Identification of alterations of immunometabolism associated with Pompe disease

Abstract

Pompe disease (PD) is a neuromuscular lysosomal storage disorder caused by mutations in the GAA gene, characterized by progressive glycogen accumulation in multiple tissues and autophagy and metabolic abnormalities. While immunological changes have largely been overlooked as part of PD's symptomatology, autophagy and metabolic regulation are crucial in immune cell function. High incidence of immune reactions against therapeutic recombinant GAA (rhGAA) in PD patients suppose an important hindrance to treatment efficacy, yet the impact of GAA deficiency on the immune system remains unclear. Here we explored the T cell phenotype in late-onset PD patient cells and in a PD mouse model, revealing heightened expression of activation markers in effector T cells compared to controls. Additionally, we observed decreased frequencies of regulatory T cells in mice. We also hypothesized that metabolic and autophagy defects in PD extend to immune cells. In fact, Gaa^−/− T cells reproduced autophagy and mitophagy defects reported in muscle cells, and upon stimulation, T cells showed impaired mitochondrial function consistent with defective mitophagy. Preliminary findings also suggest that alterations translate to a subset of CD24⁺CD172⁻ conventional dendritic cells with regulatory function, which could indirectly contribute to higher T cell activation. Our observations indicate that immune homeostasis is altered in PD, offering new insights on immune dysfunction in the context of lysosomal impairment and providing the rationale for investigating its potential role in the pathogenesis of PD.

Supplementary Information

The online version contains supplementary material available at 10.1007/s00018-026-06135-y.

Introduction

Pompe disease (PD), also known as glycogen storage disease type II (GSDII) or maltase deficiency, is a rare lysosomal storage disorder (LSD) caused by mutations on the GAA gene that encodes the lysosomal enzyme acid α-glucosidase (GAA, acid maltase) [1], an ubiquitous enzyme responsible for the breakdown of glycogen into glucose in the lysosomes [2]. GAA deficiency is linked to progressive muscle degeneration [3–6], cardiac hypertrophy, respiratory insufficiency and cognitive alterations [7–9]. Although there is a continuous gradient of disease severity, PD is classified into two main subtypes, classical infantile-onset Pompe disease (IOPD) and late-onset Pompe disease (LOPD) [5, 10], with the age of disease onset directly correlated to the residual levels of GAA activity [11].

Studies in skeletal muscle fibers show that progressive glycogen buildup in lysosomes ultimately impairs their fusion with autophagosomes to degrade their content [12, 13]. This blockade of the autophagic flux leads to the accumulation of autophagy substrates, as evidenced by increased levels of the ubiquitin-binding protein p62 in muscle biopsies [12, 14–17] and a high mitochondrial mass compared to healthy controls due to impaired mitophagy [16, 18]. Mitophagy defects can be detected by the accumulation of mitochondrial markers such as the subunit A of the succinate dehydrogenase complex (SDHA) or complex IV (COX IV) of the electron transport chain (ETC), both found on the inner mitochondrial membrane [16, 19].

Enzyme replacement therapy (ERT) based on regular infusions of recombinant human GAA protein (rhGAA) is available since 2006 [20–22]. ERT has been a breakthrough thanks to its efficacy at reverting the cardiomyopathy and improving survival in infantile patients [21, 23], and stabilizing the disease progression in late-onset patients [24, 25]. However, the efficacy of ERT has turned out to be highly variable and dependent on the stage of disease progression at which ERT is initiated [3]. Immune responses against the infused protein are another important parameter affecting the outcome of ERT, sometimes leading to loss of treatment efficacy and to poor disease prognosis. rhGAA treatment often triggers humoral responses [26], and can be accompanied by allergic reactions [21], especially in patients who have null mutations. These patients, known as cross-reactive immunologic material (CRIM)-negative, present no central immune tolerance to the protein [27] and are more prone to developing immune reactions against ERT [28–30]. In contrast, CRIM-positive patients have detectable but defective GAA protein forms with reduced function, yet an important fraction of CRIM-positive IOPD and LOPD patients still develop antibodies at variable titers [31, 32]. Given the determining impact of immune responses on the success of ERT, preventive immunosuppression regimens are currently administered concomitantly with rhGAA [27, 33, 34], often combined with anti-histaminic drugs to prevent hypersensitivity reactions [35, 36]. This highlights the challenge of immunological reactions in PD patients independently of residual endogenous GAA expression, with ongoing efforts to improve the efficacy of current immunomodulatory strategies [37].

Whereas most research addressing the PD phenotype is focused on the impact of glycogen accumulation and autophagic buildup on muscle impairment [12, 38], there are no reports on how pathological glycogen accumulation affects the regulation of immune cells. Glycogen synthesis upon T cell activation has been shown to be important for the formation and survival of memory CD8⁺ T cells [39], and several studies show the essential role of autophagy and mitochondrial integrity in controlling metabolism, activation and differentiation of effector and regulatory T cells (Teffs and Tregs, respectively) [40–43], as well as dendritic cells (DCs) [44]. Importantly, glycogen accumulation in lymphocytes from PD patients has been reported and is used to diagnose PD [45, 46]. In a previous study, we showed that culturing peripheral blood mononuclear cells (PBMCs) from LOPD patients naive to ERT in the presence of rhGAA resulted in the secretion of high levels of pro-inflammatory cytokines in vitro [47]. This excessive response was unexpected given that patients were CRIM-positive and were never exposed to rhGAA. These results revealed an unexpectedly high immune response against the protein, setting the rationale for further immunological studies in PD.

In this study we focused on characterizing the phenotype of T cells isolated from LOPD patients [8, 48], and performed a detailed immune phenotyping of the Gaa^−/− mouse model (B6;129Gaatm1Rabn/J) first described by Raben and colleagues [49]. While these mice harbor a null mutation in the Gaa gene, they exhibit an intermediate phenotype between infantile and late-onset PD [17, 49]. Gaa^−/− mice present glycogen accumulation in all tissues since birth, cardiomegaly and progressive muscle weakness detectable from 4 months of age [17, 49], becoming severe to lethal by 7–9 months of age [17, 49]. Given the reproducibility of symptoms of the human pathology, this model is often used to study disease mechanisms and for pre-clinical evaluation of therapeutic approaches.

Our present work identifies alterations in T cell homeostasis, characterized by a tendency for a high activation phenotype of Teffs from patients and PD mice together with a defect in the induction and suppressive function of murine Tregs. Importantly, T cells isolated from Gaa^−/− mice presented similar autophagy and mitochondrial defects than those reported in muscle cells, revealing the accumulation of autophagy markers and a diminished oxidative phosphorylation (OXPHOS) rate. Finally, given the upstream role of DCs in regulating T cell functions, we provided a preliminary characterization of murine DCs. Our studies showed altered frequencies in the conventional cDC1 subset, characterized by CD24 expression [50, 51], together with altered regulatory DC phenotype shown by reduced IDO1 expression and increased production of inflammatory cytokines. Altogether, our observations may provide a rationale for stratifying PD patients based on the predicted immune reactivity against ERT.

