Gmppb-mutant mice exhibit dystroglycanopathy symptoms that are rescued with GSK3β inhibition or AAV-mediated GMPPB gene replacement

Domain: neuroscience
Published: 2026-04-09
Editorial outcome: major_revision

Abstract
Mutations in GDP-mannose pyrophosphorylase B (GMPPB) cause dystroglycanopathy, a
rare neuromuscular disorder characterized by α-dystroglycan hypoglycosylation, yet the
pathogenic mechanisms and therapeutic options remain poorly defined. To dissect the molecular
basis of dystroglycanopathy, we generate Gmppb knockout and knock-in (P32L and R287Q)
mice. We show that homozygous Gmppb knockout and P32L mutant mice (both male and female)
display embryonic lethality, while heterozygous Gmppb-P32L (GmppbP32L/+) mice (both male
and female) develop progressive muscular dystrophy accompanied by Purkinje cell loss,
peripheral demyelination, and impaired nerve conduction. Integrated biochemical, transcriptomic,
metabolomic and glycoproteomic analyses reveal widespread protein hypoglycosylation,
metabolic dysregulation and suppressed Wnt/β-catenin signaling, resulting in defective
differentiation and regeneration of muscle stem cells. Pharmacological activation of Wnt
signaling with CHIR-99021 restores myogenic capacity and improves regeneration after injury.
Furthermore, AAV-mediated GMPPB gene replacement reinstates α-dystroglycan glycosylation,
normalizes GDP-mannose levels, and rescues motor and electrophysiological defects.
Collectively, our findings establish GmppbP32L/+ mice as a faithful model of GMPPB-associated
dystroglycanopathy and demonstrate that Wnt pathway activation and AAV-based gene therapy
represent promising strategies for treating glycosylation-defective muscular dystrophies.

Introduction
A group of pathological conditions in the neuromuscular system, including Fukuyama-type
congenital muscular dystrophy (FCMD), muscle-eye-brain (MEB) disease, Walker–Warburg
syndrome (WWS)1,2, congenital muscular dystrophy (CMD) 1C/D, and limb-girdle muscular
dystrophy (LGMD) 2I/K/M/N3-5, collectively fall under the classification of Dystroglycanopathy
(DGP) based on their uniform feature in abnormal glycosylation of α-dystroglycan (α-DG).
Dystroglycan (DG) is a highly glycosylated membrane protein and a key component of the
dystrophin-glycoprotein complex (DGC)6, functioning as a cell-surface receptor for proteins in
the extracellular matrix and synapses7,8. α-DG is anchored to the cell membrane and stabilizes
the structural framework of the cell by binding to the extracellular matrix. In skeletal muscle, this
binding function is vital for the protection and stabilization of the muscle fibers8. Moreover, the
DGC also functions as a platform for signal transduction, recruiting molecules such as nNOS and
modulating pathways including nitric oxide, MAPK/ERK, and PI3K/Akt to influence vascular
responses, calcium homeostasis, and muscle regeneration9.

Despite the heterogeneous genetic background of these conditions, which was historically
challenging to ascertain due to limited clinically available genomic tools, clinical classification
has been primarily based on symptoms, with the specific pathogenesis remaining largely elusive.
To date, genomic studies have identified at least 19 genes associated with DGP10. The majority
of these genes encode enzymes involved in either glycosylating α-DG or providing substrates for
post-translational glycosylation modifications of DG11-13. Among these genes, GMPPB, coding
for GDP-mannose pyrophosphorylase B, catalyzes the conversion of GTP and mannose-1-
phosphate into GDP-mannose, a key mannose donor for multiple glycosylation pathways,
including O-mannosylation, C-mannosylation, N-glycosylation, and glycosyl-
phosphatidylinositol (GPI) anchor formation14,15. GMPPB mutations are inherited in an
autosomal-recessive manner. Patients with compound mutations in GMPPB display a spectrum
of symptoms, ranging from a mild, predominant proximal weakness primarily affecting the lower
limbs, to severe, structural brain defects, intellectual disability, epilepsy, and ocular
abnormalities16. Creatine kinase levels are typically elevated, ranging from 2 to 50 times higher
than normal17. Muscle biopsies frequently show myopathic changes with varying degrees of
decreased α-DG glycosylation18,19. The severe phenotypes significantly affect physiological
functions and shorten life expectancy, with many affected patients dying before reaching
adulthood12,18. The enzymatic activity of GMPPB is negatively regulated by its allosteric
feedback inhibitor GMPPA. Mutations in GMPPA result in hyperglycosylation of α-DG and a
distinct neuromuscular syndrome20,21, reflecting the vital role of GDP-mannose homeostasis in
the disease context. A recent study shows that E3 ligase TRIM67-mediated GMPPB
ubiquitination reduces its activity, with evidence showing that particular patient-derived
mutations alter this ubiquitination 22. Moreover, GMPPB is predicted to undergo methylation and
O-GlcNacylation, though specific experimental evidence is lacking23. Gmppb-deficient cell,
zebrafish and mouse models have been generated to establish its contribution to disease
pathogenesis12,24. However, animal models that faithfully recapitulate the clinical manifestations
of GMPPB-related DGP remain lacking.

Muscle satellite cells (MuSCs), or muscle stem cells, reside between the basal lamina and the
sarcolemma of muscle fibers and are essential for postnatal muscle growth and regeneration 25,26.
Upon activation after injury, MuSCs re-enter the cell cycle and differentiate into myoblasts,
which undergo reprogramming of gene expression patterns including Pax7 and Myogenin27,
thereby initiating the differentiation phase and giving rise to myocytes. Defects in MuSC
differentiation and self-renewal are increasingly recognized as major contributors to muscular
dystrophies28,29, including Duchenne muscular dystrophy, where defects in polarity, asymmetric
division, and myogenic programming limit regeneration30,31. In contrast, the role of MuSCs in
DGP remains poorly understood. Although GMPPB mutations are linked to hypoglycosylation
of α-DG, the mechanisms by which these mutations impair muscle regeneration and contribute
to disease progression are not fully defined.

In this study, to investigate molecular pathogenesis and explore therapeutic strategies for
GMPPB-DGP, we generated genetically engineered mouse strains carrying Gmppb mutations
and identified neuromuscular phenotypes closely resembling the features of GMPPB-related
muscular dystrophy, thereby providing direct evidence of GMPPB mutations contributing to
muscular dystrophy. This characterization involved assessing parameters such as motor
performance, histological changes in muscle and brain tissues, serum creatine kinase levels, and
the glycosylation status of α-DG. We established Gmppb-knockout myoblast cells and isolated
MuSCs from dystrophic mice, to investigate the impact of Gmppb mutations on muscle
differentiation and regenerative capacity. Transcriptomic, metabolomic and glycoproteomic
analyses in these models revealed that Gmppb P32L mutation leads to impaired protein
glycosylation overall and downregulation of the Wnt signaling pathway, consequently
influencing muscle differentiation and regenerative capacity. Pharmacological activation of the
Wnt pathway successfully mitigated the phenotypes of GMPPB-DGP. Furthermore, we
determined the efficacy of AAV gene replacement therapies, which have shown promise in
treating diseases such as DMD32,33. We showed that AAV-mediated GMPPB expression in the
neuromuscular system was sufficient to reverse Gmppb mutational phenotypes. This work offers
a platform for preclinical investigations on GMPPB-DGP and presents promising therapeutic
avenues for addressing DGP diseases.

Results
Homozygous Gmppb knockout and P32L mutations result in embryonic lethality not
rescued by mannose supplementation
First, we integrated published clinical cases of patients with GMPPB mutations to catalog
the types and frequencies of variants, identifying ten GMPPB gene point mutations as the most
frequently reported (Fig. 1a)11-13,16,18,34-39. Notably, patients carrying the P32L and R287Q
mutations often present with severe pathological phenotypes, marked by congenital muscular
dystrophy along with developmental abnormalities in the brain16,18,40. Conversely, those with the
D27H mutation typically manifest with milder limb-girdle muscular dystrophy (LGMD)
phenotypes. Furthermore, our analysis of the protein structure41 (Fig. 1b) revealed that mutations
P32L, D27H, and T153I reside in the Rossman fold region, while mutations V330I, R357H, and
R287Q/W are situated in the β-strand region. These regions are critical for the catalytic activity
and stability of GMPPB42. Therefore, we generated two Gmppb knock-in mouse models, namely
GmppbP32L and GmppbR287Q mice, as well as one Gmppb knockout (Gmppb-KO) mouse model
(Fig. 1c). Specifically, the GmppbP32L and GmppbR287Q mice were generated using a
CRISPR/Cas9-mediated homology-directed repair (HDR) approach. For the P32L allele, a
single-guide RNA (sgRNA) was designed to target a sequence near nucleotide position c.95 of
the Gmppb coding region, and a single-stranded oligodeoxynucleotide (ssODN) donor template
was provided containing the desired c.95C>T substitution. Similarly, for the R287Q allele, an
sgRNA was designed to target the region encompassing nucleotide c.860, and an ssODN donor
harboring the c.860G>A substitution was used to introduce the arginine-to-glutamine change at
residue 287. For the Gmppb-KO line, two sgRNAs were designed to target the intronic regions
flanking exons 2 and 3 of Gmppb. Simultaneous Cas9-mediated double-strand breaks followed
by non-homologous end joining (NHEJ) resulted in the deletion of a 303 bp fragment, leading to
a frame shift and functional loss of the gene. In all cases, sgRNAs and donor templates were co-
injected with Cas9 mRNA into fertilized C57BL/6J zygotes. Founder mice carrying the intended
mutations were identified by Sanger sequencing of PCR-amplified genomic DNA surrounding
the target sites (Supplementary Fig. 1a, b and c). Correctly targeted founders were backcrossed
to C57BL/6J mice three times to eliminate potential off-target effects and establish stable lines.
The litter size and genotype data indicated that homozygous GmppbR287Q/R287Q mice are viable
(Supplementary Fig. 1a and 2a), whereas no homozygous Gmppb-/- or GmppbP32L/P32L mice are
present (Supplementary Fig. 1b, c and 2d, e), suggesting that they may have experienced
embryonic lethality. Next, we examined mouse embryos obtained from crossbreeding
heterozygous Gmppb+/- and GmppbP32L/+ mice. The data indicate a significant reduction in embryo
counts by E14.5 and evident embryo resorption were observed (Fig. 1d, 1f), in contrast to the
normal appearance of developing embryos in the Gmppb+/+ x Gmppb+/- (Supplementary Fig. 1d)
and Gmppb+/+ x Gmppb+/+ (Supplementary Fig. 1e) crossings. While the development of
heterozygous Gmppb+/- and GmppbP32L/+ was relatively comparable to their respective wild-type
littermates at E10.5 and E12.5, with mild to moderate level of edema and local hemorrhage
present in a portion of embryos at E12.5 (Fig. 1e and g), both homozygous Gmppb-/- and
GmppbP32L/P32L embryos exhibited remarkable developmental failure, structural abnormalities and
fusion with the maternal tissues (Fig. 1e and g). These results suggest that the homozygous
embryos failed to develop before E10.5. Evidently, allele ablation, P32L, and R287Q mutations
resulted in varying degrees of functional impairment, with R287Q phenotypes being relatively
mild. To determine whether the absence of homozygous offspring resulted from preimplantation
defects, we performed genotyping on isolated blastocysts. Consistent with the previous Gmppb
knockout model43, homozygous Gmppb–/– blastocysts were readily detected (Supplementary
Fig. 2b), indicating that complete loss of Gmppb permits early embryonic development up to the
blastocyst stage. In contrast, GmppbP32L/P32L blastocysts were not observed (Supplementary Fig.
2c), suggesting that embryos carrying the P32L substitution fail to reach or maintain this stage.
These findings indicate that while Gmppb loss-of-function allows limited pre-implantation
development, the P32L mutation causes an earlier or more severe defect. Future studies involving
blastocyst transfer or implantation assays will be valuable to determine whether the P32L
embryos can form morphologically normal blastocysts but fail during implantation, or whether
defects of the heterozygous carriers contribute to developmental arrest occurs prior to this stage.
Previous studies in Pmm2 mutant mice have shown that mannose supplementation can
potentially overcome abnormal glycosylation and embryonic lethality44,45. To determine whether
mannose supplementation can correct the lethality in Gmppb -/- and GmppbP32L/P32L embryos,
heterozygous females were provided with drinking water containing 9 mg/mL mannose starting
1 week before mating and throughout mating and gestation45 to ensure stable embryonic exposure
to mannose. The results showed that in Gmppb+/- or GmppbP32L/+ models, continuous mannose
supplementation failed to rescue embryonic lethality (Supplementary Fig. 2d and e).

Gmppb-P32L mutation leads to haploinsufficiency of the wild type alleles, resulting in
progressive muscular dystrophic phenotypes
That Gmppb+/- and GmppbP32L/+ models exhibited a more severe phenotype compared to
human patients prompted us to investigate the effect of gene dosages in these mouse models.
First, to examine histological alterations, we performed Hematoxylin and Eosin (HE) staining on
tissues obtained at 32 weeks including the tibialis anterior muscle (TA), gastrocnemius muscle
(GA), quadriceps muscle (Qu), heart, diaphragm, and brain tissue. Tissues of Gmppb+/- and
GmppbR287Q/R287Q mice did not present obvious pathological changes (Supplementary Fig. 3a).
However, in GmppbP32L/+ mice, centrally nucleated muscle fibers were evident in the TA, GA and
Qu muscles, indicating ongoing muscle tissue damage (Supplementary Fig. 3a). We did not
observe significant morphological changes in non-striated muscle tissues such as the heart or
diaphragm in these models (Supplementary Fig. 3a).

