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Keywords:

  • Calcium;
  • cerebellum;
  • degeneration;
  • Niemann-Pick;
  • sphingomyelin

Abstract

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

We recently demonstrated that calcium homeostasis is altered in mouse models of two sphingolipid storage diseases, Gaucher and Sandhoff diseases, owing to modulation of the activities of a calcium-release channel (the ryanodine receptor) and of the sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) respectively, by the accumulating sphingolipids. We now demonstrate that calcium homeostasis is also altered in a mouse model of Niemann–Pick A disease, the acid sphingomyelinase (A-SMase)-deficient mouse (ASM–/–), with reduced rates of calcium uptake via SERCA in the cerebellum of 6–7-month-old mice. However, the mechanism responsible for defective calcium homeostasis is completely different from that observed in the other two disease models. Thus, levels of SERCA expression are significantly reduced in the ASM–/– cerebellum by 6–7 months of age, immediately before death of the mice, as are levels of the inositol 1,4,5-triphosphate receptor (IP3R), the major calcium-release channel in the cerebellum. Systematic analyses of the time course of loss of SERCA and IP3R expression revealed that loss of the IP3R preceeded that of SERCA, with essentially no IP3R remaining by 4 months of age, whereas SERCA was still present even after 6 months. Expression of zebrin II (aldolase C), a protein found in about half of the Purkinje cells in the adult mouse cerebellum, was essentially unchanged during development. We discuss possible pathological mechanisms related to calcium dysfunction that may cause Purkinje cell degeneration, and as a result, the onset of neuropathology in Niemann–Pick A disease.

Abbreviations used
A-SMase

acid sphingomyelinase

CAMKII

calcium calmodulin-dependent kinase II

CHOP

Ccaat/enhancer-binding protein homologous protein

C6-NBD-SM

N-{6-[(7-nitrobenzo-2-oxa-1,3-diazol-4-yl)amino]hexanoyl}-d-erythro-sphingosylphosphorylcholine

ER

endoplasmic reticulum

GRP78

Glucose Regulated Protein 78

IP3

inositol 1,4,5-trisphosphate

IP3R

inositol 1,4,5-triphosphate receptor

NPD-A

Niemann–Pick disease type A

PBS

phosphate-buffered saline

SERCA

sarco/endoplasmic reticulum Ca2+-ATPase

SM

sphingomyelin

SPC

sphingosylphosphorylcholine

Niemann–Pick A (NPD-A) and B diseases are caused by an inherited deficiency in lysosomal acid sphingomyelinase (A-SMase) (Schuchman and Desnick 2001). As in most lysosomal storage diseases, the biochemical and cellular pathways leading to cell dysfunction and disease pathology have not been elucidated (Futerman and van Meer 2004). Moreover, in two-thirds of the lysosomal storage diseases, severe neuropathology is observed but there is no obvious correlation between the type of storage material and neurological symptoms (Raas-Rothschild et al. 2004). In NPD-A, severe phenotypes are characterized by a progressive neurodegenerative course that leads to death in early infancy.

A mouse model of NPD-A has been generated by targeted disruption of the A-SMase gene (Horinouchi et al. 1995). These mice (ASM–/–) mimic the human disease phenotype inasmuch as they die at a young age (∼7–8 months), display ataxia and tremors, and show visceral symptoms similar to those observed in human patients (Schuchman and Desnick 2001).

We have previously shown defective neuronal Ca2+ homeostasis in mouse models of two other sphingolipid storage diseases, Sandhoff disease (Pelled et al. 2003) and Gaucher disease (Korkotian et al. 1999; Lloyd-Evans et al. 2003a), and in post-mortem human brain tissue from patients with Gaucher disease (Pelled et al. 2005); likewise, Ca2+ homeostasis has recently been shown to be altered in a mouse model of a GM1 gangliosidosis (Tessitore et al. 2004). In Sandhoff and Gaucher diseases, defective Ca2+ homeostasis was caused by interaction of the primary storage lipids, ganglioside GM2 and glucosylceramide respectively, with proteins involved in regulating Ca2+ homeostasis. Thus, glucosylceramide stimulated Ca2+ release via the ryanodine receptor, whereas GM2 inhibited the activity of the sarco/endoplasmic reticulum Ca2+-ATPase (SERCA).