By providing mechanistic insights into immune cell alterations associated with lysosomal and metabolic impairments in PD, our work opens venues for the optimization of immunomodulatory strategies aimed at delaying the disease progression while improving the efficacy and safety of ERT, through targeting of altered immune cell populations such as Tregs or regulatory DCs.

Materials and methods

Human samples

De-identified healthy donor PBMCs were obtained from the French blood bank (Etablissement Française du Sang, EFS). The study was approved by the competent health authorities and by the Ethics committee of the Pitié-Salpêtrière Hospital in Paris (CNIL N/Ref MMS/CWR/AR155497; CCTIRS N: 14.520; CCP approval 25/06/2014). Peripheral blood mononuclear cells (PBMCs) from untreated LOPD patients were obtained from the French registry of adult Pompe disease patients [48]. Subjects were male and female, and ages ranged between 37 and 79 years. Participants gave their informed consent to be included in the study. Exclusion criteria were age older than 80 years, ongoing or recent (less than 3 months) immunosuppressive treatment, malignancy not in remission, immunodeficiency or autoimmunity. Blood samples were collected via venipuncture in heparin tubes and used to isolate serum and PBMCs.

Mice

Mouse studies were performed according to the French and European legislation on animal care and experimentation (2010/63/EU) and approved by Genethon’s ethical committee (protocol 2015–008 #3547). Male and female Pompe disease mice (Gaa^−/−, B6;129-Gaatm1Rabn/J, stock no. 004154) were purchased from the Jackson Laboratory. Gaa^+/+ mice with the same genetic background (JAX #004154) were used as healthy, wild-type (WT) controls. Genotypes were confirmed by polymerase chain reaction (PCR) performed on genomic DNA with oligonucleotides specific for the mutated region: GAA mutant forward 5’ CGTTGGCTACCCGTGATATT 3’, GAA wild type forward 5’ TCCTGAGCCCAAACACTTCT 3’, GAA common reverse 5’ ATTGTTGCACAACGCTCTTG 3’. Mice were sacrificed at 4–5 months of age for the isolation of spleen and bone marrow.

Splenocyte isolation

For studies in Gaa^−/− mice, we analyzed freshly isolated splenocytes from 4–5-month-old mice, which is the reported onset age of PD symptoms in this model [17, 49]. Spleens were homogenized with a 70 μm nylon cell-strainer by grinding the organ against the strainer with the use of a syringe plunger. Filters were washed with 10 ml RPMI, and cells were then pelleted at 400 × g for 5 min. The supernatant was discarded, and red blood cells were lysed with 4 ml ACK buffer (Thermo Fisher Scientific) for 5 min at room temperature. The reaction was stopped with 5 ml RPMI supplemented with 10% fetal calf serum (FCS, Gibco, Thermo Fisher Scientific), previously decomplemented by heating for 30 min at 56 °C. Cells were pelleted and resuspended in 5 ml RPMI (Gibco, Thermo Fisher Scientific) 10% FCS before continuing with experiments.

Glycogen assay

Pelleted splenocytes were lysed in distilled water and frozen at −80 °C before the start of the assay, followed by thawing and vortex. Part of the lysate was used to quantify protein content by BCA Protein Assay (Thermo Fisher Scientific, Waltham, MA). Then, 20 μl of lysate properly diluted was mixed with 55 μl of distilled water, incubated for 5 min at 95 °C, and cooled at 4 °C. Next, 25 μl of amyloglucosidase from Aspergillus Niger (Sigma-Aldrich, Saint-Louis, MO) diluted 1:50 in 0.1 M potassium acetate pH 5.5 was added to each sample. A control reaction without amyloglucosidase was done in parallel. Both digestion and control samples were incubated at 37 °C for 90 min, followed by 5 min at 95 °C. Glycogen content was determined indirectly by quantifying the number of glucose molecules released per gram of protein, with the use of a colorimetric glucose assay kit (Sigma-Aldrich). Glucose assay was performed according to the manufacturer’s instructions. Briefly, 25 μl of samples or glucose standard properly diluted were plated in a transparent 96-well plate with flat bottom, together with 50 μl of working reagent. Samples were incubated for 30 min at 37 °C. The reaction was stopped with 50 μl H2SO4 12N. Absorbance was measured with an EnSpire alpha plate reader (PerkinElmer, Waltham, MA) at 540 nm.

Western blot analyses

Homogenates were prepared as described for the analysis of glycogen content from total splenocytes or from total T cells sorted using the Dynabeads™ Untouched™ Mouse T Cells Kit (Thermo Fisher Scientific). SDS-page electrophoresis was performed with NuPAGE 4–12% Bis–Tris protein gels (Life technologies, Carlsbad, CA). After transfer, membranes were blocked with Odyssey buffer (Li-Cor Biosciences, Lincoln, NE) and incubated with primary antibodies: anti- SQSTM1/p62 (mouse monoclonal, Abcam, Cambridge, MA, USA), anti-vinculin (mouse monoclonal, clone hVIN-1, Sigma-Aldrich), anti-LAMP1 (rabbit polyclonal; Abcam), anti-COX IV (rabbit polyclonal; Abcam), and anti-SDHA (mouse monoclonal; Invitrogen, Waltham, MA). The membrane was washed and incubated with the appropriate secondary antibody (LI-COR Biosciences), conjugated with fluorescent probes with extended linear range, and visualized by the Odyssey imaging system (LI-COR Biosciences), which allows identification and exclusion of saturated signals during band quantification.

In vitro stimulation of PBMCs and murine splenocytes

Cells were incubated at 37 °C with 5% CO2 between 24 and 72 h. Human PBMCs were maintained in serum-free AIM-V Glutamax medium (Gibco, Thermo Fisher Scientific), whereas murine splenocytes were cultured in RPMI medium (Gibco, Thermo Fisher Scientific) supplemented with 10% FCS. For the stimulation of T cells, anti-human CD3e monoclonal antibody (0.5 µg/ml) (clone OKT3, Functional grade, Thermo Fisher Scientific) or anti-mouse CD3e monoclonal antibody (1.5 µg/ml) (clone 45-2C11, Functional grade, Thermo Fisher Scientific) were coated on sterile 96-well tissue culture plates one day before seeding the cells. Anti-human CD28 monoclonal antibody (clone 28.2, Thermo Fisher Scientific) or anti-mouse CD28 monoclonal antibody (clone 37.51, Functional grade, Thermo Fisher Scientific) was added at 2 μg/ml. For stimulation of PBMCs with CpG-ODN, cells were incubated with CpG-ODN 2006 (human TLR9 agonist) (Invivogen, San Diego, CA) at a concentration of 3 µg/ml, in 96-well plates. Non-stimulated controls from each subject were included in all experiments.

