Ethics statement

All experimental procedures involving animals were conducted in strict accordance with Brazilian legislation (Law No. 11794/2008) governing the use of animals in scientific research. The experimental protocol was reviewed and approved by the Committee on Ethics of Animal Experiments of the Federal University of Minas Gerais (UFMG) (Protocol Number 347/2025). Wild-type C57BL/6 mice were obtained from the Federal University of Minas Gerais (UFMG, Brazil). All animals were housed under specific-pathogen-free conditions at UFMG and used at 6–8 weeks of age. S. mansoni (LE strain) cercariae were provided by the Oswaldo Cruz Foundation – René Rachou Research Center (FIOCRUZ-CPqRR, Brazil) and obtained from infected Biomphalaria glabrata snails by exposure to light to induce parasite shedding.

Sm29 gene design

The gene (based on Smp_072190) encoding the S. mansoni Sm29 antigen was synthetically produced and codon-optimized for expression in Escherichia coli by GenScript (Piscataway, NJ, USA) to maximize translational efficiency and protein yield (S1 Fig). The recombinant construct was cloned into the pET-24a expression vector under control of the T7 promoter. To enhance solubility and reduce aggregation, three additional aspartic acid residues were strategically introduced at the C-terminus of the protein sequence [26,27]. No polyhistidine (His-tag) or other affinity-purification tag was included in the design, enabling downstream applications that require tag-free protein formulations. The construct was sequence-verified, and chemically competent SHuffle T7 E. coli cells (New England Biolabs) were transformed via the standard heat shock method [28,29].

In silico physicochemical characterization. The primary structure of the rSm29 recombinant construct (141 residues) was analyzed using sequence-based computational tools. Molecular weight, isoelectric point (pI), and net charge as a function of pH were calculated using standard amino acid residue masses and the Henderson–Hasselbalch equation with Lehninger pKa values. The molar extinction coefficient at 280 nm (ε₂₈₀) was estimated according to [30]. Hydrophobicity was assessed using the Kyte–Doolittle scale [31] with a sliding window of 9 residues, and the protein stability index was computed according to [32]. All analyses were performed using custom Python scripts with NumPy and Matplotlib libraries.

Expression, cell lysis, inclusion body washing steps, and solubilization

Recombinant Sm29 protein was expressed in E. coli and cultured in 1 L of Luria–Bertani (LB) medium under aerobic conditions. Induction was initiated at an optical density (OD₆₀₀) of 0.44 by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM. Following induction, cultures were maintained at 20 °C for 18 h to promote protein expression. E. coli cells expressing rSm29 were harvested by centrifugation at 10000 × g for 10 minutes at 4 °C, resuspended in PBS pH 7.4 (Gibco), and supplemented with Phenylmethylsulfonyl fluoride (PMSF) 1mM (Sigma), Dithiothreitol (DTT) 5mM (Roche), and Benzamidine hydrochloride hydrate 5mM (Sigma). Cell pellets were subjected to mechanical disruption using a French Press (Brand) for 15 minutes to ensure complete lysis. Lysates were clarified by centrifugation at 40,000 × g for 30 minutes at 4 °C to separate the soluble fraction from insoluble inclusion bodies. The resulting pellet, enriched in inclusion bodies, was sequentially washed (three times) with wash buffer (100 mM Tris-HCl, 10 mM EDTA, 2 M urea, 10% glycerol, 2% Triton X-100, pH 7.0) to eliminate host-derived contaminants. A final wash was performed using phosphate-buffered saline (PBS, Gibco) to remove residual detergents before solubilization. Each wash step involved resuspension by pipetting, vortexing, and centrifugation at 40.000 × g for 30 minutes at 4 °C. Washed inclusion bodies were resuspended in denaturing buffer (20 mM sodium phosphate, 8 M urea, pH 6.7) and manually solubilized at room temperature through gentle pipetting and controlled homogenization. The solubilized protein solution was centrifuged at 40,000 × g for 30 minutes at 4 °C, and the supernatant was collected for a subsequent purification step.

Purification by ion exchange chromatography and endotoxin removal

The clarified lysate was loaded onto a pre-equilibrated HiTrap Q XL anion exchange column (Cytiva) connected to an ÄKTA Pure system (Cytiva). The column was equilibrated with Buffer A (20 mM sodium phosphate, 8 M Urea, pH 6.7), and the protein sample was loaded at a flow rate of 5 mL/min. Unbound material was removed by washing with 10 column volumes of buffer A. Bound proteins were eluted using a linear gradient of NaCl (0-0.35 M) in Buffer B (20 mM sodium phosphate, 8M Urea, 1 M NaCl, pH 6.7) with 70 column volumes. Eluted fractions were monitored at 280 nm, collected, and analyzed by SDS-PAGE. Before additional chromatographic steps aimed at optimizing purification, the protein sample was dialyzed against Buffer A (20 mM sodium phosphate, 8 M urea, pH 6.7) to remove residual NaCl and equilibrate the ionic strength. An additional anion exchange chromatography was performed to further reduce contaminating host-derived proteins, following the same conditions as the initial purification, except for a modified elution profile. At this point in the chromatography workflow, Buffer A (20 mM sodium phosphate, 8 M urea, pH 6.7) was employed for column equilibration and washing. Specifically, the gradient of buffer B was adjusted to 0- 0.4 M NaCl, and the linear elution gradient was shortened from 70 to 30 column volumes (CV), thereby reducing the total elution volume required for protein recovery. Fractions were collected, analyzed by SDS-PAGE, and pooled based on purity. Following SDS-PAGE analysis, the protein concentration was determined using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, USA) according to the manufacturer’s instructions. Next, the protein sample was dialyzed overnight against an alternative Buffer A (50 mMacetic acid, 8 M urea, pH 4.3) to promote endotoxin (LPS) removal. Samples were dialyzed against 1 L of appropriate buffer at 4 °C using a Spectra/Por membrane (MWCO 6–8.000 kDa; Spectrum Medical Industries, Inc., Laguna Hills, CA), with gentle stirring using a magnetic stir bar. Subsequently, the dialyzed sample was applied to an anion exchange chromatography column for further purification. The rSm29 was applied to the column at a flow rate of 1 mL/min under these conditions and collected in the flow-through, as it does not bind significantly at this pH and conductivity. LPS molecules remained bound to the resin and were later removed during column regeneration. The flow-through fractions were pooled and quantified using Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, Massachusetts, USA), following the manufacturer’s protocol. Endotoxin removal efficiency was confirmed by the ToxinSensor Chromogenic LAL Endotoxin Assay Kit (GeneScript, Piscataway, NJ, USA), performed in triplicate. rSm29, initially solubilized in 8 M urea, was subjected to a dialysis protocol to reduce the urea concentration to 1 M, thereby equilibrating the sample in a buffer compatible with downstream applications. Dialysis was carried out at 4 °C under gentle agitation using a 10 kDa molecular weight cut-off (MWCO) membrane, against 1 L of buffer composed of 20 mM sodium phosphate, 1 M urea, and 150 mM NaCl at pH 7.4.