To determine whether Ca2+ homeostasis is also altered in the ASM mouse, we have now analyzed rates of Ca2+ uptake and release in the cerebellum and cerebral cortex. Ca2+ homeostasis in the cerebellum was indeed significantly different from that in wild-type mice but, in contrast to Gaucher and Sandhoff diseases and to the GM1 gangliosidosis, was caused by reduced levels of expression of SERCA and of the inositol 1,4,5-triphosphate receptor (IP3R), rather than by a direct effect of the accumulating lipids on the proteins.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Materials

Goat polyclonal anti-SERCA2 (N-19), goat polyclonal anti-IP3R1 (C-20), goat polyclonal anti-aldolase C (zebrin II) (D-14), goat polyclonal anti-calbindin D28K (C-20), and biotin-conjugated and horseradish peroxidase-conjugated rabbit anti-goat secondary antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Texas Red-conjugated streptavidin was from Vector Laboratories (Burlingame, CA, USA). N-{6-[(7-nitrobenzo-2-oxa-1,3-diazol-4-yl)amino]hexanoyl}-d-erythro-sphingosylphosphorylcholine (C6-NBD-SM), ganglioside GM2, sphingomyelin (SM), sphingosylphosphorylcholine (SPC) and ceramide were from Matreya (Pleasant Gap, PA, USA). Antipyrylazo III, A23187, thapsigargin, palmitoyl co-enzyme A, inositol 1,4,5-trisphosphate (IP3), creatine phosphokinase, phosphocreatine, ATP, Sephadex G25 and other chemicals were from Sigma (St. Louis, MO, USA). Silica gel 60 TLC plates were from Merck (Darmstadt, Germany). All solvents were of analytical grade and purchased from Biolab (Jerusalem, Israel).

ASM colony

A colony of ASM mice (Horinouchi et al. 1995) was maintained in the Experimental Animal Center of the Weizmann Institute of Science as a heterozygous breeding colony. For the experiments in this study, wild-type (CBL56J/OLA; ASM+/+) mice were bred with each other, as were the knockout mice (ASM–/–), to obtain homozygous offspring. The genotype of the mice was determined by the PCR using genomic DNA extracted from mouse tails (Buccoliero et al. 2004b).

Human tissue samples

A human brain sample from one patient with NPD-A, as well as an age-matched control, was kindly provided by Raphael Schiffman, Developmental and Metabolic Neurology Branch, National Institute of Neurological Disorders and Strokes (NINDS), National Institutes of Health, Bethesda, USA. The NPD-A sample had been stored at − 80°C for several years.

Preparation of brain microsomes and spectrophotometric assay of Ca2+ homeostasis

Microsomes were prepared from 18 to 30 cortices or from ∼25–50 cerebella of ASM+/+ and ASM–/– mice, or from 8–11 g human brain. Mice were killed at various ages and their brains removed. Brains were separated into cerebral cortex and cerebellum, rapidly frozen in liquid N2, and stored at − 80°C. Microsomes were prepared, and Ca2+ uptake and release were measured spectrophotometrically using the Ca2+-sensitive dye, antipyrylazo III, as described previously (Lloyd-Evans et al. 2003a, 2003b; Pelled et al. 2003). The rate of Ca2+ uptake into microsomes was calculated by measuring the linear portion of the slope following Ca2+ addition. The amount of Ca2+ released from microsomes was expressed as a percentage of the total Ca2+ in the microsomes, which was obtained by summing Ca2+ taken up during the Ca2+-loading period together with endogenous Ca2+ from the microsomal preparation (measured separately after addition of a Ca2+ ionophore, A23187 (2 µm), with and without Ca2+ loading). The effect of thapsigargin (dissolved in absolute ethanol) on Ca2+ uptake, and of palmitoyl-CoA or IP3 on Ca2+ release, was tested after preloading the microsomes with 2 additions of 25 nmol CaCl2. In some experiments, GM2 (dissolved in absolute ethanol) and SM or SPC [dissolved either in dimethylsulfoxide, ethanol or ethanol/dodecane (98 : 2, v/v)] were added to microsomes 10–15 min before Ca2+ addition. The concentration of the solvents did not exceed 1% (v/v), and the solvents by themselves had no effect on Ca2+ uptake.

