Human Molecular Genetics, 2008, Vol. 17, No. 24
doi:10.1093/hmg/ddn290
Advance Access published on September 9, 2008
3876–3886
Modulation of glycogen synthesis by RNA
interference: towards a new therapeutic
approach for glycogenosis type II
Gaelle Douillard-Guilloux1,2, Nina Raben3, Shoichi Takikita3, Lionel Batista1,2,
Catherine Caillaud1,2, and Emmanuel Richard1,2,{
1
Institut Cochin, Université Paris Descartes, CNRS (UMR 8104), Paris, France, 2Inserm, U567, Paris, France and
Musculoskeletal and Skin Diseases, National Institutes of Health, Bethesda, MD 20892, USA
3
Received July 21, 2008; Revised and Accepted September 8, 2008
Glycogen storage disease type II (GSDII) or Pompe disease is an autosomal recessive disorder caused by
defects in the acid a-glucosidase gene, which leads to lysosomal glycogen accumulation and enlargement
of the lysosomes mainly in cardiac and muscle tissues, resulting in fatal hypertrophic cardiomyopathy and
respiratory failure in the most severely affected patients. Enzyme replacement therapy has already proven
to be beneficial in this disease, but correction of pathology in skeletal muscle still remains a challenge. As
substrate deprivation was successfully used to improve the phenotype in other lysosomal storage disorders,
we explore here a novel therapeutic approach for GSDII based on a modulation of muscle glycogen synthesis. Short hairpin ribonucleic acids (shRNAs) targeted to the two major enzymes involved in glycogen
synthesis, i.e. glycogenin (shGYG) and glycogen synthase (shGYS), were selected. C2C12 cells and primary
myoblasts from GSDII mice were stably transduced with lentiviral vectors expressing both the shRNAs and
the enhanced green fluorescent protein (EGFP) reporter gene. Efficient and specific inhibition of GYG and
GYS was associated not only with a decrease in cytoplasmic and lysosomal glycogen accumulation in transduced cells, but also with a strong reduction in the lysosomal size, as demonstrated by confocal microscopy
analysis. A single intramuscular injection of recombinant AAV-1 (adeno-associated virus-1) vectors expressing shGYS into newborn GSDII mice led to a significant reduction in glycogen accumulation, demonstrating
the in vivo therapeutic efficiency. These data offer new perspectives for the treatment of GSDII and could be
relevant to other muscle glycogenoses.
INTRODUCTION
Glycogen storage disease type II (GSDII) also termed Pompe
disease (MIM 232300), is an autosomal recessive disorder
caused by a deficiency of the lysosomal enzyme, acid
maltase or a-glucosidase (GAA; EC 3.2.1.20). This enzyme
catalyzes the complete hydrolysis of its natural substrate glycogen by cleaving both a-1,4 and a-1,6 glucosidic linkages at
an acidic pH, allowing glucose to be liberated into the cytoplasm. GAA is synthesized as a 110 kDa precursor form,
which is glycosylated and phosphorylated on mannose residues in the endoplasmic reticulum and Golgi apparatus,
which allows the enzyme to be transported to the lysosomes
by the mannose-6-phosphate receptors (MPRs) pathway,
which is common to most lysosomal enzymes. This GAA precursor can also be secreted into the extracellular space where it
is taken up by the ubiquitous cell surface cation-independent
MPR (CI-MPR) on surrounding or distant cells. In the lysosome, the enzyme is further modified by limited proteolysis,
resulting in the formation of two mature forms of 76 and
70 kDa (1). GAA deficiency results in massive glycogen
accumulation in various tissues, among which skeletal and
cardiac muscle are the most affected (2). Clinically, GSDII
manifests as a continuum of phenotypes; the severity of the
disease largely correlates with the level of residual enzyme
To whom correspondence should be addressed at: Institut Cochin, Département Génétique et Développement, 24 rue du Faubourg St Jacques, 75014
Paris, France. Tel: þ33 144412402; Fax: þ33 144412446; Email: [email protected]
†
Present address: INSERM U876, IFR 66, Université Bordeaux 2, 146 rue Léo Saignat, 33076 Bordeaux; Email: [email protected]
# The Author 2008. Published by Oxford University Press. All rights reserved.
For Permissions, please email: [email protected]
Human Molecular Genetics, 2008, Vol. 17, No. 24
activity. Near complete or total loss of GAA activity results in
severe myopathy and cardiomyopathy in infants leading to
death before 2 years of life due to cardiorespiratory failure.
Partial enzyme deficiency results in late-onset forms of
GSDII with slowly progressive muscle weakness without
cardiac involvement and death occurring by respiratory
failure. Animal models mimicking the human disease have
been created by targeted disruption of the GAA gene (3,4).
The models have facilitated the development of different
therapeutic approaches for GSDII. Enzyme replacement
therapy (ERT) has been successfully applied for a number of
lysosomal storage disorders and is now available for Pompe
patients. Both preclinical and clinical studies (5 – 9) have
shown that cardiac muscle responds well to ERT. However,
most of the recombinant GAA ends up in the liver, with
little or no clearing of glycogen in skeletal muscle, particularly
in the advanced stage of the disease (10 – 12). Furthermore, a
selective resistance of type II muscle fibers to ERT was
demonstrated in mice, although this issue is still debated in
humans (13). Other therapeutic approaches such as gene
therapy have been tested in the knockout mice with different
vectors. Peripheral injection of adeno-associated virus
(AAV) vectors with high muscle tropism, expressing human
GAA, results in efficient muscle transduction associated with
sustained GAA expression and glycogen clearance in GSDII
mice (14 – 16). However, a strong and rapid humoral
immune response against human acid alpha glucosidase was
observed requiring the development of immunodeficient
GSDII mice models to evaluate the long-term efficiency of
the treatment (17). Success of both ERT and gene therapy
relies on MPR-mediated delivery of enzyme to the lysosome,
a pathway which is relatively inefficient in muscle due to the
low abundance of these receptors in skeletal muscle (18).
Based on these observations, alternative strategies independent
or complementary to ERT should be warranted.
Substrate reduction therapy (SRT) is another therapeutic
approach for lysosomal storage disorders, based on the inhibition of the biosynthesis of the storage material. It is already
available for type 1 Gaucher disease (19,20), while clinical
trials are ongoing for type 3 Gaucher disease, Niemann-Pick
disease type C and GM2 gangliosidoses (21). Therefore, we
explored the feasibility of an SRT approach in the GSDII
model by modulating glycogen synthesis, which requires
several reactions (22). The initiation step is mediated by glycogenin (GYG; EC 2.4.1.186) which is a self-glycosylating
protein primer that initiates glycogen granule formation (23).
Then, glycogen synthase (GYS; EC 2.4.1.11) catalyzes the
addition of glucose residues to the growing glycogen molecules
through the formation of a-1,4-glucosidic linkages, and finally
the branching enzyme (EC 2.4.1.18) forms the a-1,6-glucosidic
linkages (24). Mammals express two isoforms of GYS and two
of GYG, encoded by genes expressed either in muscle or liver
(23 –25). Extensive efforts to identify the mouse and rat versions of the liver GYG have failed and it appears that this
gene may be absent in non-primates.
