Address correspondence and reprint requests to Or Kakhlon, Department of Neurology, Hadassah-Hebrew University Medical Center, Ein Kerem, Jerusalem 91120 Israel. E-mail: email@example.com
Uncontrolled elongation of glycogen chains, not adequately balanced by their branching, leads to the formation of an insoluble, presumably neurotoxic, form of glycogen called polyglucosan. To test the suspected pathogenicity of polyglucosans in neurological glycogenoses, we have modeled the typical glycogenosis Adult Polyglucosan Body Disease (APBD) by suppressing glycogen branching enzyme 1 (GBE1, EC 22.214.171.124) expression using lentiviruses harboring short hairpin RNA (shRNA). GBE1 suppression in embryonic cortical neurons led to polyglucosan accumulation and associated apoptosis, which were reversible by rapamycin or starvation treatments. Further analysis revealed that rapamycin and starvation led to phosphorylation and inactivation of glycogen synthase (GS, EC 126.96.36.199), dephosphorylated and activated in the GBE1-suppressed neurons. These protective effects of rapamycin and starvation were reversed by overexpression of phosphorylation site mutant GS only if its glycogen binding site was intact. While rapamycin and starvation induce autophagy, autophagic maturation was not required for their corrective effects, which prevailed even if autophagic flux was inhibited by vinblastine. Furthermore, polyglucosans were not observed in any compartment along the autophagic pathway. Our data suggest that glycogen branching enzyme repression in glycogenoses can cause pathogenic polyglucosan buildup, which might be corrected by GS inhibition.
Knockdown of glycogen branching enzyme in neurons led to accumulation of an insoluble form of glycogen called polyglucosan, to apoptosis and to activation of glycogen synthase. These effects were reversed by glycogen synthase inhibition through starvation and rapamycin treatments, suggesting a potential therapeutic value of glycogen synthase inhibition for treating glycogen storage disorders.
sodium dodecyl sulfate polyacrylamide gel electrophoresis
short helical RNA
The brain has extremely high energy demand, but relatively restricted energy storage capacity. Moreover, although neurons have the highest energy requirement in the brain, they do not store or turn over glycogen because of glycogen synthase (GS) inactivation and glycogen phosphorylase deficiency (Benarroch 2010). Rather than storing their own fuel, neurons rely on astrocytes, the brain's energy sensors. Astrocytes can respond to insulin by increasing glycogen stores (Heni et al. 2011) and to neuronal neurotransmitters, secreted during energy deficit, by furnishing neurons with energy substrates. These substrates are produced by glycogen mobilization, or by glycolysis, both down-modulated in neurons (Belanger et al. 2011). It is still debated which energy substrate is delivered to neurons by astrocytes. Recent computational studies maintain that both lactate, the long-accepted form of energy transported to neurons (Belanger et al. 2011; Choi et al. 2012), and glucose (DiNuzzo et al. 2010; Chander and Chakravarthy 2012), which also dictates neuronal lactate uptake (Calvetti and Somersalo 2012), fuel neurons.
As a consequence of their inactivated glycogen metabolism, the only form of glycogen that neurons synthesize is the abnormal polyglucosan, associated with various neurological disorders. Polyglucosan is amylopectin-like, poorly branched, and insoluble. It is the major constituent of insoluble deposits called polyglucosan bodies (PBs), also containing damaging ubiquitinated proteins and long-lived advanced glycated end-products (Cavanagh 1999). Polyglucosan forms whenever glycogen branching activity cannot keep pace with glycogen synthetase activity. Thus, in APBD (Lossos et al. 1991), a variant of glycogen branching enzyme 1 (GBE1) deficiency (glycogen storage disease type IV (Chen and Bucherell 1995), PBs accumulate because of GBE1 deficiency (Magoulas et al. 2012). On the other hand, in Lafora disease (LD) (Andrade et al. 2007), excessive glycogen phosphorylation (Striano et al. 2008; Turnbull et al. 2010), or insufficient GS degradation (Vilchez et al. 2007), leads to excessive glycogen elongation and accumulation of PBs called Lafora bodies. PB accumulation causes neuronal-selective apoptosis which spares astrocytes (Cavanagh 1999; Magistretti and Allaman 2007; Vilchez et al. 2007). Despite the neurotoxic potential of PBs, neurons cannot dispose of them, even though they are endowed with robust rapamycin-inducible autophagy and microtubular transport, which can clear insoluble inclusion bodies.
