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Biological Chemistry

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Volume 396, Issue 6-7


Secretory sphingomyelinase in health and disease

Johannes Kornhuber
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  • Department of Psychiatry and Psychotherapy, Friedrich Alexander University of Erlangen-Nürnberg (FAU), Schwabachanlage 6, D-91054 Erlangen, Germany
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/ Cosima Rhein
  • Department of Psychiatry and Psychotherapy, Friedrich Alexander University of Erlangen-Nürnberg (FAU), Schwabachanlage 6, D-91054 Erlangen, Germany
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/ Christian P. Müller
  • Department of Psychiatry and Psychotherapy, Friedrich Alexander University of Erlangen-Nürnberg (FAU), Schwabachanlage 6, D-91054 Erlangen, Germany
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/ Christiane Mühle
  • Department of Psychiatry and Psychotherapy, Friedrich Alexander University of Erlangen-Nürnberg (FAU), Schwabachanlage 6, D-91054 Erlangen, Germany
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Published Online: 2015-03-24 | DOI: https://doi.org/10.1515/hsz-2015-0109


Acid sphingomyelinase (ASM), a key enzyme in sphingolipid metabolism, hydrolyzes sphingomyelin to ceramide and phosphorylcholine. In mammals, the expression of a single gene, SMPD1, results in two forms of the enzyme that differ in several characteristics. Lysosomal ASM (L-ASM) is located within the lysosome, requires no additional Zn2+ ions for activation and is glycosylated mainly with high-mannose oligosaccharides. By contrast, the secretory ASM (S-ASM) is located extracellularly, requires Zn2+ ions for activation, has a complex glycosylation pattern and has a longer in vivo half-life. In this review, we summarize current knowledge regarding the physiology and pathophysiology of S-ASM, including its sources and distribution, molecular and cellular mechanisms of generation and regulation and relevant in vitro and in vivo studies. Polymorphisms or mutations of SMPD1 lead to decreased S-ASM activity, as detected in patients with Niemann-Pick disease B. Thus, lower serum/plasma activities of S-ASM are trait markers. No genetic causes of increased S-ASM activity have been identified. Instead, elevated activity is the result of enhanced release (e.g., induced by lipopolysaccharide and cytokine stimulation) or increased enzyme activation (e.g., induced by oxidative stress). Increased S-ASM activity in serum or plasma is a state marker of a wide range of diseases. In particular, high S-ASM activity occurs in inflammation of the endothelium and liver. Several studies have demonstrated a correlation between S-ASM activity and mortality induced by severe inflammatory diseases. Serial measurements of S-ASM reveal prolonged activation and, therefore, the measurement of this enzyme may also provide information on past inflammatory processes. Thus, S-ASM may be both a promising clinical chemistry marker and a therapeutic target.

Keywords: ceramide; inflammation; lipids; secretory sphingomyelinase; sphingomyelin; sphingomyelinase


Acid sphingomyelinase (ASM, EC plays a major role in sphingolipid metabolism because it catalyzes the hydrolysis of sphingomyelin (SM) to ceramide and phosphorylcholine. Ceramide and related products, such as spingosine-1-phosphate, are important lipid signaling molecules. These molecules are involved in a variety of molecular and cellular processes and play a central role in a growing number of human diseases. The multiple physiological sources of ceramide in mammalian cells (de novo synthesis from palmitoyl-CoA and serine, synthesis from sphingosine and fatty acid, SM catabolism, hydrolysis of glycosylceramide and galactosylceramide, dephosphorylation of ceramide-1-phosphate) do not contribute equally to the pool, but the degradation of SM by the ASM is an important source of ceramide. The prominent position of sphingomyelinases is mainly attributed to the abundance of their substrate SM in cell membranes (van Meer et al., 2008). Because of its spatially bulky head group, SM is restricted to the membrane leaflet where it is generated, i.e., the luminal Golgi leaflet or – after vesicular transport – the outer leaflet of the plasma membrane (Gault et al., 2010), unless flipping is aided by a specific flippase (Sharom, 2011). However, of the known flippases, none is active toward SM (Takatsu et al., 2014). Mammalian cells contain a single gene for ASM (SMPD1), which is responsible for the generation of both the secretory (S-ASM) and lysosomal (L-ASM) forms (Schissel et al., 1996a). S- and L-ASM originate from the same gene without the involvement of alternative RNA splicing as they can be produced from the same full-length cDNA (Schissel et al., 1998b). The common mRNA is also translated in the same reading frame because the resulting L- and S-ASM proteins are similar in size and are both recognized by antibodies prepared against L-ASM (Schissel et al., 1996a, 1998b). By contrast, in the nematode Caenorhabditis elegans, three genes have been identified (Lin et al., 1998; Kim and Sun, 2012) on different chromosomes (Yook et al., 2012): the product of asm-1 is almost entirely secreted but is Zn2+-independent, whereas only 20% of the asm-2 product is secreted and requires Zn2+. The asm-3 protein with the strongest homology to human ASM and sensitivity to desipramine and clomipramine (Lin et al., 1998; Kim and Sun, 2012) may correspond to L-ASM. The presence of separate genes for the secreted sphingomyelinase in C. elegans with developmentally regulated expression emphasizes the critical importance of the secreted enzyme. However, in mammalian organisms the role, function and regulation of S-ASM are not as well understood as those of L-ASM. Several previous reviews cover various aspects of S-ASM physiology and pathophysiology (Tabas, 1999; Goni and Alonso, 2002; Smith and Schuchman, 2008; Jenkins et al., 2009; Pavoine and Pecker, 2009; He and Schuchman, 2012). In this review, we focus on the progress made in recent years specifically on mammalian S-ASM and some aspects of secreted L-ASM, and we examine novel studies investigating S-ASM as a mediator of pathogenic processes, a potential therapeutic target and a biomarker in human diseases. The sequence numbering at the DNA and protein level in this review is based on the standard database reference sequence NM_000543.4 with 631 amino acids, which leads to a shift of +2 amino acids when referring to some published positions because of an additional leucine-alanine dipeptide within the polymorphic signal peptide.