Bacterial endotoxin quantification

Residual endotoxin quantification was performed at two distinct stages of the production process. For in-process control and rapid monitoring of the efficiency of LPS removal steps, the ToxinSensor Chromogenic LAL Endotoxin Assay Kit (GenScript, Piscataway, NJ, USA) was used according to the manufacturer's instructions.

For the final characterization of the Active Pharmaceutical Ingredient (API) and to ensure regulatory compliance, rigorous endotoxin quantification was conducted using the kinetic turbidimetric method, as recommended by the Pharmacopoeia. Briefly, samples were diluted in endotoxin-free water to prevent interference, as required to comply with the assay operating range. Potential matrix interference was assessed through the positive product control recovery according to the manufacturer's instructions.

The analysis was based on the reaction time required to reach a predetermined level of turbidity in the mixture of the sample and the Limulus Amebocyte Lysate (LAL) reagent. Concentrations were calculated by interpolation from a Reference Standard Endotoxin curve, and the results were expressed in Endotoxin Units per microgram of protein (EU/µg).

Acceptance criteria for residual endotoxins were established based on the maximum human dose and average body weight, as defined by pharmacopeial guidelines. The endotoxin limit (EL) was calculated using the formula EL = K/M, where K represents the threshold pyrogenic dose (5.0 EU/kg), and M is the maximum dose administered per kg per hour. Based on this calculation, the safety threshold for the rSm29 vaccine was defined as < 3.5 EU/µg, ensuring the product's safety for clinical use.

SDS-PAGE and western blot analysis

The pooled fractions containing purified rSm29 were subjected to SDS-PAGE analysis and western blotting. Western blotting was employed to confirm the identity and immunoreactivity of the rSm29. Protein samples (0.1 to 0.5 µg) were resolved on a 15% SDS-PAGE gel as previously described by Laemmli [33] and electrotransferred to nitrocellulose membranes (Amersham Biosciences, Uppsala, Sweden) using a semidry system (Bio-Rad) at 25V/5mA for 25 minutes. Membranes were blocked for 1 hour at room temperature in TBS (Tris-buffered saline containing 0.05% Tween-20, pH 7.2), supplemented with 5% non-fat dry milk. Following the blocking step, membranes were incubated overnight at 4 °C with anti-Sm29 polyclonal antibodies (1:2000) previously produced in mouse. In the next step, membranes were washed three times with TBS-T and incubated for 1 hour at room temperature with goat anti-mouse IgG conjugated to horseradish peroxidase (HRP) (1:2000) - Invitrogen. Immunoreactive bands were detected using ECL western blotting detection reagents (Amersham Biosciences, Piscataway, USA) according to the manufacturer’s instructions and visualized on an Amersham Imager 600 (GE Healthcare).

Circular dichroism spectroscopy and secondary structure analysis

Far-UV CD spectra were recorded on a Jasco spectropolarimeter (600 MHz, IME/EB) using a 0.1 mm pathlength quartz cuvette. The protein sample (rSm29, 0.8 mg/mL in 20 mM sodium phosphate, 1 M urea, 150 mM NaCl, pH 7.4) was measured in duplicate over the range 200–260 nm. Buffer baseline spectra were recorded under identical conditions using the same buffer diluted 10-fold and subtracted from the sample spectra prior to conversion to mean residue ellipticity (MRE). Spectra were smoothed using the Savitzky–Golay algorithm (window = 25 points, polynomial order = 3). The high tension (HT) voltage was monitored throughout acquisition as a criterion of signal reliability, with values below 600 V considered acceptable. Secondary structure content was estimated by non-negative least squares (NNLS) deconvolution using reference spectra for α-helix, β-sheet, and random coil derived from [34].

Quantification of residual host cell DNA using a fluorescence-based assay

Residual double-stranded DNA (dsDNA) in protein samples was quantified using a high-sensitivity fluorescent assay (Quant-iT PicoGreen dsDNA Assay Kit, Thermo Fisher Scientific), following the manufacturer's protocol. Succinctly, the working reagent was freshly prepared by diluting the PicoGreendye 1:200 in 1 × TE buffer. In a black 96-well flat-bottom microplate (low-binding, fluorescence-compatible), 100 μL of each sample or DNA standard (λ DNA in TE buffer, provided in the kit) was pipetted in triplicate. An equal volume (100 μL) of PicoGreen working solution was added to each well, mixed gently, and incubated for 5 minutes at room temperature protected from light. Fluorescence was measured using a microplate reader (SpectraMax) with excitation at 480 nm and emission at 520 nm. A standard curve was generated using serial dilutions of λ DNA, and DNA concentrations in samples were interpolated using linear regression. Residual DNA concentrations were reported in ng per dose (based on previously determined protein concentration by BCA assay). Acceptance criteria followed regulatory guidelines for biopharmaceuticals, typically <10 ng/dose.

Quantification of host cell proteins (HCP)

The presence of residual HCPs in purified recombinant protein samples was quantified using a commercial ELISA-based assay (Cygnus Technologies, USA), following the manufacturer’s instructions. Briefly, standards and appropriately diluted samples were added to 96-well microplates pre-coated with anti-HCPs polyclonal antibodies. After incubation, plates were washed to remove unbound material, and a horseradish peroxidase–conjugated secondary antibody specific for HCPs was added. Following a second wash step, the substrate solution (TMB) was added, and the reaction was allowed to develop for the recommended period. The enzymatic reaction was then stopped with acidic stop solution, and absorbance was measured at 450 nm with reference to 650 nm using a microplate reader. Residual HCPs were reported in ng per mg (based on previously determined protein concentration by BCA assay). Acceptance criteria followed regulatory guidelines for biopharmaceuticals, typically <100 ng/mg.