Our study is aimed at elucidating why polyglucosan accumulation is pathogenic in neurological glycogenoses (represented by APBD) and which measures might circumvent polyglucosan neurotoxicity. Using transduction of neurons with lentiviruses harboring shRNA directed against GBE1, we generated a GBE1-knocked down neuronal model of APBD, which featured increased GS activity, polyglucosan accumulation and associated apoptosis. We then attempted to clear the PBs and reverse apoptosis by rapamycin and starvation, able to phosphorylate and inactivate GS. We further showed that these protective effects depend on GS phosphorylation/inactivation and not on autophagy, which is also stimulated by rapamycin and starvation and which can degrade glycogen (Kotoulas et al. 2004). Our results suggest that future therapeutic endeavors in neurological glycogenoses should be based on down-modulation of the ratio between glycogen synthesis and branching via manipulation of GS or GBE1 activities, or on direct polyglucosan degradation.
All reagents were from Sigma-Aldrich (Rehovot, Israel), except for radionuclides and vinblastine, which were from Perkin-Elmer (Waltham, MA, USA) and Teva (Netanya, Israel), respectively. 5-Aminoimidazole-4-carboxamide ribonucleotide (AICAR) was a kind gift from Dr. Ann Saada-Reisch (Hadassah-Hebrew University Medical Center). Periodic acid-Schiff reagent kit was from Merck (Darmstadt, Germany).
Monoclonal antibodies to GBE1 were from Abnova (Taipei, Taiwan), those to tubulins and GS from Abcam (Cambridge, UK), those to LC3 from Nanotools (Teningen, Germany), those to phospho-GS (Ser641) from Cell Signaling Technology (Danvers, MA, USA) and those to c-myc (9E10) from the hybridoma bank. The monoclonal antibody to glycogen was prepared as previously described (Nakamura-Tsuruta et al. 2012) and used for immunofluorescence (Wang et al. 2013). Secondary antibodies, conjugated either to Dylight 549 (for immunofluorescence and flow cytometry) or to peroxidase (for immunoblotting), were from KPL (Gaithersburg, MD, USA). Antibodies to LC3 and c-myc used for immunofluorescence and flow cytometry were conjugated to the FluoProbes 647H (ex. 653 nm, em. 674 nm) and Phycoerythrin (PE)-Cy5 (ex. 488 nm, em. 679 nm), respectively, using the Lightning-Link conjugation kit from Innova Biosciences (Cambridge, UK). All antibodies were used according to manufacturers’ instructions.
Primary neurons from E18 rat cortex were obtained from Brainbits (Springfield, IL, USA) and cultured according to the manufacturer's instructions. The content of these cultures was practically neuronal (96%), as confirmed by flow cytometry using antibodies against the respective neuron and glia markers β-3 tubulin and glial fibrillary acidic protein (Figure S1). Neuronal transfection was done by electroporation using Microporator MP-100 (NanoEn Tek, Seoul, Korea) and took place after lentiviral transduction.
Lymphoblasts and fibroblasts from a 65 years old female APBD patient were used with the understanding and written consent of the patient and following the ARRIVE guidelines. Derivation of these cells was approved by the Hadassah Institutional Review Board according to The Code of Ethics of the World Medical Association (Declaration of Helsinki), printed in the British Medical Journal (18 July 1964).