Fixation, sectioning and immunofluorescence

ASM+/+ and ASM–/– mice were anesthetized with sodium pentobarbital before transcardiac perfusion with phosphate-buffered saline (PBS) followed by 2.5% (w/v) paraformaldehyde in PBS. Brains were incubated for 72 h at 4°C in 15% sucrose/1.25% paraformaldehyde/PBS. Free-floating microtome sections were cut at 16 µm thickness. The sections were permeabilized in 1% Triton X-100 in 20% normal horse serum/PBS for 1 h at 25°C, and then incubated overnight at 4°C with either anti-SERCA2, anti-IP3R1, anti-calbindin or anti-zebrin II antibodies, diluted 1 : 50 in 2% normal horse serum/PBS. Sections were incubated with a biotinylated rabbit anti-goat secondary antibody (diluted 1 : 200) for 90 min at 25°C, and subsequently with Texas Red-conjugated streptavidin at a dilution of 1 : 100 in PBS for 30 min at 25°C. Images were acquired using a Nikon E600 fluorescence microscope with a Nikon DXM 1200F digital camera and analyzed using Nikon ACT-1 software (Tokyo, Japan).

Sodium dodecyl sulfate–polyacrylamide gel electrophoresis and western blotting

Sodium dodecyl sulfate–polyacrylamide gel electrophoresis and western blotting were performed using a 7% separating gel. After transfer to nitrocellulose membranes, the blot was incubated with blocking buffer [Tris-buffered saline containing 3% (w/v) non-fat dried milk, 1% (w/v) bovine serum albumin and 0.1% Tween-20] for 30 min and then incubated with either anti-SERCA2 antibody at a dilution of 1 : 500, anti-IP3R1 antibody at a 1 : 100 dilution, or anti-zebrin II antibody diluted 1 : 500, in Tris-buffered saline containing 1% (w/v) bovine serum albumin and 0.1% Tween-20. Following incubation with horseradish peroxidase-conjugated rabbit anti-goat secondary antibody at a dilution of 1 : 10 000 in blocking buffer for 1 h at 25°C, bound antibodies were detected using the SuperSignal chemiluminescent detection reagent (Pierce, Rockford, IL, USA). Protein levels were quantified by densitometry using MultiAnalyst software (Bio-Rad Laboratories, Hercules, CA, USA).

A-SMase activity

A-SMase activity was determined in human brain samples using C6-NBD-SM. Tissue homogenates (50 µg protein) were incubated for 1–2 h at 37°C with C6-NBD-SM (20 µm) in 100 µL 0.1 m sodium acetate buffer (pH 4.7) containing 0.5 mm EDTA. Lipids were extracted (Bligh and Dyer 1959) and NBD-SM and NBD-ceramide were quantified by densitometry using a Fluor-S MAX Multi-Imaging System (Bio-Rad, Hercules, CA, USA) and QuantityOne software (Bio-Rad).

Statistical analysis

Statistical analyses were performed using the one-tailed Student's t-test.