BMDC differentiation

Bone marrow was harvested from femur, tibia and pelvis using mortar and pestle in 1 × PBS supplemented with 0.5% BSA and 2 mM EDTA (MACS buffer), passed through a 70 μm cell strainer and centrifuged at 1400 rpm for 5 min. Red blood cells were lysed with ACK lysis buffer (Ammonium Chloride 0.15 M, Potassium Carbonate 10 mM) and debris were removed by a gradient centrifugation using Histopaque1119 (Sigma-Aldrich) prior to culture. Cells were resuspended at 2 × 10⁶ cells/ml in Iscove’s Modified Dulbecco’s Media (IMDM, Thermo Fisher) supplemented with 0.1 mM Non-essential Amminoacids (Thermo Fisher), 1 mM Sodium Pyruvate (Thermo Fisher), 5 mM glutamine (Thermo Fisher), 50 μM 2-Mercaptoethanol (Thermo Fisher), 100 U/ml penicillin, 100 g/ml streptomycin (Thermo Fisher) and 10% FBS (Thermo Fisher) (complete IMDM) containing 5% murine Flt3-L and were seed 5 ml/well in 6-plate tissue culture plates at 37 °C for 9 days.

In vitro antigen presentation assay using OT-I cells

OT-I CD8⁺ T cells were isolated from spleens as previously described [50]. Briefly, spleens were excised and digested in complete IMDM containing 250 μg/mL collagenase B (#11088815001, Roche) and 30 U/mL DNase I (#D4527, Sigma-Aldrich) at 37 °C for 30 min. After red blood cell lysis with ACK buffer, OT-I cells were identified and sorted as B220⁻ CD11c⁻CD4⁻CD8⁺TCR-Vα2⁺ by flow cytometry. Purified OT-I cells were then labeled with 1 μM CFSE (#C34554, Thermo Fisher Scientific) to monitor proliferation. For the antigen presentation assay, conventional dendritic cells (cDCs) were sorted from day 9 FLT3-L cultures of WT or Gaa⁻^/⁻ bone marrow precursors (BMDCs) and cultured in complete IMDM for 36 h. Subsequently, 1 × 10⁴ cDCs were co-cultured with various concentrations of soluble ovalbumin (OVA; #A5503, Sigma-Aldrich) and 2.5 × 10⁴ CFSE-labeled OT-I CD8⁺ T cells for 72 h. T proliferation was evaluated by flow cytometry based on CFSE staining gated on CD8⁺CD44⁺ cells.

Staining of PBMCs and murine cells

For intracellular staining (ICS) of cytokines, human PBMCs were stimulated overnight with plate-bound anti-CD3 and anti-CD28 as previously described. After 18 h stimulation, cytokine secretion was blocked by adding Brefeldin A (Golgi Plug, Sigma Aldrich) and monensin (GolgiStop, BD Biosciences, Franklin Lakes, NJ) at 1 μg/ml and 2 μM respectively. After 6 additional hours of incubation, cells were stained for human T cell surface markers CD3 (ECD, Beckman Coulter, Brea, CA), CD4 (PE-Cy7, eBioscience), CD8 (PercP-Cy5.5, eBioscience), CD25 (Alexa Fluor 700, Biolegend, San Diego, CA), CD69 (Brilliant violet 786, BD Biosciences) and labelled with Zombie Yellow Fixable Viability Kit (Biolegend). Next, intracellular staining was performed with antibodies against IL-2 (APC, eBioscience), IFNγ (PE, eBioscience) and TNFα (Brilliant Violet 510, Biolegend) diluted in Perm/Wash buffer (BD Biosciences).

Murine splenocytes were stained with anti-murine CD3 (APC-eF780, eBioscience), CD3 (APC, Biolegend), CD4 (V500 and BV510, BD Biosciences) CD8 (Alexa Fluor 700, eBioscience), CD8 (PercP-Cy5.5, BD Biosciences), Annexin V (PerCP-eF710, eBiosciences), CD25 (PE and PE-Cy7, eBiosience), CD69 (BV786, BD Bioscience), CD62L (BV605, BD Biosciences), CD44 (BV421, BD Biosciences), B220 (BV786, BD Biosciences), Bst2 (APC, eBioscience), MHC-II (BV510, BD Biosciences), CD11c (APC-Cy7, eBioscience), CD24 (PE-Cy7, Biolegend) or CD172 (PercP-Cy5.5, eBioscience). Viability staining was performed with Zombie Yellow Fixable Viability Kit (Biolegend) or with Fixable Viability Dye eF780 (FVD, eBioscience). Intracellular staining of Foxp3 (Alexa Fluor 647, BD Biosciences) was performed after fixing and permeabilizing cells with eBioscience Foxp3/Transcription factor staining buffer set (Invitrogen, Thermo Fisher Scientific). For the analysis of cell proliferation, cryopreserved human PBMCs or freshly isolated murine splenocytes were labelled with Cell Proliferation Dye eFluor450 20 μM (affymetrix, eBioscience) before plating them on anti-CD3 coated plates, and harvested after 72 h. Mitochondrial mass was analyzed by staining fresh splenocytes with Mitotracker Green 100 nM (Thermo Fisher Scientific). For flow cytometry analyses, cells were acquired on a Cytoflex flow cytometer (Beckman Coulter). Data analysis was performed using Flowjo software version 10.6 (Tree star, Ashland, OR, United States). For confocal imaging, stained splenocytes in suspension were cytospun onto glass slides and acquired with LAS X 3.5.5.1 Software (Leica Microsystems, Germany). Images were analyzed with ImageJ 1.54p.

Treg suppression assay

Tconv and Tregs cells were purified with the use of CD4⁺CD25⁺ Regulatory T cell isolation kit (Miltenyi Biotec, Bergisch Gladbach, Germany). Next, naive CD4⁺CD25⁻ cells from WT mice were labelled with CellTrace CFSE Cell Proliferation Kit (Thermo Fisher Scientific), and cultured for 2 days with various numbers of CD4⁺CD25^hi Tregs from WT or Gaa^−/− mice (from 50,000 to 0), in the presence of soluble anti-CD3 (1 μg/ml, affinity-purified clone 2C11) and anti-CD28 (1 μg/ml, affinity-purified clone PV-1) (both from Thermo Fisher Scientific). Proliferation was determined by analysis of CFSE staining in T cells by flow cytometry.