Suppression of GBE1 expression
After 5 days in culture, neurons were transduced with SMARTvector 2.0 lentiviral particles (Thermo Scientific, Lafayette, CO, USA) bearing shRNA against GBE1, or a non-targeting control. Transduction was done at a titer of 40 multiplicities of infection in half the volume of growth medium. 12 h post-transduction, neurons were washed twice with phosphate-buffered saline and the medium was replaced with half preconditioned/half fresh medium. Neurons were assayed 4 days after lentiviral transduction. Lentiviral constructs included a turboGFP reporter gene to enable detection of transduced cells.
Real-time Reverse Transcription PCR
Total RNA was purified from cells using the PureLink kit (Invitrogen, Carlsbad, CA, USA) and reverse-transcribed using the High-Capacity cDNA Reverse Transcription kit (Applied Biosystems, Foster City, CA, USA). GBE1 mRNA content, normalized to the endogenous control Hprt, was determined by real-time RT-PCR using the pre-designed TaqMan Gene Expression system and the StepOnePlus thermocycler and analyzer (Applied Biosystems). Site directed mutagenesis was performed on mouse GS1 cDNA using the Quick Change kit (Stratagene, Santa Clara, CA, USA).
ATP was determined by a luciferase-based assay (Sigma-Aldrich). Luminescence was read with a 10-second integration time in the luminescence mode of DTX 880 Multimode Detector (Beckman Coulter, Indianapolis, IN, USA). Phosphofructokinase activity was measured using BioVision's (Milpitas, CA, USA) specialized kit.
GBE1 and GS activity assays
GBE1 activity was assayed based on (Lossos et al. 1991). This assay is based on the difference in the rate of incorporation of 14C-glucose-1-phosphate to endogenous glycogen between samples where glycogen branching enzyme (GBE) is denatured by boiling and untreated samples. The lower GBE activity, the lower this difference. GS activity was determined based on the rate of 14C-UDP-glucose incorporation to exogenous glycogen (Akman et al. 2011).
For indirect immunofluorescence, neurons were fixed with 3% paraformaldehyde, permeabilized with 0.2% Triton X-100, stained with antibodies, and analyzed by confocal microscopy using a Zeiss LSM 710 microscope (Jena, Germany) with a 60X/1.35 NA PlanApochromat oil-immersion lens. In Fig. 5a, neurons were permeabilized by saponin before fixation, using 5 min exposure to 0.5% saponin in 80 mM Piperanzine-N-N′-bis[2-ethane]sulfonic acid pH 6.8, 1 mM CaCl2, 5 mM EGTA. LC3-positive structures were quantified by the Image-Pro software (Media Cybernetics, Bethesda, MD). Electron microscopy was performed as described in (Zahavi et al., 2011), with uranyl-ethanol staining of the thin sections.
Flow cytometry and quantification of apoptosis
Apoptosis was quantified using the FC 500 flow cytometer (Beckman Coulter). In Figs 3-5 and 7 and Figure S2, neurons were co-stained with Invitrogen's Annexin-V-PE, and 7-Amino-actinomycin D (7-AAD), for detection of apoptotic and necrotic cells, respectively. Cells (10 000) were measured using the 488 nm excitation line and PE (FL-2) and 7-AAD (FL-4) emission detectors. FL-1/FL-2 compensation [all cells analyzed were green fluorescence protein (GFP) positives] and FL-2/FL-4 compensation were set during acquisition using single dyes controls and confirmed post-acquisition by the FC 500 QuickComp feature. In Fig. 8, neurons transduced and transfected with c-myc-tagged GS constructs were permeabilized and co-stained with Annexin-V-PE, PE-Cy5-conjugated anti-myc antibody (FL3) and the TO-PRO-3 DNA dye (FL4, Invitrogen). Gated FL1 (GFP) and FL3 (c-myc tagged GS) positive cells were measured using the 488 nm laser for the GFP (FL1), PE (FL2), and PE-Cy5 (FL3) emission detectors and the 635 nm laser for the TO-PRO-3 (FL4) emission detector. Compensations between all combinations of FL-1 to FL-4 emissions were set prior to gating and confirmed as above. For microscopic detection of apoptosis, cells were stained with Annexin V-Pacific Blue and 7-AAD.
