C57BL/6 mice were purchased from Envigo (Germany) or bred in-house and maintained under specific pathogen-free conditions at the Institute of Medical Microbiology, Immunology, and Hygiene (MIH), Animal facility, TU Munich. All experiments were performed in accordance with and approved by local government authorities of Regierung von Oberbayern (Az. 55.2-1-54-2532-145-17). Cyst vesicular fluid (CVF) was prepared as described previously (3). T. solium cysts were collected from the muscle tissue of a heavily infected pig at the University of Zambia School of Veterinary Medicine (ethical approval Ref. 2018-Mar-002/0005948). Following collection, the cysts were washed with phosphate-buffered saline (PBS) (Merck, Cat. No. D8662). For CVF preparation, the cysts were transferred to a sterile dish, cut open with a scalpel, and the released fluid was collected. The collected fluid was sterile filtered through 0.45 µm size filters, centrifuged at 15.000 g for 60 min at 4 °C. Then, materials were immediately treated with Phenylmethylsulphonyl fluoride (PMSF, final concentration 5 mM) and leupeptin (final concentration 2.5 µM), and aliquoted at a concentration of 1 mg/ml in PBS. All prepared materials were tested for endotoxin (levels < 0.05 EU/ml) and stored at -80 °C until use.

Human plasma from healthy controls and persons with NCC was examined for soluble apoptosis-inducing mediators. The patients were recruited within the TOPANA study, a prospective cohort study implemented in rural Tanzania. The study received ethical approval by the Klinikum Rechts der Isar (Technical University of Munich, Germany) Ethical Committee (Reference 33/19S) and by the National Ethics Health Research Committee (NatREC) of Tanzania (Reference NIMR/HQ/R.8c/Vol.I/1808 for TOPANA and NIMR/HQ/R.8a/Vol.IX/2597 for SOLID). The protocol and study design of the study have been published elsewhere (28, 29). In brief, patients were recruited at three district hospitals (Tosamaganga, Chunya, Vwawa). NCC diagnosis was done in July 2021 according to the latest Del Brutto criteria, using computer tomography (CT) scan and laboratory testing (LDBio cysticercosis Western Blot IgG, apDia Cysticercosis Ag-ELISA) (30, 31). Patients with NCC were classified as symptomatic through the presence or history of severe progressive headaches and/or epileptic seizures and having parenchymal and extraparenchymal NCC with active lesions and mostly degenerating/calcified cysts (28). Asymptomatic NCC patients had no history of epileptic seizures and/or severe progressive headaches but had a mix of viable and degenerating cysts, with most being viable. Participants were tested for HIV by serology, Mycobacterium tuberculosis in the sputum, Strongyloides spp. in the feces, and underwent fundoscopy for the exclusion of ocular NCC according to the IDSA/ASTMH recommendations (32). Blood samples were taken from the study participants and after giving informed consent. Peripheral blood mononuclear cells (PBMC) were isolated as previously described (33). Plasma was separated from the cellular components of the blood using density gradient centrifugation (700 x g, 25 min, 20 °C), and stored at -80 °C until usage. The cohort included NCC non-infected Tanzanian individuals as controls (group 1), NCC asymptomatic individuals (group 2) and symptomatic NCC patients with epilepsy and/or severe progressive headaches (group 3). The participants included in this study were age and sex-matched (see detail in Table 1 ).

The study was approved by the local ethical committee of the Technical University of Munich (Reference: 215/18S), and all individuals included in the study consented to enrollment. Human PBMC were isolated from whole blood samples of healthy volunteers from mixed genders using density gradient centrifugation and stored in liquid nitrogen as described in (33). Human unlabeled monocytes were purified from freshly isolated human PBMCs using the Miltenyi Pan Monocyte Isolation Kit, human (Miltenyi, Germany, Cat. No. 130-096-537), according to the manufacturer’s protocol. The purity of monocytes was routinely ≥ 80%. 2.5x10⁵ cells per well were plated in 96 U-bottom well plates and stimulated with CVF (0.01 – 5 µg/ml) and incubated for 0–72 h at 37 °C, 5% CO₂. Staurosporine (TOCRIS, Bio-techne, Cat. No. 1285) (100 nM or 500 nM) was used to induce apoptosis. After incubation period, surface markers (CD3, CD4, CD8, CD14, CD16 (all antibodies from Biolegend: Cat. No. 317331, 317433, 344723, 325607, 302025), were analyzed by flow cytometry on a CytoFLEX S (Beckman Coulter). Early apoptotic (Annexin V⁺/PI^-, late apoptotic (Annexin V⁺/PI⁺), and necrotic (Annexin V^-/PI⁺) cell populations were determined by Annexin V/PI kit (Biolegend, Cat. No. 640914). For intracellular staining, cells were fixed and permeabilized with BD Cytofix/Cytoperm (BD Biosciences, Cat. No. 554714) for detection of active caspase 3 (BD Pharmingen, Cat. No. 570332). FAM FLICA™ Caspase-8 and Caspase-9 Kits (Bio-Rad, Cat. No. ICT099, ICT912) were used for detection of active caspases 8 and 9.