Protein profiling using the bioanalyzer system

Protein sizing and purity assessment were performed using the Agilent 2100 Bioanalyzer system (Agilent Technologies, Santa Clara, CA, USA) in combination with the Protein 230 LabChip Kit, following the manufacturer’s instructions. Briefly, protein samples were heated at 95 °C for 5 minutes in a denaturing buffer containing SDS to achieve denaturation, enabling linearization and uniform charge-to-mass ratio of the polypeptides. Subsequently, 4 µL of each denatured sample was loaded into individual wells of the microfluidic chip, along with the appropriate molecular weight ladder and gel-dye mix. The chip was placed in the Bioanalyzer, where microfluidic capillary electrophoresis was automatically performed. Protein separation was based on size, under denaturing conditions, and fluorescence signals were detected and analyzed by the integrated software. Each electropherogram and virtual gel image was evaluated to determine the apparent molecular weights, relative abundance, and purity of each sample. The estimated molecular weights were calculated by comparison with the internal standard ladder, and peak areas were used for semi-quantitative assessment of relative protein concentration. The integrity of the proteins was confirmed by distinct, well-defined peaks at the expected molecular weights, with no evidence of smearing or degradation. Samples exhibiting multiple peaks or high-molecular-weight aggregates were flagged for potential contamination or incomplete denaturation.

All experimental activities were performed at CT Vacinas (Federal University of Minas Gerais, Brazil), a facility certified for Good Laboratory Practice (GLP/BPL) by the Brazilian National Institute of Metrology, Quality and Technology (INMETRO). Analytical assays and process development activities were conducted according to established standard operating procedures, ensuring traceability, quality control, and data integrity throughout the study.

Mass spectrometry-based protein identity confirmation

Purified rSm29 was buffer-exchanged by methanol/chloroform precipitation and resuspended in 50 mM ammonium bicarbonate. Two aliquots of 20 µg were prepared for parallel digestion with trypsin or chymotrypsin (enzyme:protein ratio 1:50, w/w) after reduction with 10 mM DTT and alkylation with 40 mM iodoacetamide. Resulting peptides were desalted and analyzed by LC-MS/MS on a Vanquish Neo system coupled to an Orbitrap Ascend Tribrid mass spectrometer (Thermo Scientific), using a reverse-phase Acclaim PepMap column (50 cm × 75 µm, 2 µm) with an acetonitrile/formic acid gradient at 300 nL/min. MS1 and MS2 spectra were acquired in data-dependent mode (DDA) at resolving powers of 60,000 and 15,000, respectively, with HCD fragmentation at 25% normalized collision energy. For protein identity confirmation, raw data were processed in Proteome Discoverer 3.2 (Sequest HT) against a database containing the rSm29 sequence and common contaminants (precursor tolerance: 10 ppm; fragment tolerance: 0.02 Da; fixed modification: carbamidomethyl-Cys; up to 4 missed cleavages; FDR 1%). All PSMs were manually curated.

Mice immunization protocol

C57BL/6 mice (6–8 weeks old) were housed under specific pathogen-free (SPF) conditions and handled following institutional animal care and guidelines. Mice were randomly assigned to experimental and control groups (n = 10 per group). Immunizations were performed subcutaneously, at the nape of the neck, using 25 μg of rSm29 formulated with Alum or CTVad1 (vol/vol). Control animals were injected with adjuvant alone at the same volume and on the same schedule. Mice received three doses at 14-day intervals (Days 0, 14, and 28). Sera were collected by submandibular bleeding at baseline (pre-immune) and two weeks after each immunization.

Specifics antibodies detection by indirect ELISA

Total IgG, IgG1, IgG2c, IgG3 and IgE antibody responses in mouse sera were quantified by indirect ELISA. Following immunization, sera were collected from mice in each experimental group at 2-week intervals. High-binding microplates (Nunc MaxiSorp, Thermo Fisher Scientific) were coated overnight at 4 °C with 100μL/well of rSm29 (10μg/mL) diluted in carbonate-bicarbonate buffer (pH 9.6). Plates were washed three times with PBS containing 0.05% Tween-20 (PBST) and blocked with 100μL/well of 5% BSA in PBS for 1 hour at room temperature (11). After blocking, serum samples diluted 1:100 in PBST containing 5% BSA were added in triplicate and incubated for 2 hours at room temperature. To determine total IgG antibody titers, pooled serum samples from each group were used in a serial dilution range from 1:20–1:1,310,720 [12]. After washing, plates were incubated for 1 hour with horseradish peroxidase (HRP)-conjugated goat anti-mouse antibodies specific for IgG (1:5,000), IgG1 (1:10,000), IgG2c (1:12,000), IgG3 (1:2.000) and IgE (1:2.000) (Southern Biotechnology, CA, USA) [35]. The enzymatic reaction was developed by adding 100μL/well of TMB substrate for 15min, stopped with sulfuric acid (H₂SO₄), and absorbance was measured at 450 nm using a microplate reader (BioRad, Hercules, CA, USA).

Cercarial challenge and parasite burden evaluation

Following standard procedures, mice were percutaneously challenged with S. mansoni cercariae (LE strain) two weeks after the final immunization. Briefly, six-to-eight-week-old wild-type mice were anesthetized with 5% ketamine, 2% xylazine, and 0.9% NaCl. Animals were individually exposed to 100 viable cercariae through the shaved abdominal skin for 1 hour using the cover slip technique. Cercarial viability and concentration were confirmed microscopically before infection. Forty-five days after challenge, mice were euthanized, and the portal and mesenteric veins were perfused with pre-warmed citrate-saline solution (0.85% NaCl, 1.5% sodium citrate) to recover adult worms [36]. The perfusate was passed through a 100 μm mesh and examined under a stereomicroscope. Worms were counted manually and classified by sex. Total worm burden per animal was recorded, and the protection level was calculated by comparing immunized groups to the control group [35].

Splenocyte culture and cytokine determination

Cytokine secretion determination was conducted using splenocyte cultures from individually immunized mice (n = 5 per group). Splenocytes were isolated from macerated spleens of individual mice 15 days after the third immunization and washed twice with sterile PBS. After washing, cells were adjusted to 1 × 10⁵ cells per in wells RPMI 1640 medium (Gibco, CA, USA) supplemented with 10% fetal bovine serum (FBS), 100 U/mL sodium penicillin G, 100 μg/mL streptomycin sulfate, and 250 ng/mL amphotericin B. Splenocytes were cultured in medium alone or stimulated with rSm29 (25 μg/mL) or concanavalin A (ConA) (5 μg/mL), as previously described (11). Cells were incubated at 37 °C with 5% CO₂. Culture supernatants were collected after 48 hours for IL-4 measurements, and after 72 hours for IFN-γ and IL-10 measurements by ELISA (R&D Diagnostics, Minneapolis, MN, USA) according to the manufacturer’s instructions.