Results

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

To determine whether Ca2+ homeostasis is defective in the ASM mouse near the endpoint of disease progression, microsomes were prepared from the cortex or cerebellum of 5–7-month-old ASM–/– mice, a stage at which the mice show severe disease symptoms and die soon afterwards, and the rates of Ca2+ release and uptake were measured. No changes were observed in the rate of Ca2+ release using 30–50 µm palmitoyl CoA as a ryanodine receptor agonist (Lloyd-Evans et al. 2003a, 2003b) in either brain region (not shown). However, Ca2+ uptake via the SERCA was reduced by ∼60% in ASM–/– cerebellum compared with ASM+/+ cerebellum, but no changes were observed in the rate of Ca2+ uptake in cortical microsomes (Fig. 1). The maximal amount of Ca2+ that could be taken up into microsomes from ASM–/– cerebellum before induction of sustained and prolonged Ca2+ release was also significantly reduced. Thus, whereas approximately six additions of Ca2+ could be taken up in cerebellar microsomes from ASM+/+ mice before induction of sustained and prolonged Ca2+ release, only about four Ca2+ additions were necessary in ASM–/– cerebellum (Figs 2b and c). A small reduction in the number of Ca2+ additions required to induce this phenomenon was observed in microsomes from ASM–/– cortex (Fig. 2a). In addition, 75 µm thapsigargin, a specific SERCA inhibitor (Treiman et al. 1998), almost completely blocked Ca2+ uptake in ASM–/– cerebellar microsomes (Fig. 2e), but blocked Ca2+ uptake in ASM–/– cortical microsomes to a smaller extent (Fig. 2d). In contrast, 150 µm thapsigargin completely blocked Ca2+ uptake in wild-type and ASM–/– microsomes, whereas neither 150 µg/mL heparin (an inhibitor of the IP3R; Ghosh et al. 1988) or 5 mm oxalate (a Ca2+ chelator; Wells and Abercrombie 1998) had any effect (data not shown), demonstrating that the Ca2+ is taken up into a thapsigargin-inhibitable pool. Together, these results demonstrate defective Ca2+ homeostasis in the cerebellum, but not the cortex, of 5–7-month-old ASM–/– mice.

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Figure 1. Rate of Ca2+ uptake into microsomes from ASM mice. Microsomes prepared from 5–7-month-old mice were loaded with two sequential additions of 25 nmol Ca2+ (loads 1 and 2). The mean ± SD rate of Ca2+ uptake for each addition was measured in (a) cortical (n = 20–21 ) and (b) cerebellar (n = 11–13) microsomes; black bars represent the rate of uptake in ASM+/+ microsomes and open bars that in ASM–/– microsomes. *p < 0.001 versus AM+/+ (one-tailed Student's t-test). (c) A representative trace from cerebellar microsomes showing absorbance change (A710–A790) of antipyrylazo III with time; an increase in absorbance demonstrates an increase in free Ca2+ in the cuvette, and a decrease in absorbance signifies a decrease in free Ca2+ due to microsomal Ca2+ uptake by the SERCA.

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Figure 2. Characterization of Ca2+ uptake and release in cortical and cerebellar microsomes. (a) Cortical (n = 20–21) and (b) cerebellar (n = 7–9) microsomes from 5–7 month-old mice were loaded by sequential addition of 25 nmol Ca2+ until sustained Ca2+ release was induced. Values are mean ± SD. *p < 0.001 versus ASM+/+ (one-tailed Student's t-test). (c) Typical traces of absorbance change (A710–A790) of antipyrylazo III with time in cerebellar microsomes; in the upper trace (ASM+/+) sustained Ca2+ release is induced after six additions of 25 nmol Ca2+, whereas in the bottom trace (ASM–/–) sustained Ca2+ release is induced after four additions of 25 nmol Ca2+. (d) Cortical and (e) cerebellar microsomes were incubated with 75 µm thapsigargin, followed by one addition of 25 nmol Ca2+. Results are mean ± SD of 10–12 (cortex) or 6–9 (cerebellum) independent experiments. **p < 0.005 versus ASM+/+ (one-tailed Student's t-test).

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We next examined expression levels of SERCA2, the predominant SERCA isoform in brain (Villa et al. 1991; Wu et al. 1995). In our previous study on Sandhoff mice near the endstage of disease, reductions in Ca2+ uptake via SERCA were caused by a decrease in the Vmax of the SERCA induced by ganglioside GM2, with no change in SERCA expression (Pelled et al. 2003). Surprisingly, SERCA2 levels were dramatically reduced in cerebellar homogenates from 6-month-old ASM–/– mice, whereas no change was observed in the cortex (Fig. 3). To determine whether expression of other proteins involved in Ca2+ homeostasis might also be altered, we further analyzed levels of the IP3R1, the major Ca2+-release channel in cerebellum (Meldolesi 2002). The IP3R1 was barely detectable in 6–7-month-old ASM–/– cerebellar homogenates (Fig. 3). In addition, IP3 did not induce Ca2+ release in ASM–/– cerebellar microsomes at concentrations as high as 200 µm, although 30 µm IP3 induced significant Ca2+ release in ASM+/+ microsomes (data not shown).