The GA muscles of 32-week GmppbP32L/+ mice exhibited reduced GDP-mannose content
assessed by liquid chromatography-mass spectrometry (LC-MS) (Fig. 2a), reduced muscle fiber
size, as well as an increased level of centrally nucleated myofibers (Fig. 2b-d). Furthermore, the
proportion of centrally nucleated myofibers increased as GmppbP32L/+ mice aged (Fig. 2e),
indicating a progressive deterioration of this pathological condition. Electron microscopy
revealed that 32-week-old GmppbP32L/+ mice showed disordered myofilament and dissolving of
focal myofilament, which was absent in wild-type muscles (Supplementary Fig. 3b).
Additionally, myoplasmic necrosis spanning multiple muscle fibers was present in the muscles
of 48-week-old GmppbP32L/+ mice (Supplementary Fig. 3c). Accumulation of scattered
autophagic vacuoles containing cytoplasmic debris and electron-dense material was prevalent,
and a significant portion of mitochondria appeared distorted in the muscle of GmppbP32L/+ mice
(Fig. 2f). Homeostasis of autophagic activities is critical for muscle development and
regeneration, and its dysregulation has been associated with pathological conditions such as
myopathies46,47. It is important to further investigate whether/how the observed increase in
autophagic vacuoles contributes to muscular defects in GmppbP32L/+ mice.

Severely affected GMPPB-DGP patients routinely display neurological abnormalities17,48.
In addition, GMPPA deficient mice display a progressive loss of Purkinje cells20. We performed
a series of analyses to determine whether GmppbP32L/+ mice develop additional phenotypes
affecting the central and peripheral nerve systems. Glycosylation of α-DG in the whole brain
lysate is markedly reduced in the GmppbP32L/+ groups, as assessed by Western blots
(Supplementary Fig. 4a). GmppbP32L/+ mice showed no notable differences in structural
organization of the brain, with no aberrant neuronal migration in the cerebral cortex and
hippocampal region observed (Supplementary Fig. 4b). Immunofluorescent staining with NeuN
revealed well-organized hippocampal formation, including clearly defined pyramidal and granule
cell layers (Supplementary Fig. 4c). However, GmppbP32L/+ mice showed significantly
impairment in spatial learning and memory as assessed by Morris water maze tests
(Supplementary Fig. 4d). However, though consistent with defective muscle functions,
GmppbP32L/+ mice showed significantly reduced average swimming speed, which confounded the
interpretation (Supplementary Fig. 4d). Additional motor-independent behavioral assays, such
as novel object recognition or fear conditioning, will be important to further confirm the
impairment in recognition. Calbindin immunofluorescent staining of the cerebellum indicated a
reduced population of Purkinje cells specifically found in 12 month GmppbP32L/+ mice (Fig. 3a).
Together with findings in Gmppa mutants20, this likely reflecting the particular vulnerability of
Purkinje cells to disturbances in GDP-mannose homeostasis due to their high metabolic and
glycosylation demands. Furthermore, the sciatic nerves of GmppbP32L/+ mice displayed marked
demyelination, as shown by H&E staining (Fig. 3b) and electron microscopy (Fig. 3c, red
arrows), with spatially disoriented Schwann cells broadly evident (Fig. 3d, yellow arrows). In
the spinal cord nerves, however, H&E staining and electron microscopy showed no significant
changes in neuronal population, myelination or organization (Supplementary Fig. 4e-g).
Because proper myelination of axons is critical for rapid conduction of electrical impulses, we
next assessed the nerve conductance velocity (NCV) and compound muscle action potential
(CMAP) with electromyography. The results indicated a significant reduction in conduction
velocity and CMAP amplitude in GmppbP32L/+ sciatic nerves (Fig. 3e and f), indicating impaired
transmission. These data show that although gross brain structure appears preserved, GmppbP32L/+
mice develop progressive cerebellar Purkinje cell loss, cognition defect, peripheral nerve
demyelination, and impaired nerve conduction, indicating that GMPPB deficiency affects both
central and peripheral nervous system integrity.

To evaluate muscle functions and motor activities, we performed rotarod and grip tests on
16-week-old Gmppb+/-, GmppbP32L/+ GmppbR287Q/R287Q and wild-type mice. The average time
spent on rotarod significantly decreased in Gmppb+/- and GmppbP32L/+ mice (Supplementary Fig.
5a), while the average grip strength significantly decreased only in GmppbP32L/+ mice
(Supplementary Fig. 5b). The muscle weakening appeared to be progressive, shown by
comparative studies of rotarod and grip test with 16- and 48-week-old mice (Fig. 3g and h).
However, the results of these analyses are dependent on multifactorial determinations, such as
muscle mass/body weight ratio, neural conductance. The lean/body weight ratio assessed by
EchoMRI displayed significant reduction in GmppbP32L/+ mice (Supplementary Fig. 5c and d),
consistent with the myopathic phenotype. This, together with the impaired nerve conductance
and motor coordination (loss of Purkinje cells) we reported above (Fig. 3a), necessitating a
specific muscle function test in this context. Thus, we performed in vivo muscle contraction test
on the GA muscle. In this artificially stimulated muscle contraction test, we found that
GmppbP32L/+ muscle generated a significantly reduced maximal tetanic force compared to WT
(Fig. 3i). These data demonstrate that GmppbP32L/+ mice present a complex phenotype involving
both neuropathic and myopathic components.

Patients with GMPPB-dystroglycanopathy generally display elevated levels of serum
creatine kinase (CK)11,40. Similarly, serum CK levels in 30-week-old GmppbP32L/+ mice were
significantly increased, while those of Gmppb+/- and GmppbR287Q/R287Q mice were comparable to
those of wild-type mice (Fig. 3j). In contrast, at 16-week serum CK levels did not differ
significantly among WT, GmppbP32L/+, Gmppb+/- and GmppbR287Q/R287Q (Supplementary Fig. 5a).
Collectively, our results suggest that GmppbP32L/+ mice developed progressive defects in muscle
development, leading to impaired muscle functions that deteriorate over time.

GmppbP32L/+ and Gmppb+/- mice showed a reduced level of α-DG glycosylation
To evaluate the impact of specific mutations on protein abundance, we assessed GMPPB
protein levels by immunofluorescence and found no significant differences among the genotypes
(Supplementary Fig. 6a). Previous research has established a strong correlation between
decreased levels of α-dystroglycan (α-DG) glycosylation and dystroglycanopathies, particularly
evident in patients with GMPPB mutations12,13,16. Therefore, we analyzed the level of α-DG
glycosylation in our mouse models. First, protein lysates extracted from skeletal muscle lysates
and enriched by wheat germ agglutinin (WGA) agarose were assessed for the levels of α-DG
glycosylation using antibodies specifically targeting α-DG glycosylation epitopes VIA4-1 and
IIH6C449,50 (Fig. 4a). The results indicated a significant reduction in α-DG glycosylation in
muscles from GmppbP32L/+ mice. However, Gmppb+/- and GmppbR287Q/R287Q mice exhibited
reduced glycosylation levels specifically for VIA4-1(Fig. 4b and c). The disparity between
VIA4-1 and IIH6C4 may be attributed to their distinct epitope specificities, warranting further
investigation. Nevertheless, there were no notable changes in the level of β-DG, nor any observed
mobility shifts, as reported in some GMPPB patients35, across all groups (Fig. 4a and d). Western
blot analyses with two independent total α-DG antibodies showed reduced signal of the mature
α-DG banding pattern, consistent with hypoglycosylation (Fig. 4e and Supplementary Fig. 6b).
To determine the protein stability, inhibition of protein turnover with cycloheximide (CHX) in in
vitro cultured muscle slices resulted in varying α-DG degradation rate, with Gmppb+/- and
GmppbP32L/+ muscles showing reduced protein stability compared to WT and GmppbR287Q/R287Q
samples (Supplementary Fig. 6c and d). In addition, combination treatment of the proteasomal
inhibitor MG-132 and CHX indicated that in GmppbP32L/+ muscles, degradation pathway is
shifted towards the autophagy- or lysosome-dependent routes compared to the relatively
preserved α-DG protein in Gmppb+/- muscles (Supplementary Fig. 6c and d). Additionally,
immunofluorescent labeling with IIH6C4 (Fig. 4f) and the core α-DG (Supplementary Fig. 6e)
in cross sections of muscle specimen confirmed the reduced level of α-DG glycosylation and α-
DG membrane abundance specifically in GmppbP32L/+ muscles. These results suggest the
potentially dominant negative role of P32L mutation in the pathogenesis of GMPPB-
dystroglycanopathy.

The physiological roles of Dystroglycan in maintaining structural stability of muscle relies
on their ability to bind to extracellular ligands such as laminin51. To investigate whether reduced
levels of α-DG glycosylation impact its binding capacity with extracellular ligands, we performed
solid-phase laminin-binding assay (Fig. 4g) and laminin overlay assay (Fig. 4h) using WGA-
enriched glycoproteins extracted from the muscles. The overall ligand-binding capacity of native
(non-denatured) α-DG was markedly diminished in GmppbP32L/+ mice (Fig. 4g, h).
Immunofluorescent staining for laminin indicated the membrane binding activity of laminin was
significantly reduced in GmppbP32L/+ muscles (Fig. 4i). Thus, the defective glycosylation of α-
DG in GmppbP32L/+ mice significantly disrupts the interactions between α-DG and extracellular
matrix ligands, such as laminin, in the muscles of these mice.

GMPPB Knockout causes aberrant muscle differentiation
Having established that GMPPB P32L mutations lead to impaired α-DG glycosylation
resulting in long-term muscle damage resembling muscular dystrophic phenotypes in GMPPB
patients, we further investigated the molecular consequences on GMPPB loss-of-function with
C2C12 myoblast cells. First, we generated GMPPB knockout C2C12 myoblast cells with
CRISPR-Cas9 and confirmed the successful knockout with Western blots and RT-qPCR (Fig.
5a). The viability of GMPPB-KO cells did not change, and the proliferation activity in GMPPB-
KO cells were similar to WT C2C12 cells (Fig. 5b). We assessed the GDP-mannose abundance
using LC-MS showing a significant reduction of GDP-mannose levels in Gmppb-KO cells (Fig.
5c, Supplementary Fig. 7a). Primary myoblasts can be induced by 2% horse serum to
differentiate into multinucleated myotubes and undergo shifts in expression patterns of muscle-
specific regulators such as Myogenin and MyHC, a process that is critical in muscle development
and regeneration52,53. FACS analyses with Annexin V/PI staining showed increased rate of
apoptosis in Gmppb-KO C2C12 cells during the time course of differentiation (Fig. 5d).
Additionally, when subjected to differentiation for 7 days with horse serum, immunofluorescent
labeling of MyHC indicated abnormal myotube morphology, reduced myotube diameter, and a
lower myotube fusion index in Gmppb-KO C2C12 cells (Fig. 5e, f, g). Together, these data
suggest that GMPPB loss-of-function disrupts the muscle differentiation and formation of
myotubes.

Myogenic regulatory factors (MRFs), such as MyoD, Myf5, Pax7 and Myogenin (MyoG),
play vital roles in regulating muscle differentiation, as their temporal expression patterns drive
myogenesis54. We next analyzed the level of MRFs in Gmppb-KO and wild-type cells during
proliferation and differentiation. In the proliferative phase, the expression levels of Pax7 and
MyoD were reduced (Fig. 5h, i, l, m), whereas the level of Myf5 is increased (Fig. 5j, n) in
Gmppb-KO cells relative to their wild-type counterparts. Meanwhile, consistent with previous
studies55,56, the expression levels of Pax7 and MyoD gradually decrease over time during the
differentiation phase in wild-type C2C12 cells (Fig. 5h, i, l, m). However, in Gmppb-KO cells,
this decline of Pax7 and MyoD is absent. Instead, their expression persistently remains at low
levels (Fig. 5 h, i, l, m). Conversely, during differentiation, the expression of Myogenin
gradually increases in Gmppb-KO cells but remains significantly lower than in wild-type cells
(Fig. 5k, o), whereas Myf5 expression remains elevated compared to wild-type cells (Fig. 5j, n).
To control for potential effects introduced during KO generation, we assessed Myf5 and
Myogenin expression in Gmppb-KO cells re-expressing GMPPB (Gmppb-KO-CDS;
Supplementary Fig. 7e); these cells showed expression patterns similar to wild-type C2C12
cells (Supplementary Fig. 7b, c and d), indicating that the shift observed in Gmppb-KO cells
was not due to the knockout procedure itself. Taken together, these findings suggest that GMPPB
deficiency disrupts the normal temporal pattern of myogenic regulatory factor expression in
C2C12 cells during proliferation and differentiation, which may underlie the impaired muscle
regeneration and myopathy observed when GMPPB activity is reduced by mutations.