OCR and ECAR measurements

OCR and ECAR were measured using the XF96 extracellular flux analyzer (Agilent Technologies Inc., Santa Clara, CA). Assays were performed with CD4⁺ and CD8⁺ T cells per well previously sorted from freshly isolated splenocytes by negative selection using the EasySep™ Mouse Naïve CD4⁺ T Cell Isolation kit or EasySep™ Mouse CD8⁺ T Cell Isolation kit (STEMCELL Technologies, St. Louis, MO) respectively. Previous to splenocyte isolation, XF96 microplates were coated with anti-mouse CD3e monoclonal antibody at 1.5 µg/ml, whereas anti-mouse CD28 was added soluble at 2 µg/ml as previously described. Seahorse tests were performed after 24 h stimulation of sorted T cells, time at which media was replaced by unbuffered XF assay media (Agilent Technologies). Biological samples were plated in experimental triplicates. Mitochondrial activity was assessed with Seahorse XF Cell Mito Stress Test kit (Agilent Technologies). For oxygen consumption rate (OCR) measurements, compounds were injected during the assay at the following final concentrations: oligomycin (ATP synthase inhibitor to measure respiration associated with cellular ATP production, 1 μM), FCCP (uncoupling agent to measure the maximal respiration capacity; 1.5 μM), rotenone and antimycin A (electron transport chain inhibitors to measure the non-mitochondrial respiration, 1 μM each). Glycolytic activity was assessed by Seahorse XF Glycolysis Stress Test (Agilent Technologies). Extracellular acidification rate (ECAR) measurements were performed in basal conditions and upon sequential injection of glucose 20 mM, 1 μM oligomycin A and 2-deoxyglucose 25 mM (2-DG) to determine the glycolytic rate and the glycolytic capacity of tested cells.

2-NBDG uptake

Splenocytes were stimulated for 24 h in a 96-well plate with plate-bound anti-CD3 (1.5 µg/ml) and soluble anti-CD28 (2 µg/ml). Then, cells were incubated in DMEM without glucose during 1 h followed by incubation in the presence of 150 µg/ml 2-NBDG during 10 min, washed two times with PBS and stained with anti-CD4 (V500, BD Biosciences) and anti-CD8 (AF700, eBioscience) for flow cytometry analysis.

Luminex assay

Analysis of cytokine concentration from supernatans was performed after 24 h stimulation of murine splenocytes with LPS at 250 ng/ml with the use of Milliplex HCYTOMAG-60 K kit (Merck-Millipore). The concentration of analytes was measured with a Bio-Plex MAGPIX Multiplex reader (Bio-rad, Hercules, CA).

Single cell RNA-seq analysis

Single-cell RNA sequencing data were downloaded from the Gene Expression Omnibus (GEO) using samples from datasets GSE226488 (GSM7077865, GSM7077866) and GSE244804 (GSM7829917), which include PBMCs from healthy donors and splenocytes from healthy C57BL/6 J mice, respectively. Data processing and analysis were performed using the Seurat package [52] following a standard pre-processing workflow. Cells with mitochondrial content exceeding 10% or fewer than 200 detected genes were excluded. Doublets and multiplets were removed using DoubletFinder [53]. The filtered data were normalized (scale factor: 10,000), regressing out the number of unique molecular identifiers (nUMI) using a negative binomial model, followed by log-transformation. Highly variable genes were identified using the FindVariableFeatures function for downstream analyses. After scaling, principal component analysis (PCA) was performed on the top 3,000 variable genes, followed by clustering with the FindClusters function in Seurat. Uniform Manifold Approximation and Projection (UMAP) was used for visualization. Batch effects in GSE226488 were corrected using Harmony [54]. Cell populations were annotated using the SingleR package [55] and manually curated. All visualizations and graphs were generated using ggplot2.

Statistical analysis

Data are expressed as mean ± SD or mean ± SEM as indicated in the figure legends. GraphPad Prism 9.5 software was used for statistical analyses. P-value < 0.05 was considered significant. Non-parametric Mann–Whitney t-test was used for two-group comparisons, whereas multiple comparisons were performed with two-way ANOVA with Bonferroni or Tukey post-hoc correction as indicated in figure legends.

Results

LOPD T cells show enhanced activation phenotype upon in vitro stimulation

We initially used publicly available data to examine GAA expression in healthy-donor (HD) human PBMCs analyzed by single-cell RNA sequencing (scRNAseq). Results in Fig. S1A, B show that GAA is expressed across different immune populations, with the highest levels found in myeloid cells (Fig. S1C). Then, given our previous observation that PBMCs derived from untreated LOPD patients become highly activated upon incubation with rhGAA [47], we set to specifically assess the activation state of LOPD T cells that were naive to ERT at the time of blood collection, to avoid confounding results associated with GAA enzyme supplementation. PBMCs from LOPD and healthy donors were labelled with proliferation dye and stimulated with plate-bound anti-CD3 and soluble anti-CD28 antibodies [56]. Cells were harvested after 24 and 72 h of stimulation to assess the expression levels of activation markers and T cell proliferation, respectively, by flow cytometry. While no significant differences were observed under basal conditions (Fig. S2), results after 24 h-stimulation revealed a higher induction of surface activation marker CD25. We also observed significantly higher expression of the memory/effector marker CD44 in CD8⁺ T cells, while no differences were detected in the expression of the early activation marker CD69 (Fig. 1A, B). Intracellular staining further demonstrated that LOPD T cells produced significantly higher levels of IL-2 in both CD4⁺ and CD8⁺ T cells, and TNFα in CD4⁺, compared to HD-derived cells (Fig. 1C, D). In line with these findings, proliferation analysis at 72 h indicated that both CD4⁺ and CD8⁺ T cells exhibited a significantly higher proliferation rate compared to HD cells (Fig. 1E, F), confirming the heightened activation and effector state of T cells derived from LOPD individuals.

Fig. 1.