To determine the role of proteins in the CVF in the induction of apoptosis on PBMCs, active proteins in the CVF were inactivated by heat-inactivation using 56 °C for 1h, 56 °C for 3h, 99 °C for 1h, and 99 °C for 3h and 10µg/ml Proteinase K (Sigma-Aldrich, Cat. No. P2308) digestion overnight at 37 °C, 5% CO₂. SDS page and Coomassie staining were performed to confirm effective protein inactivation. CVF was additionally treated with the cysteine protease inhibitor E-64 (Sigma-Aldrich, Cat. No. E3132) 10µM for 1 hour to inhibit cysteine proteases (e.g. Cathepsins). After protein inactivation and protease inhibition, PBMCs were stimulated with 2.5 µg/ml pretreated CVF for 72 h, and apoptosis induction was evaluated using Annexin V/PI. Furthermore, CVF was separated according to molecular size using 30 kDa column fractionation (GE Healthcare, Cat. No. 17VS50039A). Flow through and retarded CVF fractions (< 30 kDa, > 30 kDa) were used for stimulation of PBMCs and apoptosis was detected.

To measure mitochondrial damage, PBMCs were stained with Tetramethylrhodamine, ethyl ester, perchlorate (TMRE) (ThermoFisher Scientific, Cat. No. T669) and MitoTracker following CVF stimulation. For TMRE staining, TMRE was diluted to 10nM in RPMI (Merck, Cat. No. R0883) (10% FCS (Merck, Cat. No. S0615) and 1% Penicillin/Streptomycin (Merck, Cat. No. P4333) and cyclosporin H (Enzo, Cat. No. 83602-39-5) (final concentration 0.5 µM), and incubated with CVF-treated PBMCs for 30 minutes at 37 °C to assess the loss of mitochondrial transmembrane potential. For MitoTracker staining, PBMC were incubated with MitoTracker Deep Red FM (ThermoFisher Scientific, Cat. No. M46753) and MitoTracker Green FM (ThermoFisher Scientific, Cat. No. M7514) in PBS for 60 minutes at 37 °C to evaluate the mitochondrial potential and mass loss, respectively (34).

Porcine blood samples were obtained from freshly slaughtered pigs at a local slaughterhouse. The blood was collected in 500 ml bottles containing 1g EDTA (Ethylenediaminetetraacetic acid) anticoagulant. Samples were stored at 4 °C within an hour of collection and processed within 24 hours.

Porcine PBMCs were isolated using SepMate™ isolation tubes (Stemcell Technologies, Cat. No. 85460) and Histopaque^®-1077 (Merck, Cat. No. 10771). The blood was first diluted 1:1 with sterile PBS/2% fetal calf serum (FCS) (Merck, Cat. No. S0615), and then layered onto Histopaque within centrifugation tubes. PBMCs were separated using a density gradient. Centrifugation was performed at 700xg for 25 min with the lowest acceleration and break. The PBMC layer was collected using a single-use pipette. PBCMs were washed twice with PBS + 2% FCS. Remaining erythrocytes were lysed using 0.5 ml erythrocyte lysis buffer (0.01 M KHCO₃, 0.155 M NH₄Cl, 0.1 mM EDTA) per 10 ml blood for 4 min at RT.