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Figure 3. Western blot of SERCA2 and IP3R1 in ASM brain. Levels of SERCA2 and IP3R1 were examined by western blotting in cortical and cerebellar homogenates (50 µg protein) from 6-month-old mice, in duplicate homogenates prepared from different mice. One representative western blot from 5–6 similar experiments is shown.

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As the mechanism of altered Ca2+ homeostasis in the ASM mouse appeared primarily to involve a reduction in protein expression, at least at the endstage of disease, we next performed a systematic analysis of changes in SERCA and IP3R expression and function during disease progression. In 1-month-old mice, there was no difference in either IP3R or SERCA expression in ASM–/– cerebellum, but by 2 months a significant reduction was observed in IP3R expression with a concomitant decrease in the rate of IP3R-induced Ca2+ release (Fig. 4); interestingly, 2-month-old ASM–/– mice show initial signs of ataxia and tremor (Horinouchi et al. 1995). By 4 months, at which stage severe ataxia is observed (Horinouchi et al. 1995), the reduction in the rate of IP3R-induced Ca2+ release was much more pronounced and the IP3R was barely detectable. The reduction in SERCA levels, and in Ca2+ uptake via SERCA, was not significantly different until 4 months of age (Fig. 4).

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Figure 4. Time course of reduction in SERCA and IP3R expression and function. (a) IP3R1 and SERCA2 levels were examined by western blotting at the indicated ages (m, month) in cerebellar homogenates. The upper panel shows representative western blots and the lower panel quantification by densitometry of 8–9 blots from two different homogenates, expressed as a percentage of protein levels in ASM+/+ homogenates. Values are mean ± SD. *p < 0.05, **p < 0.001 (Student's t-test). (b) IP3R-induced Ca2+ release (left-hand panel) and SERCA-mediated Ca2+ uptake (right-hand panel) was measured in microsomes prepared from the same homogenates as used for western blotting. Values are mean ± SD. ▪, ASM+/+; ○, ASM–/–. *p < 0.005 versus ASM–/– (Student's t-test).

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We next examined, by immunohistochemical staining of cerebellar slices, which cell types, i.e. neurons or glia, were affected in the ASM–/– mice at 5 months of age. The IP3R1 was expressed at high levels in a specific cell layer of the cerebellum in ASM+/+ mice, but its levels were dramatically reduced in the ASM–/– mice (Fig. 5). This layer corresponded to the Purkinje cell layer, as it was labeled by an antibody against calbindin, a specific Purkinje cell marker (De Camilli et al. 1984); calbindin levels were also dramatically reduced in the ASM–/– mice (Fig. 5). We therefore conclude that the reduction in IP3R1 is due to loss of Purkinje cells in the cerebellum. Moreover, levels of SERCA2 expression were also significantly reduced in the same Purkinje cell layer (Fig. 6), where SERCA2 expression was particularly high in the cell bodies and dendrites, as was that of the IP3R1 (Fig. 6).

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Figure 5. Immunolocalization of the IP3R1 and calbindin in mouse cerebellum. The upper panels are from ASM+/+ mice and the lower panels from ASM–/– mice. The bar represents 500 µm. This panel is representative of 2–3 independent analyses.

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Figure 6. Immunolocalization of the IP3R1, SERCA2, calbindin and zebrin II in mouse cerebellum. The bar represents 50 µm. This panel is representative of 2–3 independent analyses.

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Previous studies have shown degeneration of a subset of Purkinje cells that stain negative for zebrin II (Sarna et al. 2001), a protein found in about half of the Purkinje cells in adult mouse cerebellum (Hawkes and Herrup 1995). Although there appeared to be a small reduction in zebrin II at all ages examined, zebrin II was still found at significant levels even after 6 months in ASM–/– cerebellum (Fig. 7), when no IP3R expression and only low levels of SERCA were detected (Fig. 3), consistent with the lack of change in zebrin II expression observed by immunofluorescence (Fig. 6). Thus, our data demonstrate that, in the cerebellum of ASM–/– mice, the reduction in IP3R expression precedes that of SERCA which precedes that of zebrin II.