Molecular and cellular regulatory mechanisms of S-ASM activity

Two distinct enzymes arise from the single SMPD1 gene because of differential modification and trafficking processes. The lysosomal form (L-ASM) is located in the endolysosomal compartment, whereas the secretory form (S-ASM) is released by the secretory pathway (Schissel et al., 1996a; Jenkins et al., 2010). S-ASM differs from the lysosomal form by its dependence on exogenously added Zn2+ ions (Schissel et al., 1998b) associated with its inhibition by ethylenediaminetetraacetic acid (EDTA), heat lability at 55°C (Spence et al., 1979) and exceptionally longer half-life (Jenkins et al., 2010), different molecular weight of the protein core due to an intact C-terminus (Jenkins et al., 2011b) and different N-terminal proteolytic processing. In addition, S-ASM has a complex N-glycosylation pattern (Ferlinz et al., 1994; Hurwitz et al., 1994b) that enables resistance to endoglycosidase H and results in a different localization (Schissel et al., 1998b) compared to L-ASM with high-mannose oligosaccharides for lysosomal targeting (Kornfeld, 1987). Whether L-ASM and S-ASM act on different SM pools in the cell, which would result in distinct effects via differential ceramide generation, remains unclear (Jenkins et al., 2010).

Genetic impact on S-ASM activity

A number of genetic sequence variations and mutations influence S-ASM activity and the process of generating L- or S-ASM. The repeat number variation of a hexanucleotide sequence – c.108GCTGGC(3_8)/p.37LA(3)_8) – that leads to the modification of the signal peptide of the pre-pro-form of ASM (Wan and Schuchman, 1995), was associated with S-ASM activity but not L-ASM activity in a cell culture model. In addition, analysis of a human sample resulted in a significant association of S-ASM activity with the number of hexanucleotide repeats; levels were highest in subjects homozygous for six repeats, intermediate in subjects homozygous for five repeats, and lowest in subjects homozygous for four repeats. One of the most frequent SMPD1 sequence variations, rs1050239/c.1522G>A/p.G508R (Rhein et al., 2014), has been associated with S-ASM activity but not L-ASM activity. S-ASM activity was measured in the blood plasma of healthy young adults and was highest in subjects homozygous for the major G allele, intermediate in heterozygous subjects and lowest in subjects homozygous for the A allele (Reichel et al., 2014). Importantly, the influences of these two polymorphic sites on S-ASM activity are independent (Rhein et al., 2014). The nonsynonymous variation rs141641266/c.1460C>T/p.A487V, previously assumed to be a missense mutation causing Niemann-Pick disease (NPD) type B (Simonaro et al., 2002), leads to normal L-ASM and only slightly lower S-ASM activity levels (Rhein et al., 2013).