PBMCs were resuspended in RPMI (Merck, Cat. No. R8758) supplemented with 10% FCS (Merck, Cat. No. S0615), 100 U/ml penicillin, 100 µg/ml streptomycin (Merck, Cat. No. P4333). Porcine monocytes were separated from freshly isolated PBMCs using the autoMACS separator (Miltenyi Biotec). PBMCs from each animal were rebuffered in autoMACS running buffer (Miltenyi Biotec, Cat. No. 130-091-221). Cells were labeled with CD14 MicroBeads (Miltenyi Biotec, clone Tük4, Cat. No. 130-050-201) and separated according to the manufacturer’s instructions. CD14⁺ monocytes were pelleted and resuspended in RPMI. Trypan blue staining and automated cell counting were used to determine the number of viable monocytes and PBMCs.

2.5–5 x 10⁵ cells of porcine PBMCs and 3 x10⁵ cells of porcine monocytes were plated per well and stimulated with 2.5 µg/ml CVF for 0–72 h at 37 °C and 5% CO₂. After incubation, porcine-specific anti-CD4α (clone 74-12-4, mouse IgG2b, inhouse + anti-mouse IgG2b, clone R12-3, BD Biosciences, Cat. No. 743177), anti-CD8α (clone 11/295/33, mouse IgG2a, inhouse + anti-mouse IgG2a, clone RMG2 A-62, BioLegend, Cat. No. 407117), anti-CD3e (clone BB-23-8E6-8C8, BD Biosciences, Cat. No. 743177), anti-CD172a (clone 74-22-15, mouse IgG1, inhouse + anti-mouse IgG1, clone X-56, Miltenyi, Cat. No. 130-096-593) and CD14 (clone Tük4, Bio-Rad, Cat. No. MCA1568P647) antibodies were used to characterize cell populations on an Attune NxT Flow cytometer (Thermo Fisher Scientific). Induction of apoptosis was detected with Annexin V/PI kit (Invitrogen, Cat. No. 13242) and Staurosporine (500 nM) (Sigma-Aldrich, Cat. No. S6942) served as a positive control.

Ten 2 - 4-year-old pigs (6 females and 4 males, 4 uninfected and 6 naturally infected with T. solium cysticercosis, were purchased from rural rearers. The animals were diagnosed for cysticercosis by tongue examination. Pigs were humanely euthanized at the Faculty of Veterinary Medicine and Zootechnics in accordance with Mexican official standard NOM-033-SAG/ZOO-2014. Blood samples were collected immediately after euthanasia, and brain tissue was dissected. Brain samples were fixed in neutral buffered 10% formalin for 48 hours. After dehydration, brain tissue was embedded in paraffin blocks.

Serial sections of 2µm slices were mounted on Superfrost (HE) or Superfrost Plus (IHC) slides using a rotary microtome (ThermoFisher Scientific, Cat. No. HM355S). The sections were stained with Hematoxylin-Eosin to evaluate general pathological changes using standard protocols. In brief, the wax was removed with xylene and then passed through different changes of alcohol to remove the xylene. After rehydration with distilled water, the sections were stained with hematoxylin for 8 minutes. Following rinsing with tab water, counterstaining with 1% alcoholic eosin was performed for 4 minutes. Slides were dehydrated by passing through different changes of alcohol and cleared with xylene.

To visualize and localize cell populations in the granulomatous regions surrounding T. solium cysts in the pig brains, immunohistochemical (IHC) staining was performed using an autostainer (Bond RXm system, Leica). In brief, the sections were deparaffinized with the Leica Kit de-wax (Leica, Cat. No. AR9222) and rehydrated by washing with decreasing alcohol concentrations. Epitope retrieval was done with heat induction using citrate buffer (pH=6) for 20 minutes (40 minutes for anti-CD79a). Endogenous peroxidase activity was blocked by incubating the sections with 3% hydrogen peroxidase (Leica) for 5 minutes, and non-specific binding sites were blocked with normal goat serum (Abcam, Cat. No. ab138478). Primary antibodies were then added to the sections for 15 minutes at room temperature (anti-CD3 (Cellmarque, Cat. No. MRQ39), anti-CD79a (ThermoFisher, Cat. No. HM57), anti-Iba1 (poly, Wako, Cat. No. 019-19741), and anti-Caspase 3 (Cell Signaling, Cat. No. 5A1E)). Antibody detection was done with a polymer refine detection kit (Leica, Cat. No. DS9800) and Diaminobenzidine (DAB) (Merck, Cat. No. D12384) was used for visualization. The sections were counterstained with hematoxylin for 10 minutes and dehydrated with increasing alcohol concentrations and xylene. Pertex mounting medium (Histolab, Cat. No. SEA-0100-00A) was used to mount the sections. All antibodies were tested for cross-reactivity in pigs and adequate positive controls were used in each staining run.