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Figure 7. Time course of zebrin II expression. The western blot is representative of three independent experiments. m, month.

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We next determined whether a similar effect is observed in a human brain sample from a patient with NPD-A. In this sample, levels of A-SMase activity were ∼12% of those in a control sample (Fig. 8a) and SM levels were raised 3.7-fold (data not shown). The mean ± SD rate of Ca2+ uptake into the NPD-A microsomes was 0.012 ± 0.005 compared with 0.26 ± 0.09 nmol/s/mg protein in the control brain (Fig. 8b), and only two additions of Ca2+ were required to induce sustained Ca2+ release in the NPD-A brain compared with five in the control. Similar to findings in the ASM–/– mouse, levels of SERCA protein were significantly reduced in the NPD-A brain homogenate (Fig. 8c).

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Figure 8. Ca2+ uptake in human NPD-A brain. (a,b) A-SMase activity and rate of Ca2+ uptake in brain tissue from a patient with NPD-A compared with that in an age-matched control. Values are mean ± SD. (c) Levels of SERCA2 protein in homogenates (50 µg protein) from the same brain samples.

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In an attempt to determine the biochemical mechanism of degeneration of cerebellar neurons, we examined changes in lipid levels during mouse development. Levels of SM, the primary storage material, increased with age in both cortex and cerebellum (Fig. 9). However, unlike in Sandhoff and Gaucher disease models, in which the primary storage material is responsible for effects on the SERCA and the ryanodine receptor respectively, exogenously added SM had no effect on either IP3-induced Ca2+ release (Fig. 10) or SERCA-mediated Ca2+-uptake (data not shown). Its lyso-derivative, SPC, which also accumulates in NPD-A (Rodriguez-Lafrasse and Vanier 1999), induced a small but statistically significant decrease in IP3-induced Ca2+ release (Fig. 10), but at much higher levels than those found endogenously (see Discussion). In addition, ganglioside GM2, which also accumulates in patients with NPD-A (Brunngraber et al. 1973; Rodriguez-Lafrasse and Vanier 1999) and in an animal (cat) model of the disease (Wenger et al. 1980), accumulated in the cerebellum (mean ± SD 0.52 ± 0.12 nmol/mg protein) and in the cortex (0.72 ± 0.18 nmol/mg protein) of 6–7-month-old ASM mice, but at ∼10-fold lower levels than in the Sandhoff mouse (∼5–10 nmol/mg protein; Buccoliero et al. 2004a), and at levels lower than those that affect SERCA-mediated Ca2+ uptake (Pelled et al. 2003). We therefore conclude that a direct effect of either primary or secondary storage lipids on the proteins involved in regulating Ca2+ homeostasis in the cerebellum is unlikely to be responsible for the initiation of the pathological pathway resulting in neuronal degeneration.

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Figure 9. SM levels in ASM mice. SM levels were measured in (a) cortex and (b) cerebellum of ASM–/– (open bars) and wild-type (black bars) mice. Results are mean ± SD of 3–4 independent analyses. E17, embryonic day 17.

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Figure 10. Effect of exogenously added lipids on IP3-induced Ca2+-release. Lipids [100 µm, except for ganglioside GM2 (10 µm)] were added to cerebellar microsomes before addition of IP3, and Ca2+ release was measured. Results are mean ± SD of 4–9 independent analyses. *p < 0.001 versus control (Student's t-test). Cer, ceramide.

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Discussion

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

In the present study we demonstrated changes in Ca2+ homeostasis in the cerebellum of a mouse model of NPD-A. These changes can be explained by the degeneration of a specific population of cerebellar neurons enriched in the IP3R1 and SERCA2, and is consistent with the observed Purkinje cell loss in the cerebellum of ASM–/– mice (Sarna et al. 2001). Thus, NPD-A can be classified as a neurodegenerative disorder of the cerebellum (Sarna and Hawkes 2003). Moreover, the progressive dysfunction in regulation of intracellular Ca2+ homeostasis in the cerebellum correlates with the time course of onset of neurological symptoms observed in ASM–/– mice (Horinouchi et al. 1995).