LOPD patient-derived T cells show an enhanced activation state upon in vitro stimulation. A Representative histograms showing flow cytometry staining of surface activation markers CD44, CD25 and CD69 gated on CD3⁺/CD4⁺ and and CD3⁺/CD8⁺ PBMCs after 24 h stimulation with anti-CD3 (0.5 µg/ml) and anti-CD28 (2 µg/ml) antibodies compared to non-stimulated cells. B Percentage of CD44⁺, CD25⁺ and CD69⁺ cells gated on CD3⁺/CD4⁺ and CD3⁺/CD8⁺ PBMCs respectively. C, D Representative flow cytometry plots and quantification of the percentage of CD4⁺ (C) and CD8⁺ (D) PBMCs positive for intracellular cytokines IL-2, IFNγ and TNFα after 24 h stimulation with anti-CD3 and anti-CD28. E Representative flow cytometry histograms showing the gating strategy on proliferation dye eF450^low CD4⁺ and CD8⁺ PBMCs from HD and LOPD donors after 72 h stimulation with anti-CD3 and anti-CD28. F Quantification of the percentage of proliferating CD4⁺ and CD8⁺ PBMCs. For B, C, D and F data are shown as mean ± SD between 6 donor samples per group. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. NS, non-stimulated; HD, healthy donor; LOPD, late-onset Pompe disease

Effector T cells from Pompe disease mice show elevated basal activation phenotype together with defective activation of CD4⁺Foxp3⁺ Tregs

Due to the scarcity of patient-derived samples from untreated PD individuals, we conducted further characterization of immune cells in Gaa^−/− mice [16, 17, 49, 57]. ScRNAseq again showed Gaa expression across different splenocyte populations in wild-type (WT) C57BL/6 J mice (Fig. S3A, B), with higher expression in certain myeloid subsets (Fig. S3C). We then examined whether T cells from Gaa-deficient mice displayed an altered activation phenotype in comparison to wild-type (WT) controls, as observed in humans.

In contrast to our observations in LOPD samples, we noted a significantly higher frequency of CD4⁺ and CD8⁺ T cells expressing the surface activation markers CD25, CD69 and CD44, already in basal condition mainly in the CD8⁺ compartment (Fig. 2A, B). Moreover, Gaa^−/− CD8⁺CD44⁺ cells showed higher expression levels of the PD1 marker (Fig. 2A), suggestive of sustained activation. These results were consistent with the identification of a higher frequency of CD4⁺ and CD8⁺ T cells presenting an active CD62L⁻CD44⁺/CD25⁺CD69⁺ effector memory (TEM) phenotype (Fig. S4A-D), as well as higher expression of late-apoptotic markers, particularly among CD8⁺ cells (Fig. S4E, F). These results support the notion of chronic activation within this T cell subset. Interestingly, we also observed a significantly lower percentage of Tregs (CD4⁺Foxp3⁺) in spleens from PD mice (Fig. 2C). Together, these findings indicate a disturbance in T cell homeostasis and Treg frequencies.

Fig. 2.

Effector T cells from GAA-deficient mice show elevated basal activation phenotype together with defective activation of CD4⁺Foxp3⁺ Tregs. A, B Flow cytometry analysis of T cell markers CD25, CD69, CD44 and PD1 in freshly isolated splenocytes gated on CD8⁺ (A) and CD4⁺ (B) T cells. C Flow cytometry analysis of the frequency of CD4⁺Foxp3⁺ T cells from fresh splenocytes. Data in A-C are shown as mean ± SD between samples from 5 wild-type (WT) and 6 Gaa^−/− female mice. Statistical significance was determined by Mann–Whitney t-test. D, E Analysis of the expression of CD69 (D) and CD25 (E) after 24 h incubation with plate-bound anti-CD3 (1.5 µg/ml) and anti-CD28 (2 µg/ml). F Representative flow cytometry plots showing the percentage of Foxp3⁺CD25⁺ and Foxp3⁻CD25⁺ cells gated on CD3⁺/CD4⁺ T cells from stimulated WT and Gaa^−/− splenocytes. F Quantification of CD25⁺ cells gated on Foxp3⁻ (left) or Foxp3⁺ (right) CD4⁺ T cells upon stimulation. Data in D, E and F are shown as mean ± SD between 11 WT and 12 Gaa^−/− female mice pooled from two independent experiments. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. NS; non-stimulated

To specifically test T cell activation, we stimulated splenocytes with plate-bound anti-CD3 and anti-CD28 antibodies for 24 h (Fig. 2D-G). Notably, upon stimulation, the expression of the early activation marker CD69 was comparable between WT and Gaa^−/− T cells (Fig. 2D). However, we observed a defective induction of CD25 in the CD4⁺, but not the CD8⁺ subset (Fig. 2E). Remarkably, this defective induction of CD25 was more pronounced within the CD4⁺Foxp3⁺ population, while the frequency of Foxp3⁻CD25⁺ cells exhibited similarity between WT and Gaa^−/− CD4⁺ T cells (Fig. 2F, G), pointing towards specific activation defects within the Foxp3⁺ Treg compartment. Nevertheless, analysis of cell proliferation after 72-h stimulation did not reveal significant differences in the proliferation levels of Foxp3⁺ cells. Instead, we observed a slightly higher percentage of total proliferating Gaa^−/− CD4⁺ T cells compared to WT controls (Fig. S5).

Given the lower frequency of CD4⁺Foxp3⁺ cells detected under basal conditions (Fig. 2C) and the defective induction of Foxp3⁺CD25⁺ cells upon stimulation (Fig. 2D-F), we also investigated whether these findings correlated with functional impairments in Tregs from PD mice. To achieve this, we sorted CD4⁺CD25^hi Tregs from fresh splenocytes of both WT and Gaa^−/− mice and subsequently co-cultured them with CD4⁺CD25^lo conventional T cells (Tconv) from WT mice, at various Tconv:Treg ratios. Results reported in Supplementary Fig. 6 show a slightly lower suppressive potential of PD Tregs compared to their WT counterparts, which only reached statistical significance at the 1:16 Treg:Tconv ratio. Altogether, our data suggest that Gaa^−/− T cells present a heightened basal activated phenotype together with a defect in the Treg compartment.

Murine Gaa^−/− T cells replicate autophagy and mitophagy defects characteristic of muscle cells

We next examined the levels of autophagy and mitophagy markers, including p62, SHDA, and COX IV (COX4I1) in T cells from WT and Gaa^−/− mice stimulated with anti-CD3 and anti-CD28 antibodies for 24 h, to specifically stimulate T cells (Fig. 3A, B). Stimulated cells from Gaa^−/− mice exhibited significantly higher levels of all three markers, similar to what is observed in PD muscle tissue [16, 17]. To confirm the presence of autophagy defects in T cells, we sorted total T cells from fresh splenocytes and examined the levels of Gaa mRNA, glycogen, and the lysosomal marker LAMP1, which is known to accumulate in muscle fibers from Gaa^−/− mice [3, 16, 58]. Our findings confirmed the expression of Gaa mRNA in WT T cells compared to Gaa^−/− (Fig. 3D), the elevated glycogen content (Fig. 3D) and significant accumulation of LAMP1 (Fig. S7A-D) in Gaa^−/− cells. Furthermore, analysis of mitochondrial content using MitoTracker Green staining in CD4⁺ and CD8⁺ T cells validated the notable mitochondrial buildup in fresh murine Pompe T cells, particularly in the CD4⁺ T cell subset (Fig. 3E, F, Fig. S7E, F), consistent with the inhibition of mitophagy. Collectively, our investigations demonstrate that T cells from Pompe mice replicate the autophagy and mitophagy defects characteristic of muscle fibers, resulting from the accumulation of lysosomal glycogen.