To elucidate the effects of allelic mutations of GMPPB, we re-expressed recombinant
GMPPB with point mutations (P32L, T153I, R185C, R287Q and V330I) into Gmppb-KO C2C12
cells. Western blots with flag antibody confirmed the expression of mutant GMPPB proteins in
these stable cell lines (Supplementary Fig. 7e). To determine whether GMPPB mutations impact
the catalytic activity, we determined the GDP-mannose concentrations using LC-MS
(Supplementary Fig. 7f). We found that both GMPPB-P32L and GMPPB-R287Q cells
displayed a significant reduction in the GDP-mannose content compared to the wild-type cells,
indicating again that P32L and R287Q mutations are hypomorphic/amorphic. Notably, the GDP-
mannose content in GMPPB-P32L cells was approximately 20% of that in wild-type cells
(Supplementary Fig. 7f). Moreover, we also determined the subcellular localization of these
GMPPB variants with immunofluorescent staining (Supplementary Fig. 7g). In wild-type cells,
GMPPB displayed a diffuse cytoplasmic localization surrounding the nuclei (Supplementary
Fig. 7g), consistent with its known role as a cytosolic enzyme involved in GDP-mannose
biosynthesis. The P32L and R287Q mutations caused protein aggregation in the cytoplasm. The
T153I mutation caused GMPPB to aggregate near membrane protrusions into the cytoplasm.
Conversely, the R185C and V330I mutations have no significant changes compared with wild-
type GMPPB (Supplementary Fig. 7g). Together, these data indicated that GMPPB is essential
for muscle differentiation and proper subcellular localization.

Gmppb Knockout and GmppbP32L/+ lead to widespread disruption of protein glycosylation
and impairment of the physiological process related to muscle functions
The varying impact of different mutations on the catalytic activity of GMPPB may offer an
explanation for the wide spectrum of disease phenotypes observed in patients. However,
examination of clinical features of patients with different mutations suggests that phenotypes of
compound GMPPB mutations may involve a more complicated mechanism that potentially
involves compensatory effect12,18. Indeed, that GMPPB-KO cells are viable, exhibiting no
obvious defects in proliferative activity (Fig. 5b) further stresses the existence of possible
compensation. GDP-mannose is a critical precursor in protein and lipid glycosylation not limited
to α-DG57, emphasizing that deficiency in GMPPB may result in global change of the proteome
and metabolome, with numerous components involved in gene transcription which promotes the
exertion of compensatory mechanisms.

To characterize the mechanisms of GMPPB-KO myoblasts on muscle development, we
performed RNA sequencing (RNA-seq) analysis on Gmppb-KO and wild-type C2C12 cells, and
subsequently investigated differentially expressed genes (DEGs) using gene ontology (GO)
enrichment analysis. 1177 differentially expressed genes (DEGs, (490 up/687 down, KO vs. WT))
were identified with the threshold set at p.adjust < 0.05 and |log2FC| > 1 (Supplementary Fig.
7h and i). We validated the DEGs expression with RT-qPCR (Supplementary Fig. 7j and k).
The results revealed a significant enrichment of genes associated with muscle organ development,
striated muscle cell differentiation, muscle contraction, and receptor regulator activity (Fig. 6a).
Similarly, GO analysis of RNA sequencing data from muscle tissue of WT, Gmppb+/-,
GmppbP32L/+ and GmppbR287Q/R287Q mice further confirmed the enrichment of pathways related to
muscle development specifically in GmppbP32L/+ mice. These pathways included response to
extracellular stimulus, skeletal muscle organ development, skeletal muscle tissue development,
and skeletal muscle cell differentiation (Fig. 6b). These pathway alterations again emphasize an
association between the GMPPB mutation and global transcriptomic changes that result in
abnormalities in muscle development, consistent with our observations of muscular dystrophy
phenotypes in GmppbP32L/+ mice. Furthermore, we observed substantial changes in the abundance
of various cytosolic metabolites through LC-MS-based untargeted metabolomics analysis with
gastrocnemius muscle of GmppbP32L/+ mice (Supplementary Fig. 8a). Pathway enrichment
analyses indicated significant alteration in the glycerophospholipid pathway (Supplementary
Fig. 8b), which plays a vital role in the development of muscular and cellular membrane
components58. Overall, these results suggest that GMPPB loss-of-function (in both GMPPB-KO
C2C12 cells and GmppbP32L/+ mouse model) disrupts the metabolism and transcriptional activities
globally. Although our transcriptomic data in mice suggest altered immune pathways, the
Gmppb-mutant animals did not display obvious immune deficiencies or increased infection under
routine conditions. A dedicated immunological evaluation will be required to determine whether
these transcriptomic changes translate into functional immune alterations.

To test whether GMPPB knockout results in changes in protein glycosylation overall, we
detected the glycosylation level of total protein extracted from GMPPB-KO cells, using
concanavalin A (ConA) that attaches to the mannose and glucose groups on glycosylated
proteins59, as well as aleuria aurantia lectin (AAL) that recognizes fucose residues in elongated
sugar chains60 (Supplementary Fig. 9a, b, c). The results showed a significant reduction in both
ConA and AAL signals upon GMPPB knockout (Supplementary Fig. 9d), suggesting that
GMPPB loss-of-function disrupted glycosylation of proteins not limited to α-DG. Gene set
enrichment analyses (GSEA) of RNAseq data showed that GMPPB knockout significantly
altered mannose-type O-glycan biosynthesis, with the pathway influencing N-glycan
biosynthesis relatively untouched (Fig. 6c). These results suggest a potential upregulation of O-
mannose biosynthesis in response to GMPPB deficiency, although further experimental
characterization is required to confirm this compensation mechanism.

Furthermore, GSEA analyses revealed a significant downregulation of pathways associated
with the neuromuscular junction (NMJ) in Gmppb-KO cells, including neuroactive ligand
receptor interaction, acetylcholine-gated channel complex, and postsynaptic neurotransmitter
receptor activity (Fig. 6d). Quantitative reverse transcription PCR also indicated that the
expression of multiple key genes participating in the NMJ signaling transduction, including musk,
ryr1 and AchR subunits (chrnd and chrna1) was significantly inhibited in Gmppb-KO cells
(Supplementary Fig. 9e). Meanwhile, it has been established that regulation of postsynaptic
calcium release at the neuromuscular junction is critical for NMJ transmission and generation of
contractile force61. Also, transcriptomic profiles suggested functional impairments in calcium
signaling pathway in Gmppb-KO cells (Fig. 6a). Therefore, to determine the consequence of
GMPPB knockout specifically on muscular calcium signaling, we assessed intracellular Ca2+
level with fluorescent Fluo-4 Ca2+ indicator. Notably, stimulation with 1 mM ATP resulted in a
hyperactive surge in cytosolic calcium influx in Gmppb-KO cells (Supplementary Fig. 9f),
suggesting dysregulated calcium release from intracellular stores via ectopic activation of a P2Y-
evoked and G-protein-mediated signaling pathway. The hyperactive response observed in
Gmppb-KO cells may indicate an increased susceptibility of dystrophic cells to such stimulation,
as has been observed in the mdx mouse model of Duchenne muscular dystrophy (DMD),
leading to sarcolemma damage62,63. These findings highlight the potential disruption of Ca2+ homeostasis
following the loss of GMPPB, where hyperactive calcium flow disrupted Ca2+ dynamics in
skeletal myogenesis, hindering muscle development. Taken together, the deficiency of GMPPB
may lead to deficient protein glycosylation not limited to dystroglycan and disruption of the NMJ
signaling pathway, both critical for maintaining normal muscle functions. The dysregulated Ca²⁺
handling observed in our cell and mouse models suggests that NMJ function may also be affected.
Detailed NMJ analyses are planned for follow-up collaborations aimed at linking our biochemical
findings to synaptic structure.

To further characterize the electron microscopic findings where scattered autophagic vacuoles
were identified in GmppbP32L/+ muscles (Fig. 2f and Supplementary Fig. 3b and c), we
performed gene set enrichment analysis using transcriptomic data from GmppbP32L/+ mutant
muscles. This analysis (Fig. 6e) revealed significant enrichment of the “unfolded protein
response” pathway and a trend toward enrichment of the “reactive oxygen species pathway” (p
= 0.0634), suggesting activation of proteostatic and oxidative stress programs. Consistent with
these findings, Western blot analyses showed a marked increase in the LC3-II/I ratio and
accumulation of p62 (Fig. 6f) in GmppbP32L/+ muscles, whereas these changes were not observed
in Gmppb+/- or GmppbR287Q/R287Q models. These results indicate that the P32L mutation leads to
autophagic flux impairment and mitochondrial stress, revealing a mutation-specific disruption of
cellular homeostasis that may contribute to the distinct pathological severity associated with this
variant. In addition, to further examine the mitochondrial defects suggested by our ultrastructural
analyses (Fig. 2f), we performed MitoTracker immunofluorescence staining in C2C12 cells (Fig.
6g). In wild-type and Gmppb-KO-R287Q cells, mitochondria displayed an interconnected
network typical of healthy mitochondrial dynamics. In contrast, Gmppb-KO cells exhibited a
punctate, fragmented mitochondrial pattern, characterized by discrete dot-like fluorescent signals
and loss of the reticular organization, whereas P32L expression (Gmppb-KO-P32L) failed to
rescue this mitochondria defect (Fig. 6g). This altered distribution suggests increased
mitochondrial fragmentation and impaired fusion–fission balance. Together, these data indicate
that GMPPB deficiency, particularly the P32L mutation, induces mitochondrial stress and
structural disorganization, contributing to impaired cellular homeostasis in affected muscle fibers.

Glycoproteomic Analyses indicates widespread protein hypoglycosylation
RNAseq results suggested dysregulation of pathways related to protein glycosylation (Fig.
6c). To directly assess this at the protein level, we performed glycoproteomic analysis in which
total protein from gastrocnemius muscles of GmppbP32L/+ and WT mice was enriched with ConA
that specifically immunoprecipitated glycoproteins and their interaction partners, and subjected
to LC-MS based identification and quantification. A greater number of peptides were identified
in the GmppbP32L/+ samples, shown by identified scatter plot (Fig. 6h), consistent with Coomassie
blue staining of pre- and post-ConA-enriched samples (Supplementary Fig. 10a). In total, 692
peptides were identified in WT only, compared to 85 peptides in P32L only (Supplementary Fig.
10b). Among the 1288 peptides identified in both groups, 275 showed significantly decreased
abundance (FC>1.5; p>0.05), compared to 148 peptides showing increased abundance in
GmppbP32L/+ muscle (Supplementary Fig . 10c). Pathway analyses of differentially expressed
peptides indicated positive correlation in pathway related to glycan degradation, and negative
correlation in glycosaminoglycan biosynthesis, cytokine-cytokine receptor interaction and the
Notch signaling pathway (Supplementary Fig. 10d). Together, these data demonstrate that the
P32L mutation induces widespread protein hypoglycosylation.

GmppbP32L/+mice exhibited reduced capacity for skeletal muscle differentiation and
regeneration.
Next, to determine whether muscle differentiation is also impaired in Gmppb mouse models,
we isolated GA muscles from GmppbP32L/+ mice and wild-type mice, prepared single-cell
suspensions, and isolated muscle satellite cells with flow cytometry using surface-specific
markers CD31, CD45, Sca1, and VCAM1 (Fig. 7a). The proportion of CD45-CD31-Sca1-
VCAM1+ MuSCs from GmppbP32L/+ muscles showed no significant difference from that of WT
muscles (Supplementary Fig. 11a and b). However, under differentiation condition they
displayed a significant decrease in the capacity to fuse into myotubes (Fig. 7b), with the fusion
index reduced by approximately 30% compared to WT (Fig. 7d). Furthermore, the expression of
MyoG during differentiation in GmppbP32L/+ satellite cells was significantly reduced compared to
wild-type satellite cells (Fig. 7c, e). Immunoblotting analyses revealed that, on the first day of
differentiation, the expression levels of MyHC and MyoG were comparably low and showed no
significant difference between GmppbP32L/+ and wild-type satellite cells (Fig. 7f). However, by
the fifth day of differentiation, induction of MyHC and MyoG expression in GmppbP32L/+ satellite
cells is significantly dampened compared to that of WT MuSCs (Fig. 7g, h).