In this paper, we have used a ribonucleic acid (RNA)
interference-based strategy to reduce glycogen synthesis in
myoblasts derived from GSDII mice by modulating muscle
GYS and GYG expression. An efficient inhibition of GYS
and GYG messenger RNA (mRNA) was obtained, resulting
3877
in a profound decrease in lysosomal glycogen accumulation
and reduction in lysosomal size in GSDII primary myoblasts.
Furthermore, efficient muscle transduction with an AAV-1
vector expressing EGFP and shGYS in newborn GSDII mice
resulted in strong inhibition of GYS expression and decrease
in cytoplasmic and vacuolar glycogen accumulation. This substrate reduction strategy could constitute a novel approach for
GSDII treatment, which may complement or be used as an
alternative to ERT.
RESULTS
Development of self-inactivated (SIN)-lentiviral vectors
expressing shRNA targeted to the muscle isoforms
of GYS and GYG
Short hairpin RNA (shRNA) targeted to the murine muscle GYS
(named hereafter shGYS1, shGYS2 and shGYS3) and GYG
(shGYG1 and shGYG2) regions were designed (Fig. 1A).
They were chosen to avoid the inhibition of hepatic glycogen
synthesis (these shRNAs had low sequence homology to liver
isoforms). These shRNAs were inserted into the 30 LTR (long
terminal repeat) of a lentiviral construct expressing the EGFP
reporter gene under the control of the ubiquitous phosphoglycerate kinase (PGK) promoter (Fig. 1C). After reverse transcription
and genomic integration, two copies of the shRNA cassette were
present in the provirus at both 50 and 30 LTR extremities ensuring
a strong shRNA expression. Vesicular stomatitis virusglycoprotein (VSV-G) pseudotyped vectors were produced as
described (26) and titers were determined by transduction of
murine myogenic C2C12 cells after serial dilutions of the
vector. The infectious titers ranged from 8 107 to 2.8 108
infectious units/ml.
Efficient shRNA-mediated inhibition of GYS and GYG
impairs glycogen synthesis in C2C12 myogenic cells
In order to analyze the effects of GYS and GYG inhibition on
glycogen synthesis, C2C12 murine myoblasts were transduced
at low (low copy, LC) or high (high copy, HC) multiplicity of
infection (m.o.i.) with the lentiviral vectors (m.o.i. 2 and 50 for
LC and HC, respectively). EGFP-positive (EGFPþ) cells were
isolated by fluorescence-activated cell-sorter (FACS) analysis
7 days after transduction followed by their expansion. Efficient
gene silencing was observed in HC and LC-transduced C2C12
myoblasts as shown by real-time quantitative polymerase chain
reaction (PCR) analysis of GYG and GYS mRNA expression
levels (Fig. 2A and B). In HC-transduced C2C12 myoblasts,
reduction in GYS and GYG mRNA reaches 89 and 75% for
the most effective shGYS2 and shGYG2, respectively. A
vector dose-dependent effect was observed on the inhibition efficiency, correlating with the increase of integrated proviral copy
number, especially for shGYS2 (1 versus 3.4 viral copy/cell for
LC versus HC shGYS2, and 1.1 versus 5.1 viral copy/cell for LC
versus HC shGYG2, respectively). To assess the specificity of
the shRNA constructs, several controls were performed: (i)
GYS and GYG mRNA were quantified in C2C12 cells transduced with lentivectors expressing EGFP without shRNA and
(ii) GYS mRNA was quantified in shGYG-transduced cells
and GYG mRNA was quantified in shGYS-transduced cells.
3878
Human Molecular Genetics, 2008, Vol. 17, No. 24
Figure 1. Short hairpin RNA (shRNA) design and lentiviral constructs. (A) shRNA sequences and their position from the start codon. (B) Representation of the
shRNA expressed from lentiviral constructs. (C) Schematic representation of lentiviral constructs driving shRNA expression. WPRE, woodchuck hepatitis posttranscriptional regulatory element; LTR, long terminal repeat; DU3, self-inactivated (SIN) lentiviral vector obtained after partial deletion of the U3 region;
shRNAs were expressed under the control of the mammalian polymerase III H1 promoter (H1p); EGFP was expressed from the phosphoglycerate kinase
(PGK) housekeeping gene promoter.
Figure 2. Expression of glycogen synthase (GYS) and glycogenin (GYG)
messenger RNA (mRNA) in shRNA(short hairpin RNA)-transduced C2C12
cells. (A and B) GYG and GYS mRNA expression level in C2C12 myoblasts.
(C and D) GYG and GYS mRNA expression level in differentiated C2C12
myotubes. Results are standardized with b-actin mRNA and expressed as a
ratio versus non-transduced (NT) cells. LC and HC are cells containing low
or high proviral copy number, respectively. They are expressed as the
mean + SEM of three independent experiments (for each experiment, values
are the average of two samples). Differences between groups were examined
using the paired t-test. P , 0.05 compared with NT cells and EGFPtransduced C2C12 cells, #P , 0.05 HC versus LC.
No significant gene silencing was found in these controls (data
not shown).
C2C12 myoblasts expressing the most efficient shGYS2 and
shGYG2 were induced to differentiate into myotubes. Cell
Figure 3. Glycogen content in differentiated C2C12 myotubes transduced
with GYS (glycogen synthase) or GYG (glycogenin)-shRNA (short hairpin
RNA). Results were expressed as mean + SEM of four independent experiments. P , 0.05 versus non-transduced (NT) C2C12 myotubes, #P , 0.05
versus shGYG-transduced C2C12 myotubes.
fusion and myotube differentiation were observed in both
shRNA-transduced and untransduced cells (data not shown).
Both untransduced and transduced C2C12 myoblasts had low
levels of glycogen ranging from 0.012 + 0.009 to 0.017 +
0.009 mg glycogen/mg protein. During myotube differentiation, stimulation of GYS and GYG gene expression was
demonstrated by RT – PCR in untransduced cells (data not
shown). The silencing of GYS and GYG genes was maintained
during differentiation, with a silencing of mRNA up to 98%
with shGYG2 and 95% with shGYS2 (Fig. 2C and D). Glycogen content was quantified after 140 h of differentiation
(Fig. 3). In untransduced cells, glycogen increased up to
26-fold in differentiated myotubes in comparison with
Human Molecular Genetics, 2008, Vol. 17, No. 24
3879
Figure 4. Short hairpin RNA (shRNA)-mediated inhibition of glycogen synthase (GYS) and glycogenin (GYG) impairs glycogen storage in murine glycogen
storage disease type II (GSDII) primary myotubes. (A and B) Quantitative real-time RT –PCR analysis of GYG and GYS messenger RNA (mRNA) content in
shRNA-transduced myotubes. Results are mean + SEM of three independent experiments (values are the average of two samples). Values were standardized
with ß-actin mRNA and expressed as a ratio versus non-transduced (NT) GSDII myotubes. P , 0.05 versus NT Gaa2/2 myotubes. (C) Western blot analysis
of GYS expression in NT and shRNA-transduced Gaa2/2 myotubes. GAPDH, glyceraldehyde 3-phosphate dehydrogenase. (D) Glycogen content in NT and
shRNA-transduced Gaa2/2 myotubes. P , 0.05 versus NT Gaa2/2 cells, #P , 0.05 versus Gaaþ/þ myotubes.
myoblasts, reaching 0.46 mg glycogen/mg protein. Glycogen
was decreased by 50% in LC shGYS2-transduced myotubes
as compared with untransduced myotubes (0.2 mg glycogen/
mg protein), while glycogen was almost undetectable in HC
shGYS2-transduced myotubes. Surprisingly, no effect on glycogen levels was observed in shGYG2-transduced cells
(0.55 mg/mg protein). These results confirmed that GYS is a
limiting step during in vitro glycogen biosynthesis in murine
C2C12 myogenic cells.