Fig. 3.

Gaa^−/− T cells present altered autophagy and mitophagy. A, B Western blots (A) and quantification (B) of SHDA, p62 and COX IV in splenocytes from 5-month-old female WT (n = 5) and Gaa^−/− mice (n = 4) stimulated for 24 h with anti-CD3 and anti-CD28. Vinculin was used as normalizer. Data are shown as mean ± SD. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. C Relative Gaa mRNA expression in sorted total T cells expressed as ΔΔCt values (n = 6). β-actin expression levels were used to normalize Gaa expression. D Glycogen content in sorted total T sorted cells expressed as nmol glucose/g protein (n = 6). E, F Representative flow cytometry histograms (E) and analysis (F) of Mitotracker green mean fluorescence intensity (MFI) in freshly isolated splenocytes, gated on CD4⁺ and CD8⁺ cells (n = 5). Data in C, D and F are shown as mean ± SD. Statistical significance was determined by Mann–Whitney t-test

Induction of oxidative phosphorylation is impaired in stimulated Gaa^−/− T cells

Because autophagy and mitochondrial metabolism are important regulators of T cell activation [59, 60], and play an important role in Treg survival and function [42, 43, 61], we next assessed the oxidative phosphorylation (OXPHOS) (Fig. 4) and glycolysis (Fig. 5) rates in sorted murine CD4⁺ and CD8⁺ T cells. Seahorse XF Cell Mito Stress Test showed that both CD4⁺ and CD8⁺ Gaa^−/− T cells have significantly lower oxygen consumption rates (OCR) compared to WT controls in terms of basal respiration (CD8⁺ only), maximal respiration and spare respiratory capacity upon stimulation (Fig. 4A-C).

Fig. 4.

Induction of oxidative phosphorylation is impaired in stimulated Gaa^−/− T cells. A Oxygen consumption rate (OCR) measurements from sorted CD4⁺ and CD8⁺ T cells after 24 h stimulation with αCD3/CD28. Data are representative of two independent experiments. B, C Mito stress test measurements in sorted CD4⁺ (B) and CD8⁺ T (C) cells after 24 h stimulation with αCD3/CD28. N = 8 mice per group from two independent experiments. Data are shown as mean ± SD. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. Oligo, oligomycin A; FCCP, Fluoro-carbonyl cynade phenylhydrazon; Rot, Rotenone; AA, antimycin A

Fig. 5.

Analysis of glycolysis rate in T cells. A Extracellular acidification rate (ECAR) measurements from sorted CD4⁺ T cells after 24 h stimulation with αCD3/CD28. Data are representative of two independent experiments and shown as mean ± SEM. B Glycolysis stress test measurements in sorted CD4⁺ cells after 24 h stimulation with αCD3/CD28. N = 8 mice per group from two independent experiments. C Extracellular acidification rate (ECAR) measurements from sorted CD8⁺ T cells after 24 h stimulation with αCD3/CD28. Data are representative of one experiment and shown as shown as mean ± SEM. D Glycolysis stress test measurements in sorted CD8⁺ T cells after 24 h stimulation with αCD3/CD28. N = 4 mice per group. For B and D, Data are shown as mean ± SD. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. E Representative flow cytometry plots showing the percentage of 2-NBDG⁺ cells in CD3/CD28-stimulated CD4⁺ and CD8⁺ T cells (gated on CD3⁺). F Analysis of fluorescently-labelled glucose uptake (2-NBDG) in WT (n = 5) and Gaa^−/− (n = 6) splenocytes stimulated for 24 h with plate-bound anti-CD3 and anti-CD28 antibodies. Data are shown as mean ± SD. Statistical significance was determined by two-way ANOVA with Bonferroni post-hoc test. Oligo, Oligomycin; 2-DG, 2-Deoxy-D-glucose

Analysis of the glycolysis rate using the Seahorse XF Glycolysis Stress Test didn’t reveal differences between Gaa^−/− and WT cells (Fig. 5A-D), suggesting a lack of a compensatory mechanism between OXPHOS and glycolysis. Nevertheless, staining of stimulated splenocytes with the fluorescently labelled glucose analog 2-NBDG showed significantly lower glucose uptake in stimulated CD4⁺ T cells but not in CD8⁺ T cells from Gaa^−/− mice compared to WT cells (Fig. 5E, F), indicating a potential energetic deficiency in these cells that has also been reported in skeletal muscle cells from PD mice [58, 62].

In all, our studies show that murine Gaa^−/− T cells exhibit autophagy and metabolic defects akin to those observed in skeletal muscle cells, which could be linked to the altered activation phenotype.

GAA-deficient DCs exhibit an increased pro-inflammatory phenotype and altered cDC1 frequencies

Because DCs are upstream of T cell activation in the antigen recognition process, and given the relatively abundant GAA expression detected in these cells (Fig. S3), we next focused on this immune cell subset and its potential contribution to the enhanced activation state of PD-derived T cells. DCs can be mainly divided into conventional cDC1, cDC2 and plasmacytoid DCs (pDCs) subtypes [50, 51, 63, 64]. We performed a preliminary assessment of DCs differentiated from hematopoietic bone marrow precursors (BMDCs) as well as those present in the spleens of PD mice (Fig. 6, Fig. S8, S9). In vitro differentiation of BMDCs with FLT3L [65] yielded similar frequencies of pDCs (B220⁺Bst2⁺) and total cDC subsets (B220⁻Bst2⁻/CD11c⁺MHC-II⁺) (Fig. S8, Fig. 6A). We found a significantly lower percentage of cDC1 (CD24⁺CD172⁻) and cDC2 (CD172⁺) subsets expressing the maturation marker CCR7 [66, 67], involved in DC migration (Fig. 6A). In spleen, we observed a lower frequency of total cDC1 cells, but similar cDC2 frequencies (Fig. 6B). We also stimulated splenocytes for 24 h with LPS and evaluated cytokine secretion in culture supernatants, revealing significantly higher levels of various cytokines and chemokines CCL5, CXCL1, IL-12p70, eotaxin, TNFα and IL-9, in supernatants from Gaa^−/− cells (Fig. 6C, Fig. S9A).

Fig. 6.