Histology was evaluated under the supervision of a board-certified veterinary Pathologist (K.S.). For digital image analysis, the software QuPath [v0.5.1] was used. To quantify CD3⁺ and CD79a⁺ cells, 8–10 rectangles (1,25 mm x 0,875 mm) covering the area around the cysts were drawn for automated cell counting, and positive and negative cells were counted automatically with QuPath software.

Newborn mice (P1-P4) were decapitated, and the olfactory bulbs and the cerebellum were removed, as well as the meninges and choroid plexus. Tissue was dissociated and filtered through 100 µl cell strainer. Cells were centrifuged for 5 min at 1500 RPM at 4 °C. The cell pellet was resuspended in 5 ml of DMEM (Dulbecco’s Modified Eagle Medium) high glucose (Gibco, Cat. No. 11965092) (10% FCS, 0.5% Ciprofloxacin) and plated on a T25 culture flask, which was previously coated overnight with 5 ug/ml Poly-L-Lysine (Merck, Cat. No. P8920). The T25 culture flasks were incubated for 12 days with repeated medium exchange at 37 °C in a 5% CO₂ humidified incubator. 48h before further analysis, 20ng/ml of M-CSF (Macrophage colony-stimulating factor) (Pepro-Tech, Cat. No. 315-02) was added to the cultures. The cultures were trypsinized in 0.05% Trypsin-EDTA for 15 minutes at 37 °C, washed with medium, scraped, and seeded to 96-well flat-bottom plate (Thermo Fisher Scientific), with a density of 1 x 10⁵ cells per well. After resting overnight, cells were stimulated with CVF (2.5 µg/ml) for 0 - 72h, and, either lysed for qPCR (quantitative polymerase chain reaction), or proceeded for active caspases detection with intracellular staining for cleaved caspase 3 (Bio-techne, R&D Systems, Cat. No. IC835G) and with FLICA caspase assays for active caspase 8 (Bio-Rad, Cat. No. ICT099) and 9 (Bio-Rad, Cat. No. ICT960) using flow cytometry.

Commercially available ELISA kits were used according to the manufacturer’s instructions. Human TNFα ELISA (R&D Systems, Cat. No. DTA00D) and FasL (R&D Systems, Cat. No. DFL00B) were used to evaluate the levels of both apoptosis mediators in the culture supernatants of CVF-stimulated PBMCs from healthy individuals and in plasma samples from NCC patients and controls from Tanzania.

An intracellular fluorometric reactive oxygen species (ROS) assay (Cell Biolabs, Cat. No. STA-342) was used according to the manufacturer’s instructions to measure the intracellular ROS activity within the CVF-stimulated PBMCs. Briefly, human PBMCs were stimulated with CVF 2.5 µg/ml. After 72 h, cells were washed twice with PBS, and 100 µl of 1X cell-permeable fluorogenic probe Dichlorodihydrofluorescin diacetate (DCFH-DA) were added to the wells and incubated at 37 °C, 5% CO₂. After 45 min, cells were washed twice with PBS, and the fluorescence intensity, proportional to intracellular ROS activity to the DCF, was measured with a CLARIOstar Plus plate reader (BMG LABTECH) at 480 nm/530 nm. 100 µM and 1000 µM H₂O₂ were added to the cells as positive controls.

For microglia RNA extraction, cells were treated with RLysis buffer containing 1% β-mercaptoethanol, and RNA was extracted using the Extractme Total RNA Micro Spin kit (BLIRT, Cat. No. EM31.1-050) according to manufacturer’s information. RNA was reversely transcribed to cDNA with the SuperScript™ IV VILO mix (ThermoFisher Scientific). qRT-PCR was performed with ABsolute qPCR SYBR Green (Thermo Fisher Scientific) at the LightCycler 480 (Roche). ΔΔCt was used to calculate relative gene expression and differences. The oligonucleotide sequences used for the qRT-PCR are detailed in Table 2 .

The mass spectrometry analysis of T. solium CVF was performed as previously described (3). The retrieved data were analyzed using the WormBase Parasite database for T. solium (Version: WS282, October 2022) to identify apoptosis-related molecules in the CVF.