ShRNA-mediated inhibition of GYS and GYG results in
decreased lysosomal glycogen accumulation in primary
myoblasts and myotubes from GSDII mice
We then investigated whether shGYS2 and shGYG2 expression
could reverse the lysosomal glycogen accumulation in myoblasts
from GSDII mice (Gaa2/2 ). Satellite cells from normal and
GSDII mice were isolated as described previously (27). Myogenic cells were transduced with shGYS2 and shGYG2-lentiviral
vectors at m.o.i. 10 and 100 and EGFPþ cells were isolated by
FACS. GYS and GYG mRNA levels were analyzed by quantitative RT–PCR in EGFPþ Gaa2/2 primary myoblasts and differentiated myotubes. As in C2C12 cells, GYS expression was
stimulated during myotube differentiation of untransduced
Gaa2/2 cells (Supplementary Material, Fig. S8A–C). Following
transduction with shGYS2 and shGYG2-expressing vectors, the
level of GYS or GYG mRNA was significantly reduced in
Gaa2/2 primary myoblasts (Supplementary Material, Fig. S9A
and B) and myotubes (98 and 99% decreased for shGYS2 HC
and shGYG2 HC, respectively) (Fig. 4A and B). The degree of
inhibition was also dependent on the integrated proviral copy
number in primary cells (1.3 versus 6.3 viral copy/cell for LC
versus HC shGYS2, and 1.3 versus 6.0 viral copy/cell for LC
versus HC shGYG2, respectively). The decrease in GYS
mRNA was accompanied by a reduced expression of the GYS
protein (87 and 48% for shGYS2 HC and shGYS2 LC, respectively), as compared with non-transduced myotubes (Fig. 4C).
In shRNA-transduced cells, the silencing efficiency was maintained during differentiation.
As lysosomal glycogen storage is a hallmark of GSDII, the
effect of shGYS2 and shGYG2 expression on the intracellular
glycogen level was investigated in differentiated myotubes.
Gaa2/2 myotubes contained a significantly higher glycogen
content as compared with Gaaþ/þ after a 140 h-differentiation
course (1.4 mg glycogen/mg protein versus 0.7 mg glycogen/
mg protein for Gaa2/2 versus Gaaþ/þ , respectively; P , 0.05).
A significant reduction in glycogen storage (below that seen
in Gaaþ/þ ) was observed in both HC and LC shGYS and
shGYG-expressing cells in differentiated Gaa2/2 myotubes
(0.12 mg glycogen/mg protein and 0.4 mg glycogen/mg
protein for HC shGYS2 and HC shGYG2, respectively)
(Fig. 4D). These results suggest that the inhibition of glycogen
synthesis can prevent glycogen accumulation in GAA-deficient
cells.
3880
Human Molecular Genetics, 2008, Vol. 17, No. 24
Figure 5. Correction of lysosomal enlargement in shRNA (short hairpin RNA)-transduced glycogen storage disease type II (GSDII) myotubes. (A –F) Representative confocal images of fixed myotubes immunostained for lysosomal-associated membrane protein 1 (LAMP1 – late endosomal/lysosomal marker):
normal (B); non-transduced (NT) GSDII (A); low-copy (C) or high-copy (D) shGYG-transduced GSDII myotubes; and low-copy (E) or high-copy (F)
shGYS-transduced GSDII myotubes. (G– L) The size (expressed in mm2, X-axis) and frequency (% of total, Y-axis) of lysosomes were analyzed in normal
(H), NT GSDII (G), low copy (I) or high copy (J) shGYG-transduced GSDII myotubes and low copy (K) or high copy (L) shGYS-transduced GSDII myotubes
using Metamorph software.
Enlargement of lysosomes is reversed in shGYS
and shGYG-expressing cells
AAV.shGYS2 reduces muscle glycogen storage
in a murine model of GSDII
The enlargement and rupture of the lysosomal/endosomal
compartment (due to excessive glycogen storage in lysosomes) along with autophagic buildup, are thought to be the
main pathogenic mechanisms responsible for muscle damage
in GSDII. Therefore, the appearance of the lysosomes was
evaluated in shGYS2 and shGYG2-transduced GSDII myotubes by confocal microscopy analysis after immunostaining
with LAMP1 (lysosomal-associated membrane protein 1), a
late endosomal/lysosomal marker (Fig. 5). The size of lysosomes in GSDII myotubes was significantly increased as compared with wild-type (Fig. 5A and G, and B and H). The mean
area of the lysosomes was 0.48 mm2 + 0.04 versus
0.21 mm2 + 0.02 with a maximum area of 18.6 mm2 versus
3.2 mm2 for Gaa2/2 versus Gaaþ/þ . Periodic acid Schiff
(PAS) staining of GSDII myotubes demonstrated that these
enlarged lysosomes are overloaded with glycogen (data not
shown). Normalization of the lysosome size was observed in
Gaa2/2 cells expressing HC and LC shGYS2 (Fig. 5E and
K, and F and L). The maximum lysosomal area was 5.9 and
4.1 in Gaa2/2 myotubes expressing LC and HC shGYS2.
LC shGYG2 expression was less efficient at reducing the lysosomal hypertrophy (maximum area: 11.7) (Fig. 5C and I), but
HC shGYG2 was quite effective (the maximum lysosomal
area was 3.5) (Fig. 5D and J).
shGYS2 expression was then tested in vivo and the impact of
GYS inhibition on muscle glycogen storage was evaluated in
GSDII mice. High muscle tropism recombinant AAV-1
vector co-expressing both EGFP and shGYS2 (AAV.shGYS2)
was developed (Fig. 6A). It was directly injected into gastrocnemius muscle of newborn GSDII mice in order to evaluate
the in vivo efficacy of the shRNA-mediated therapeutic
approach, using non-injected contralateral limbs as Controls.
Three weeks after injection, mice were sacrificed and
gastrocnemius muscles were harvested for GYS mRNA and
glycogen storage analysis. EGFP microscopic visualization
showed that transduction efficiency was 100%, even
though expression in individual muscle fibers was heterogeneous (Fig. 6B). The quantification of GYS mRNA revealed
a 50% decrease in injected muscle as compared with noninjected contralateral muscle (Fig. 6C).
In GSDII mice, muscle glycogen was increased after 2
weeks and it was easily quantified after 3 weeks (1.2 mg/mg
of protein in GSDII muscle versus 0.24 mg/mg of protein in
wild-type muscle). A 62% decrease in glycogen accumulation
was observed 4 weeks post-injection in AAV.shGYS2 injected
muscle as compared with the contralateral muscle (0.46 mg/
mg of protein), indicating that GYS inhibition can prevent
in vivo glycogen accumulation (Fig. 7A).