The significance of differences (p < 0.05) between pairs of treatments and among multiple treatments was confirmed by Student's t-test and one-way anova with Tukey post hoc test, respectively.
Generation and characterization of a neuronal model for APBD
To reproduce polyglucosan deposition, the hallmark of APBD, and ensuing neurotoxicity in cultured neurons, we generated a neuronal GBE1 knockdown model. This model is based on the observation that GBE1 levels (Assereto et al. 2007) and activities (Lossos et al. 1991) are reduced in APBD, which is, in fact, a clinical variant of glycogen storage disease type IV. GBE1 suppression in our model was confirmed at the mRNA and enzyme activity levels (Fig. 1a), and at the protein level by immunoblotting (Fig. 1b) and indirect immunofluorescence (Fig. 1c). This reduction is similar in degree to that observed in lymphoblasts from an APBD patient homozygous for the GBE1 Y329S mutation (Fig. 1b).
We next tested the ability of our model to reproduce polyglucosan accumulation and neurotoxicity. As our indirect immunofluorescence data show, no glycogen is detected in control neurons transduced with lentiviral particles containing non-targeting shRNA (Fig. 2a). In contrast, GBE1-knocked down neurons stained for glycogen show punctate cytosolic structures, similar to those observed in neurons in which GS activity had been induced (Vilchez et al. 2007, Fig. 2b). These structures are purportedly polyglucosans, being larger than soluble glycogen and showing typical fibrillar structure in the EM (Fig. 6d). Unfortunately, we were not able to extract enough glycogen to further confirm the polyglucosan nature of these deposits by iodine adsorption. Interestingly, neurons transduced with relatively lower level of shGBE1 lentiviruses (as documented by the lower GFP expression) also had lower levels of polyglucosan and lesser cell rounding, typical of the early stages of apoptosis (Fig. 2c). This observation suggests that polyglucosan accumulation is responsible for neuronal damage in our model.
GBE1-knocked down neurons also manifested apoptotic blebbing (Fig. 3a) and stained positive for Annexin V, a marker of apoptosis, and the DNA stain 7-AAD, a marker of necrosis, similar to neurons treated with the apoptosis-inducer sodium nitroprusside (Fig. 3b). Confocal staining of fixed neurons with Annexin V and 7-AAD (Fig. 3c) confirmed that increased apoptotic indices (scored in Fig. 3d) were not simply caused by increased susceptibility to flow cytometry sample preparation. These results suggest that polyglucosan accumulation in APBD may be pathogenic because it triggers apoptosis.
The mTOR inhibitors rapamycin and starvation can reverse polyglucosan accumulation and damage
Having established and validated a neuronal model for APBD, we used it to test possible therapeutic avenues. As shown in Fig. 2, GBE1 knockdown led to accumulation of the polyglucosan form of glycogen, while glycogen was not observed at all in control neurons. This finding suggested that de novo glycogen synthesis by GS had to be activated by the GBE1 knockdown and that the adverse effects of this knockdown might be corrected by GS inhibition. We therefore opted to test GS inhibition as a strategy for correcting GBE knockdown-mediated damage. Using shRNA as a tool for GS inhibition was unsuccessful because the yield of viable neurons where both GBE1 and GS were knocked down was too low for performing experiments. Therefore, as an alternative tool for inhibiting GS, we capitalized on inhibition of mTOR, a major switch of cell anabolism which can activate GS via phosphorylative inhibition of glycogen synthase kinase3ß. Treatment of GBE1-knocked down neurons with the mTOR inhibitors rapamycin or starvation reduced polyglucosan accumulation (Fig. 4a) and counteracted its associated apoptosis (Fig. 4b and c, cf Fig. 3b).