Liver pathology

After euthanasia, the left liver lobe was fixed in 10% buffered formaldehyde in PBS. Histological sections were prepared at 6 μm using a microtome and stained with Hematoxylin-Eosin (HE). For granuloma size, images were captured using a Michrome 20 digital camera coupled to a microscope (20 × objective) and the Mosaic 3.0 software (Tucsen Imaging Technology Co., Ltd). Analyses were performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Granuloma size was measured in mm² for all granulomas observed in liver sections. The number of eggs per granuloma was also counted.

Statistical analysis

Data normality was tested using D’Agostino-Pearson omnibus tests. Statistical analyses were performed using ANOVA and Dunn’s multiple comparison test for the antibody measurements. The percentage of granuloma/liver areas was plotted in box plots (median, maximum, and minimum) and analyzed via the Kruskal-Wallis test followed by Dunn’s multiple comparison test. The number of granulomas per liver area and the number of eggs per granuloma were evaluated using one-way ANOVA followed by Tukey's multiple comparison test. The level of protection in infected mice was determined using Student’s t-test. All calculations were performed using GraphPad Prism 6 (GraphPad Software, La Jolla, CA, United States). The p-values were considered significant when < 0.05.

Design and in silico characterization of the optimized Sm29 construct

The gene encoding the Sm29 target protein was successfully cloned into the pET24a(+) expression vector under the control of the T7 promoter (GenScript), enabling high-level, inducible expression in the E. coli Shuffle strain. Codon optimization was performed to enhance translational efficiency in E. coli, by reducing the frequency of rare codons and maximizing predicted mRNA stability without altering the amino acid sequence. The expression cassette was engineered without affinity tags to comply with regulatory requirements and preserve the native biochemical and immunological properties of the protein. The final construct therefore expresses the recombinant protein in its native form, without a C-terminal or N-terminal His-tag or other fusion domains, ensuring compatibility with pre-clinical development and analytical characterization workflows. To enhance solubility during protein expression in E. coli, three aspartic acid residues (DDD) were rationally introduced at the C-terminus of the protein (S1 Fig). Incorporation of negatively charged residues at the C-terminus has been associated with reduced aggregation propensity and enhanced solubility through increased electrostatic repulsion and disruption of hydrophobic aggregation interfaces. In silico solubility prediction demonstrated an increase in solubility scores relative to the native sequence, supporting this engineering strategy [37–40].

Sequence confirmation by Sanger sequencing validated the integrity of the transgene and the successful integration of the DDD motif. Alignment of sequencing chromatograms with the reference plasmid map demonstrated 100% identity with the designed construct, with no frameshift mutations, premature stop codons or unexpected sequence alterations. Restriction digestion analysis using NdeI and XhoI yielded the expected fragment pattern, further confirming proper assembly of the recombinant plasmid. Additionally, the recombinant plasmid was confirmed to maintain the kanamycin resistance cassette and origin of replication associated with pET-24a (+), ensuring stable selection during bacterial amplification and protein expression. The final construct was designated pET-24a-Sm29-DDD. This native recombinant protein expression construct incorporating a C-terminal DDD solubility enhancement motif was successfully engineered, sequence-validated, and was suitable for downstream expression, purification, and pre-clinical quality assessment.

The primary structure analysis of Sm29 reveals several physicochemical features consistent with its classification as a member of the Ly6/uPAR (three-finger fold) protein family [41]. The recombinant construct comprises 141 amino acid residues with a calculated molecular weight of 15.39 kDa and an isoelectric point (pI) of 4.66, conferring a net negative charge of −5.6 at physiological pH (7.4), which is consistent with the anion-exchange chromatography purification strategy employed. The most striking feature of the primary structure is the exceptionally high cysteine content — 17 residues comprising 12.1% of the sequence — arranged in a characteristic spacing pattern (CxxC and CxxxC motifs) that supports the formation of up to eight disulfide bonds, the hallmark of the Ly6/uPAR LU domain [42,43]Notably, the protein lacks tryptophan residues entirely, resulting in a relatively low molar extinction coefficient at 280 nm (ε₂₈₀ = 8,940–9,940 M⁻¹cm⁻¹, depending on the oxidation state of cysteine residues), calculated according to [30]. The overall hydrophilicity (GRAVY index = −0.119) is consistent with a surface-exposed protein, and the computed instability index of 21.77 predicts conformational stability in solution, likely stabilized by the extensive disulfide bond network. The C-terminal DDD motif, rationally introduced to enhance solubility during recombinant expression in E. coli, contributes three additional aspartate residues to the overall negative charge of the protein at physiological pH (S1 Fig).

Anion exchange chromatography purification strategy

All stages of the rSm29 expression and optimization process are described in detail in Fig 1A-1C. The purification of the rSm29 was conducted using an anion exchange chromatography strategy employing a QXL column functionalized with quaternary amine ligands. This resin provides a positively charged matrix that selectively binds negatively charged biomolecules under controlled ionic conditions. The first QXL chromatography step was primarily designed as a capture step for bulk impurity reduction and target protein enrichment.

Download:

• PNG
  larger image
• TIFF
  original image

Fig 1. Workflow diagram of the expression, purification process, and sample quality control steps for Sm29 recombinant protein.