More than 100 clinically relevant missense mutations in the SMPD1 gene leading to enzymes with decreased catalytic activity, the cause of autosomal recessive NPD types A and B (Brady et al., 1966; Schuchman, 2010), have been deposited in the Human Gene Mutation Database (www.hgmd.cf.ac.uk). While genotype/phenotype correlations are typically complex, the DR608 deletion mutation appears to protect patients against the development of the more severe neuropathic NPD form type A (Schuchman and Miranda, 1997). L-ASM activity in cultured skin fibroblasts of patients correlates with the respective phenotype, with residual L-ASM activity of <5% corresponding to the more severe NPD type A (Desnick et al., 2010), for which – similar to other neurodegenerative sphingolipidoses – effective therapeutic options are missing (Eckhardt, 2010). The determination of S-ASM activity in blood plasma can be used diagnostically, particularly to discriminate between NPD patients and NPD carriers (He et al., 2003). However, clinical data on S-ASM activity in NPD patients remain limited because in most studies only cellular L-ASM activity is determined (Vanier et al., 1980; Pavlu-Pereira et al., 2005). Because lysosomal hypertrophy is a hallmark of lysosomal storage disorders, a parallel assessment of the acidic compartment volume combined with L- and S-ASM activities could provide a basis for genotype-phenotype correlations, particularly to explain the large heterogeneity of symptom severity observed in NPD patients with an intermediate phenotype (Pavlu-Pereira et al., 2005; Wasserstein et al., 2006). Interestingly, there is growing support for a genetic convergence of lysosomal storage disorders and Parkinson’s disease (Deng et al., 2014). Mutations such as p.L304P (Gan-Or et al., 2013), which may be limited to Ashkenazi Jews (Wu et al., 2014), or variants in SMPD1, e.g., p.R591C, p.P533L (Foo et al., 2013), have not only been associated with altered ASM activities but also appear to be linked to an up to 9-fold increased risk for Parkinson’s disease (Spatola and Wider, 2014). This effect is likely mediated via ceramide because plasma ceramide and monohexosylceramide levels are increased and associated with cognitive impairment in Parkinson’s disease patients lacking a risk-conveying glucocerebrosidase mutation (Mielke et al., 2013).

Transcriptional and translational effects on ASM activity

ASM is up-regulated at the transcriptional level in different cell culture models, but only effects on L-ASM activity have been investigated (Langmann et al., 1999; Murate et al., 2002). A variety of alternatively spliced ASM transcripts have been identified, of which only one, ASM-1, encodes an enzyme with complete protein domains and L-ASM activity. The isoforms ASM-2 to -7 lack protein domains that are essential for ASM activity. Some ASM isoforms have a dominant-negative effect on cellular L-ASM activity levels (Rhein et al., 2012). However, whether and how S-ASM activity is influenced by alternative splicing has not been investigated. The unexpectedly mild NPD-B phenotype and 20–25% residual L-ASM activity observed in two patients homozygous for either p.M1_W32del or p.W32X and, consequently, complete absence of full-length ASM, suggests that in the absence of a functional first initiation codon, the second initiation codon (ATG coding for p.33M) may still produce a relatively functional enzyme (Pittis et al., 2004).

Trafficking and post-translational processing relevant for S-ASM and L-ASM activity

The generation of L-ASM versus S-ASM is determined by the differential processing and trafficking of the common precursor ASM protein (Figure 1). Upon entry of the nascent ASM polypeptide (631 amino acids corresponding to 70 kDa) into the rough endoplasmic reticulum (ER) (Hasilik, 1992), cleavage of the putative N-terminal signal peptide (Wan and Schuchman, 1995), which is postulated to extend to p.A48 (Schuchman et al., 1991) yields the pre-pro-form with a molecular weight of 75 kDa after glycosylation (65 kDa protein core). The signal peptide is essential for the production of enzymatically active mature L-ASM. ASM constructs with N-terminal truncations of the signal sequence lack catalytic L-ASM activity (Ferlinz et al., 1994). Despite the predicted absence of the signal peptide in the mature enzyme, the length of the hydrophobic core of the polymorphic ASM signal peptide affects the secretion and activity of S-ASM, whereas L-ASM activity is unaffected (Rhein et al., 2014).

L- and S-ASM arise from a common ASM precursor protein. The ASM-encoding gene SMPD1 gives rise to a common mannosylated precursor protein, pre-pro-ASM, which is cleaved to yield pro-ASM. Differential glycosylation and N-terminal as well as C-terminal processing inside the Golgi then lead to the generation of two distinct ASM forms. Whereas S-ASM is released into the extracellular space via the constitutive secretory pathway and requires Zn2+ ions for activation, L-ASM is shuttled into the lysosomal trafficking pathway via its mannose 6-phosphate groups and encounters Zn2+ ions on its way to the endolysosomal compartment. The details of the exocytotic and endocytotic steps have not been fully elucidated (dotted lines). Data regarding the molecular weight of the mature enzyme with the deglycosylated protein core in brackets are based on the most recently reported data (Edelmann et al., 2011; Jenkins et al., 2011b) and protein molecular weight calculations (protcalc.sourceforge.net) after modifications cited in the text; they may vary from other reports.
Figure 1:

L- and S-ASM arise from a common ASM precursor protein.