Upon muscle damage, satellite cells undergo a process of activation, initiated by progressive
differentiation to produce muscle cells that fuse into multinucleated myotubes and/or integrate
with damaged muscle fibers, facilitating rapid regeneration of mature muscle tissue64,65. Since
GmppbP32L/+ muscle satellite cells exhibit defects in differentiation, next, we further determined
whether such defects lead to impairment in muscle regeneration. The cardiotoxin (CTX)-induced
muscle injury model has been widely used to investigate capability in skeletal muscle
regeneration for its transient and replicable characteristics66. We administered CTX to the tibialis
anterior (TA) muscles of GmppbP32L/+ mice and conducted histological determination at 3, 5 and
14 day post CTX-induced muscle damage (Fig. 8a). At 3 days post-injury, wild type TA muscles
exhibited a notable presence of newly formed muscle fibers, marked by increased level of
centralized nuclei and increased proportion of muscle cells with 2 or more nuclei, as well as
increased average cross-sectional area (CSA) of muscle fibers, while GmppbP32L/+ mice displayed a
limited number of regenerating muscle fibers (Fig. 8b). By day 5 post-injury, control mice
demonstrated abundant multinucleated myofibers, whereas GmppbP32L/+ mice predominantly
showed mononucleated fibers, with myofibers being smaller in their CSA (Fig. 8b). By day 14
post-injury, control mice presented tightly packed, well-structured muscle fibers, whereas
GmppbP32L/+ mice displayed regenerating muscle fibers of varying sizes (Fig. 8b). Moreover, by
five days post-injury, GmppbP32L/+ mice exhibited a marked reduction in the proportion of
myofibers containing two or more central nuclei (Fig. 8b, c). Similarly, the average CSA of
muscle fibers in GmppbP32L/+ mice decreased by approximately 40% compared to control mice
(Fig. 8b, d). Very few of the GmppbP32L/+ muscle fibers exceeding 2000 μm2 in CSA (Fig. 8e).
The serum CK level in WT mice following CTX treatment spiked at day 3, gradually declined
and normalized at day 14 (Fig. 8f), while the CK level in GmppbP32L/+ mice displayed similar
pattern for day 3 and 5, yet failed to normalize (compared to day 0 in GmppbP32L/+ mice) at day
14, indicating incomplete regeneration (Fig. 8f). Meanwhile, immunofluorescent labeling of
embryonic MyHC showed that immediately following CTX treatment, eMyHC was robustly
induced at day 3 and 5 in WT mice (Fig. 8g, Supplementary Fig. 11c), and declined to the
baseline at day 14 (Supplementary Fig. 11c), indicating successful tissue repair despite
centralized nuclei still evident in some muscle fibers (Fig. 8b). However, eMyHC expression
was at a consistently low level at day 3, 5 and 14 in GmppbP32L/+ mice, indicating delayed and
inefficient myofiber regeneration, consistent with incomplete tissue repair observed in HE
staining (Fig. 8g, Supplementary Fig. 11 c). These findings indicate a crucial role for Gmppb in
adult skeletal muscle regeneration and suggest that Gmppb P32L mutation disrupts skeletal
muscle regeneration.

Pharmacological activation of Wnt signaling rescued impairment of myogenesis
The process of myoblasts fusion into myotubes represents a critical step in embryonic and
adult skeletal myogenesis necessitating precise temporal and spatial regulation by multiple
signaling pathways67. To explore the molecular mechanisms underlying GMPPB's involvement
in muscle development, we analyzed RNA-seq data from Gmppb-KO and wild-type C2C12 cells.
KEGG enrichment analyses revealed the downregulation of nineteen signaling pathways, in
which the Wnt signaling pathway is closely related to myoblast differentiation (Supplementary
Fig. 11f). Wnt family proteins are involved in a variety of embryonic and adult processes
including proliferation, differentiation and cell fate determination68. Specific glycosylation
patterns can dictate the spatial and temporal activation of Wnt signaling during myogenesis,
whereas aberrant expression of Wnt proteins or their receptors has been associated with muscle-
related disorders and impaired regeneration processes69,70. The genes associated with the
canonical Wnt pathway exhibited significant downregulation in Gmppb-KO cells, accompanied
by substantial downregulation of genes linked to muscle development (Fig. 9a), consistent with
the disruption of expression pattern during induced differentiation (Pax7 and MyoD, Fig. 5h and
i). Validation through quantitative real-time PCR further confirmed the downregulation of Wnt
signaling pathway genes and myogenic genes (Fig. 9c). Furthermore, KEGG enrichment of
lectin-enriched proteomic data also support alteration in the Wnt signaling pathway (Fig. 9b).
The activation of Wnt signaling cascade involves the engagement of Wnt ligands with their
receptor complexes that leads to receptor activation and subsequent inhibition of β-catenin
phosphorylation and degradation, thereby inducing translocation of β-catenin into the nucleus
where it promotes the transcription of effector genes68. In C2C12 cells, we assessed the
abundance of nuclear activated β-catenin during the proliferation and differentiation phase. The
data indicated a significant decrease in the level of active β-catenin in the nuclear fractions of
Gmppb-KO C2C12 cells during differentiation, suggesting attenuated Wnt signaling (Fig. 9d).
Previous studies have demonstrated that reduced nuclear abundance of β-catenin results in
reduced differentiation efficiency of muscle progenitor cells (MPCs), while its constitutive
presence in the nucleus induces premature differentiation of MPCs, leading to a strong induction
of Myogenin expression in primary myoblasts71. Multiple studies have also suggested that β-
catenin directly interacts with the myogenic factor MyoD during muscle development, thus
facilitating subsequent muscle differentiation processes72,73. Thus, the impaired Wnt signaling
cascade in Gmppb-KO cells may underlie defects in muscle differentiation and regeneration.
To test the hypothesis, we administered CHIR-99021, a GSK3β inhibitor that stabilizes β-
catenin and thereby activating the Wnt pathway independently of Wnt ligand-receptor
interactions, during the differentiation in C2C12 myoblasts, as well as in FACS-isolated muscle
satellite cells. The results showed that compared to DMSO-treated cells, CHIR-99021
significantly enhanced myotube formation and fusion indices in Gmppb-KO C2C12 cells (Fig.
9e) and GmppbP32L/+satellite cells (Fig. 9f). Furthermore, treating the mice post CTX-induced
muscle injuries with CHIR-99021 led to significant increases in the cross-sectional area of
regenerating muscle fibers in GmppbP32L/+ mice (Fig. 9g). 48-week-old GmppbP32L/+ mice treated
with CHIR-99021 for 7 consecutive days successfully restored motor ability assessed by rotarod
and grip tests (Supplementary Fig. 11d and e). Although pharmacological activation of Wnt
signaling with CHIR promotes differentiation even in WT cells, the pronounced rescue in
Gmppb-KO cells indicates that reduced β-catenin activity is a major downstream consequence of
GMPPB loss. Thus, while the primary defect in GMPPB deficiency lies in protein
hypoglycosylation, secondary attenuation of Wnt/β-catenin signaling likely contributes to the
impaired regenerative response. Enhancing this pathway may therefore represent a rational
strategy to ameliorate myogenic defects associated with disrupted glycosylation.

AAV-GMPPB treatment ameliorated the myopathy phenotype of Gmppb mutant mice
Most muscular dystrophies are monogenic diseases, and gene therapy mediated by adeno-
associated virus (AAV) to replace defective genes holds promise as a potentially curative
treatment, as a single injection may achieve therapeutic effects74. Early studies have shown that
AAV-mediated overexpression of LARGE improved the hypoglycosylation of α-dystroglycan (α-
DG) and pathological status of muscles in LARGEmyd muscular dystrophy mice. Additionally,
LARGE overexpression was able to compensate for the α-DG glycosylation defects caused by
mutations in FKTN and POMGnT175,76. Preclinical data suggests that intravascular AAV micro-
dystrophin delivery can significantly ameliorate muscle pathology, enhance muscle force, and
attenuate dystrophic cardiomyopathy in DMD animal models33, as well as improving disease
symptoms in DMD patients77. Therefore, we investigated whether AAV-mediated expression of
GMPPB ameliorates the muscular phenotypes in GmppbP32L/+ mice. In our studies, GMPPB
expression was achieved using the serotype Myo2A adeno-associated virus (Myo2A-AAV),
which has exhibited favorable muscle transduction efficiency compared to the serotype AAV978.
Based on the phenotypic characterization of GmppbP32L/+ mice in our prior findings where we
observed a delayed onset of disease in these mice, we administered systemic delivery of the
Myo2A-Gmppb virus via tail vein injection into 12-week-old GmppbP32L/+ mice to assess the
efficacy of AAV therapy (Supplementary Fig. 12a).
After 3 months of AAV injection, Western blots with the GA muscle samples indicated
durable GMPPB expression (Supplementary Fig. 12b). In these AAV treated GmppbP32L/+ 
muscle samples, a notable reduction in centrally nucleated muscle fibers was evident, with
subsequent restoration of muscle morphology (Fig. 10a, c). Moreover, IIH6C4-positive α-DG
was restored, indicating that AAV therapy elevated the α-DG glycosylation levels of GmppbP32L/+ 
mice muscles (Fig. 10a, b). Meanwhile, Western blots showed increased level of IIH6C4 and
VIA4-1 in the AAV-treated GmppbP32L/+ mice, suggesting that GMPPB overexpression rescues
the α-DG glycosylation (Fig. 10d). Measurement with LC-MS showed restored GDP-mannose
level in GmppbP32L/+-AAV muscles (Supplementary Fig. 12c), while Western blots indicated
restored β-catenin nuclear abundance in GmppbP32L/+-CHIR and GmppbP32L/+-AAV groups (Fig.
10e). Next, we examined the improvement of motor function in mice post-AAV therapy using
rotarod test and grip test. Following gene transfer, GmppbP32L/+ mice in the AAV treatment group
had significantly increased time spent on the rotarod, to a level comparable to those of wild-type
mice (Fig. 10g). The grip strength of the AAV-treated GmppbP32L/+ mice exhibited a significant
improvement, suggesting the restoration of muscle physiological function (Fig. 10h).
Additionally, the serum CK levels in these mice decreased to levels comparable to those seen in
wild-type mice (Fig. 10i). Meanwhile, we induced muscle injuries with CTX 4 months post AAV
administration, to determine the regenerative capacity of muscles. Five days after CTX injection,
GmppbP32L/+ mice treated with AAV showed a significant increase in the cross-sectional areas of
regenerating muscle fibers compared to untreated GmppbP32L/+ mice, with no statistically
significant difference observed relative to wild-type mice (Supplementary Fig. 12d). The
number of eMyHC+ muscle fibers, demonstrated by eMyHC immunofluorescent labeling,
increased by approximately 50% in GmppbP32L/+ mice (Fig. 10j and k). As shown above, we
observed reduced nerve conductance in GmppbP32L/+ mice compared to wild-type controls (Fig.
3e and f). To evaluate the functional impact of therapeutic interventions, we next measured NCV
following AAV-GMPPB or CHIR treatment. AAV-GMPPB–treated GmppbP32L/+ mice showed a
marked improvement in both conduction velocity and CMAP amplitude, reaching levels
comparable to wild-type animals. In contrast, CHIR-treated GmppbP32L/+ mice exhibited only
marginal changes, with NCV and CMAP remaining significantly lower than that of wild-type or
AAV-treated groups (Fig. 3e and f). These results indicate that direct restoration of GMPPB
expression effectively rescues nerve transmission defects, whereas pharmacological modulation
by CHIR provides limited benefit at the functional level. This discrepancy likely reflects the
difference in therapeutic mechanisms: AAV delivery reinstates GMPPB expression and thereby
corrects the underlying glycosylation defect, whereas CHIR acts indirectly through GSK3
inhibition and may only modulate downstream signaling or metabolic compensation. Because
proper glycosylation of α-dystroglycan and other neuronal glycoproteins is essential for efficient
neuromuscular transmission, direct restoration of GMPPB function appears necessary for
functional rescue of nerve conduction.

In conclusion, AAV-mediated GMPPB overexpression significantly enhances the muscle
regeneration capacity and nerve conductance of GmppbP32L+/- mice. These results underscore the
effectiveness of AAV-GMPPB gene therapy in alleviating the disease phenotype in GmppbP32L/+ 
mice.

Collectively, our data reveal that GMPPB deficiency leads to widespread hypoglycosylation,
impaired Wnt/β-catenin signaling, and defective differentiation and regeneration of skeletal
muscle stem cells. The P32L mutation results in more pronounced phenotypes than simple
haploinsufficiency, manifesting as progressive muscular dystrophy accompanied by neurological
involvement, including Purkinje cell loss and peripheral demyelination. These cellular and tissue
defects converge on disrupted glycosylation-dependent signaling and proteostasis stress.
Accordingly pharmacological activation of Wnt signaling partially rescued the myogenic defects,
while AAV-mediated Gmppb expression effectively restored α-dystroglycan glycosylation,
improved neuromuscular performance, and normalized muscle regeneration. Together, these
findings establish GmppbP32L/+ mice as a faithful model of GMPPB-associated
dystroglycanopathy and provide proof-of-principle that both pathway modulation and gene
replacement represent viable therapeutic strategies.

Methods
Ethical statement
All animal experiments were carried out in accordance with ethical regulations regarding
animal research and were approved by the Experimental Animal Ethics Committee of the
School of Basic Medicine of FudanUniversity under license number IACUC20211018-001.
Generation and genotyping of Gmppb mutant mice. The Gmppb knock-out (Gmppb-KO) and
knock-in mice (GmppbP32L and GmppbR287Q mice) were generated by the CRISPR-cas9 system,
in collaboration with the National Resource Center for Mutant Mice, Nanjing University. Gmppb
knock-out mice were generated in C57/BL6 mice and deleted 303 bp genomic sequence. The
GmppbP32L and GmppbR287Q mutations were introduced into the mouse gmppb orthologue to
generate GmppbP32L mice and GmppbR287Q mice using the CRISPR system and donor template
in C57/BL6 mice. The following genotyping primers were designed for testing Gmppb-KO mice:
P1: 5’-AACATCTGCAGACGCCAT-TGA-3’; P2: 5’-AGGGTGGGAATGTCAGAGGA-3’; P3:
5’-GGGAGGGTATGGGACAICACGG-3’; P4：5’-AGGGACCTAAAGAAAG-
TAAAGGGCTAAGT-3’, Gmppb knock-out mice genotyping can be tested by analyzing PCR
products. The PCR results from the GmppbP32L and GmppbR287Q mice should be sequenced using
the primers listed below: P32L-F: 5’-GGGCGTTGAAATCTAATTGG-3’；P32L-R: 5’-
CATGTAGCTCACTGCCAGGATCA-3’; R287Q-F: 5’-TGCCTCTTCCTACAATCGCTGA-3’;
R287Q-R: 5’-CTATGACATCTTCACCCAGCACTG-3’. Mice embryos from intercrosses of
heterozygotes were also genotyped using the above primer. For embryo genotyping, pregnant
females were euthanized at the indicated gestational stage, and embryos were dissected from the
uterus under a stereomicroscope. Extraembryonic membranes (yolk sac and placenta) were
carefully removed to avoid maternal tissue contamination. Embryos were briefly rinsed in sterile
phosphate-buffered saline (PBS) and placed in fresh tubes for DNA extraction. Genomic DNA
was isolated from embryonic tissue (head or trunk) using standard lysis buffer and proteinase K
digestion, followed by PCR-based genotyping using allele-specific primers. All procedures
followed established protocols for embryo handling and genotyping103. In the case of partially
resorbed embryos, only clearly identifiable embryonic tissue was dissected free from maternal
decidual tissue under a stereomicroscope. Samples were extensively rinsed in fresh PBS prior to
DNA extraction. Resorption sites without discernible embryonic structures were not used for
genotyping to avoid maternal contamination.
Sex was not considered in the study design as no previous report indicated obvious association
of sex and GMPPB phenotypes. Therefore, the analyses did not involve sex-specific
considerations.