For statistical analysis, the Prism software (Version 10.1.2) (GraphPad Software, LLC, San Diego, CA, USA) was used. For direct comparisons between two groups, a Mann-Whitney test was employed. For more than two groups, we applied a Kruskal-Wallis test followed by Dunn’s multiple comparison test and a two-way ANOVA test followed by Tukey’s multiple comparison test if more time points were observed. P values < 0.05 were considered statistically significant.

Our previous findings indicated that cyst-derived vesicular fluid (CVF) from T. solium cysts promotes apoptosis in alveolar macrophages (3). Here, we extended our investigation to assess the pro-apoptotic effects of CVF on various peripheral immune cell populations. To evaluate the impact of CVF on PBMCs, we analyzed both early apoptotic (Annexin V⁺/PI^-) and late apoptotic (Annexin V⁺/PI⁺) cell populations by Annexin V/PI staining. Our data revealed that CVF induced significant apoptosis in PBMCs ( Figure 1A ; Supplementary Figure S1 ). This effect was concentration-dependent and predominantly restricted to apoptosis rather than necrosis, affecting both CD3⁺ T cells ( Figure 1B ) and CD3^- cells (non-T cells) ( Figure 1C ) in a dose-dependent manner ( Supplementary Figure S2 ). Notably, CD3^- immune cells exhibited a more pronounced apoptotic response compared to CD3⁺ T cells ( Figure 1D ). Furthermore, the apoptotic effect intensified over time, particularly in the late apoptotic CD3⁺ and CD3^- cell populations ( Figure 1E ) as well as in CD3⁺CD4⁺ and CD3⁺CD8⁺ T cell subsets ( Figure 1F ). Interestingly, while CD3^- and CD8⁺ early apoptotic cells decreased over time, late apoptotic populations increased, suggesting a dynamic progression from early to late apoptosis upon CVF exposure ( Figures 1E, F ). Given that CD3^- cells were significantly affected by CVF-induced apoptosis, we further investigated monocyte subpopulations to determine their susceptibility, since monocytes are also the second largest cell population in PBMCs after lymphocytes (35). Monocytes were purified from PBMCs using magnetic-activated cell sorting (MACS) and categorized into classical (CD14⁺CD16^-) and non-classical (CD14^dimCD16⁺) monocytes. Our findings confirmed the pro-apoptotic potential of CVF, demonstrating a time-dependent shift from early to late apoptosis in both monocyte subsets ( Figure 1G ). Of note, monocytes were very sensitive to spontaneous cell apoptosis, especially in the first 24 h of culture as reported elsewhere (36, 37).

In summary, our findings demonstrate that degenerating T. solium cysts actively drive apoptosis, rather than non-apoptotic death (necrosis), in peripheral immune cell populations. This effect was particularly pronounced in CD8⁺ T cells and monocytes. These results underscore the selective targeting of immune cells by degenerating cyst-derived molecules, further implicating their role in NCC-associated systemic immune inflammation and dysregulation.

Pigs serve as natural hosts for T. solium cysts and their infection mirrors in many ways human NCC in terms of immunopathological responses (9, 15, 38). To extend our findings on CVF-induced apoptosis in host immune cells, we examined the effects of CVF on porcine peripheral immune cells and assessed its relevance to the CNS by analyzing immune responses in naturally infected pigs harboring granulomas with intact and degenerating T. solium cysts in the brain. Porcine PBMCs were treated with CVF under the same conditions as human PBMCs, and apoptosis was assessed in CD3⁺ T cells, including major subsets (CD4⁺CD8α^-, CD4^-CD8α⁺, CD4⁺CD8α⁺), as well as in CD172a⁺ monocytes ( Figures 2A-C ). Unlike humans, pigs possess a unique population of CD4⁺CD8α⁺ double-positive T cells, which are thought to originate from CD4⁺ helper T cells that acquire CD8α expression following antigen recognition (39, 40). Similar to human cells, early apoptosis in porcine PBMCs declined over time, promoting late stages of apoptosis, which peaked at 24 h post-CVF treatment, with a delayed response in CD4^-CD8α⁺ T cells ( Figure 2A ). However, in contrast to human T lymphocytes, CVF-dependent necrosis increased significantly over time in all porcine lymphocyte populations ( Figure 2B ). In addition, late apoptotic cells steadily increased in the CD3^- compartment over time, while CD3⁺ lymphocytes displayed a reduction in late apoptosis after 48 h of treatment ( Figures 2A–B ). Porcine monocyte subsets are primarily identified by CD172a, CD14, and CD163 expression (41). Given that all porcine monocytes express CD172a (41), we focused on CD172a⁺ monocytes for better comparability with human monocytes. Similar to porcine T lymphocytes, CD172a⁺ monocytes showed a peak in late apoptosis at 24 h post-CVF treatment, which declined at later time points ( Figure 2C ). Necrosis also increased over time in monocytes, although this change was not statistically significant ( Figure 2C ). These results suggest that while CVF predominantly induces apoptosis in human immune cells, it promotes a substantial degree of necrosis in porcine immune cells. This indicates a species-specific variation in the mechanisms of degenerated cyst-induced cell death.