However, a caveat of this approach is its lack of specificity: As an anabolic switch, mTOR does not only activate GS and glycogenesis but also inhibits catabolic autophagy. Inhibition of mTOR, an established inhibitor of autophagy, is therefore also expected to induce autophagy. Indeed both rapamycin and starvation treatments induced autophagy, as shown by the accumulation of autophagosomes, detected by their marker LC3 (Fig. 4a). We therefore had to test the possibility that autophagy induction is epiphenomenal to GS inhibition as a mechanism by which mTOR inhibitors restored GBE knockdown damage.
Autophagy, induced by mTOR inhibitors, is dispensable for correcting GBE knockdown damage
Theoretically, autophagy could have led to sequestration and auto-lysosomal digestion of polyglucosan by acid maltase and thus mediate the observed beneficial effects of the mTOR inhibitors. Using saponin permeabilization, we showed that the LC3 positive structures accumulated by rapamycin and starvation treatments (Fig. 4a) are autophagic vesicles [membrane associated, rather than cytosolic, entities (Fig. 5a)]. The increased autophagic flux, induced by rapamycin and starvation, presents maximal number of autophagosomes to the autophagic maturation process in which these autophagosomes mature into degradative autolysosomes. Hence, LC3 positive autophagosomes accumulate, as autolysosomal degradation is unable to keep pace with the overwhelming autophagic flux (Fig. 5a and (Boland et al. 2008)). If autolysosomal degradation was responsible for polyglucosan clearance, rapamycin and starvation treatments would have maximized it.
To test the dependence of polyglucosan purging on autophagic maturation and autolysosomal degradation, we blocked both processes pharmacologically (double knockdown of GBE1 and autophagy genes is unfeasible, as explained above) using vinblastine, an established inhibitor of autophagic maturation (Kochl et al. 2006). The vinblastine block was manifested by further increase in the number and intensity of LC3 positive vesicles (Fig. 5a) and in the ratio of lipidated to cytosolic LC3 (LC3II/LC3I, Fig. 5b), which is inversely correlated with autophagic flux. We confirmed vinblastine's effect by showing that it reproduced the block in autophagic flux induced by lysosomal protease inhibitors (Fig. 5b, cf lipidated LC3 (LC3 II) to cytosolic LC3 (LC3 I) ratio between ‘Rap+PI’ and ‘Rap+Vin’ lanes), and also blunted the sensitivity of rapamycin induced neurons to them (Fig. 5b, cf ‘Rap+Vin’ lane to ‘Rap+Vin+PI’ lane). Even though it blocked autophagic maturation, vinblastine did not reverse the rapamycin (Fig. 5c, upper panel), or starvation (Fig. 5c, lower panel)-mediated down-modulation of polyglucosan accumulation, or did it reverse their anti-apoptotic protection (Fig. 5d). Interestingly, in contrast to vinblastine, 3-MA, an inhibitor of autophagosome formation, did blunt the reversal of apoptosis by rapamycin and starvation (Figure S2a and b). However, we presume this reversal depended not on inhibition of autophagy, but on the 3-MA-mediated GS activation (Fig. 7a and (Das and Hollinger 2012)) through dephosphorylation (Figure S2c and (Das and Hollinger 2012)), or phosphofructokinase1 inhibition (Figure S2c), which would slightly increase glycogen-6-phosphate (G-6-P). This assumption is corroborated by the observation that, in contrast to 3-MA, vinblastine affected only autophagy and not GS phosphorylation (Fig. 5e). The data presented in Fig. 5 thus suggest that protection of GBE1-knocked down neurons by rapamycin or starvation was independent of autophagic maturation and autolysosomal degradation.