Cell lines and Gmppb mutant C2C12 cells. C2C12 cell lines were purchased from the American
Type Culture Collection (ATCC). Cell lines were authenticated by short tandem repeat analyses,
and routinely monitored for mycoplasma contamination every month. To knock out Gmppb in
C2C12 cells, we transfected C2C12 cells with CRISPR plasmids PX458-GFP and PX459-
puromycine with two sgRNA. Two pairs of sgRNAs were designed to target exons 1 and 5 of
Gmppb gene: exon 1 (F: 5’-CACCGLAGCACTCCGAAGCCATTGG-3’; R: 5’-AAACCCAA-
TGGCTTCGGAGTGCTC-3’); exon5 (F: 5’- CACCGAGAGCCTTCCAAGTATG-
GTG-3’; R: 5’- AAACCACCATACTTGGAAGGCTCTC-3’). We tested transfection efficiency
through PX458 plasmids with GFP. At 48 h after transfection, the cells were diluted and cultured
with puromycine (4 μg/ml) for a two-day resistance screening. The clones were then diluted to a
96-well plate to expand the culture. Finally, expanded monoclonal cells were confirmed by PCR
sequencing. The efficacy of guide RNA was determined by western blotting. In addition, we
constructed GMPPB mutant plasmids in Gmppb-KO cells. To acquire the Gmppb open reading
frame, we amplified Gmppb CDS (coding DNA sequence) using C2C12 cDNA as a template and
cloned Gmppb CDS into pCDH-3xflag-GFP-puro (Addgene, #167463). Then, the allelic
mutations c.95C>T, c.485C>T, c.553C>T, c.860G>A and c.988G>A were introduced into the
wild-type plasmid using the NEBuilder HiFi DNA assembly master mix (New England Biolabs,
MA, cat. # E2621S). All plasmids were verified by Sanger sequencing, and overexpressed
GMPPB mutant proteins were also verified by Western blots.

Western blot analysis. For total protein extraction, C2C12 and satellite cells were lysed in RIPA
buffer (Beyotime Biotechnology) and then centrifuged at 12000 rpm for 15 minutes at 4 ℃. The
supernatant was collected in a new tube. Skeletal muscle tissue protein extraction using Minute™
Total Protein Extraction Kit (Invent Biotechnologies). Nuclear protein extraction using Nuclear
and Cytoplasmic Protein Extraction Kit (KeyGEN BioTECH). For glycoprotein extraction.
100mg skeletal muscles were added to 1ml cold Tris-buffered saline [TBS,50 mM Tris-Hcl(PH
7.4),100 mM NaCl] plus 1% Triton-X100 and protease inhibitors, and then homogenized for 1
hour. Glycoproteins were captured with 200ul wheat-germ agglutinin (WGA) agarose (Vector
Laboratories) beads overnight. The next day, the homogenate was washed three times with TBS
buffer containing 0.1 % 1% Triton-X100 and protease inhibitors and then eluted with SDS-
loading buffer. Next, 30μg of the total protein was separated by SDS-PAGE and
electrotransferred onto a polyvinylidene fluoride (PVDF) (Millipore) membrane. Membranes
were blocked in 5% skim milk powder for 1 hour at room temperature, then incubated with
primary antibodies at 4 ℃ overnight. The next day, The PVDF membrane was washed with TBST
three times for 5 minutes and stained with the HRP-conjugated secondary antibodies for 1 hour
at room temperature. Membranes were washed with TBST three times for 5 minutes and imaged
using the Tanon 5200 Series Chemiluminescence Image Analysis System. Band intensities were
analyzed with ImageJ software. The primary antibodies used were as follows: mouse anti-
GMPPB (Proteintech, 15094-1-AP) 1:500, mouse anti-α-DG IIH6C4 (MilliporeSigma, 05-593)
1:250, mouse anti-α-DG VIA4 (MilliporeSigma, 05-298) 1:250, mouse anti-RL2 antibody
(Abcam, ab2739), sheep anti-core DG (R&D Systems, cat# HAF016) 1:1,000, mouse anti-
MyHC (DSHB, MF20) 1:500, mouse anti-β-DG (DSHB, 7D11) 1:500, mouse anti-PAX7 (DSHB,
1-9) 1:500, mouse anti-MyoG antibody (DSHB, F5D) 1:500, mouse anti-vinculin
(MilliporeSigma, V4505) 1:5000, mouse anti-MyoD1(Dako, M3512) 1:500, mouse anti-
myogenin (Santa Cruz,sc-12732) 1:500, mouse anti-GAPDH(Santa Cruz, sc32233)1:1000,
rabbit anti-Myf5 (Abcam, ab125301) 1:500, rabbit anti-Histone H3 (Abcam, ab1791) 1:1000,
rabbit anti-β-tubulin (Abcam, ab6046) 1:500, rabbit Non-phospho (Active) β-catenin (Abclonal,
A22180) 1:2000, rabbit Lamin B1 antibody (Abclonal, A11495) 1:1000. All images of
uncropped blots are available in the Supplementary Information file.

Laminin overlay assay. Laminin overlay assay was performed as previously reported104. Briefly,
protein bands were transferred onto the PVDF membrane. The PVDF membranes were blocked
in laminin binding buffer (LBB: 10 mM triethanolamine, 140 mM NaCl, 1 mM MgCl2, 1 mM
CaCl2, pH 7.6) containing 5% nonfat dry milk, further incubated 10nM mouse EHS laminin
(L2020, Sigma) overnight at 4℃ in LBB. Membranes were washed and incubated with anti-
laminin (ab11575; Abcam) followed by anti-rabbit IgG-HRP. Blots were exposed by
chemiluminescence.

Solid-phase laminin binding assay. Solid-phase assay was also performed as previously
reported104. Wheat germ agglutinin (WGA)-purified skeletal muscle samples were diluted 1:100
in TBS buffer and coated onto 96-well polystyrene ELISA microplates (Costar) overnight at 4℃.
After washing in LBB four times the next day, plates were coated with 3%BSA in LBB for 2
hours. Wells were rinsed with 3%BSA in LBB and added with dilutions of 2 nM-30 nM mouse
EHS laminin (L2020A, Sigma) for 2 hours at room temperature. After washing four times with
3%BSA in LBB, Wells were incubated for 1 hour with anti-laminin (1:5000; ab11575; Abcam)
followed by anti-rabbit HRP. Plates were developed with o-phenylenediamine dihydrochloride
and H2O2, reactions were stopped with 2M H2SO4, and values were obtained on a microplate
reader at 495nm.

Isolation and culture of muscle satellite cells. Muscle satellite cells (MuSCs) isolation was
performed as previously described105. Briefly, hind-limb skeletal muscles were dissected,
digested by collagenase type II, and dissociated using a gentleMACS Dissociator (Miltenyi
Biotec). Digested muscle suspensions were subsequently filtered through a 70-μm cell strainer
and 40-μm cell strainer and then washed to yield a mononuclear cell suspension. The cells were
stained with CD45-BV650(1:75; Clone I3/2.3; BD Bioscience); CD31-BV650 (1:75; clone
390(RUO); BD Biosciences); Sca-1-FITC (1:75; 553335; BD Biosciences; VCAM1-biotin (1:75;
553331; BD Biosciences) and Streptavidin-APC (1:300; 405207; BioLegend). Streptavidin-APC
(1:300; 405207; BioLegend) was used to amplify the VCAM1 signal. Cells were sorted using a
FACSAria II cell sorter (BD Biosciences) equipped with 488-nm, 633-nm, and 405-nm lasers.
Isolated MuSC were either directly used for protein extraction or cultured for proliferation and
differentiation assays. MuSC were cultured in 6 cm plates with growth medium (30% fetal bovine
serum (FBS), 1% chick embryo extract,10 ng/ml basic fibroblast growth factor (bFGF), and 1%
PS in DMEM). To induce MuSC differentiation, cells were incubated with DMEM supplemented
with 2% horse serum.

Histology and Immunofluorescence. Mouse skeletal muscles were collected and embedded in
OCT embedding matrix and frozen in liquid N2-cooled isopentane. 6 μm thick cryosections were
then prepared at -80 ℃ to maintain muscle morphology. To visualize muscle morphology,
Hematoxylin and eosin(H&E) staining was performed in muscle sections. For muscle fiber
diameter quantification, the borders of fibers were manually outlined in given Area of Interest
(AOI) and fibers cut at the edges were excluded. The Hue, Saturation, Intensity (HSI) was
adjusted to effectively separate the stained muscle fibers from the interstitial space. The total
cross-section AOI was calculated and average fiber radius generated by r = √[Area/π. All images
used for quantification for the gastrocnemius were taken from the mid-belly region at a
comparable anatomical level in each animal, and the same selection criteria and magnification
were applied across genotypes. The analyses were double blinded to minimize bias. For the
Immunofluorescence assay, sections were fixed with 4%PFA for 15 minutes at RT and
subsequently permeabilized using 0.5% Triton X-100 in PBS. The sections were blocked for 1
hour at RT in PBS containing 3%BSA. After blocking, sections were stained overnight at 4℃
with primary antibodies. Subsequently, the samples were washed with PBS and stained with the
appropriate secondary antibodies (Alexa Fluor 647, Alexa Fluor 488) for 1h at RT with
avoiding the light. After washing with PBS, 1 μg/ml DAPI was used to stain nuclei for 5 min.
The primary antibodies used were as follows: anti-GMPPB (Proteintech, 15094-1-AP) 1:200,
mouse anti-MyHC (DSHB, MF20)1:300, mouse anti-eMyHC (DSHB, F1.652) 1:50, mouse anti-
MyoG antibody (DSHB, F5D) 1:200, rabbit anti-Laminin (Abcam, ab11575)1:500. Analysis of
myotube diameter and fusion index was performed on images after MyHC immunofluorescence
staining. At least 10 non-overlapping high-power fields were randomly selected for analysis. The
measurement of myotube diameter was defined as follows: for each selected myotube, the width
at the narrowest point was measured perpendicular to its long axis, and each myotube was
measured three times to take the average value. Only muscle-like structures containing 3 or more
nuclei were identified as myotubes. The calculation formula for the fusion index is: Fusion Index
= (Total number of nuclei fused into myotubes / Total number of nuclei in the field of view) ×
100%.

Transmission Electron Microscopy. We examined the ultrastructural alterations in the
gastrocnemius muscles of both 32-week-old and 48-week-old GmppbP32L/+ mice and WT mice.
The muscle tissues were sliced into 1mm × 1mm × 3mm sections with a sharp blade and
subsequently immersed in 2.5% glutaraldehyde fixative solution, where they were fixed at 4°C
for 24 hours. Then, the samples were dehydrated in a series of ethanol and embedded in resin.
The ultra-thin sections were stained with uranyl acetate and lead citrate. The sections were
imaged using a PHILIPS transmission electron microscope (PHILIPS, CM120, Netherlands).
The sciatic and spinal cord nerves were isolated from 32-week mice and underwent similar
processing for EM determination of the cross sections.

Quantitative PCR with reverse transcription. Total RNA was isolated by trizol-chloroform
extraction. 3μg of RNA was reverse-transcribed using the SuperScript III Transcriptase kit
(Invitrogen, #18080093), and qPCR was performed using Taq Pro Universal SYBR qPCR
Master Mix (Vazyme, #Q712). The primers used were as follows: Gapdh (5′-
GTCAAGGCCGAGAATGGGAA-3′, and 5′-CTCGTGGTTCAC
ACCCATCA-3′), Wnt7b (5′- CAATGGTGGTCTGGTACCCAA-3′, and 5′-AGTCT-
CATGGTCCCTTTGTGGTT-3′), Wnt6 (5′-CTTGGTCATGGATCCTACCAGCAT-
CT-3′, and 5′- TCGTTGTTGTGCAGTTGCACCAATGC-3′), Lef1 (5′-ACAGCGA-
CCCGTACATGTCAAA-3′), FZD4 (5′-GACAACTTTCACGCCGCTCATC-3′, and 5′-
CAGGCAAACCCAAATTCTCTCAG-3′), MyoD1 (5′-AGCACTACAGTGGCG-
ACTCA-3′, and 5′-GGCCGCTGTAATCCATCA-3′), Pax7 (5′-GGCACAGAGGAC-
CAAGCTC-3′, and 5′-GCACGCCGGTTACTGAAC-3′), Musk (5′-AGCCGATGTG-
TCTGCTCTTT-3′, and 5′-ACAGGACAGTGGTGGAGGAC-3′), Chrnd (5′-CTAAG-
TCTCACCCTCTCCAACCTCATCTC-3′, and 5′-GCACATTGCAGGCATAAGAA-
ATCTGGAAC-3′), Ryr1 (5′-TTTCCTGGNACCGAGTGTATGGC-3′, and 5′-CAGAC-
AGAGGTAGCGGTTCAGT-3′), Chrna1 (5′- TCATCATTCCCTGCCTGCTCTTCT-3′, and 5′-
TCTCTGCAATGTACTTCACGCCLCT-3′). Experiments were performed in biological
replicates as indicated in figure legends. Gapdh RNA was used to normalize data from RT-qPCR
mouse samples.