In humans, the most severe clinical manifestations of NCC, including recurrent seizures and progressive headaches, are often associated with degenerating cysts encased within granulomatous structures in the brain (8). To investigate the immunopathological relevance of CVF-induced apoptosis in vivo, we first analyzed immune cell infiltrates in brain sections from five NCC-infected pigs, comparing regions surrounding degenerating “inflammatory” T. solium cysts to those surrounding viable “non-inflammatory” cysts (38, 42). Employing a combination of HE staining with spatial analysis of CD3⁺ T and CD79⁺ B lymphocytes and the Iba1⁺ activated macrophages/microglia, we identified 18 cysts classified into three stages according to the classification used by Gutierrez and Singh (38, 42). In brief, viable cysts (noninflammatory) were classified as cysts with a well-stainable parasite with intact scolex and bladder wall and no edema or inflammation in the surrounding tissue. Degenerating cysts (inflammatory) had an intact bladder wall and scolex, but nuclear disintegration and a mixed inflammatory or granulomatous reaction around the cysts were observed. Although calcified cysts were also detected, no defined structure was found in these cysts. Of the 18 cysts, 1 was a viable cyst (noninflammatory), 11 were degenerating (inflammatory), and 6 were calcified or unclassifiable cysts. Out of the 11 inflammatory cysts, 6 had clearly visible parasites with scolex and are thus included in the present investigation. Representative images of a noninflammatory and an inflammatory cyst from the same pig are shown in Figure 2D . Immunohistochemical staining for CD3, CD79a, and Iba1 revealed a distinctive localization of immune cells within granulomatous structures. Iba1⁺ activated macrophages/microglia were predominantly located on the inner side of the granuloma, adjacent to degenerating cysts ( Figures 2E, H ). In contrast, CD3⁺ T cells and CD79a⁺ B cells were primarily found in the outer layers of the granuloma ( Figures 2E, G ). Quantitative analysis indicated an approximately three-fold increase in CD3⁺ T cell infiltration ( Figures 2E, F ) and a ten-fold increase in CD79a⁺ B cell infiltration ( Figures 2E, G ) around inflammatory cysts compared to non-inflammatory cysts ( Figures 2E, G ; Supplementary Figures S3A, B ).

These findings provide compelling evidence that cyst-derived vesicular fluid products contribute to peripheral immune cell apoptosis and inflammation. Moreover, cyst degeneration is associated with significant immune cell infiltration in the CNS, highlighting its potential role in NCC immunopathogenesis.

Several signaling pathways contribute to apoptosis, with the extrinsic pathway initiated by death receptors, and the intrinsic pathway that occurs through the mitochondria, being the main ones (43). To determine which apoptotic pathways are activated by CVF, we assessed caspase activity, given that caspase 3 serves as a key effector in both apoptotic mechanism pathways (44). CVF treatment led to a significant increase in CD3⁺ T cell-expressing active caspase 3, including both CD4⁺ and CD8⁺ subsets, compared to untreated and staurosporine-treated positive controls ( Figure 3A ). To distinguish between the extrinsic and intrinsic apoptosis pathways, we evaluated the activation of initiator caspases for both pathways: caspase 8 (extrinsic) and caspase 9 (intrinsic) (44). Using fluorochrome-labeled inhibitor of caspase (FLICA) assays, we observed significant activation of both caspase 8 and caspase 9 in CD3⁺, CD4⁺, and CD8⁺ T cells, as well as in CD3^- non-T cells ( Figure 3B ). However, in monocytes, only caspase 9 was activated, with no detectable caspase 8 activity ( Figure 3B ). Of note, CVF treatment shifted the entire monocyte compartment toward CD16⁺-expressing monocytes. Despite the involvement of both pathways in lymphocytes, caspase 9 activation was significantly higher than caspase 8, suggesting a predominant role of the intrinsic mitochondrial pathway in CVF-induced apoptosis. An analogous pattern was found in microglia, where CVF-induced caspase 3 activation ( Figure 3C ) correlated with a progressive increase in cleaved caspase 9 levels, whereas cleaved caspase 8 decreased sharply over time ( Figure 3D ).