Schematic representation of the production process of recombinant Sm29 expressed in Escherichia coli, including bacterial cultivation, cell disruption by high-pressure homogenization, clarification of the soluble and insoluble fractions, recovery of inclusion bodies, solubilization under denaturing conditions, and sequential purification by anion-exchange chromatography. The downstream process comprised an initial capture step, an intermediate purification step to reduce host cell-derived contaminants, and a polishing step dedicated to endotoxin removal, followed by gradual reduction of the chaotropic agent by stepwise dialysis and formulation in the final storage buffer. Analytical characterization included determination of protein concentration by BCA assay, assessment of purity by SDS-PAGE and microfluidic electrophoresis, immunoreactivity by Western blot and ELISA, quantification of residual host-cell proteins (HCPs), residual host-cell DNA, and endotoxin levels. Each purification stage was designed to progressively reduce process-related impurities while maximizing recombinant protein recovery and maintaining product quality attributes relevant to future vaccine development.

https://doi.org/10.1371/journal.pntd.0014617.g001

Following the first purification step, 15% SDS-PAGE analysis was performed to evaluate the expression profile and purity of the recombinant Sm29 protein (Fig 2). Based on the electrophoretic profiles (11 a 178) obtained from the loaded fractions (30 μL per lane), purified fractions exhibiting the highest purity (Fig 2A and 2B) and recovery were pooled for further processing (Fig 2C). The total volume of the pooled fractions was recorded before dialysis. Fractions 55–126 were combined, resulting in a final pool volume of 142 mL. To remove residual salt and prepare the sample for the subsequent repurification step aimed at further reducing host cell-derived contaminants, the pooled protein solution was dialyzed overnight against 20 mM sodium phosphate buffer containing 8 M urea (pH 6.7). After dialysis, the pool volume remained unchanged at 142 mL, indicating that no significant sample loss occurred during buffer exchange, at a concentration of 0.74 mg/mL, corresponding to a total protein recovery of 105.1 mg of rSm29.

Download:

• PNG
  larger image
• TIFF
  original image

Fig 2. SDS-PAGE analysis of chromatographic fractions obtained during recombinant Sm29 purification.

A and B- SDS-PAGE (15%) analysis of fractions obtained during the purification of 80 mL of solubilized recombinant Sm29. MW: molecular weight marker (kDa); PC: solubilized sample prior to column loading; FT: flow-through fraction; lanes 11 to 178 correspond to 2-mL fractions collected during the purification process. A total of 30 μL of each sample was loaded per lane. C- SDS-PAGE (15%) analysis of pooled recombinant Sm29 fractions (55 to 126) before and after dialysis. The predominant protein band migrated at the expected molecular weight of approximately 15.34 kDa, indicating enrichment of recombinant Sm29 and preservation of protein purity throughout the dialysis process. D and E- Electrophoretic profile of fractions collected during the repurification of Sm29. MW: molecular weight marker (kDa); PC: sample prior to column loading; FT: flow-through fraction; lanes 08 to 80 correspond to 2-mL fractions collected during the repurification process. F- SDS-PAGE (15%) analysis of pooled recombinant Sm29 fractions (09 a 78) before and after dialysis. A total of 30 μL of each sample was loaded per lane. G and H- Electrophoretic profile of fractions collected during the endotoxin removal purification. MW: molecular weight marker (kDa); PC: sample prior to column loading; lanes 02–75 correspond to sequential 2-mL fractions collected during the purification process. I- Electrophoretic profile of fractions collected during the endotoxin removal purification. MW: molecular weight marker (kDa); PC: sample prior to column loading; lanes 10 a 75 correspond to sequential 2-mL fractions collected during the purification process. A total of 30 μL of each sample was loaded per lane. *Pre-dialysis: pooled elution fractions prior to buffer exchange; post-dialysis: pooled fractions following dialysis.

https://doi.org/10.1371/journal.pntd.0014617.g002

The purification process was performed at pH 6.7, a value above the theoretical isoelectric point (pI = 4.96) of rSm29. Under these conditions, the recombinant protein carried an overall negative net charge, enabling binding to the anion-exchange matrix. Consistent with the chromatographic behavior observed during process optimization, this interaction was sufficient to retain the protein on the column but was readily disrupted by relatively low NaCl concentrations during elution.

The second QXL step functioned as an intermediate purification stage, improving protein purity and reducing residual host-cell contaminants. During the loading phase, the target protein was adsorbed onto the resin while unbound impurities were removed in the flow-through Controlled elution was achieved by gradually increasing the ionic strength through a NaCl gradient, which weakened the electrostatic interactions and promoted desorption of the protein. This purification approach provided effective removal of HCPs and other process-related impurities, while maintaining structural integrity and activity of the recombinant protein. The use of a QXL strong anion exchanger at optimized pH conditions has been widely reported as a robust and scalable method for downstream purification of acidic recombinant proteins, ensuring both high recovery and regulatory-grade purity.

For the second chromatography step (intermediate purification), the entire dialyzed pool (144 mL) was loaded onto the QXL column. Based on the SDS-PAGE profiles of the collected fractions (08–80) (Fig 2D and 2E) and fractions 09–78 were selected and combined (Fig 2F). After the intermediate purification step, the pooled fractions contained 140 mL at 0.77 mg/mL, yielding 107.8 mg of recombinant protein. Before the endotoxin removal chromatography step, the pooled sample (140 mL) was dialyzed overnight against 50 mM acetic acid buffer containing 8 M urea (pH 4.3) to perform buffer exchange and prepare the sample for the subsequent purification stage. Following dialysis, the pool volume increased from 140 mL to 147 mL. This increase is consistent with osmotic effects arising from differences in buffer composition and ionic properties between the sodium phosphate and acetic acid dialysis buffers. The lower ionic strength and altered physicochemical characteristics of the acetic acid buffer may have promoted water influx across the dialysis membrane, resulting in a modest increase in sample volume. Despite this volumetric variation, no visible protein precipitation was observed, and the sample remained suitable for the subsequent chromatography step.

Additionally, a third anion exchange chromatography strategy was employed using a QXL strong anion exchange column to efficiently remove endotoxin contaminants from the recombinant protein preparation. The third QXL chromatography step acted as a polishing stage specifically optimized for lipopolysaccharide (LPS) reduction and final impurity clearance.

The QXL resin provides a positively charged matrix that binds negatively charged molecules, such as lipopolysaccharides (LPS). Purification was performed at pH 4.3, a condition in which the LPS molecules (pI ≈ 2.0) carry a strong negative charge, thereby exhibiting high affinity for the positively charged resin. In contrast, the target protein rSm29, with a theoretical isoelectric point (pI = 4.96), remains near its net neutral charge under these conditions and consequently displays minimal interaction with the matrix. As a result, the protein of interest was efficiently recovered in the flow-through fraction, while endotoxins were retained on the column This selective separation mechanism effectively reduced endotoxin burden to levels compliant with pharmacopeial and regulatory limits for parenteral biologics. The approach demonstrates a practical application of charge-based differential binding for endotoxin clearance, providing a scalable, non-denaturing purification step that is compatible with GLP bioprocessing standards.