GAA-deficient DCs exhibit an increased pro-inflammatory phenotype and altered cDC1 frequencies. A Flow cytometry analysis of the frequency of pDCs, cDC1s and cDC2s in BMDCs differentiated in vitro from bone-marrow progenitors of 5-month-old WT and Gaa^−/− female mice. Data are shown as mean ± SD (n = 6). B Flow cytometry analysis of the frequency of pDCs, cDC1s and cDC2s in fresh splenocytes from 5-month-old WT and Gaa^−/− male mice. Data are shown as mean ± SD (n = 6). C Cytokine analysis on supernatants from splenocytes in B stimulated for 24 h with LPS 250 ng/ml. Data are shown as mean ± SD. Statistical significance was determined by 2-way ANOVA with Bonferroni post-hoc test. Western blot D and quantification E of IDO1 in freshly isolated splenocytes normalized to β-Tubulin. Statistical significance in A, B and D was determined by Mann–Whitney t-test. F Percent of proliferating OT-I T cells co-cultured with pooled BMDC-derived cDCs from WT or Gaa^−/− mice in the presence of increasing concentrations of OVA protein for 72 h. Data are shown as mean ± SD between samples from two independent replicates. Statistical significance was determined by 2-way ANOVA with Bonferroni post-hoc test

The immunometabolic pathway of indoleamine 2,3-dioxygenase 1 (IDO1) regulates the suppression of proinflammatory cytokines [68]. Interestingly, we found that splenocytes from PD mice expressed significantly lower levels of IDO1 protein compared to controls (Fig. 6D, E). IDO1⁺CCR7⁺ cDC1 cells can educate other DCs to dampen inflammatory cytokine production [50]. Given that IDO1 is primarily induced upon DC maturation, we used flow cytometry to assess IDO1 expression specifically in mature cDCs, confirming a trend for decreased expression particularly in cDC1, as well as cDC2 and pDCs (Fig. S9B).

We then assessed potential functional effects of the detected DC pro-inflammatory phenotype on T cell activation. To that aim, OT-I CD8⁺ T cells (WT) were cultured with cDCs from WT and Gaa^⁻/⁻ mice. Results in Fig. 6F indicate that CD8⁺ T cells cultured with Gaa^⁻/⁻ cDCs proliferated more robustly than those cultured with WT cells, across multiple tested OVA concentrations. Given these data, we also wondered whether stimulating human LOPD APCs could indirectly lead to higher T cell activation. Specific APC subtypes including pDCs and B cells are reported to be high responders to CpG oligonucleotides (CpG-ODNs) through engagement of TLR9 [69–72]. We therefore stimulated human PBMCs from HDs (n = 4), untreated LOPD patients (n = 4) and ERT-treated patients (n = 2) with CpG-ODNs during 72 h, and assessed the expression of T cell surface activation markers as well as their proliferation (Fig. S10). We found that incubation of untreated LOPD PBMCs with CpG led to higher upregulation of activation markers on both CD4⁺ (Fig. S10A) and CD8⁺ (Fig. S10B) T cells as well as to higher proliferation (Fig. S10C-E), compared to HD and ERT-treated controls, indicating a potential benefit of the enzyme therapy.

Together, these data indicate that GAA deficiency has a negative impact on tolerogenic cDC maturation and migration, leading to defective induction of IDO1 and a heightened proinflammatory phenotype in splenocyte- and bone marrow-derived DCs. This dysregulation may contribute to the steady-state inflammatory profile of PD T cells.

Discussion

Pompe disease is a multi-system disorder caused by the accumulation of glycogen in virtually any cell of the body. In the current study, we have evaluated the potential impact of GAA deficiency on subsequent lysosomal impairments and the phenotype and function of immune cells. We focused our studies on T cells, which are key effectors of inflammatory and tolerogenic responses, and on DCs, critical mediators of T cell activation. Importantly, the differentiation and function of T cells and DCs is influenced by the regulation of autophagy and glucose metabolism [44, 56, 73], two pathways shown to be altered in PD. Glycogen is naturally stored in muscle, explaining why GAA deficiency leads to severe muscle impairment [10, 49]. However, the role of GAA and lysosomal glycogen storage in immune-cell function is largely unknown, with the exception of few studies reporting a role for cytosolic glycogen in the activation of DCs [74] and in the formation of memory CD8⁺ T cells [39]. Of note, a recent study described the presence of immune-cell infiltrates in skeletal muscles of IOPD patients, identifying the expression of key immune-related genes as potential biomarkers of PD progression [75].

We first investigated whether T cells from untreated LOPD patients had a different activation profile than healthy donor cells when stimulated in vitro. Although clinical manifestations among LOPD patients vary widely, we found a consistent and significant higher degree of activation of patient-derived T cells compared to those from healthy donors. Increased expression of surface activation markers correlated with higher secretion of IL-2, and trends towards higher IFN-γ and TNF-α production, with significantly higher frequencies of TNF-α – producing CD4⁺ T cells. These results are in agreement with our previous work showing that PBMCs from untreated LOPD patients secrete high levels of pro-inflammatory cytokines upon exposure to rhGAA [47]. Notably, our cytokine analyses indicate that the rise in total cytokine output is primarily driven by an increased number of cytokine-producing CD8⁺ cells rather than by enhanced per-cell secretion. Given the expansion of activated CD8⁺ cells, it is plausible that absolute IFN-γ levels increase accordingly, even in the absence of a clear change in per-cell production. The limited sample size likely contributed to the lack of statistical significance for IFN-γ and TNF-α, and these trends should therefore be interpreted with caution. Larger cohorts will be essential to confirm these immunological changes and to distinguish increased cell frequency from true increases in per-cell cytokine production. Considering that LOPD patients and PD mice have a relatively mild disease phenotype, it is noteworthy that significant changes in T cell activation were detected in both cases. The immune cell phenotype of severe IOPD patients carrying null mutations in the GAA gene remains to be addressed, given the particularly enhanced immune responses to ERT observed in these patients, often requiring immunosuppression regimens to ensure efficacy [30, 34]. Studies involving a larger cohort, and particularly of IOPD patients, would allow for a deeper understanding of T cell responses in PD and confirm whether there is an effect of ERT treatment on the observed phenotype. No IOPD patient samples were analyzed in the current study, which represent a limitation of the work presented.

To further investigate the underlying mechanisms of immune alterations in PD, we performed our studies in GAA knockout mice. As previously described in human blood cells [45, 46], T cells from Gaa^−/− mice had a higher glycogen content compared to WT cells, although with variability possibly due to T cells being a dynamic and heterogeneous cell population. In contrast, glycogen breakdown is impaired in Gaa^−/− cells, leading to glycogen buildup [45, 46]. Further studies using specific inhibitors of glycogen synthesis [39], along with glycogen and lysosome labelling would be important to further address the role of glycogen metabolism in T cells at baseline and upon activation. However, it is essential to note that these investigations exceeded the scope of the current study.