To determine whether CVF stimulation modulated gene expression of caspases, we performed qRT-PCR analysis ( Figure 3E ). Caspase 9 expression increased over time, although not statistically significant, while caspase 3 and caspase 8 expression declined. Additionally, we analyzed key mediators influencing caspase 8 and caspase 9 activation. TNFα and FasL, two primary inducers of the extrinsic apoptotic pathway (23), showed no significant changes in mRNA levels. Similarly, the expression of Bcl-2, an anti-apoptotic protein (45), remained unchanged. In contrast, Bid exhibited a marked decline. Of note, Bid is a pro-apoptotic protein and active inducer of mitochondrial outer membrane permeabilization that bridges caspase 8 activation to the extrinsic pathway (45). This suggests that the caspase 8 - Bid axis is not a major contributor to CVF-induced apoptosis. To explore the relevance of these findings in vivo, we stained brain tissue sections from NCC-infected pigs for caspase 3. Apoptotic caspase 3⁺ cells were detected exclusively in areas surrounding degenerating inflammatory cysts ( Figure 3F ).

Taken together, these findings indicate that CVF predominantly activates the mitochondrial apoptosis pathway via caspase 9, leading to caspase 3 activation and immune cell death. Furthermore, the presence of apoptotic cells in the CNS near degenerating cysts underscores the potential role of CVF-induced apoptosis in NCC immunopathogenesis.

Our data demonstrate that T. solium CVF preferentially activates the intrinsic apoptosis pathway to promote cell death. A hallmark of this pathway is mitochondrial dysfunction, characterized by the release of cytochrome C from the mitochondria and a loss of mitochondrial transmembrane potential (44, 46). We thus next assessed the mitochondrial integrity following CVF exposure using the TMRE and MitoTracker assays in human PBMC. As depicted in Figure 4A , CVF-treated cells exhibited a significant reduction in TMRE fluorescence and signal, indicating mitochondrial transmembrane potential loss and depolarization across CD3⁺ T cell and CD3^- non-T cell compartments, as well as CD4⁺ and CD8⁺ T cells ( Figure 4B ), and CD16⁺ monocytes ( Figure 4C ). MitoTrackers are cationic fluorescent probes that accumulate inside mitochondria due to the high mitochondrial transmembrane potential and are useful tools to visualize mitochondrial mass, integrity, and morphology (47). We used MitoTracker Deep Red (MDR) and MitoTracker Green (MG) to assess mitochondrial membrane potential and mitochondrial mass, respectively (34). The MDR/MG ratio is considered a parameter for mitochondrial activity per mass (34). As Figures 4D, F show, we observed a marked decrease in the MDR/MG ratio, suggesting an impairment of mitochondrial activity relative to mass for the lymphocyte ( Figures 4D, E ) and CD16⁺ monocyte ( Figure 4F ) populations treated with CVF and Staurosporine as compared to the untreated controls. This was accompanied by an increase in MDR/MG^lo populations ( Figure 4G ), further supporting mitochondrial dysfunction.

Mitochondrial impairment involves many upstream mediators and cellular stress signals in the induction of the intrinsic apoptosis pathway such as mitochondrial DNA damage, heat shock, hypoxia, ER stress, and reactive oxygen species (ROS) (23, 45, 48–51). We tested the involvement of intracellular ROS activity using a fluorometric ROS assay in human PBMC as compared to TNFα, a marker of the extrinsic pathway ( Figure 4H ), and to provide further mechanistic insights. We detected no difference in intracellular ROS levels between the CVF-treated and control cells, while TNFα decreased over time in CVF-stimulated PBMCs. Thus, treatment with CVF led to cellular mitochondrial damage and depolarization, associated with impaired mitochondrial function.