Muscle injury. Muscle regeneration was induced by the injection of cardiotoxin (CTX) into TA
muscle. Briefly, a total of 100 μl of 20 μM CTX was injected into TA muscle with three injections.
The 28-gauge needles were inserted parallel to the muscle fiber and injected into each end of the
TA muscle and the middle of the TA muscle separately. CTX-injured muscles were dissected on
3 days, 5 days, and 14 days following injury. Every experiment included at least three animals to
assess muscle regeneration and repair.

GDP-mannose measurement. GDP-mannose measurement was performed as previously
reported42. C2C12 cells (1×107) with different GMPPB mutations were collected and washed
three times with PBS. After that, cells were put into a 15-ml tube and chilled with liquid nitrogen
for 20 seconds. The cell mixture was given a pre-chill of -80 °C and methanol (2 ml, 80%
(vol/vol)). The tube was then kept at 80 °C overnight for incubation. The supernatant was
collected after centrifugation at 12,000g for 20 minutes at 4°C. A speed vacuum dried the samples
at RT for two hours. GDP-mannose was analyzed using a TSQ Quantiva mass spectrometer
(Thermo Fisher) coupled with reverse-phase chromatography48. Mobile phases A and B
contained water with 10mM tributylamine and 15mM acetate and 100% methanol, respectively.
GDP-mannose (purity ≥99.9%) were used as standard samples. Ion transitions of 604/362nm and
588/442nm were used for GDP-mannose. The resolution for Q1 and Q3 was set at 0.7 full-width
at half-maximum (FWHM). The source parameters were as follows: spray voltage, 2,500V; ion
transfer tube temperature, 320°C; vaporizer temperature, 300°C; sheath gas flow rate, 30
(arbitrary units); auxiliary gas flow rate, 10 (arbitrary units).

Ex Vivo muscle culture and CHX/MG-132 treatment. GA muscles were rapidly dissected into
ice-cold DMEM, fascia and tendinous ends were trimmed to improve reagent penetration and
mounted in a chilled slicing chamber and cut into 200–300 µm transverse slices. Only slices with
uniform thickness and intact architecture were used. Slices were equilibrated for 15 min at 37 °C
in DMEM supplemented with 10% FBS and 1% penicillin/streptomycin, with gentle orbital
agitation (50–70 rpm). CHX (50 µg/mL) or MG132 (10 µM) was added to block biosynthesis or
prevent proteasomal turnover. Medium was refreshed every 60–90 min to maintain oxygenation
and remove debris. Slices were harvested at 6 h and protein lysates were isolated by grinding
with liquid nitrogen. Protein lysates were subjected to Western blots using α-DG core antibody
(Millipore, 3D7).

Morris water maze test. The Morris water maze apparatus consisted of a circular pool (120 cm
diameter, 50 cm height) filled with opaque water maintained at 22 °C. A transparent escape
platform (10 cm diameter) was submerged 1 cm below the surface in a fixed location. Mice were
subjected to four trials per day for six consecutive training days, with a maximum trial duration
of 60 s and an intertrial interval of approximately 15 min. If a mouse failed to locate the platform
within 60 s, it was gently guided to the platform and allowed to remain there for 10 s. The starting
position was varied pseudorandomly among four quadrants for each trial. On day 7, a probe trial
was performed in which the platform was removed, and the mouse was allowed to swim freely
for 60 s. Escape latency, swimming path, and time spent in each quadrant were recorded and
analyzed using an automated video tracking system (EthoVision XT, Noldus).

Serum creatine kinase (CK) activity assay. Fresh serum was collected from the orbital sinus
of mice. According to the manufacturer’s guidelines, the CK activity was determined using the
Creatine Kinase Activity Assay Kit (Sigma, MAK116). CK activity was expressed as units/liter.

Untargeted Metabolomics in Mouse Muscle Tissue. Three biological replicates of muscle
tissue samples of Gmppb+/-, GmppbP32L/+, GmppbR287Q/R287Q and WT were subjected to untargeted
metabolomic profiling. Muscle tissue samples were rinsed with pre-chilled PBS, snap-frozen in
liquid nitrogen. Frozen tissues (50 mg) were homogenized with steel balls at 60 Hz for 120 s
under liquid nitrogen cooling. Metabolites were extracted with 1 mL of pre-chilled
methanol:acetonitrile:water (2:2:1, v/v/v ) containing 0.1 μg/mL L-2-chlorophenylalanine,
vortexed, ice-water sonicated for 10 min, incubated at −20 °C for 1 h, and centrifuged at 13,000×g
for 15 min at 4 °C. The extraction was repeated 1 time, supernatants combined, and vacuum-
dried. Dried extracts were reconstituted in 100 μL acetonitrile:water (1:1, v/v), filtered through a
0.22 μm membrane, and transferred to LC-MS vials. Quality control (QC) samples were prepared
by pooling equal volumes of all sample supernatants. Metabolite separation was performed on
an ACQUITY UPLC HSS T3 column (2.1 mm × 100 mm, 1.8 μm) at 40 °C with a mobile phase
of 0.1% formic acid in water (A) and 0.1% formic acid in acetonitrile (B), using a gradient elution
(0–2 min, 2% B; 2–12 min, 2%–98% B; 12–15 min, 98% B; 15–15.1 min, 98%–2% B; 15.1–18
min, 2% B) at 0.3 mL/min, with a 2 μL injection volume. Mass spectrometric detection was
conducted on a Q Exactive HF-X Orbitrap mass spectrometer in positive/negative ESI modes
(spray voltage: 3.5 kV/2.8 kV; capillary temperature: 320 °C). Full-scan MS data (m/z 70–1050,
60,000 resolution) and data-dependent MS/MS spectra (15,000 resolution, 20–40 eV collision
energy) were acquired for non-targeted metabolomics analysis. Metabolites with VIP > 1
(variable importance in the projection of the first principal component in OPLS-DA) and p < 0.05
(Student's t-test) were considered significantly changed. Pathway enrichment analysis was
performed using KEGG and MetaboAnalyst databases.

Bulk RNA Sequencing of Cell and Muscle Tissue Samples. For cell samples, adherent cells
were washed with pre-chilled PBS, dissociated with trypsin, centrifuged at 300×g for 5 min at
4 °C, and the pellet was lysed with RNAiso Plus (Takara) and stored at −80 °C. Muscle tissue
samples were rinsed with pre-chilled PBS, snap-frozen in liquid nitrogen, ground into powder
under liquid nitrogen, and lysed with RNAiso Plus. Total RNA was extracted using RNAiso Plus
following the manufacturer’s protocol. RNA concentration and purity were determined with a
NanoDrop ND-2000 spectrophotometer (A260/A280: 1.8–2.1; A260/A230 ≥ 2.0), and integrity
was verified using an Agilent 2100 Bioanalyzer (RIN ≥ 7.0). Qualified RNA was used for library
construction. mRNA was enriched via oligo (dT) magnetic beads, fragmented into 200–300 bp
fragments, and reverse-transcribed into double-stranded cDNA. After end repair, A-tailing,
adapter ligation, and PCR amplification, libraries were purified with AMPure XP beads and
validated using an Agilent 2100 Bioanalyzer and qPCR. Qualified libraries were sequenced on
an Illumina NovaSeq 6000 platform with the PE150 mode. Raw reads were filtered to obtain
clean reads for subsequent genome alignment, gene expression quantification (FPKM/TPM), and
differential gene expression analysis.

Lectin-based glycoprotein enrichment and LC–MS/MS analysis. GA muscles were ground
in liquid nitrogen and lysed with protease inhibitors. Equal amounts (2mg) of clarified protein
lysates were incubated with ConA-agarose beads (Vector Laboratories, AL-1003-10) at 4°C
overnight. After extensive washing, bound glycoproteins were eluted with 200 mM methyl-α-D-
mannopyranoside. Protein samples were transferred to the filter (Millipore, MRCPRT010) and
washed with UA buffer three times and 25mM NH4HCO3 buffer twice. Trypsin was added (1:50,
wt/wt)) and incubated at 37℃ overnight, and the resulting peptides were collected as a filtrate.
The peptides were analyzed by OrbitrapTM AstralTM mass spectrometer (Thermo Scientific)
connected to an Vanquish Neo system liquid chromatography (Thermo Scientific) in the data
independent acquisition (DIA) mode. Precursor ions were scanned at a mass range of 380-980m/z,
MS1 resolution was 240000 at 200 m/z, Normalized AGC Target: 500%, Maximum IT: 5ms. 299
windows were set for DIA mode in MS2 scanning, Isolation Window: 2m/z, HCD Collision
Energy: 25ev, Normalized AGC Target: 500%, Maximum IT: 3ms. iRT (indexed retention time)
calibration peptides were spiked into the sample. DIA data was analyzed with DIA-NN 1.8.1.
Main software parameters were set as follows: enzyme is trypsin, max missed cleavages is 1,
fixed modification is carbamidomethyl(C), dynamic modification is oxidation(M) and acetyl
(Protein N-term). All reported data were based on 99% confidence for protein identification as
determined by false discovery rate (FDR) ≤ 1%.

Electromyography and nerve conduction velocity test. Mice were anesthetized with 2%
isoflurane, and body temperature was maintained at 37 °C using a heating pad. CMAPs were
recorded using a needle electrode inserted into the gastrocnemius muscle, while the sciatic nerve
was stimulated proximally at the sciatic notch and distally at the ankle using bipolar stimulating
electrodes. Stimulation was delivered as supramaximal square-wave pulses (0.1 ms duration),
and CMAP amplitude and latency were measured using an EMG recorder (Yuyan Instruments,
Shanghai, cat# IX-RA-834). Motor nerve conduction velocity was calculated as the distance
between the two stimulation sites divided by the latency difference.

In vivo muscle contraction test. Muscle contractile performance was assessed in vivo using a
nerve-stimulated gastrocnemius contraction assay. Mice were anesthetized with 2% isoflurane,
and body temperature was maintained at 37 °C using a thermostatic heating pad. The hindlimb
was secured to the platform of an in vivo muscle testing system (Aurora Scientific, #1300A),
with the foot fixed to a footplate connected to a force transducer. The sciatic nerve was exposed
and stimulated via electrodes using 1 ms pulses of supramaximal intensity delivered by a
stimulator (Aurora Scientific, #701C). Isometric contractions of the gastrocnemius muscle were
recorded in response to stimulation frequencies ranging from 10 Hz to 150 Hz to generate a
force–frequency curve. Each contraction was separated by at least 1 min to avoid fatigue.

Grip test. Forelimb muscle strength was measured with a grip strength meter. To assess grip
strength, each mouse was held by the tail and its forelimbs firmly grabbed a metal grid. The
mouse was then pulled backward at an even speed until it released its grip. Each mouse was
assessed ten times, and the average force in each trial was taken as the grip strength.

Rotarod test. The motor coordination of the mice was analyzed by a rotarod test. Three days
before the test, subject mice were exercised for 5min, three times per day. The mice that fell off
the rod were put back again to adapt to the test. On the fourth day, mice were subjected to three
trials mice reached 40 rpm during 300s, with a 30-minute rest between each trial. The Latency
to fall was recorded and 300s is the maximum retention time.

AAV production and injection. For muscle-targeted gene therapy, the MyoAAV 2A-CMV-
GMPPB was generated by PackGene and systemically injected into 12-week-old GmppbP32L+/-
mice. This AAV construct carries the expression cassette of Gmppb, which is driven by the
cytomegalovirus (CMV) promoter. It was reported that MyoAAV administration at low doses
results in therapeutic efficacy in disease models and systemically administrated MyoAAV
transduces primate muscles highly efficiently78. For i.v. injection, a total of 5×1011 genome
copies (GC) particles of AAV (in a volume of 100μl) were injected into adult GmppbP32L+/- mice
(12-week-old) through the tail vein.
MyoAAV viruses were produced by triple-plasmid transfection in human embryonic kidney
(HEK) 293 cells and three rounds of CsCl ultracentrifugation purification. The AAV titer was
quantitated by real-time PCR using a Fast SYBR Green master mix kit (Applied Biosystems,
Foster City, CA, USA) with a pair of primers that amplify a fragment in the CMV promoter. The
primers were as follows: forward, 5′- TGGCACCAAAATCAACGGGAC -3′; reverse, 5′-
CGTCCAGCTCGACCAGGAT -3′.