Human Molecular Genetics, 2008, Vol. 17, No. 24
Figure 6. Adeno-associated virus (AAV2/1) vector construction and muscle
transduction efficiency in glycogen storage disease type II (GSDII) mice.
(A) Schematic representation of the AAV vector construct. LITR, left inverted
terminal repeat; RITR, right inverted terminal repeat; shGYS2 was expressed
from the human Pol III (polymerase III) promoter H1; the cytomegalovirus
(CMV) early promoter drives the EGFP expression; pA, polyadenylation
signal. (B) EGFP expression on a cross-section of non-injected (left panel)
and shGYS2-AAV2/1 injected (right panel) GSDII gastrocnemius muscle.
(C) GYS mRNA (glycogen synthase messenger RNA) expression in noninjected and shGYS2-AAV2/1 injected GSDII gastrocnemius muscle.
Results are mean + SEM of five mice. Analyses were performed 4 weeks
after muscle injection.
As expected, PAS staining of untreated gastrocnemius
muscle from GSDII mice demonstrated massive glycogen
storage (Fig. 7C and D). Consistent with the biochemical
data, a significant reduction in PAS-positive material was
observed on sections from the AAV.shGYS2 injected
muscle. Some myofibers had nearly normal appearance
(Fig. 7C). The number and size of PAS-positive vacuoles
were also greatly reduced in AAV.shGYS2 injected muscle
as compared with contralateral muscle (Fig. 7B).
DISCUSSION
Pompe disease or glycogenosis type II is an inherited metabolic disorder characterized by a lysosomal a-glucosidase
deficiency resulting in massive glycogen storage in muscle
cells. We modulated the degree of muscle glycogen accumulation by reducing the expression of the major enzymes
involved in its biosynthesis and evaluated the benefits of this
approach on the reversal of pathology in GSDII. Initial experiments demonstrated that differentiation of C2C12 as well as
normal and GSDII primary myoblasts into multinucleate myotubes resulted in a significant increase of GYS and GYG
mRNA content and subsequently a stimulation of glycogen
synthesis. These results suggest that glycogen synthesis
3881
could be modulated by reducing the abundance of GYS and
GYG mRNA in the cell. Therefore, an RNA interferencebased strategy was used to decrease the expression of these
genes. We have demonstrated that lentiviral vectors expressing shRNA targeted to murine GYS and GYG mRNA
sequences, result in efficient and specific gene silencing in
transduced C2C12 cells as well as in GSDII primary myoblasts. Furthermore, in the diseased cells this manipulation
resulted in a significant reduction in glycogen synthesis.
These data confirm that GYS and GYG are key enzymes of
glycogen biosynthesis and that their inhibition is associated
with severe glycogen deprivation.
However, it is important to note that glycogen synthesis is
affected more by the inhibition of GYS gene expression than
by the inhibition of GYG, especially in C2C12 cells. This
finding could be explained by the recycling of pre-existing
GYG, since even a limited amount of the initiator molecule
could be sufficient to start new glycogen synthesis (28,29).
Mature glycogen particles in mammalian muscle have a molecular weight between 103 and 104 kDa and can be visualized
by electron microscopy (29). Even a small amount of GYG
protein could store a large amount of carbohydrate. Previous
reports have also speculated on the presence of alternative
primers for glycogen initiation (30 – 32). In C2C12 cells, the
initiator would either be a oligosaccharide containing a-1,4
or a-1,6 glucosyl linkages or a glucosylated protein. In
Gaa2/2 , where lysosomal glycogen degradation is impaired,
the de novo synthesis of glycogen requires new GYG proteins.
In shGYG-transduced cells, the low abundance of GYG proteins due to the degradation of GYG mRNA could be a limiting step in glycogen synthesis. Two different pathways have
been hypothesized for glycogen formation: in a normal state,
the pre-existing GYG or proglycogen molecules (small glycogen granules with low carbohydrate content) could be recycled
whereas a de novo formation of the initiator is required during
stress or supplementary needs (30,31). These data and ours
suggest that glycogen storage could exist even with a low
GYG level and that GYS is the best candidate to modulate glycogen synthesis and the level of glycogen.
The efficiency of the shRNAs used in this study was first
assessed by the reduction in the normal amount of glycogen
in C2C12. More importantly, shRNAs were efficient in the
reversal of the pathological accumulation of glycogen in
murine GSDII myotubes. A similar effect on glycogen
accumulation was shown in vivo after the injection of
AAV2/1-shGYS in the gastrocnemius of Gaa2/2 mice. The
area of glycogen granules was reduced by shRNA injection
without altering the muscle fibrils, which suggests that inhibition of GYS could reduce the pathological enlargement of
the lysosome/endosome compartment. GYS and GYG are in
charge of glycogen synthesis in the cytoplasm, which contains
two spatially distinct pools of glycogen, i.e. cytosolic and
vacuolar (lysosomal and autophagosomal). The mechanism
of glycogen uptake from the cytoplasm to the lysosome is
still unknown, but strong evidence suggests that a feedback
mechanism links the vacuolar and cytoplasmic glycogen
(33,34). Consistent with this notion, the reduction in cytoplasmic glycogen synthesis, as shown in our experiments, is
followed by a decline in the lysosomal glycogen content in
GSDII myoblasts.
3882
Human Molecular Genetics, 2008, Vol. 17, No. 24
Figure 7. Partial metabolic correction of lysosomal glycogen storage after AAV.shGYS2 vector injection in the gastrocnemius of Gaa2/2 mice. (A) Glycogen
content in wild-type (Gaaþ/þ ), non-injected and AAV.shGYS2(AAV-1 vector co-expressing both EGFP and shGYS2)-injected Gaa2/2 gastrocnemius. P ,
0.05 versus wild-type muscle, #P , 0.05 versus non-injected GSDII muscle. Results are mean + SEM of four to six mice. (B) The size (expressed in mm2,
X-axis) and frequency (% of total, Y-axis) of periodic acid Schiff (PAS)-positive granules were analyzed in myofibers of injected (gray) and non-injected
(black) Gaa2/2 muscle cross-sections using Metamorph software. Granules of ,24 mm2 were not considered because a similar distribution was found
between injected and non-injected Gaa2/2 muscle. (C) Cross and (D) longitudinal sections of Gaaþ/þ , non-injected Gaa2/2 and AAV.shGYS2-injected
Gaa2/2 gastrocnemius were analyzed for glycogen storage after PAS staining.
ERT was successfully used in treating infantile forms of
Pompe disease whereas a moderate effect was observed in
late-onset forms. The resistance of muscle fibers to ERT is
mainly due to the low density of CI-MPR in skeletal muscle
compared with cardiac muscle, and to the altered trafficking
of the recombinant enzyme along the endocytic pathway
(10,35). Previous studies demonstrated that fast-twitch type
II myofibers [gastrocnemius, tibialis anterior (TA)] are resistant to ERT as compared with slow-twitch type I (soleus) (10 –
12). It has been shown that GSDII myoblasts contain a subset
of late endosome/lysosomes with the acidification defect; in
addition, the levels of proteins (CI-MPR, clathrin, AP-2
complex, TfR) involved in receptor-mediated endocytosis
and trafficking of lysosomal enzymes are reduced in muscle
cells (10,35). Although a high dosage of recombinant GAA
enzyme is administered to GSDII patients during ERT, only
a fraction of the enzyme is targeted to muscle lysosomes;
80% is mistargeted and ends up in the liver (10). Therapeutic doses of recombinant human acid alpha glucosidase
(rhGAA) are significantly higher than those used for therapy
of other lysosomal storage diseases. For example, in Gaucher
disease glucocerebrosidase is administered at a dosage of
1 mg/kg while in Pompe disease rhGAA is administered at a
dosage of 20 mg/kg. Furthermore, a humoral immune response
was described after ERT in both Gaa2/2 mice and GSDII
patients, especially in CRIM negative subjects. Raben et al.