For the anion-exchange chromatography step dedicated to endotoxin (LPS) removal, 147 mL of the Sm29-containing pool was loaded onto the QXL column. Under these conditions, the recombinant protein did not bind to the chromatographic matrix, and the entire flow-through fraction was collected individually throughout the purification process. Following sample collection, the chromatographic run was completed, and the endotoxin content of the recovered fractions was subsequently determined. Fraction quality and protein integrity were evaluated (02–75) (Fig 2G and 2H) by 15% SDS-PAGE analysis, which was used to identify the fractions containing the target protein and assess the effectiveness of the purification strategy (10–75) (Fig 2I). Following the endotoxin removal step, the final pool consisted of 104 mL at 0,88 mg/mL, corresponding to 91,5 mg of purified rSm29.

Throughout the purification process, 8 M urea was maintained during all three anion-exchange chromatography steps and during the first two intermediate dialysis procedures. This strategy was adopted to preserve Sm29 in a soluble state after recovery from inclusion bodies and to minimize protein aggregation and precipitation during purification. Because Sm29 contains multiple cysteine residues and was initially solubilized under strongly denaturing conditions, premature removal of urea could promote intermolecular interactions and irreversible aggregation, leading to product loss and reduced process robustness. Therefore, urea concentration was reduced gradually only during the final refolding stage through sequential dialysis steps, allowing controlled protein renaturation while maintaining high recovery yields.

Following the final dialysis step in buffer containing 1 M urea, the final preparation volume was 100 mL with a protein concentration of 1.06 mg/mL, as determined by the BCA assay. This corresponded to a total recovery of approximately 100 mg of purified rSm29, yielding approximately 100 mg of recombinant protein per liter of bacterial culture. The overall protein recovery throughout the purification process remained above 98%, indicating that the sequential chromatographic steps did not significantly compromise protein yield.

Recombinant protein yield, purity, and identification by SDS-PAGE and western blot

The recombinant protein was successfully expressed and purified under optimized induction and downstream conditions, yielding consistent production across independent fermentation batches. From a 1 L bacterial culture, the final recovery of purified target protein averaged 100 mg (mean ± SD, n = 3), as determined using the BCA assay. Process reproducibility was demonstrated by low inter-batch variation (<10%), indicating robustness of the expression and purification strategy. The achieved yield aligns with the expected productivity range for soluble recombinant proteins expressed in E. coli under standard laboratory-scale fermentation parameters (0.1-5 g/L, depending on construct and solubility tag) [44–46]. This performance reflects the effectiveness of vector design and culture optimization parameters, particularly induction temperature, IPTG concentration, and buffer composition, which collectively enhanced solubility and minimized inclusion body formation. The obtained yield is suitable for subsequent analytical characterization, pre-formulation, and preclinical testing stages of the active pharmaceutical ingredient (API) development.

Pooled fractions obtained after purification steps were analyzed by SDS-PAGE to evaluate protein purity and recovery. The electrophoretic profile revealed a predominant band at the expected molecular mass of approximately 15 kDa, corresponding to the recombinant protein of interest, with minimal detectable impurities after each purification step (Fig 2C, 2F and 2I).

Densitometric analysis of SDS-PAGE profiles demonstrated a high degree of sample purity both before and after dialysis, with no detectable degradation or formation of secondary bands, indicating structural stability of the recombinant protein. Comparative quantification revealed consistent band intensity corresponding to the target molecular weight, confirming that the dialysis process did not lead to significant protein loss or aggregation. The calculated purity exceeded 95%, consistent with preparative chromatography performance and appropriate buffer exchange conditions (Fig 3B). Furthermore, the overall yield after dialysis remained above 98% of the pre-dialysis protein concentration, supporting the robustness of the purification and formulation procedures in maintaining protein integrity and recovery. These results confirm that the selected dialysis parameters ensured efficient removal of low-molecular-weight impurities while preserving protein stability and purity in accordance with regulatory expectations for preclinical-grade biopharmaceutical materials.

Download:

• PNG
  larger image
• TIFF
  original image

Fig 3. Sm29 recombinant protein Western blotting and Bioanalyzer.

(A) Western blot analysis of recombinant Sm29 loaded at different concentrations (0.1–0.5 μg). An immunoreactive band at the expected molecular weight of approximately 15.34 kDa was observed across all protein loads, confirming the detection of recombinant Sm29 by the anti-Sm29 antibody. (B) Microfluidic electrophoresis analysis of recombinant Sm29 using the Agilent 2100 Bioanalyzer. The electropherogram and corresponding virtual gel revealed a predominant protein species with an apparent molecular weight of approximately 15.34 kDa, consistent with the expected size of recombinant Sm29. Quantitative analysis indicated an estimated protein purity of approximately 95%, supporting the effectiveness of the purification process and the enrichment of the target protein.

https://doi.org/10.1371/journal.pntd.0014617.g003

Protein identity was further confirmed by Western blotting using a specific polyclonal antibody raised against the recombinant Sm29 (Fig 3A). A single immunoreactive band was detected at the same molecular weight observed by SDS-PAGE, confirming antigenic integrity and absence of degradation products or cross-reactive impurities. Together, these results validate the effectiveness of the purification strategy in producing a homogeneous and immunoreactive recombinant protein suitable for analytical characterization, formulation, and preclinical evaluation.

Analytical strategies for detection and quantification of process-related impurities during recombinant protein production

The analytical characterization of process-related impurities is a critical component of the quality assessment of recombinant protein products. These impurities, originating from the host cell or manufacturing process, included host cell proteins (HCPs), residual host cell DNA, endotoxins, and protein integrity, which were assessed by microfluidic electrophoresis. Their identification and quantification ensure that the API meets safety and purity criteria established by international regulatory guidelines. Due to the relatively weak interaction of recombinant Sm29 with the anion-exchange matrix under the selected operating conditions, the presence of NaCl in the washing buffer promoted premature target protein desorption. Therefore, a salt-free washing step was adopted to preserve product recovery while maintaining satisfactory impurity removal.

Host cell proteins (HCPs)

The levels of HCPs were quantified using a validated ELISA specific for the E. coli expression system. Across production batches, there was consistent HCP clearance throughout downstream purification. Initial harvest material contained elevated concentrations of HCPs, which were progressively reduced following chromatographic purification.

The HCP content measured for the rSm29 protein was 140 ng/mg (Table 1). While this value is near the established target of ≤100 ng/mg, it remains within a range consistently associated with high-purity biologicals and aligns with levels often accepted in early-stage vaccine development. The observed consistency across multiple batches demonstrates the efficiency of the downstream process in managing host cell impurities. Moreover, this level of residual HCP did not interfere with the protein's safety profile in the subsequent preclinical evaluations, confirming the suitability of the purified rSm29 for its intended applications.