Intracellular Ca2+ measurements. C2C12 cells were incubated with fluo-4-AM at 37°C in a
buffer containing 137 mM NaCl, 4 mM KCl, 1 mM MgCl2,1 mM CaCl2, 10 mM glucose and 10
mM HEPES-NaOH (pH 7.4) for 60 minutes. The tube was maintained in the assay buffer at room
temperature in a SpectraMax i3X Microplate reader. Measure fluorescence using instrument
settings appropriate for excitation at 494 nm and emission at 516 nm. The effects of ATP were
also measured as above but in the CaCl2-free buffer with the addition of 0.05 mM EGTA. The
detailed experiment procedures can be seen in the guidelines of Fluo-4 Direct™ Calcium Assay
Kits (Invitrogen, #F10471).

Treatment with CHIR-99021 in vitro and in vivo. In order to verify the effect of canonical
Wnt signaling on muscle differentiation, C2C12 myoblasts and Satellite cells were treated with
3 μM CHIR-99021 (Selleck, #S1263) during differentiation. Immuno-fluorescence analysis was
performed after 4 days of differentiation. To activate canonical Wnt signaling in vivo, the TA
muscles of WT and GmppbP32L/+ mice were injected with 10 μM CHIR-99021 (20μl) or saline at
2.5d post injury. The injured TA muscles were then collected for H&E analysis at 5 d post-injury.
For continuous treatment, mice received 10 μM CHIR-99021 (20μl) each day at the TA muscles
for 7 days before rotarod/grip tests.