(10) showed that GSDII mice treated by ERT developed
signs of anaphylaxis (such as labored breathing and collapse)
and none of them survived beyond the seventh injection. The
induction of immune tolerance to ERT for Pompe disease is
currently under intensive investigation (36,37). Strategies
aimed to decrease the therapeutic dosage of the recombinant
enzyme could contribute to the safety of the treatment by reducing the immune response.
Overexpression of the muscle GYS gene in GSDII mice
leads to the accumulation of abnormal polysaccharides and
to a severe early-onset muscle wasting disorder (38). The
knockout of muscle GYS is lethal in 90% of null pups
which die soon after birth due to impaired cardiac function
(39). In the surviving 10% of GYS null mice, GYS and glycogen are undetectable in cardiac and skeletal muscle. It is
widely accepted that muscle is an important site for glucose
disposal and one might expect that, in the absence of muscle
glycogen, glucose clearance would be impaired. Surprisingly,
despite impaired glucose uptake into skeletal muscle, glucose
disposal is either normal or improved in muscle of GYS null
mice (40). GYS-deficient mice present with normal exercise
capacity suggesting that at least in mice, muscle glycogen is
not essential for exercise (41). However, in humans, mutations
in the GYS gene resulting in decreased muscle GYS activity
have been implicated in certain diabetic populations (42).
Recently, a homozygous mutation was described in the
muscle GYS gene in children with severe cardiomyopathy
and exercise intolerance (43). Taken together, these results
indicate that a complete inactivation of the GYS enzyme in
muscle can lead to unexpected adverse effects. In view of a
potential clinical application of GYS modulation for GSDII,
one should be aware that it is important to reduce glycogen
storage without inducing major disturbances in glucose or
energetic metabolism.
The aim of this study was to demonstrate the feasibility of
an SRT in Pompe disease using a gene transfer approach.
Human Molecular Genetics, 2008, Vol. 17, No. 24
Further development of a hydrosoluble pharmacological agent
allowing partial inactivation of GYS enzyme needs to be
explored. There are several advantages associated with this
approach: (i) an orally available drug could be used, allowing
for a non-invasive treatment of the disease, (ii) as small molecules are non-immunogenic, host immune responses will not
be induced, (iii) such a molecule does not need to be taken by
the CI-MPR-mediated pathway, which is impaired in GSDII.
An additional therapy that does not rely on CI-MPR-mediated
delivery of the recombinant GAA may decrease lysosomal
glycogen load, improve the lysosomal function, and possibly
even facilitate the trafficking of the therapeutic enzyme to
the lysosomes. Combined therapies may be necessary for
other lysosomal diseases as well. For example, a combination
of ERT (imiglucerase) with SRT (miglustat) has already
demonstrated its efficacy in type 3 Gaucher patients by
improving the neurological symptoms (44). In the murine
model of Sandhoff disease (another lysosomal disorder affecting brain), a combination of SRT with bone-marrow transplantation enhanced the survival of the animals (45). In
glycogenosis type II, involvement of the central nervous
system has recently been confirmed in the murine model of
the disease (46). It has been well established that recombinant
GAA does not cross the blood–brain barrier. However, the
small molecules (utilized in SRT) do cross the blood–brain
barrier and may alleviate glycogen accumulation in the
central nervous system; thus a combination of these therapeutic
approaches may prove extremely beneficial. In conclusion,
the approach used in this paper may become a significant
component of the comprehensive therapy for GSDII.
MATERIALS AND METHODS
Lentiviral vector construction and production
Different lentiviral vectors co-expressing EGFP and shRNAs,
respectively, under the control of the human PGK promoter
and the polymerase III (Pol III) H1 promoter were constructed.
Briefly, three shRNA sequences targeting the muscular GYS
mRNA (shGYS) (NM 030678) and two shRNA targeting the
GYG mRNA (shGYG) (NM 013755) were designed and
cloned downstream the H1 promoter using Sal1-HindIII
restriction enzymes (Fig. 1A). A Xho1-Nhe1 fragment containing the H1 promoter and shRNA was introduced in the
30 U3 LTR region of the SIN-U3-PGK-EGFP-WPRE (woodchuck hepatitis post-transcriptional regulatory element)
vector (kindly provided by Dr P. Charneau, Institut Pasteur,
Paris, France) in order to obtain a dual shRNA expression cassette from the integrated provirus. Lentiviral vectors were produced by a triple transient transfection (26). Human kidney
293T cells were plated at a density of 5 106 per 10 cm
tissue culture plates. The following day, they were transfected
with the pCMVD8.91 packaging construct (10 ml), the VSV-G
pMD.G envelope plasmid (3 mg), and the specific constructs
(10 mg), using the calcium phosphate precipitation method.
Thirty-six hours after the transfection, viral supernatants
were collected, passed through 0.22 mm filters, centrifuged
at 30 000g (90 min, 48C) and immediately cryopreserved at
2808C. Viral titers were determined by transducing C2C12
cells with serial dilutions of viral supernatants. EGFP
3883
expression was quantified 7 days after transfection by flow
cytometry analysis.
Primary culture and differentiation of myoblasts
TA muscle was removed from 4 –6-week-old Gaa2/2 and
wild-type mice. Myoblasts were isolated as described by
Ohanna et al. (27). Briefly, TA was digested for 1 h with a protease from Streptomyces griseus (Sigma-Aldrich, Saint Louis,
MO, USA), followed by filtration through 100 mm nylon
gauze. Cells were then plated at a low density in 12-well
gelatin-coated (0.02%) plates, containing Ham F12/DMEM
medium (v/v) supplemented with 20% fetal calf serum,
5 mM L-glutamine, 100 IU/ml penicillin, 100 mg streptomycin
(Invitrogen, Carlsbad, CA, USA) and Ultroser G 2% (Pall Corporation, East Hills, NY, USA). Five days after, wells containing myoblasts were pooled and seeded in 6-well gelatin-coated
plates. Medium was changed every 2 days and cells were trypsinized and seeded every 4 days at a density of 2 – 5 104
cells/flask (75 cm2). Myotube differentiation was induced by
culturing myoblasts at a density of 3 105 cells/well in
DMEM containing 2% horse serum (differentiation medium)
in plates pre-coated with Matrigel (BD Biosciences,
Bedford, MA, USA). Cells were maintained in this medium
for the duration of the experiment.