Download:

• PNG
  larger image
• TIFF
  original image

Table 1. Analytical characterization and quality attributes of the purified rSm29 protein.

https://doi.org/10.1371/journal.pntd.0014617.t001

Residual DNA

Residual host cell DNA content was evaluated using a high-sensitivity fluorescent assay (Quant-iT PicoGreen dsDNA Assay Kit, Thermo Fisher Scientific). The measured concentration of residual DNA in the final purified recombinant protein preparation was 3.5 ng DNA per therapeutic dose as demonstrated in Table 1. This value is well below the international regulatory limit of <10 ng DNA per therapeutic dose, as established by the World Health Organization [47] and ANVISA [21] for recombinant biopharmaceuticals. These results confirm that the purification process efficiently removed host-derived nucleic acids, ensuring compliance with global quality and safety standards for preclinical and toxicological applications.

Endotoxin quantification

Endotoxin concentrations were measured by the kinetic turbidimetric method using the Limulus Amebocyte Lysate (LAL) reagent. Endotoxin analysis performed after the second QXL chromatography revealed a residual endotoxin concentration of 58 EU/mg of protein. This result demonstrated that, despite the effectiveness of the capture and intermediate purification steps, additional endotoxin clearance was required. Consequently, a third QXL chromatography step was introduced as a polishing stage specifically aimed at reducing endotoxin levels. The substantial reduction achieved after this final chromatographic step confirmed the necessity and effectiveness of this additional purification stage. Process intermediates demonstrated a marked decrease in endotoxin levels. Final endotoxin concentrations were consistently below 0.154 EU/µg total protein (Table 1), well within the specific safety limit calculated for rSm29 (3.5 EU/µg total protein). The observed results indicate adequate control of upstream bacterial lysis and effective downstream endotoxin removal capability. The validity of the LAL assay was confirmed through the acceptance criteria of the positive product control, indicating that residual buffer components, including urea, did not significantly interfere with endotoxin quantification.

Bioanalyzer protein integrity profile

Microfluidic electrophoresis (Bioanalyzer) confirmed protein purity and structural integrity at critical stages of manufacturing. The purified final product displayed a predominant band at the expected molecular weight (15 kDa), with purity exceeding 95% (Table 1), as determined by peak area integration of the main protein peak. No fragmentation or aggregation species were detected. Batch-to-batch profiles were highly comparable, supporting manufacturing consistency. Absence of degradation species indicates effective protein stabilization, appropriate chromatographic selection, and consistent product quality (Fig 3B).

Mass spectrometry-based identity confirmation of rSm29

To unambiguously confirm the identity and assess the protein composition of the purified rSm29 preparation, the final product was subjected to LC-MS/MS analysis following proteolytic digestion with two complementary enzymes, trypsin and chymotrypsin (Fig 4A). This orthogonal dual-enzyme strategy was used to maximize sequence coverage and ensure that all regions of the recombinant construct, including cysteine-rich segments and regions refractory to a single protease, were represented in the peptide map.

Download:

• PNG
  larger image
• TIFF
  original image

Fig 4. LC-MS/MS-based sequence coverage and identity confirmation of rSm29.

(A) Peptide coverage map of the 141-amino acid rSm29 recombinant construct obtained by LC-MS/MS following enzymatic digestion with trypsin and chymotrypsin. Each residue is color-coded according to the enzyme generating the covering peptide: teal, trypsin only; purple, chymotrypsin only; dark green, covered by both enzymes; light grey, not covered. Trypsin-derived peptides (T1–T10, bars below the sequence) achieved 87% sequence coverage, chymotrypsin-derived peptides (Q1–Q6, bars above the sequence) achieved 100% coverage, and the combined dual-enzyme strategy yielded complete 100% coverage of the full-length construct including the engineered C-terminal DDD motif. (B) Experimental parameters of trypsin- and Chymotripsin-derived peptides identified by LC-MS/MS: peptide sequences are reported with carbamidomethylated cysteines (+57.021 Da); charge state, monoisotopic m/z, protonated molecular mass (MH⁺), chromatographic retention time (RT), and Sequest HT XCorr score are shown for each peptide. All peptides were detected at MS1 and MS2 levels and manually curated. Data were acquired on an Orbitrap Ascend Tribrid mass spectrometer and processed with Proteome Discoverer 3.2 (precursor tolerance: 10 ppm; fragment tolerance: 0.02 Da; FDR: 1%).

https://doi.org/10.1371/journal.pntd.0014617.g004

Trypsin digestion generated ten peptides (T1–T10) covering 87% of the rSm29 sequence, while chymotrypsin digestion generated six peptides (Q1–Q6) independently achieving 100% coverage (Fig 4B). The combined analysis yielded complete 100% sequence coverage of the 141-amino acid construct, including the engineered C-terminal DDD motif (Fig 4). All identified peptides were detected at both MS1 and MS2 levels and manually curated, with XCorr scores ranging from 2.75 to 8.45 for tryptic peptides and 2.93 to 8.35 for chymotryptic peptides, reflecting high-confidence spectral matches. The complete sequence identity of the recombinant protein was thus confirmed at the peptide level, providing stronger evidence of protein identity than immunoblotting alone.

Circular dichroism reveals a three-finger fold spectral signature in refolded rSm29

The far-UV CD spectrum of rSm29 displays a broad negative band centered at ~205 nm. The signal-to-noise ratio at lower wavelengths is partially affected by the buffer composition, which contains residual urea and salt that absorb in the far-UV region; however, the high tension (HT) voltage remained below 600 V throughout the entire spectral range (200–260 nm), confirming that the acquired signal is reliable and that the data are within acceptable quality thresholds for spectral interpretation [48]. This spectral profile is consistent with the structural characteristics of Ly6/uPAR (three-finger fold) family members, which are dominated by short β-strands, abundant β-turns, and loops projecting from a disulfide-stabilized core [42,43]. In such proteins, the classical β-sheet minimum at ~217 nm is shifted and broadened due to contributions from β-turns and disulfide bonds to the far-UV CD signal [49,50], resulting in a spectral envelope distinct from canonical antiparallel β-sheet proteins. These data provide experimental evidence that the refolding process from inclusion bodies yielded a structurally competent rSm29 protein [48] (Fig 5A-5D).

Download:

• PNG
  larger image
• TIFF
  original image

Fig 5. Circular dichroism analysis of recombinant Sm29.

