZnMP under standard conditions. The ZnMP fluorescence intensity was analyzed as previously described.
In vivo bafilomycin A1 treatment of T. spiralis-infected mice
BALB/c mice (6–8 weeks old, n = 6 per group) were orally infected with 200 T. spiralis ML and divided into seven groups. Three groups received daily oral administration of BafA1 (0.1, 0.5, or 1 mg/kg, MCE, USA) from 0–7 days post infection (dpi); another three groups were treated with the same BafA1 doses from 21–29 dpi. The control mice received PBS (5% DMSO vehicle) during both treatment windows. At 6 and 35 dpi, the mice were euthanized for parasite collection. AD6 were isolated from the intestinal contents at 6 dpi (37°C saline incubation), and the muscle larva (ML) burden was assessed at 35 dpi using pepsin-HCl digestion. Concurrently collected diaphragm tissues underwent histological processing: they were fixed in 4% paraformaldehyde, paraffin-embedded, sectioned at a thickness of 5 μm, and stained with H&E for microscopic analysis (Nikon Eclipse E100).
Supporting information
S1 Fig. Developmental stage-specific mRNA expression in T. spiralis and hem1 knockout PCR validation in BY4741 yeast.
(A) Muscle larvae (ML), newborn larvae (NBL), adult worms at 3 days post infection (AD3), and adult worms at 6 days post infection (AD6) were analyzed. T. spiralis 18s rRNA was used as the internal reference gene. The data are presented as the means ± SEMs. Statistical significance was determined by one-way ANOVA (n = 4). (B) Schematic of primer design for HEM1 deletion via homologous recombination. The target HEM1 sequence (yellow) is flanked by 5’ and 3’ homologous arms (brown). The primers used included Flanking-F (binding upstream of the 5’arm), Flanking-R (binding downstream of the 3’arm), and Inside-F (binding within the HEM1 target sequence). (C) PCR verification of the wild-type (BY4741) and Δhem1 strains. Lane 1: WT with Flanking-F + Flanking-R (native HEM1 locus, 2600 bp); Lane 2: Δhem1 with Flanking-F + Flanking-R (deletion band, 1000 bp); Lane 3: WT with Inside-F + Flanking-R (1600 bp fragment); Lane 4: Δhem1 with Inside-F + Flanking-R (no product). M: 2000 Plus II DNA ladder. (D) Immunofluorescence localization of Ts-HRG-1 in AD6 using a Ts-HRG-1 polyclonal antibody (red; Alexa Fluor 647-conjugated secondary antibody) and DAPI (blue, nuclei). The negative control serum (preimmune serum with identical secondary antibodies) showed no specific signal. Ts-HRG-1 localized in the stichosome. Scale bars: 100 μm.
https://doi.org/10.1371/journal.ppat.1014042.s001
(TIF)
S2 Fig. siRNA screening and optimal concentration determination were performed for Ts-HRG-1 knockdown in T. spiralis larvae.
(A) Validation of the transfection efficiency of CY3-labeled negative control (NC-siRNA) in muscle larvae (ML). (B) Dose‒response evaluation of the effects of selected siRNA-2 (0–6 μM) on Ts-HRG-1 mRNA levels. Statistical analysis was performed using one-way ANOVA (n = 3). (C) Ts-HRG-1 mRNA expression analysis by qPCR following treatment with three siRNA candidates (siRNAs 1–3); 18s rRNA was used as an internal control. Statistical analysis was performed using one-way ANOVA (n = 3). (D) Western blot verification of Ts-HRG-1 protein knockdown; α-tubulin was used as a loading control. (E) Grayscale quantification of the WB bands (n = 3). Statistical analysis was performed using one-way ANOVA.
https://doi.org/10.1371/journal.ppat.1014042.s002
(TIF)
S3 Fig. Ts-HRG-1 does not interact with Ts-V-ATPase_V1_A and Ts-ATP6V0C knockdown is confirmed by RNAi.
(A) Yeast two-hybrid assay. AD-Ts-V-ATPase_V1_A and BD-Ts-HRG-1 were cotransformed into yeast. The lack of growth in the selective medium confirmed that there was no interaction, with the controls showing no autoactivation (AD/BD empty vectors). (B) Subcellular localization in HeLa cells. Ts-HRG-1-GFP (green) and Ts-V-ATPase_V1_A-mCherry (red) showed no colocalization intracellularly. Scale bar: 5 μm. (C) Colocalization analysis. Pearson’s correlation coefficient (Rr = 0.153) and distance-dependent intensity profile analysis confirmed the absence of spatial proximity between Ts-HRG-1 and Ts-V-ATPase_V1_A. (D) Coimmunoprecipitation (Co-IP). Lysates from HeLa cells coexpressing Ts-HRG-1-GFP and Ts-V-ATPase_V1_A-mCherry were immunoprecipitated with anti-GFP. Ts-V-ATPase_V1_A-mCherry was not detected in the IP fractions (anti-mCherry blot), confirming that there was no physical interaction. No signal was observed in the controls. (E) Ts-ATP6V0C mRNA expression analysis by qPCR following treatment with three siRNA candidates; 18s rRNA was used as an internal control. Statistical analysis was performed using one-way ANOVA, (n = 3). (F) Western blot verification of Ts-ATP6V0C protein knockdown; α-tubulin was used as a loading control. (G) Grayscale quantification of the WB bands (n = 3). Statistical analysis was performed using one-way ANOVA.
https://doi.org/10.1371/journal.ppat.1014042.s003
(TIF)
S4 Fig. Effects of BafA1 and Ts-HRG-1 knockdown on larval survival and molting in T. spiralis.
(A) ML molting responses following 24h treatments with: CON, Ts-HRG-1 RNAi, Ts-ATP6V0C RNAi and BafA1 (7.5 μg/mL). (B) Viability of NBL cultured with increasing BafA1 concentrations (0–7.5 μg/mL) for 24 hours. (C) Survival of AD6 treated with BafA1 (0–25 μg/mL) for 24 hours. (D) Molting status of ML exposed to stepwise BafA1 concentrations (0–10 μg/mL) for 24 hours.
https://doi.org/10.1371/journal.ppat.1014042.s004
(TIF)
S1 Table. Proteomic profiling of Ts-HRG-1-associated complexes via co-immunoprecipitation and LC-MS/MS.
https://doi.org/10.1371/journal.ppat.1014042.s005
(XLSX)
S2 Table. PCR primers for gene cloning, heterologous expression, Y2H, Cas9 KO validation, qPCR and RNAi.
https://doi.org/10.1371/journal.ppat.1014042.s006
(XLSX)
Acknowledgments
We would like to thank the State Key Laboratory for Diagnosis and Treatment of Severe Zoonotic Infectious Diseases for providing access to instruments. We thank Researcher Xinrui Wang for performing the tissue sectioning. We also thank Researcher Li Yang and Researcher Yuanyuan Zhang for assisting with the operation of instruments.
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