Lentiviral transduction of C2C12 and primary myoblasts
Cells were transduced in 6-well plates with each viral supernatant with an estimated m.o.i. of 2 and 50 (C2C12 cells) or
10 and 100 (GSDII myoblasts), in order to select cells respectively expressing HC or LC number of shRNA. EGFP-positive
cells were isolated using a FACS, and cell populations expressing either high or low level of shRNA products were isolated
and expanded.
The average integrated proviral copy number was determined
by real-time quantitative PCR. Total deoxyribonucleic acid
(DNA) was extracted using a DNA extraction kit (Qiagen,
Hilden, Germany) and amplified in the WPRE region of the
SIN-U3-PGK-WPRE-shRNA integrated sequence. Samples
were normalized to ß-actin DNA levels. Primers and PCR conditions were as follows: for WPRE, forward 50 -GCTATGTG
GATACGCTGC-30 , reverse 50 -GTTGCGGCAAACACATCA30 , annealing temperature 568C, 2 mM MgCl2, fragment size
160 bp; for ß-actin: forward: 50 -CAGGATTCCATACCTA
AGAG-30 , reverse 50 -CAGGATTCCATACCTAAGAG-30 ;
annealing temperature 568C, 2 mM MgCl2. The vector copy
number was calculated using standards containing 0.5, 1, 2, 3,
4 and 6 copies of the WPRE plasmid.
RT – PCR analysis of GYS and GYG expression
Total RNAs from EGFP-positive C2C12 cells, primary myoblasts and muscle tissues were extracted using the mRNA
mini kit (Qiagen) according to standard procedures. A quantitative real-time PCR assay was developed to detect GYS
and GYG mRNA levels; ß-actin was used as an endogenous
Control. PCR was carried out using a LightCyclerw CarouselBased System (Roche, Mannheim, Germany) and
SYBR Green PCR Master Mix Reagent (LightCyclerw RNA
3884
Human Molecular Genetics, 2008, Vol. 17, No. 24
Amplification Kit SYBR Green, Roche). Results are expressed
as a ratio versus negative Control (untransduced C2C12 or
Gaa2/2 primary myoblasts). Primers used were as follows:
for murine muscle GYS (249 bp), forward 50 -TATCGCTG
GCCGCTATGAGTT-30 and reverse 50 -CACTAAAAGGGA
TTCATAGAG-30 ; for murine GYG (204 bp): forward
50 -ACTCAGTATTCCAAATGTGTG-30 and reverse 50 -ATC
AAAACTACCTTGCTCAGA-30 ; for actin (210 bp), forward
50 -GGCATAGAGGTCTTTACGG-30 and reverse 50 -GTGGC
ATCCATGAAACTACAT-30 .
fixed in a 2% paraformaldehyde solution for 30 min at 378C,
followed by 1 h incubation in a blocking solution. Cells
were stained with primary rat anti-mouse LAMP1 antibody
(dilution 1/500; BD Pharmingen, San Diego, CA, USA), followed by AlexaFluor 680-labeled anti-rat secondary antibody
(dilution 1/5000; Invitrogen). Slides were mounted in Vectashield mounting medium (Vector laboratories, Burlingame,
CA, USA) and analyzed by confocal microscopy (Zeiss
LSM 510 META; Zeiss, Thornwood, NY, USA).
AAV construction and injection in Gaa2/2 mice
Western blot analysis
Protein lysates from cells were prepared in radioimmunoprecipitation assay buffer containing 50 mM Tris – HCl pH 7.4,
150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate and
0.1% SDS (sodium dodecyl sulfate). The following antibodies
were used: rabbit anti-mouse muscle GYS monoclonal antibody, dilution: 1/500 (Cell Signaling Technology, Danvers,
MA, USA), rabbit anti-mouse GAPDH (glyceraldehyde
3-phosphate dehydrogenase) monoclonal antibody, dilution:
1/3000 (Abcam, Cambridge, MA, USA), and goat anti-rabbit
horseradish peroxidase monoclonal antibody, dilution:
1/3000 (BioRad, Hercules, CA, USA).
Determination of glycogen concentration
Biochemical measurement of glycogen content was performed
on cultured cells. Hundred microliter of acetate buffer (0.2 M,
pH 4.5) was added to frozen cells on dry ice and the mix was
then heated to 908C for 15 min and homogenized. For glycogen measurement in muscle, heart or liver, tissues were
removed from mice and immediately frozen. They were then
homogenized at 2808C in acetate buffer (0.2 M, pH 4.5).
The cellular or tissue extracts were incubated at 548C for
1 h in the presence or absence of Aspergillus niger
amylo-a-1,4-a-1,6 glucosidase (5 U/ml; Roche, Mannheim,
Germany) which converts glycogen to glucose. Samples
were centrifuged and glucose level was determined in the
supernatant using Glucose Assay Reagent (Sigma-Aldrich)
according to the manufacturer’s instructions.
Histochemical analysis of muscle sections
Glycogen storage was determined by PAS staining. Muscle
tissues were frozen and 10 mm cryosections were kept at
2808C. Sections were hydrated in deionized water for 5 min
followed by incubation in 0.5% periodic acid solution at
room temperature for 5 min. They were then rinsed under
tap water and incubated in Schiff reagent for 15 min at room
temperature. They were finally dehydrated, coverslipped in a
xylene-based mounting media (Eukitt; Sigma), and analyzed
by light microscopy (Nikon Eclipse E 600; Nikon, Thornwood, NY, USA).
LAMP1 staining and confocal microscopy analysis
Normal and GSDII myoblasts were plated in Permanox
chamber slides (33 000 cells/cm2) for 24 h and induced to
differentiate into myotubes for 8 days. Myotubes were then
Recombinant AAV2/1 vectors co-expressing EGFP and shGYS2
respectively from the PGK promoter and the human H1 promoter were produced by the Laboratoire de Thérapie Génique
(Inserm U649, Nantes, France). AAV vectors were diluted in
phosphate buffer saline (PBS) and injected (6.5 106 UI/kg)
into the gastrocnemius (left leg) of 2-day-old Gaa2/2 mice
(3). PBS alone was injected into the contralateral gastrocnemius
(right leg) to serve as Control. Mice were sacrificed 4 weeks
after injection and biochemical and histological analyses were
performed on injected and non-injected muscles.
Metamorph analysis
Size of lysosomes and PAS-positive areas were determined by
Metamorph analysis. Representative fields were photographed
with a digital camera on a Jeol 1200 confocal microscope
(JEOL, Peabody, MA, USA) and with a DXM 1200 camera
on a Nikon Eclipse 600 light microscope.
Statistical analysis
All the data are presented as means + standard error of the
mean (SEM). One-way ANOVA (analysis of variance) were
performed on each group. P-value ,0.05 was considered as
statistically significant.
SUPPLEMENTARY MATERIAL
Supplementary Material is available at HMG Online.
FUNDING
This work was supported by INSERM and the Association
Vaincre les Maladies Lysosomales (VML). E.R. was supported by post-doctoral fellowships from VML and the
Association Française contre les Myopathies (AFM). G.D.
was supported by doctoral fellowship from Genzyme
(France) and AFM.
ACKNOWLEDGEMENTS
We thank Dr Patrice Petit for critical reading of the manuscript. We also thank the Vector Core Facility of the University Hospital of Nantes supported by the Association Française
contre les Myopathies for providing the AAV vectors.