(A) Raw circular dichroism (CD) spectra of recombinant Sm29 obtained from two independent measurements, showing the reproducibility of the spectral profile between runs. (B) Mean residue ellipticity (MRE) spectra calculated from the experimental data and compared with the Chou-Fasman secondary structure prediction and the non-negative least squares (NNLS) deconvolution fit. The experimental spectrum exhibited a minimum around 205 nm, suggestive of the presence of mixed secondary structural elements. (C) Secondary structure composition estimated from the amino acid sequence using the Chou-Fasman algorithm and from CD spectral deconvolution by the NNLS method, showing the relative contributions of α-helices, β-sheets, and random coil structures. (D) High-tension (HT) voltage profiles recorded during CD measurements, demonstrating that the signal quality remained within the recommended instrumental limits (<600 V) throughout the analyzed spectral range. CD measurements were performed using recombinant Sm29 at a concentration of 0.8 mg/mL in a 0.1-mm pathlength quartz cuvette.

https://doi.org/10.1371/journal.pntd.0014617.g005

Specific humoral response in mice after immunization with rSm29 formulations

The humoral immune response specific to the rSm29 protein was evaluated two weeks after the last immunization. Total IgG levels were determined by titration across a serial dilutions rage (Fig 6A), whereas specific IgG1, IgG2C, IgG3, and IgE levels were evaluated at a single serum dilution (Fig 6B-6E). This analysis aimed to compare the ability of vaccine formulations containing different adjuvants (Alum or CTVad1) to induce specific antibodies to rSm29 and suggest effector functions of immunoglobulin isotypes. Both immunization regimes (Sm29 + Alum and Sm29 + CTVad1) induced higher total IgG, IgG1, IgG2c, and moderate levels of IgG3, and IgE anti-rSm29 when compared to the control groups (Fig 6A-6E).

Download:

• PNG
  larger image
• TIFF
  original image

Fig 6. Quantification of antigen-specific antibody responses by ELISA.

(A) Serum titration for total IgG levels. Serum levels of IgG1 (B), IgG2c (C), IgG3 (D) and IgE (E). Antibody levels were measured by indirect ELISA two weeks after the final immunization. Plates were coated with recombinant Sm29 protein (10 μg/mL). Data are presented as individual values with mean ± SD. Statistical analyses were performed using ANOVA and Dunn’s multiple comparison test. P-values are indicated in the graph.

https://doi.org/10.1371/journal.pntd.0014617.g006

Cytokine production by splenocytes

To evaluate the cellular immune response profile induced by the different vaccine formulations, cytokines produced by mouse splenocytes were quantified 15 days after the last immunization, following in vitro stimulation with the rSm29 protein. The cytokines IFN-γ and IL-10 were quantified after 72 hours, while IL-4 and TNF were evaluated after 48 hours (Table 2). Regarding the cytokine profile, vaccination with rSm29 + Alum induced a mixed Th1/Th2 profile with significant production of IFN-γ, IL-10, and IL-4, while the rSm29 + CTVad1 immunized group showed a Th2-like pattern with production of IL-10 and IL-4. TNF was not detected in rSm29-stimulated cells, indicating that this cytokine is not relevant in the vaccine formulations tested (S2 Fig).

Download:

• PNG
  larger image
• TIFF
  original image

Table 2. Modulation of Sm29-Specific Cytokine Response by Different Adjuvant Formulations: Alum vs CTVad1.

https://doi.org/10.1371/journal.pntd.0014617.t002

rSm29 vaccination significantly reduces worm burden in mice

To evaluate the protective efficacy of the rSm29 formulated with either Alum or CTVad1, mice were challenged with 100 S. mansoni cercariae 15 days after the last immunization and perfused 45 days post-challenge to recover adult worms. Both vaccine formulations induced significant reductions in worm burden when compared with their respective adjuvant controls, as depicted in Table 3. Immunization with Alum+Sm29 resulted in a 34.1% reduction in worm recovery relative to the Alum group, while CTVad1 + Sm29 induced a 36.9% reduction compared with the CTVad1 group. These findings demonstrate that both formulations containing rSm29 elicited significant and comparable levels of protection against S. mansoni infection in mice.

Download:

• PNG
  larger image
• TIFF
  original image

Table 3. Protection level induced in C57BL⁄6 mice by immunization with recombinant Sm29 vaccine.

https://doi.org/10.1371/journal.pntd.0014617.t003

Histopathological evaluation of the liver

Histopathological evaluation was performed to investigate the impact of different vaccine formulations on the formation and organization of hepatic granulomas induced by S. mansoni infection. The number of concentric (well-defined) granulomas, and the number of eggs per granuloma were analyzed to correlate immune responses with the degree of tissue pathology.

The Sm29 + Alum group exhibited a 37% reduction in the number of granulomas compared to Alum alone, and the Sm29 + CTVad1 group showed a decrease in 39.2% in granulomas number compared to CTVad1 alone (Fig 7A). Additionally, a similar response was observed when analyzing the number of eggs per granuloma. The Sm29 + Alum immunized group showed a reduction in 33.1% in egg numbers and the Sm29 + CTVad1 vaccinated group presented 47.2% reduction compared to the respective controls (Fig 7B). The groups immunized with the Sm29 protein combined with adjuvants (Alum or CTVad1) exhibited lower number of concentric granulomas and schistosome eggs (Fig 7C) compared to their respective adjuvant-only control groups, suggesting that immunization with the recombinant Sm29 protein was effective in modulating the inflammatory response and mitigating infection-associated tissue damage.

Download:

• PNG
  larger image
• TIFF
  original image

Fig 7. Schistosoma-induced liver pathology is influenced by protein Sm29.

Pathological and molecular parameters were evaluated, after 6 weeks of infection in mice. (A) Number of granulomas per liver area, (B) number of eggs per gram of liver and (C) Representative images of granulomas detected in hematoxylin-eosin. The percentage of granuloma/liver areas were plotted in box plots (median, maximum and minimum) and analyzed via the Kruskal–Wallis test followed by Dunn’s multiple comparison test. The number of granulomas per liver area and the number of eggs per granuloma were evaluated using one-way ANOVA followed by Tukey's multiple comparison test. An asterisk denotes statistically significant differences when *p < 0.05 and ns indicates no statistical difference. Scale bars: 500µm (upper panels) and 100 µm (lower panels).

https://doi.org/10.1371/journal.pntd.0014617.g007