To investigate whether markers for apoptosis such as FasL or TNFα would be detectable in symptomatic NCC patients, we made use of samples from our previous study (28). In this study, we had recruited a cohort of Tanzanian participants, including healthy controls and NCC-infected asymptomatic and symptomatic patients harboring active lesions with degenerating cysts in the brain (28) ( Figure 5A ). To assess the involvement of apoptosis-associated soluble mediators in NCC, we measured plasma levels of TNFα and FasL in the different patient groups ( Figure 5B ). TNFα levels did not significantly differ across the cohort groups. However, symptomatic NCC patients exhibited significantly higher plasma levels of FasL compared to asymptomatic patients and healthy controls ( Figure 5B ). Notably, asymptomatic patients had a trend of lower FasL levels than healthy controls. These findings suggested that degenerating cysts may contribute to the elevated levels of the pro-apoptotic mediator FasL.

To further explore the relevance of caspase 3-driven apoptosis in symptomatic NCC, we assessed caspase 3 activity in PBMCs from patients after stimulation with CVF ex vivo ( Figures 5C, D ). In the absence of stimulation, caspase 3 activity was similar between the different groups. However, upon CVF exposure, PBMCs from symptomatic NCC patients exhibited a marked increase in caspase 3 activity across all cell populations, except CD4⁺ T cells, when compared to healthy controls. In contrast, PBMCs from asymptomatic NCC patients demonstrated a relatively weaker response to CVF, except for CD3^- cells, when compared to the controls ( Figures 5C, D ).

Taken together, these results indicate a strong association between elevated levels of the soluble inflammatory mediator FasL and caspase-driven apoptosis with symptomatic disease in NCC.

Having demonstrated that T. solium CVF induced apoptosis preferentially through the caspase 9-mediated mitochondrial pathway in various immune cell populations, we next sought to identify specific CVF molecules responsible for this pro-apoptotic effect. To achieve this, we conducted a differential mass spectrometry analysis of CVF components. Among the identified CVF molecules (3), we detected 12 proteins, namely cysteine proteases, known to be associated with apoptotic pathways, including the caspases (2, 3, 6, and 7), cathepsins (B, D, L, O, and W), and calpains (1, 2, and 3) (52, 53) ( Figure 6A ). To determine the role of cysteine proteases in CVF-induced apoptosis, we pretreated PBMCs with E-64, an irreversible cysteine protease inhibitor (54), and assessed apoptosis levels. Surprisingly, E-64 pretreatment did not prevent CVF-induced early or late apoptosis in CD3⁺ and CD3^- cells ( Figure 6B ), nor in CD4⁺ and CD8⁺ T cells ( Figure 6C ), indicating that cysteine proteases may not be essential mediators of CVF-driven apoptosis.

We next evaluated the biochemical properties of the active pro-apoptotic molecules in CVF by subjecting CVF to heat treatment (56 °C and 99 °C), as well as proteinase K digestion, to assess the involvement of protein-based factors ( Figure 6D ; Supplementary Figure S4 ). While heat treatment at 99 °C for 3 hours mildly reduced early apoptosis induction in CD3⁺, CD4⁺ ( Figure 6D ), and CD8⁺ T cells ( Supplementary Figure S4 ), overall, treatment at lower temperature and with proteinase K did not significantly alter CVF-induced apoptosis. Accordingly, cysteine proteases and heat- or proteinase K-sensitive proteins did not appear to be the primary drivers of CVF-mediated late-stage apoptosis.

Next, we investigated the molecular size of the active pro-apoptotic factors. We fractionated CVF into molecules smaller or larger than 30 kDa and assessed the pro-apoptotic potency. We found that molecular size with the chosen cut-off did not significantly impact the apoptotic potency of CVF. However, we observed a trend to stronger pro-apoptotic effects, mostly late apoptosis, by molecules < 30 kDa, as compared to molecules > 30 kDa in CD3⁺, CD3^- cells ( Figure 6E ), and CD4⁺ and CD8⁺ T cells ( Figure 6F ).

These results indicated that CVF-mediated apoptosis was driven by small or a combination of small molecular weight components rather than heat- and proteinase-sensitive proteins. The identification of these pro-apoptotic molecules may provide insights into immune-mediated inflammation and disease exacerbation during NCC. A proposed model of CVF-induced apoptosis mechanisms and implications during NCC is presented in Figure 7 .