Conflict of Interest statement. None declared.
Human Molecular Genetics, 2008, Vol. 17, No. 24
REFERENCES
1. Reuser, A.J., Kroos, M., Oude Elferink, R.P. and Tager, J.M. (1985)
Defects in synthesis, phosphorylation, and maturation of acid
alpha-glucosidase in glycogenosis type II. J. Biol. Chem., 260,
8336– 8341.
2. Kishnani, P.S., Steiner, R.D., Bali, D., Berger, K., Byrne, B.J., Case, L.E.,
Crowley, J.F., Downs, S., Howell, R.R., Kravitz, R.M. et al. (2006)
Pompe disease diagnosis and management guideline. Genet. Med., 8,
267–288.
3. Raben, N., Nagaraju, K., Lee, E., Kessler, P., Byrne, B., Lee, L., LaMarca,
M., King, C., Ward, J., Sauer, B. et al. (1998) Targeted disruption of the
acid alpha-glucosidase gene in mice causes an illness with critical features
of both infantile and adult human glycogen storage disease type II. J. Biol.
Chem., 7, 53–62.
4. Bijvoet, A.G., van de Kamp, E.H., Kroos, M.A., Ding, J.H., Yang, B.Z.,
Visser, P., Bakker, C.E., Verbeet, M.P., Oostra, B.A., Reuser, A.J. and van
der Ploeg, A.T. (1998) Generalized glycogen storage and cardiomegaly in
a knockout mouse model of Pompe disease. Hum. Mol. Genet., 7, 53– 62.
5. Klinge, L., Straub, V., Neudorf, U. and Voit, T. (2005) Enzyme
replacement therapy in classical infantile Pompe disease: results of a
ten-month follow-up study. Neuropediatrics, 36, 6 –11.
6. Van den Hout, J.M., Reuser, A.J., de Klerk, J.B., Arts, W.F., Smeitink,
J.A. and Van der Ploeg, A.T. (2001) Enzyme therapy for Pompe disease
with recombinant human alpha-glucosidase from rabbit milk. J. Inherit.
Metab. Dis., 24, 266 –274.
7. Amalfitano, A., Bengur, A.R., Morse, R.P., Majure, J.M., Case, L.E.,
Veerling, D.L., Mackey, J., Kishnani, P., Smith, W., McVie-Wylie, A.
et al. (2001) Recombinant human acid alpha-glucosidase enzyme therapy
for infantile glycogen storage disease type II: results of a phase I/II
clinical trial. Genet. Med., 3, 132–138.
8. Koeberl, D.D., Kishnani, P.S. and Chen, Y.T. (2007) Glycogen storage
disease types I and II: treatment updates. J. Inherit. Metab. Dis., 30,
159–164.
9. Reuser, A.J., Van Den Hout, H., Bijvoet, A.G., Kroos, M.A., Verbeet,
M.P. and Van Der Ploeg, A.T. (2002) Enzyme therapy for Pompe disease:
from science to industrial enterprise. Eur. J. Pediatr., 161 (Suppl. 1),
S106– S111.
10. Raben, N., Danon, M., Gilbert, A.L., Dwivedi, S., Collins, B., Thurberg,
B.L., Mattaliano, R.J., Nagaraju, K. and Plotz, P.H. (2003) Enzyme
replacement therapy in the mouse model of Pompe disease. Mol. Genet.
Metab., 80, 159– 169.
11. Raben, N., Jatkar, T., Lee, A., Lu, N., Dwivedi, S., Nagaraju, K. and Plotz,
P.H. (2002) Glycogen stored in skeletal but not in cardiac muscle in acid
alpha-glucosidase mutant (Pompe) mice is highly resistant to
transgene-encoded human enzyme. Mol. Ther., 6, 601–608.
12. Raben, N., Fukuda, T., Gilbert, A.L., de Jong, D., Thurberg, B.L.,
Mattaliano, R.J., Meikle, P., Hopwood, J.J., Nagashima, K., Nagaraju, K.
et al. (2005) Replacing acid alpha-glucosidase in Pompe disease:
recombinant and transgenic enzymes are equipotent, but neither
completely clears glycogen from type II muscle fibers. Mol. Ther., 11,
48– 56.
13. Drost, M.R., Schaart, G., van Dijk, P., van Capelle, C.I., van der Vusse,
G.J., Delhaas, T., van der Ploeg, A.T. and Reuser, A.J. (2008) Both type 1
and type 2a muscle fibers can respond to enzyme therapy in Pompe
disease. Muscle Nerve, 37, 251–255.
14. Sun, B., Zhang, H., Franco, L.M., Brown, T., Bird, A., Schneider, A. and
Koeberl, D.D. (2005) Correction of glycogen storage disease type II by an
adeno-associated virus vector containing a muscle-specific promoter. Mol.
Ther., 11, 889– 898.
15. Sun, B., Zhang, H., Franco, L.M., Young, S.P., Schneider, A., Bird, A.,
Amalfitano, A., Chen, Y.T. and Koeberl, D.D. (2005) Efficacy of an
adeno-associated virus 8-pseudotyped vector in glycogen storage disease
type II. Mol. Ther., 11, 57– 65.
16. Ziegler, R.J., Bercury, S.D., Fidler, J., Zhao, M.A., Foley, J., Taksir, T.V.,
Ryan, S., Hodges, B.L., Scheule, R.K., Shihabuddin, L.S. et al. (2008)
Ability of adeno-associated virus serotype 8-mediated hepatic expression
of acid alpha-glucosidase to correct the biochemical and motor function
deficits of presymptomatic and symptomatic Pompe mice. Hum. Gene
Ther., 19, 609– 621.
17. Xu, F., Ding, E., Liao, S.X., Migone, F., Dai, J., Schneider, A., Serra, D.,
Chen, Y.T. and Amalfitano, A. (2004) Improved efficacy of gene therapy
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
3885
approaches for Pompe disease using a new, immune-deficient GSD-II
mouse model. Gene Ther., 11, 1590–1598.
Fukuda, T., Ewan, L., Bauer, M., Mattaliano, R.J., Zaal, K., Ralston, E.,
Plotz, P.H. and Raben, N. (2005) Dysfunction of endocytic and
autophagic pathways in a lysosomal storage disease. Ann. Neurol., 52,
1824–1835.
Elstein, D., Dweck, A., Attias, D., Hadas-Halpern, I., Zevin, S., Altarescu,
G., Aerts, J.F., van Weely, S. and Zimran, A. (2007) Oral maintenance
clinical trial with miglustat for type I Gaucher disease: switch from or
combination with intravenous enzyme replacement. Blood, 110,
2296–2311.
Cox, T.M., Aerts, J.M., Andria, G., Beck, M., Belmatoug, N., Bembi, B.,
Chertkoff, R., Vom Dahl, S., Elstein, D., Erikson, A. et al. (2003) The role
of the iminosugar N-butyldeoxynojirimycin (miglustat) in the
management of type I (non-neuronopathic) Gaucher disease: a position
statement. J. Inherit. Metab. Dis., 26, 513–526.
Patterson, M.C., Vecchio, D., Prady, H., Abel, L. and Wraith, J.E. (2007)
Miglustat for treatment of Niemann-Pick C disease: a randomised
controlled study. Lancet Neurol., 6, 765– 772.