A similar loss of IP3R1 and SERCA2 was reported in a mouse model of spinocerebellar ataxia type 1 (Lin et al. 2000). In this case, IP3R1 and SERCA2 mRNA levels were decreased by 2 weeks of age. These reductions were attributed to Purkinje cell degeneration, although no reduction was seen in eight other genes expressed at high levels in Purkinje neurons, even by 3 months of age, suggesting specificity of protein or neuronal loss. Previously, a reduction in Purkinje cell-specific Ca2+-binding proteins was shown by 6 weeks of age, the time of onset of neurological symptoms (Vig et al. 1998), and down-regulation of IP3R1 and SERCA3 in the cerebellum was observed before the onset of a neurological phenotype (Vig et al. 1998; Serra et al. 2004). Likewise, levels of other proteins involved in Ca2+ homeostasis and glutamate signaling in Purkinje neurons were also reduced.

As in NPD-A, the biochemical pathways involved in the pathological cascade causing spinocerebellar ataxia type 1 are not known. Likewise, how changes in these pathways affect neuronal viability is not known for either disease, but the similarity between NPD-A and spinocerebellar ataxia type 1, at least with respect to IP3R1 and SERCA2 expression, is striking. The IP3R1, the major endoplasmic reticulum (ER) Ca2+-release channel, is found in cerebellar Purkinje neurons at levels hundreds of times higher than in other cell types in the brain (Meldolesi 2002), and SERCA, which pumps Ca2+ back into the ER from the cytosol (Berridge et al. 2003), is also expressed at high levels in cerebellar Purkinje neurons (Villa et al. 1991; Wu et al. 1995). The abundance of these two proteins suggests their physiological importance in Purkinje cells, and a reduction in their levels would presumably have severe effects on Ca2+ homeostasis, which could lead to the dysfunction and degeneration of these neurons. The ER lumen also contains other proteins, such as calreticulin and calnexin that act to buffer luminal Ca2+ and interact with Ca2+ channels and pumps to modulate their function (Michalak et al. 2002); these proteins could conceivably be involved in the pathogenesis of NPD-A. As a result of changes in intracellular Ca2+ concentrations, important events in signal transduction pathways, such as activation of calcium-calmodulin-dependent kinase II (CAMKII) and protein kinase C cascades, and opening of ion channels at the plasma membrane, might be initiated (Berridge et al. 2003). High cytosolic Ca2+ concentrations can also lead to cell death owing to induction of oxidative stress (Mattson and Chan 2003).

Interestingly, a recent study has shown cell loss in a particular population of Purkinje neurons in the ASM mouse (Sarna et al. 2001). These cells, which stain negative for the cerebellar Purkinje neuron marker zebrin II (aldolase C) begin to degenerate at ∼2 months of age, leaving a subset of zebrin II-positive neurons intact. This latter cell population also degenerates, but at a much later time, with complete cell loss only occuring after ∼6 months. The relevance of the sequential loss of zebrin II-negative cells might be related to the proposed role of zebrin II as an IP3-buffering protein (Baron et al. 1995). Similarly, one of the Purkinje cell-specific proteins that was down-regulated in the mouse model of spinocerebellar ataxia type 1 was a carbonic anhydrase-related protein (Serra et al. 2004), which reduced levels of IP3 binding to the IP3R1 (Hirota et al. 2003). Therefore we suggest a possible role for the IP3R1, and the loss of neurons in which it is expressed at high levels, in initiation of the neuropathological cascade in NPD-A.

Our data based on the one human brain NPD-A sample available to us are supportive of our hypothesis. However, we remain somewhat cautious about the interpretation of the human brain data, as no clinical history was available for the patient with NPD-A and the brain region from which the sample was taken is ill defined. However, the reduction in SERCA levels is remarkable.