In skeletal muscle from Gaa^−/− mice, Lim and colleagues observed an increased mitochondrial mass due to defective autophagy, with compromised ability to produce ATP as shown by the reduced OCR [38]. Our studies revealed similar metabolic defects in both CD4⁺ and CD8⁺ murine T cells related to OCR, yet the impact of these changes on their respective phenotypes differed. Gaa^−/− CD4⁺ T cells showed lower glucose uptake than WT counterparts upon stimulation, which may contribute to the impaired CD25 upregulation on stimulated CD4⁺Foxp3⁺ Tregs. Given that Tregs are highly dependent on IL-2 for survival [76], decreased CD25 expression on this cell subset could be linked to lower Treg frequencies. Of note, low CD25 expression on Foxp3⁺ Tregs has been associated with autoimmune conditions [77]. In contrast, we found a tendency toward a higher baseline expression of activation, effector memory and apoptosis markers in the CD8⁺ T cell subset. Further studies will be required to determine how chronic expression of activation, exhaustion and apoptosis markers impacts on the effector function and cytokine production of Gaa^−/− T cells.

Overall, our observations align with previous works showing that autophagy and OXPHOS defects lead to loss of T cell quiescence, impaired Treg function and systemic inflammation in mice [42, 61, 78], which is remarkable given that previous reports were based on direct knockouts on autophagy or mitochondrial proteins. Our findings are also in line with reports of autoimmunity incidence in patients with glycogen storage disease type Ib caused by Treg defects and alterations of glucose metabolism in T cells from these patients [79], although autoimmunity has not been reported in PD, and studies are lacking. Of note, characterization of autophagic buildup in specific T cell subsets such as Tregs is lacking in the context of our studies, and the link between autophagy defects in and the impaired expansion and suppressor function of GAA-deficient Tregs needs to be confirmed. The anti-inflammatory role of Tregs makes them important mediators in muscle-repair processes [80], for which reduced Treg function could contribute to muscle damage progression in PD. Our studies on murine Gaa^−/− Treg suppressive function will require further confirmation, given the mild difference observed between PD and WT cells, and will be more relevant on human samples from more severe IOPD patients.

Finally, we presented preliminary data suggesting that intrinsic defects in myeloid cells, in particular in cDC1s, and a defect in the expression of the immunoregulatory IDO1 protein, may contribute to the increased pro-inflammatory state detected in mice and LOPD patients. Our observation of decreased cDC1 maturation and peripheral frequencies aligns with works indicating that mitochondrial defects particularly impact the formation and function of cDC1s compared to pDCs and cDC2s [81], yet the mitochondrial integrity in this cell subset has yet to be investigated in the context of PD. Cytokine analysis supports that Gaa^−/− splenocytes are prone to producing higher levels of pro-inflammatory signals upon LPS stimulation, including IL-9, which is linked to allergic hypersensitivity and T cell responses [82], which are observed in some rhGAA-infused patients [35]. We hypothesize that this pro-inflammatory microenvironment could have paracrine effects on T cells, potentially contributing to the elevated basal T cell activation state observed in freshly isolated Gaa^−/− splenocytes. In agreement with this hypothesis, TLR stimulation in PBMCs with CpG led to higher activation status of T cells from LOPD patients, although we cannot rule out that results are due to the intrinsic alterations occurring in T cells and not to enhanced DC activation.

Several gene therapeutic approaches based on adeno-associated virus (AAV) vectors have been proposed as alternative to ERT to treat PD, with ongoing efforts to achieve whole body correction [17, 83–87]. In this setting, pro-inflammatory alterations of the immune system could result in enhanced immune responses to the AAV vector capsid [88], or the therapeutic transgene product [89]. Of note, establishment of immune tolerance to a transgene in the setting of gene therapy is key to achieving long-term efficacy in AAV gene therapy [90], and impairment of Treg induction has been associated with enhanced immunogenicity [91]. Our findings underscore the importance of understanding the immune system of PD and of other lysosomal and metabolic disorders, and potentially focusing on strategies to harness Treg induction with the aim of improving treatment outcomes of ERT and emerging gene therapies [92–94]. Nevertheless, the presented results do not provide a conclusive causative relationship between the observed immune phenotype and the occurrence of immune reactions to rhGAA, future studies will be needed to address this point.

Our studies provide the first evidence that autophagy and metabolic defects reported in PD muscle tissue also occur in T cells, and possibly in other immune cells such as DCs. These findings pave the way for future studies that address the existence of a link between autophagy impairments, metabolic defects and enhanced immune activation. Although findings were limited to murine cells and our observations should be confirmed in patient samples, this sets a rationale for a broader investigation of impacted cell populations and for further addressing the potential role of cDC1s on immune tolerance induction in PD [50]. Immune alterations may be caused by a combination of excessive activation of effector subsets together with insufficient activity of regulatory T and DC subsets, crucial in limiting peripheral T cell responses and promoting tolerance to self-antigens. Future studies will be aimed at determining the role of immune cell changes on PD progression and the potential of anti-inflammatory treatments to promote muscle repair, as well as to improve tolerance towards therapeutic strategies.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Helena Costa-Verdera and Marco Gargaro designed and performed experiments, analysed data and wrote the manuscript. Umut Cagin, Giorgia Manni, Giulia Scalisi, Philippe Veron and Ivan Nemazanyy contributed to experimental design and execution. Estevão Carlos Silva Barcelos, Benedetta Pieroni, Giulia Mencarelli, Doriana Ricciuti and Peggy Sanatine contributed to biological data acquisition and analysis. Laetitia van Wittenberghe contributed to harvesting of mouse samples. Pascal Laforêt contributed to the acquisition of clinical data. David-Alexandre Gross and Giuseppe Ronzitti contributed to experimental design and data interpretation. Francesca Fallarino and Federico Mingozzi directed the studies.

Funding

This work was supported by Genethon and the French Muscular Dystrophy Association (AFM), and part by research grants awarded to Prof. Francesca Fallarino from the Telethon Foundation (grant number GMR22T1081), the Italian Association for Cancer Research (AIRC 2025, grant number 30318), the Italian National Recovery and Resilience Plan (PNRR) – Bando a Cascata (project code CN00000041), and the Italian Ministry of University and Research (PRIN 2022, project code 2022CAWRK5). Additional funding was awarded to Marco Gargaro by the Italian Ministry of University and Research (PRIN 2022, project code 2022NBKCWP) and by the European Union (ERC Starting Grant REACT-DC, grant number 101078646). David-Alexandre Gross was supported by an ATIGE grant from Genopole and from the SPARK Competitive Research Grant Program on Pompe disease.

Data availability

All datasets generated during this study are available from the corresponding author upon request.

Declarations

Competing interests

Federico Mingozzi and Giuseppe Ronzitti are inventors in patents applications concerning the treatment of Pompe disease by AAV licensed to Spark Therapeutics (WO2018046774, WO2018046775, WO2018046775). Federico Mingozzi is a former employee of Spark Therapeutics. Helena Costa-Verdera is a current employee of Spark Therapeutics. All other authors have no relevant interests to disclose.