In support of this conclusion, polyglucosans in rapamycin-treated or starved GBE-knocked down neurons were not observed in autophagosomes (Fig. 4a) and autolysosomes (Fig. 6c and Figure S3a), or were they observed in organelles partaking in the autophagic pathway: multivesicular body (Fig. 6a and Figure S3a) and amphisomes (the fusion product of autophagosomes and multivesicular bodies, Fig. 6b and c, sup. Fig. 3b). Apparent exosomes, formed by exocytotic diversion of the amphisome internal vesicles, also did not contain polyglucosans in rapamycin-treated (Fig. 6c) or starved (Figure S3c) GBE-knocked down neurons.
In contrast to their rapamycin-treated and starved counterparts, the untreated, apoptotic GBE1-knocked down neurons did contain non-membrane-associated polyglucosans smaller than 200 nm (Fig. 6d). This finding suggests that neurons that are not in contact with other cell types, as they are in the brain, might succumb to cell death once polyglucosan deposits appear and before they are able to grow in size to the > 1 μm PBs observed in APBD. This would explain why the average size of glycogen deposits (see also Figs 2 and 4) was smaller than in APBD affected brains, where PBs can occlude axons.
Correction of GBE knockdown damage by mTOR inhibitors is mediated by GS inhibition
We next aimed at confirming that the mode of action of rapamycin and starvation was through GS inhibition, as also observed in HepG2 cells (Varma et al. 2008), and implemented in rescue of a Pompe disease mouse model (Ashe et al. 2010). We show that both rapamycin treatment and starvation reduced GS activity both in control and in GBE1-knocked down neurons (Fig. 7a). GS basal activity, up-modulated 4-fold in GBE1-knocked down neurons, was lowered to control level when GBE1-knocked down neurons were treated with rapamycin, or starved (Fig. 7a). This action of both treatments was 3-MA reversible, suggesting that GS is rendered constitutively active by 3-MA, as corroborated by the observation that 3-MA rendered GS activity refractory to G-6-P stimulation. None of the treatments affected G-6-P-stimulated GS activity, suggesting it overrode GS phosphorylation state, in accordance with (Bouskila et al. 2010). GS protein levels were not affected by any of the treatments as our immunoblotting data show (Fig. 7a). To confirm that rapamycin and starvation mediated their effect via GS repression, we treated GBE1-knocked down neurons with the GS inactivator (AICAR) (Bultot et al. 2012). Like rapamycin and starvation treatments, AICAR action in GBE1-knocked down neurons led to reduction in GS basal activity without affecting the G-6-P-stimulated one (Fig. 7a), induction of autophagy (Fig. 7b, also observed in Lee et al. 2010), clearing of polyglucosans (Fig. 7b), and rescue of apoptosis (Fig. 7c). These results further corroborate our main concept that induction of autophagy is epiphenomenal to GS inactivation as a neuroprotective strategy against polyglucosan-mediated damage. Our observation that only GS activity but not its levels where modified in our APBD neuronal model is in apparent contrast with the results of (Valles-Ortega et al. 2011) who showed that in a mouse model of another neurological glycogenosis, LD, total GS levels were actually increased, while its activity did not significantly change. This apparent discrepancy demonstrates that increase in polyglucosan levels on its own does not necessarily lead to increase and accumulation of glycogen binding proteins in insoluble PB, as shown by the persistence of cytosolic GS (alongside GS in PB) in GBE1-knocked down neurons (Fig. 7d). Accumulation of glycogen binding proteins in PB is expected; however, if their levels are independently increased, as might be the case for GS when its E3 ubiquitin ligase malin is knocked down (Valles-Ortega et al. 2011).