NPD-A is the fourth sphingolipid storage disease in which defective Ca2+ homeostasis has been reported and implicated in the progression of disease pathology. Our previous studies demonstrated defective Ca2+ homeostasis in neuronal forms of Gaucher disease (Korkotian et al. 1999; Lloyd-Evans et al. 2003b; Pelled et al. 2005), and in Sandhoff disease (Pelled et al. 2003). Defective Ca2+ homeostasis was also reported in a mouse model of a GM1 gangliosidosis (Tessitore et al. 2004). However, in these three studies, the neuropathology appeared to be caused by a direct effect of the accumulating sphingolipids on the proteins involved in regulating Ca2+ homeostasis, whereas we have no evidence that the accumulating lipids in NPD-A directly affect either SERCA or the IP3R. Indeed, the primary accumulating lipid in the ASM mouse, SM, did not affect either SERCA-mediated Ca2+ uptake or IP3R-mediated Ca2+ release. Likewise, GM2, whose levels were also raised in ASM mice, had no effect on the IP3R and, in any case, accumulated at levels that are probably too low to affect the SERCA (Pelled et al. 2003). SPC, the lyso-derivative of SM, and a second messenger in its own right (Meyer zu Heringdorf et al. 2002), had a small but statistically significant effect on IP3-mediated Ca2+ release; however, the levels of exogenously added SPC (100 µm) needed to induce this small effect were significantly higher than the SPC levels observed in NPD-A brain (Rodriguez-Lafrasse and Vanier 1999). Similarly, the levels of exogenously added lyso-glycosphingolipids, such as glucosylsphingosine, required to modulate Ca2+ homeostasis were at least an order of magnitude higher (Lloyd-Evans et al. 2003a) than those observed in the diseases (i.e. Gaucher disease) in which they accumulate. However, we cannot formally exclude the possibility that the local concentration of GM2 or SPC might be high enough to affect either the SERCA or the IP3R. Moreover, SPC also acts as an extracellular second messenger, and could induce changes in IP3-mediated Ca2+-release via a G protein-coupled receptor-dependent mechanism (Meyer zu Heringdorf et al. 2002), which would not be detectable in the in vitro microsomal analyses used in the present study.

At this stage, we do not know whether any other functions of the ER, other than regulation of Ca2+ homeostasis, are modified in NPD-A. However, it seems likely that there will be other changes, based both on our own work (Futerman and van Meer 2004; Ginzburg et al. 2004) and also a recent study on the GM1 gangliosidosis mouse (Tessitore et al. 2004) showing induction of ER stress, which leads to the unfolded protein response. However, we observed no change in ER stress proteins, such as Glucose Regulated Protein 78 (GRP78), GRP94 or Ccaat/enhancer-binding protein homologous protein (CHOP), in ASM–/– brain (data not shown). Another important function of the ER is phospholipid biosynthesis, and we observed that the rate of phosphatidylserine synthesis was slightly decreased in cerebellum, but not in cortex, of ASM–/– mice, as were levels of phosphatidylserine synthase mRNA and protein (Y. Kacher and A. H. Futerman, unpublished observation), supporting the idea that other ER functions will indeed be compromised in NPD-A.

In summary, our results suggest that dysregulated Ca2+ homeostasis plays a role in the neurodegeneration observed in a specific Purkinje cell population in NPD-A. Altered Ca2+ homeostasis and raised cytosolic Ca2+ levels can lead to cell death by multiple pathways (Mattson and Chan 2003), and further investigation is required to delineate which of these are involved in Purkinje cell death in ASM–/– mice, and presumably also in NPD-A.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

This work was supported by the National Niemann–Pick Disease Foundation, and by the Benoziyo center for neurological diseases in the Weizmann Institute of Science. We thank Dr Raya Eilam, Unit of Veterinary Resources, Weizmann Institute of Science, for help with immunohistochemical staining, Emyr Lloyd-Evans and Dori Pelled for help in establishing the Ca2+ uptake and release assays, and Raphael Schiffman, Developmental and Metabolic Neurology Branch, NINDS, National Institutes of Health, Bethesda, USA, for providing the human brain samples. Anthony H. Futerman is the Joseph Meyerhoff Professor of Biochemistry at the Weizmann Institute of Science.

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  6. Acknowledgements
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