Statistical analysis. All experiments included at least three biological replicates. Prism 9.0
(GraphPad) was used for analysis. The areas of fluorescent staining for muscle fibers and nuclei
were determined using ImageJ software. Statistical analyses were based on Student’s t-test or
analysis of variance (ANOVA) with Bonferroni post hoc test or Tukey post-hoc test.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Revision „Genetically engineered mouse models unveil mechanisms and therapeutic strategies for GMPPB-associated dystroglycanopathy”\nThe manuscript „Genetically engineered mouse models unveil mechanisms and therapeutic strategies for GMPPB-associated dystroglycanopathy” assesses new therapeutic strategies for GMPPB-associated muscle weakness. The data provided in this study might serve as a basis for future gene therapy for patients. The authors provide interesting insights into the mechanisms of GMPPB-associated muscle disorders. However, some findings have already been reported by other groups and a deeper analysis of the mechanisms is missing. I recommend major revision for this manuscript. Please find my comments below.\nMinor points:\n• Grammar and spelling should be revised. Please use abbreviations (e.g. DPG vs. DGP), capitalization and italicisation of words consistently. Please use the same designation of genotypes throughout (e.g. Gmppb+/- vs. KO+/- or P32L vs P32l). Please follow either the 1- or 3-letter amino acid code throughout the paper and make sure that the labeling of graphs is self explaining (see Fig 7e).\n• Please number main figures and pages.\n• Authors should discuss published studies related to the topic.\n• Introduction:\no Quite often references are not given.\no Authors did not comment on other studies already performed in mice, cell culture, or zebrafish.\no The DGC is also important for cell signaling.\no This study is not the first one that shows that GMPPB mutations contribute to muscular dystrophy – other studies already provided direct evidence, either in zebrafish or in cell culture models.\n• Results:\no Authors should comment if there is anything known about post-translational protein modifications, especially in regard of patient mutations since this may affect protein stability, interaction, localization or activity.\no Then layout of graphs differs in shape, data points and color between figures. Authors should provide graphs with individual data points.\no Generation of mutant mouse models: Authors should clearly describe how they created these mouse models (not only in material and methods section).\no Figure 1: Please perform a PCR on isolated blastocysts to show whether you find homozygous blastocysts for assessed mutations or not. If so, blastocyst implantation/ hatching studies would be recommended. Figure 1d + f: Please provide images with the same magnification. Please provide images for WT mice as well. Please clarify whether you mean GmppbR287Q+/+ or GmppbR287Q-/-. Figure 1e + g: Why are only WT mice shown/ labeled? Please show homozygous and heterozygous P23L and KO mice as well. Please explain how you genotyped the lost embryos without contamination by maternal tissue.\no Figure 2: Please also show CK activity for younger mouse cohorts. It would be nice to see additional data regarding the analysis of autophagy in myoblasts and murine tissue lysates. Mitochondrial function should be assessed in more detail (e.g. in myoblasts). Please explain how the fiber diameter was measured.\no Figure S2: How much mannose did you administer and in which way (orally via food or water, i.p., etc.)? When did you start to administer the mannose? Are mannose contents of the food known?\no Figure 3: It is not clear whether the VIA4 and IIH6CV4 immunoblots were performed after WGA enrichment or without enrichment – the main text is confusing. If the blots are performed after WGA enrichment, authors should normalize VIA4 and IIH6C4 intensities to the α-DG core peptide and should show α-DG glycosylation without enrichment.\no Figure 4: Authors see an increase in Myf5 in GMPPB KO cells compared to WT cells. What does this mean? Figure 4f-g: Authors should describe how they measured the diameter of myotubes as well as the fusion index. Figure 4i-o: Authors should show p-values and individual data points. At best, they should provide at least 3 biological replicates.\no Figure S5d: Please show images with a higher magnification.\no Figure 5: Glycoproteomics would give more insights into affected glycoproteins. NMJ morphology analysis is recommended. Please provide a list with affected genes and verify some by qPCR and immunoblot analysis. In mice, RNA-seq data suggest that the immune system seems to be affected. Authors should state if GMPPB mutant mice are more prone to get diseases/ sepsis and compare their data to patients. Authors should show heatmaps or volcano plots with affected genes.\n• Discussion:\no Authors should cite more the related studies already published.\no How do authors explain that in GMPPA KO mice mannose depletion worked? Authors should discuss why they didn´t try mannose supplementation after birth for heterozygous GMPPB KO and P32L mice.\no Authors should discuss their findings in the context of other studies about dietary intervention in CDGs (e.g. glucose).\no Authors should discuss their findings in myoblasts with other studies\no Authors should comment on cognitive functions upon disruption of GMPPB\no Please discuss to what extent the generated models are able to replicate the entire spectrum of disease phenotypes seen in human patients.\nMajor points:\n• Animal numbers should be increased, especially for behavioral tests (minimum: 10 mice per cohort). Animal numbers should always be indicated.\n• HE stained skeletal muscle images: Cell borders are not visible. I wonder how authors were able to measure the fiber diameter in view of this image quality. Please provide images with a better resolution or higher magnification.\n• It is puzzling that GMPPB +/- mice show a much milder phenotype than GMPPB P32L +/- mice. As discussed, this suggests a dominant negative effect of the point mutation on the remaining WT allele. Additional experiments should be conducted to confirm this hypothesis. The authors should measure GMPPB enzyme activity and GDP-mannose levels in tissues for the different mouse models. Does the P32L mutant inhibit WT activity? In this regard, it would be interesting to assess the effect of the point mutations on GMPPB protein abundance in vivo in WB and histology. Dis authors observe GMPPB aggregate formation in vivo? Assuming a dominant negative effect of the P32L mutant: Why does this not interfere with the gene replacement therapy?\n• Figure 1; S1: How many pups/ embryos were assessed? Normally you need to have at least 5 breeding pairs to get valid data. How often were mice backcrossed (should be more than 3 times at least)? Include data on how many breedings were analysed and provide exact numbers of the off-spring. Breeding +/- with +/+ mice or +/+ with +/+ and repeating the same analysis would help to verify that embryo resorption is indeed due to homozygosity. Embryonic lethality, as seen in the pictures provided, might be not related to the genotype. Authors did not provide data for R287Q embryos.\n• Figure S3: Authors should also assess the cerebellum of GMPPB mutant mice since GMPPA KO mice show a progressive loss of Purkinje cells. Brain sections should be assessed at different time points (e.g. 3 and 12 months of age) to make a clear statement. Authors should assess spinal cord neurons and sciatic nerve axons too, since neurons with long extensions are usually more sensitive. Authors should assess cognitive functions of GMPPB mutant mice as well since most patients present with additional neurological symptoms. The authors should also assess the glycosylation of α-DG in brain tissue.\n• Figure 3: Since authors claim that hypoglycosylation of α-DG might affect α-DG stability, they should perform turnover experiments (e.g. with cycloheximide). Laminin OL should be normalized to α-DG core peptide and VIA4 as well as IIH6C4 bands should be shown. A deglycosylation experiment of α-DG would show if hypoglycosylation does indeed lead to a decreased protein stability and laminin binding. Authors should perform α-DG core immunofluorescence staining if possible or at least DAG1 staining. Authors should assess the distribution of laminin by immunofluorescence stainings.\n• Figure 4: Because GMPPB KO cells still show a band at the molecular weight of GMPPB, authors should assess GMPPB RNA levels.\n• Figure S7/8: Nerve conductance velocity measurements with and without treatment (CHIR and AAV) should be performed since most patients show differences here.\n• Figure 7: Authors should show data for centralized nuclei and fiber diameter for all time-points after and before CTX injection. Additionally, authors should provide immunofluorescence images with according analysis for developmental markers for all time points before and after CTX injection. Is CK activity increased after CTX injection and does it normalize?\n• Figure 8: Authors should provide data for behavioral tests after CHIR treatment. Authors should assess glycosylation of Wnt proteins before and after CHIR treatment. Authors should discuss the exact mechanisms connecting GMPPB mutations and Wnt signaling.\n• Figure 3 and 9: In skeletal muscle, α-DG glycosylation should be over 150 kda (up to 280 kDa). How do authors explain this? Which gel percentage did authors use? Authors should assess α-DG core peptide by immunoblot as well.\n• Figure 9: Authors should assess GDP-mannose levels or whole tissue glycosylation and Wnt signaling before and after gene therapy in skeletal muscle and brain. At best, authors should assess other organs to judge about possible side effects. Authors should also show data for WT mice injected with the AAV vector in order to judge potential side effects. Authors should verify GMPPB protein expression after gene replacement therapy.", "reviewer_2": "Dystroglycanopathy (DGpathy) is a group of muscular dystrophies that are caused by abnormal glycosylation of dystroglycan (DG), which serves a cell surface laminin receptor. DGpathy is accompanied by central nervous system disorders in addition to muscular dystrophy. At least 18 responsible genes for DGpathy are known. GMPPB is an enzyme that synthesizes GDP-mannose, which is used in the synthesis of various glycans. Modifications of O-mannose-type glycans, which are found in DG and other proteins, and GPI anchors use Dol-P-mannose as a donor substrate and Dol-P-mannose is synthesized from GDP-mannose and dolichol phosphate. GDP-mannose is also used in the process of N-glycosylation that are found in many glycoproteins. Therefore, it is thought that the effects of GMPPB mutations also occur in glycoproteins other than DG, and that this is involved in the pathology, but the details are not clear.\nThis paper contains extensive data showing that GMPPB knockout and mutant mice show muscular dystrophic pathology, that GMPPB mutations cause defects in muscle differentiation and regeneration, that GSK inhibitors can suppress the decline in muscle regeneration, and that AAV gene therapy can rescue the pathology. However, most of the data presented by the authors are confirmations what has already been clarified in other studies using other DGpathy models. Although the suppression of the decline in muscle regeneration by the GSK inhibitor is a new finding in DGpathy models, it is known that GSK are involved in myogenic differentiation. In addition, the reason why P32L/+ heterozygous mice show muscular dystrophic phenotype needs to be solved because DGpathy is a recessive genetic disease.\nSpecific Comments\n1. DGpathy is a recessive genetic disease, but the reason why P32L/+ heterozygous mice show dystrophic pathology is unclear. In the Discussion, the authors stated that the P32L mutation has a potential dominant negative effect, and its biochemical elucidation is necessary. However, this point should be clarified in the present paper. Do the parents of patients with the P32L mutation show mild muscle symptoms like the heterozygous mice? If so, there should be a dominant form of GMPPB-DGpathy, but as far as I know, there have been no reports of such a condition.\n2. In relation to the above comment, although a decrease in the reactivity of the antibody IIH6, which recognizes laminin-binding glycans and its reactivity is absent in DGpahty, is observed in Figure 3, the reactivity itself remains to some extent, and quantitative measurements show that more than half of the laminin-binding activity remains. On the other hand, other reports have shown that the disease does not develop unless the IIH6 reactivity and laminin-binding activity are almost completely lost. An experimental explanation is needed for the pathology in P32L/+ mice despite a considerable amount of glycans and laminin-binding activity remaining. For example, modification of glycoproteins with O-mannose glycans other than DG may be affected. What about the amount of GDP-mannose in the tissues of P32L/+ heterozygous mice? Have the authors tried any methods other than lectin blotting to identify molecules whose glycosylation is affected? Has N-glycosylation changed (since the band position of βDG does not change, how about treating other N-glycosylated proteins with PNGaseF?)\n3. In Fig. 3e, the authors mentioned that the protein amount of αDG is decreased. In the case of DGpathy, the molecular size of αDG is dramatically decreased due to lack of O-mannosyl glycans. The authors should use better antibodies (e.g., Millipore, 3D7) to examine whether the protein amount is decreased or the glycosylation is abnormal. If the molecular size is not decreased, it cannot be said that abnormal DG glycosylation is the cause of the pathology in P32L/+ mice. In addition, immunostaining using an antibody against αDG core protein should be performed to examine the possibility that the decreased amount of αDG protein on the plasma membrane is involved in the pathology. If the decreased localization of αDG core protein on the cell membrane is the cause of the pathology, cell biological experiments are required to clarify the mechanism leading to the decreased localization (intracellular trafficking and proteostasis).\n4. Patients with GMPPB mutations show a wide clinical spectrum, including muscle and central nervous system lesions, but P32L/+ heterozygous mice do not show significant brain structural abnormalities. Have the authors confirmed the DG glycosylation status in the brain?\n5. Figure 4: Among D1, 3, 5, and 7, by when is proliferation, and from when is differentiation? In Gmppb-KO C2C12, the expression of Pax7 and MyoD is dramatically decreased at D0. Therefore, it is not appropriate to say that the decline of Pax7 and MyoD observed during differentiation in normal cells is absent in Gmppb-KO cells. Rather, is it possible that myogenicity has changed during the genome editing experiment independent of genome editing? Since the myogenicity of C2C12 cells is easily changed depending on culture conditions, Gmppb rescue cells should also be used as a control group. Is myogenicity improved in the cells shown in Fig S5?\n6. In the experiment with isolated satellite cells (Fig. 6), are there any differences between WT and P32L/+ in the number and proliferation rate of isolated satellite cells? Differentiation in P32L/+ heterozygous cells seems to be reduced, but no data is present to show that this effect is due to abnormal glycosylation of DG? Have the authors confirmed the DG status in isolated satellite cells and myotubes?\n7. Regarding the effect of GSK3b inhibitor, since the regeneration rate is increased even in wild type condition, it is difficult to say that Gmppb is involved in myogenesis through modulation of Wnt signaling.\nMinor Comments\nOverall, I think the way of mouse genotype description should be reconsidered. R287Q/R287Q is easier to understand than R287Q-/-, and P32L/+ is easier to understand than P32L+/-, and I think these are more commonly used notations.\nFig. 4h MyOD should be MyoD\nSupplementary Fig1: the resolution of tissue staining images is low.\nSupplementary Fig5: Is WT GMPPB localized in the cytoplasm? Please state this in the text.", "reviewer_3": "The manuscript “Genetically Engineered Mouse Models Unveil Mechanisms and Therapeutic Strategies for GMPPB-Associated Dystroglycanopathy” presents work on mouse models that attempt to recapitulate human models of DGP using point mutations and GMPPB KO. Overall, the study design is quite complicated and hard to follow with various models intertwined throughout the manuscript. There are very novel and important findings, but the translational aspect of some of their results are questionable due to the disjointed presentation. Also, I find the therapeutic potential of their studies in cells and mice loose, given that they use small sample sizes, a small molecule that inhibits a vast signaling pathway (not only Wnt), and AAV models reinserting a gene that they specifically mutated in the same mouse (can we be certain in a human that this is the only gene affecting the phenotype?). I think more clear presentation of the data can result in a very convincing conclusion, but as it stands I recommend substantial revisions.\nSpecific comments are listed below:\n- The link between mechanistic link between MuSC and GMPPB is unclear in the intro, other than that they both are related to DMD. Does GMPPB mutation affect MuSC somehow? What is the mechanistic connection between GMPPB and MuSC function?\n- There is no indication of any of the clinical participant study characteristics in Fig 1a.\n- The data in figures 1-2 are nice and show a good recapitulation of human symptoms in the P32L mutants, but this represents a mutation that occurs in ~9 of 109 patients (~8%) based on the data in Fig 1a. Thus, as stated above in my comment on the introduction section, therapeutic potential based on these data are limited to a specific mutation in humans.\n- Quantification of Gastroc cross sections is unclear (throughout multiple figures). The authors show a very zoomed in image of small area of the gastroc cross section and the methods describe now specific criteria for how certain zoomed regions were selected for quantification. The gastroc is a muscle that has differing muscle fiber type proportion in different regions of the muscle. If the authors did not quantify the entire cross section then there can be significant bias in the results based on what area of the muscle was quantified for different genotypes.\n- The muscle function analyses (rotarod and grip test) chosen are dependent on multiple factors, such as mouse body composition/weight, neural connectivity, etc. Do the general body mass or composition change in these mice? Do the authors have any muscle specific function tests (such as in vivo or ex-vivo contraction) that are more indicative of function at the muscle only?\n- No indication of what stains/colors represent in ECM staining images or legends. Further, quantification of fluorescence intensity in histological sections is extremely subject to microscope settings and without proper controls cannot be confirmed by a reviewer. How can the authors justify the possibility that different exposure settings were not used for each of these images?\n- Some bar graphs do not show individual values. This should be shown.\n- It is unclear why such emphasis is placed on GMPPB KO in myotubes when it appears that in humans the GMPPB gene has mutations which result in phenotypes (not KO), and in the mice the genotype that most recapitulated human symptoms was the P32L mutant and that KO is lethal. It was also incredibly frustrating and difficult to understand as a reader when the authors finessed their wording. Sometimes they say “GMPPB loss of function” when they are referring to P32L mutant and other times they us the phrase “GMPPB loss” when referring to the KO. However, the translational impact of these two are very different, and the results from these two are intertwined throughout the results and not always immediately recognizable in the figures. This convolutes what appears to be a fairly compelling story (with regard to the P32L mutation).\n- GSK3B inhibition (by CHOR-99021) will influence a host of pathways aside from WNT signaling. Though Wnt signaling is a great hypothesis, how can the authors be sure that the Wnt pathway is what influence their results and not other GSK3B related effects?"}, "author_response": "We sincerely thank the reviewers for their thoughtful and constructive feedback on our manuscript. Their comments have been instrumental in guiding us to substantially strengthen both the depth and scope of our study.\nIn this revision, we expanded the work to provide a more comprehensive characterization of GMPPB deficiency in vivo. Beyond confirming the muscular abnormalities, our analyses now reveal clear neurological involvement, including structural and behavioral alterations consistent with impaired motor coordination and cognitive performance. Proteomic profiling uncovered dysregulation of pathways related to glycosylation, mitochondrial function, and autophagy, aligning with ultrastructural evidence of autophagosome accumulation in mutant tissues. Together, these findings indicate that GMPPB mutations exert systemic effects extending beyond muscle pathology.\nWe also revisited the P32L variant, which produced more pronounced phenotypic consequences than reported in the previous version. While the precise mechanism remains under investigation, our results suggest that loss of GMPPB function may exert broader cellular impacts than initially recognized (for example, the hypoglycosylation of α-DG). In addition, we provide multi-level evidence that modulation of the GSK3 pathway can restore aspects of differentiation and muscle regeneration, highlighting its potential therapeutic relevance. In addition, our extended analyses uncovered previously unrecognized neurological phenotypes in the Gmppb mutant mice, including abnormalities in neuronal morphology and function, which further broaden the disease spectrum beyond skeletal muscle pathology. These findings strengthen our understanding of the biological consequences of impaired protein glycosylation and indicate that GMPPB deficiency exerts systemic effects on both muscular and neural tissues. By integrating these observations with our molecular and cellular analyses, we provide a more comprehensive view of how GMPPB dysfunction perturbs glycoprotein processing, cellular homeostasis, and ultimately tissue integrity.\nThrough this revision process, we recognized that our findings collectively point to a broader and still unresolved question in the molecular pathology of GMPPB-related dystroglycanopathy: how does a defect in GDP-mannose production lead from impaired protein glycosylation to cellular degeneration? While α-DG hypoglycosylation has been considered the major downstream event, our results showing alterations in the autophagic pathways, abnormal calcium responses, mitochondrial fragmentation suggest that additional glycoprotein substrates and intracellular stress pathways may play a crucial role. This emerging view reframes the disease not only as a defect in extracellular adhesion but also as a disorder of proteostasis and organelle quality control, highlighting an important direction for future research.\nThis study represents our first steps into this field, and in many ways, the journey has been both challenging and enlightening. We are deeply grateful to the reviewers for their thoughtful and rigorous feedback, which guided us to look beyond our initial assumptions. Although the scope of the requested work was substantial, the additional experiments and analyses they encouraged led us to recognize that the locomotor defect arises not only from muscular dysfunction but also from neurological involvement. These findings have reshaped our understanding of the disease as a more systemic manifestation. Through this process, we have not only strengthened the manuscript but also developed the conceptual and technical foundation needed to pursue the next, more mature questions that emerged from this work. We are sincerely thankful to the reviewers for their mentorship and for helping us advance both scientifically and personally through this revision. Below we outline the point-by-point response to their comments:\n[...]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors did a great revision making this manuscript now acceptable for publication.\nI only have some minor concerns:\n- Authors still use \"Gmppb\" and \"GMPPB\" sometimes for mouse models/cells. This should be avoided.\n- Authors should also comment on other PTMs reported for Gmppb. For example, Gmppb itself is thought to be glycosylated (O-GlcNAc) and methylated. Mutations in Gmppb might also affect these PTMs.\n- Figure S1 d-e: If possible, authors should show E14.5 or E16.5 for both genotypes.\n- Figure 6f: If possible, authors should select another Blot with a higher exposition time so that one can see a band in wt lysates for p62 too (there is plenty p62 in skm lysates to detect).\n- Figure 6g: Magnifications are really blurry, pictures with a higher resolution a recommended.\n- alpha-DG glycosylation blots after WGA enrichment: Authors really should show alpha-DG core peptide probing for these membranes so that one can see if alpha-DG enrichment was similar between genotypes. Beta-DG glycosylation/interaction might be affected too (although Blots suggest otherwise). IIH6C4 membrane probing of total skm lysates should normally give good signal intensities. Authors might consider checking their secondary antibody (should be IgM).\n- Glycoproteomics: Authors should note that ConA enrichment will enrich glycoproteins and their interaction partners. This should be mentioned.\n- Morris Water Maze test: this test is not suited for mice with muscle problems (mice have to swim towards a platform). It would be better to perform \"novel object test\" or \"fear conditioning\". If performing another test is not possible, authors should at least comment this.\nDid authors access depression-like symptoms in these mice? One study linked Gmppb alterations with depression. This should be at least discussed.\n- Turnover experiments: Did authors probe membranes with IIH6C4 or alpha-DG core peptide antibody? This was not clear for me. Best would be if authors would lable alpha-DG membranes with the antibody used.\n- Figure 4e: alpha-DG core peptide seems to have a higher molecular weight in P32L lysates compared to WT lysates although one would assume a lower molecular weight. How is this possible?\n- Authors did not comment on Rev3 question regarding Gmppb Ko myoblasts: Why did authors use Gmppb Ko instead of P32L myoblasts? This should be explained in the text.", "reviewer_2": "The authors have added substantial new data to the revised manuscript, which further deepens our understanding of the pathogenesis of GMPPB-associated DGpathy. In addition, they have adequately addressed my previous concerns. I have only a few additional comments that the authors may consider incorporating into the manuscript.\nAlthough P32L/+ mice exhibit dystrophic pathology, the phenotype is considerably milder than that observed in previously reported recessive DGpathy models such as Large-myd and skeletal muscle selective Myf5-Fktn-cKO mice (Nature 2002, 418:417-22; Hum Mol Genet 2013, 22:3003-15). In contrast, POMGNT1-KO mice and Fktn-transposon knock-in mice showed almost no dystrophic phenotype, despite having more severely hypoglycosylated a-DG compared with P32L/+ mice (Mech Dev 2009, 126, 107-16; Hum Mol Genet 2009,18, 621-31). Taken together, these reports and the present findings strongly suggest the presence of molecular and/or cellular abnormalities, beyond a-DG hypoglycosylation, that contribute to the dystrophic phenotype of P32L/+ mice. The current study therefore provides important insights into the broader pathomechanisms underlying GMPPB-associated DGpathy.\nConcerning Fig 4e and Supplementary Fig 6b, the authors stated that “Western blot analyses with two independent total α-DG antibodies suggested a potential impact of α-DG glycosylation on protein stability, as evidenced by a significant overall reduction in α-DG levels (Fig. 4e and Supplementary Fig. 6b).” However, this description is not correct. As discussed in the rebuttal letter (page 49), these Western blot data indicate abnormal a-DG glycosylation, as evident by reduced reactivity to the IIH6 antibody, and the presence of hypoglycosylated a-DG species (lower band in Supplementary Fig 6b). The reduction of a-DG at the plasma membrane is instead demonstrated in Supplementary Fig 6e. I recommend that the authors revise this description accordingly.\nIn line 653-655, the authors state that “GmppbP32L/+ mice treated with CHIR-99021 for 7 consecutive days successfully restored motor ability assessed by rotarod and grip tests.” However, this observation is difficult to interpret. P32L/+ mice exhibit only a very minor muscle phenotype at 16 weeks of age, suggesting that little muscle regeneration is occurring at this stage. Under such conditions, it is unclear why CHIR treatment, which is shown in this manuscript to enhance regeneration, would lead to improved muscle strength and rotor rod performance. This raises the possibility that CHIR may exert effects beyond promoting regeneration. I encourage the authors to discuss potential alternative mechanisms.", "reviewer_3": "All of my previous comments have been adequately addressed. I feel that the manuscript is improved from the previous version."}, "author_response": "We sincerely thank the reviewers for their careful evaluation of our revised manuscript and for their constructive and insightful comments. We are particularly grateful for the positive assessment that the manuscript is now suitable for publication, and for the remaining suggestions that helped us further improve clarity, rigor, and presentation.\nIn this final revision, we have addressed all outstanding points raised by the reviewers as detailed below.\n[...]"}]

Decision letter