Yeaman, S.J., Armstrong, J.L., Bonavaud, S.M., Poinasamy, D.,
Pickersgill, L. and Halse, R. (2001) Regulation of glycogen synthesis in
human muscle cells. Biochem. Soc. Trans., 29, 537–541.
Lomako, J., Lomako, W.M. and Whelan, W.J. (2004) Glycogenin: the
primer for mammalian and yeast glycogen synthesis. Biochim. Biophys.
Acta, 1673, 45– 55.
Shulman, R.G., Bloch, G. and Rothman, D.L. (1995) In vivo regulation of
muscle glycogen synthase and the control of glycogen synthesis. Proc.
Natl Acad. Sci. USA, 92, 8535– 8542.
Katz, A. (2006) Glycogenin, proglycogen, and glycogen biogenesis:
what’s the story? Am. J. Physiol. Endocrinol. Metab., 290, E757–E758.
Richard, E., Robert, E., Cario-Andre, M., Ged, C., Geronimi, F., Gerson,
S.L., de Verneuil, H. and Moreau-Gaudry, F. (2004) Hematopoietic stem
cell gene therapy of murine protoporphyria by
methylguanine-DNA-methyltransferase-mediated in vivo drug selection.
Gene Ther., 11, 1638–1647.
Ohanna, M., Sobering, A.K., Lapointe, T., Lorenzo, L., Praud, C.,
Petroulakis, E., Sonenberg, N., Kelly, P.A., Sotiropoulos, A. and Pende, M.
(2005) Atrophy of S6K1(2/2) skeletal muscle cells reveals distinct
mTOR effectors for cell cycle and size control. Nat. Cell Biol., 7, 286 –294.
Shearer, J., Wilson, R.J., Battram, D.S., Richter, E.A., Robinson, D.L.,
Bakovic, M. and Graham, T.E. (2005) Increases in glycogenin and
glycogenin mRNA accompany glycogen resynthesis in human skeletal
muscle. Am. J. Physiol. Endocrinol. Metab., 289, E508– E514.
Skurat, A.V., Lim, S.S. and Roach, P.J. (1997) Glycogen biogenesis in rat
1 fibroblasts expressing rabbit muscle glycogenin. Eur. J. Biochem., 245,
147– 155.
Ugalde, J.E., Parodi, A.J. and Ugalde, R.A. (2003) De novo synthesis of
bacterial glycogen: agrobacterium tumefaciens glycogen synthase is
involved in glucan initiation and elongation. Proc. Natl Acad. Sci. USA,
100, 10659– 10663.
Meezan, E., Meezan, E., Meezan, J., Manzella, S. and Roden, L. (1997)
Alkylglycosides as artificial primers for glycogen biosynthesis. Cell Mol.
Biol., 43, 369 –381.
Mu, J., Cheng, C. and Roach, P.J. (1996) Initiation of glycogen synthesis in
yeast. Requirement of multiple tyrosine residues for function of the
self-glucosylating Glg proteins in vivo. J. Biol. Chem., 271, 26554–26560.
Kotoulas, O.B. and Phillips, M.J. (1971) Fine structural aspects of the
mobilization of hepatic glycogen. I. Acceleration of glycogen breakdown.
Am. J. Pathol., 63, 1 –22.
Kotoulas, O.B., Ho, J., Adachi, F., Weigensberg, B.I. and Phillips, M.J.
(1971) Fine structural aspects of the mobilization of hepatic glycogen. II.
Inhibition of glycogen breakdown. Am. J. Pathol., 63, 23– 36.
Fukuda, T., Roberts, A., Ahearn, M., Zaal, K., Ralston, E., Plotz, P.H. and
Raben, N. (2006) Autophagy and lysosomes in Pompe disease.
Autophagy, 2, 318– 320.
Joseph, A., Munroe, K., Housman, M., Garman, R. and Richards, S.
(2008) Immune tolerance induction to enzyme-replacement therapy by
co-administration of short-term, low-dose methotrexate in a murine
Pompe disease model. Clin. Exp. Immunol., 152, 138–146.
Sun, B., Bird, A., Young, S.P., Kishnani, P.S., Chen, Y.T. and Koeberl,
D.D. (2007) Enhanced response to enzyme replacement therapy in Pompe
disease after the induction of immune tolerance. Am. J. Hum. Genet., 81,
1042–1049.
3886
Human Molecular Genetics, 2008, Vol. 17, No. 24
38. Raben, N., Danon, M., Lu, N., Lee, E., Shliselfeld, L., Skurat, A.V.,
Roach, P.J., Lawrence, J.C. Jr, Musumeci, O., Shanske, S. et al. (2001)
Surprises of genetic engineering: a possible model of polyglucosan body
disease. Neurology, 56, 1739–1745.
39. Pederson, B.A., Chen, H., Schroeder, J.M., Shou, W., DePaoli-Roach,
A.A. and Roach, P.J. (2004) Abnormal cardiac development in the
absence of heart glycogen. Mol. Cell Biol., 24, 7179– 7187.
40. Pederson, B.A., Cope, C.R., Schroeder, J.M., Smith, M.W., Irimia, J.M.,
Thurberg, B.L., DePaoli-Roach, A.A. and Roach, P.J. (2005) Exercise
capacity of mice genetically lacking muscle glycogen synthase: in mice,
muscle glycogen is not essential for exercise. J. Biol. Chem., 280,
17260–17365.
41. Pederson, B.A., Schroeder, J.M., Parker, G.E., Smith, M.W.,
DePaoli-Roach, A.A. and Roach, P.J. (2005) Glucose metabolism in mice
lacking muscle glycogen synthase. Diabetes, 54, 3466– 3473.
42. Groop, L. and Orho-Melander, M. (2008) New insights into impaired
muscle glycogen synthesis. PLoS Med., 5, e25.
43. Kollberg, G., Tulinius, M., Gilljam, T., Ostman-Smith, I., Forsander, G.,
Jotorp, P., Oldfors, A. and Holme, E. (2007) Cardiomyopathy and
exercise intolerance in muscle glycogen storage disease. N. Engl. J. Med.,
357, 1507– 1514.
44. Capablo, J.L., Franco, R., de Cabezón, A.S., Alfonso, P., Pocovi, M. and
Giraldo, P. (2007) Neurologic improvement in a type 3 Gaucher disease
patient treated with imiglucerase/miglustat combination. Epilepsia, 48,
1406– 1408.
45. Jeyakumar, M., Norflus, F., Tifft, C.J., Cortina-Borja, M., Butters, T.D.,
Proia, R.L., Perry, V.H., Dwek, R.A. and Platt, F.M. (2001) Enhanced
survival in Sandhoff disease mice receiving a combination of substrate
deprivation therapy and bone marrow transplantation. Blood, 97,
327–329.
46. Sidman, R.L., Taksir, T., Fidler, J., Zhao, M., Dodge, J.C., Passini, M.A.,
Raben, N., Thurberg, B.L., Cheng, S.H. and Shihabuddin, L.S. (2008)
Temporal neuropathologic and behavioral phenotype of 6neo/6neo Pompe
disease mice. J. Neuropathol. Exp. Neurol., 67, 3– 818.