Having demonstrated that mTOR inhibitors exerted their action via GS inhibition, we set out to find a molecular mechanism for this activity. As Fig. 8a, shows, treating neurons with both rapamycin and starvation restored the inhibitory GS phosphorylation at Ser641 (Skurat and Roach 1995), dephosphorylated by GBE knockdown. In parallel, we reconfirmed, as in Fig. 7a, that rapamycin and starvation also lowered GS activity, up-modulated in GBE1-knocked down neurons, while none of the treatments affected total GS levels. This GS inhibition also coincided with restoration of ATP to control levels. Depletion of ATP as a kinase substrate, or as an AMP source for the GS inactivator AMP-activated protein kinase (AMPK) (Bultot et al. 2012), can possibly explain why GS was less phosphorylated and responsive to G-6-P activation in the GBE1-knocked down neurons, which undergo ATP-consuming apoptosis (Fig. 3). Further support for the notion that rapamycin and starvation acted via GS inhibition can be found in our observation that over-expression of a S641A, S645A, S649A, S653A, S657A phosphorylation sites-mutated and activated GS (Skurat and Roach 1995) overturned their ameliorating effects on polyglucosan accumulation and apoptosis (Fig. 8b). On the other hand, the beneficial effects of rapamycin and starvation were refractory to over-expression of a dual-site mutated GS where the quintuple phosphorylation mutations were supplemented by the D450A, R461A, F466A glycogen binding site mutations (Baskaran et al. 2011). These observations demonstrate that only an active (dephosphorylated) and glycogen binding GS can reverse the beneficial effects of rapamycin and starvation, which deducibly act by GS inhibition.
This study was aimed at providing a proof of principle for developing therapeutics against APBD and other glycogenoses resulting from PB accumulation. To test the pathomechanism of PB accumulation, we opted for a neuronal model where formation of polyglucosans can be induced and the consequences followed. This system consisted of naïve neurons where the GBE/GS activity ratio (inversely proportional to the rate of polyglucosan buildup) was down-modulated by transduction with lentiviruses harboring shRNA against GBE1. While the relatively low cell number of transduced neurons limits the range of experiments that can be carried out using this system (e.g., fractionation is not feasible), the system is conceptually preferred over neurons derived from GBE knockdown or knockout mice (Akman et al. 2011) where neuronal polyglucosans pre-exist and therefore the effects following their formation cannot be followed.
Fig. 7 describes the effect of GBE1 on GS activation. In GBE1-knocked down neurons, basal GS activity increased 3-4-fold and became refractory to G-6-P activation. The results of (Akman et al. 2011) in embryonic mouse muscle show a similar, albeit significantly less pronounced, trend on GS activity in GBE1 knockdown mice (1.4-fold increase in basal activity and change in G-6-P activity ratio from 0.27 in Gbe1+/+ to 0.6 in Gbe1−/−, as compared to a change from 0.27 in control to 1 in GBE1-knocked down neurons). In contrast, however, (Lamperti et al. 2009) showed a lack of GS activity in the heart and liver of a GBE1 deficient neonate, possibly attributable to low availability of free glucosyl ends in larger PBs. A possible explanation of GBE1 control over GS activity in relative values (the absolute values are expectedly lower in neurons) is that GBE1 suppression leads to ATP deficit (Fig. 8a), probably more pronounced in isolated neurons, which undergo ATP-consuming apoptosis (Fig. 3). This ATP deficit might inhibit kinases and AMPK and render GS both less phosphorylated (more active) and less responsive to G-6-P activation. Hence, the differences between the three studies might be explained in terms of different ATP availabilities. Another explanation is based on the rate of glycogen2 synthesis. Reduced rate of glycogen synthesis could theoretically increase glycolytic flux, generating more G-6-P and rendering GS less G-6-P responsive. Although we could not determine the rate of glycogen synthesis in neurons, we presume it would have been higher in GBE1-knocked down neurons, since they contained glycogen in contrast to control neurons. Therefore, we hypothesize that reduced rate of glycogen synthesis contributed to the decreased GS G-6-P responsiveness only in Gbe1−/− embryonic muscle (Akman et al. 2011) and not in GBE1-knocked down neurons. In general, the observation that GBE knockdown can increase GS activity, induce de novo formation of polyglucosans (Fig. 2) and therefore be rescued by GS inhibition is important therapeutically as GS inhibition is more feasible as a prospective treatment than restoration of aberrant GBE activity. However, therapeutic implementation of this principle is still far.
Our results show that GS inhibition reversed polyglucosan accumulation and ensuing apoptosis (Fig. 7). On the basis of these results, we propose a therapeutic strategy aimed at alleviating polyglucosan-mediated damage in neurological glycogenoses. This strategy is based either on increasing GBE/GS activity ratio, and thus arresting further buildup of polyglucosans, or, preferably, on polyglucosan degradation, which eliminates pre-existing polyglucosans. We observed pre-existing polyglucosans in an APBD patient skin fibroblasts grown in ketogenic medium, where no de novo polyglucosans could have formed. As Figure S4 shows, polyglucosan accumulation induced by glucose can be reduced by rapamycin, which could not remove pre-existing polyglucosans. These data are consistent with the data in Fig. 4 where rapamycin removed polyglucosans from GBE-knocked down neurons, when it was introduced in parallel to the shGBE lentivirus, so that it inhibited de novo synthesis of polyglucosans, by counteracting GS activation (Figs 7 and 8). Therefore, rapamycin and other GS inhibitors are only expected to ameliorate glycogenoses, or slow down their progress, rather than eradicate polyglucosans which underlie their pathogenesis. GS inhibition has already been shown effective in the treatment of a murine model of LD via prtotein targeting to glycogen knockout (Turnbull et al. 2011). This approach is also clinically practical since GS deficiency is relatively tolerable in both mice (Pederson et al. 2005; Douillard-Guilloux et al. 2010) and humans (Kollberg et al. 2007), especially if blood glucose is monitored closely to avoid hypoglycemia.
A complementary approach to GS inhibition, also aimed at increasing GBE/GS activity ratio, would be up-modulation of GBE1 activity. We currently employ high-throughput screening of small molecule libraries to identify molecules with acceptable pharmacokinetic and pharmacodynamic profiles and minimal side effects which reduce polyglucosan levels. These molecules might be novel GS inhibitors, or enhancers of endogenous GS inhibitors, such as AMPK. Alternatively, these molecules may be GBE1 stabilizers, or more potent drugs promoting polyglucosan degradation.
Our results (Figs 5 and 6) suggest that, while induced, autophagy was not responsible for the improvement of GBE knockdown damage mediated by GS inhibition. Nevertheless, it is still possible that glycogen-selective autophagy might be effective in removing polyglucosans and relieving their damage. Specifically, glycogen autophagy could be induced by laforin, associated with both polyglucosan [not soluble glycogen, (Chan et al. 2004; Tiberia et al. 2012)] and the ER, the initiation site for autophagosome biogenesis (Hayashi-Nishino et al. 2009). These two associations suggest that laforin may be a primer for selective glycogen autophagy, especially as it was also shown to induce cytoprotective autophagy (Aguado et al. 2010). We conjecture that glycogen autophagy might have a therapeutic potential in glycogenoses based on observations in non-neuronal cells, where autophagy was shown to sequester and metabolize glycogen (Kotoulas et al. 2004; Schiaffino et al. 2008). Thus, glycogen autophagy might benefit APBD and LD patients both by PB disposal and as energy source through glycogen degradation in astrocytes.
In conclusion, we propose that GS inhibition, GBE1 activation, polyglucosan degradation and possibly glycogen autophagy are all avenues worth pursuing for the treatment of APBD and other neurological glycogenoses.
This study was supported by a research grant from the APBD Research Foundation to which we are greatly indebted. O. K. and H. O. A. designed research; O. K., H. G., N. F. and Y. L. performed research; O. B., T. T., Y. L. and H. O. A. contributed vital new reagents and analytical tools; A. L. provided clinical advice and patient samples; O. K. and H. O. A. analyzed and interpreted data; O. K. performed statistical analysis; O. K., H. O. A. and S. D. wrote the manuscript. The authors declare no conflict of interests.