The ASM-encoding gene SMPD1 gives rise to a common mannosylated precursor protein, pre-pro-ASM, which is cleaved to yield pro-ASM. Differential glycosylation and N-terminal as well as C-terminal processing inside the Golgi then lead to the generation of two distinct ASM forms. Whereas S-ASM is released into the extracellular space via the constitutive secretory pathway and requires Zn2+ ions for activation, L-ASM is shuttled into the lysosomal trafficking pathway via its mannose 6-phosphate groups and encounters Zn2+ ions on its way to the endolysosomal compartment. The details of the exocytotic and endocytotic steps have not been fully elucidated (dotted lines). Data regarding the molecular weight of the mature enzyme with the deglycosylated protein core in brackets are based on the most recently reported data (Edelmann et al., 2011; Jenkins et al., 2011b) and protein molecular weight calculations (protcalc.sourceforge.net) after modifications cited in the text; they may vary from other reports.

L- and S-ASM undergo differential glycosylation processing inside the Golgi giving rise to the ASM pro-form of 72–75 kDa (63–64 kDa protein core). L-ASM has a high mannose N-glycan composition, which provides stabilization and protection against proteolysis within the lysosome, whereas S-ASM possesses a complex N-glycosylation pattern (Hurwitz et al., 1994b; Schissel et al., 1998b). This differential glycosylation is evident in the susceptibility of L-ASM to endoglycosidase H, which is specific for high mannose-type N-linked oligosaccharides (Yamamoto, 1994). In contrast, S-ASM is completely resistant to endoglycosidase H, and both enzymes are susceptible to peptide-N-glycosidase F (Schissel et al., 1998b). In the cis-Golgi, L-ASM acquires mannose-phosphate residues via the typical sequential action of N-acetylglucosamine-1-phosphotransferase (Reitman and Kornfeld, 1981) and N-acetylglucosamine phosphodiesterase on the mannose residues of the precursor (Hurwitz et al., 1994b; Schissel et al., 1998b), whereas S-ASM is spared of this modification. Bovine DNase I is an example of a suboptimal substrate for the phosphotransferase that is required to confer the mannose 6-phosphate moieties for the lysosomal pathway, and this feature is assumed to give rise to an intralysosomal as well as a secretory form of bovine DNaseI. A similar phenomenon may occur for ASM (Nishikawa et al., 1997; Schissel et al., 1998b). The ASM pro-form contains six potential N-glycosylation sites (N88, N177, N337, N397, N505, N522), five of which appear to be occupied in purified ASM (Ferlinz et al., 1997). The two C-terminal N-glycosylation sites (N505 and N522) are essential for proper trafficking and affect S-ASM secretion and the enzymatic activity of L-ASM (Ferlinz et al., 1997). Although presumably not glycosylated, removal of the fifth site reduces enzymatic activity to <20% due to protein misfolding (Newrzella and Stoffel, 1996; Ferlinz et al., 1997).

The S-ASM pro-form is constitutively secreted via the default Golgi secretory pathway resulting in a 75–80 kDa extracellular enzyme with a protein core of 64 kDa (Jenkins et al., 2010, 2011b). The L-ASM pro-form is targeted mostly via cation-dependent and cation-independent mannose 6-phosphate receptors and to a lesser extent via sortilin-mediated transport to the endolysosomal compartment (Ni and Morales, 2006). The importance of the mannose 6-phosphate recognition system is evident in the 8-fold increase in the ratio of secreted to intracellular ASM activity in cells deficient in mannose phosphorylation. Alanine scanning mutagenesis of the 13 lysine residues revealed that p.K95 is critical for the specific three-dimensional arrangement required for mannose phosphorylation, and p.K95A replacement results in an overall 2-fold increase in the ratio of secreted to intracellular ASM activity via a reduction of intracellular activity and an increase in Zn2+-dependent secretory activity (Takahashi et al., 2005). Similarly, deglycosylation treatment or expression of a dominant-negative sortilin variant results in trapping of the enzyme within the Golgi in its degradation-vulnerable form (Bartelsen et al., 1998; He et al., 1999; Ni and Morales, 2006).

Sequencing of purified enzymes has revealed that the N-terminus of S-ASM begins at the amino acid p.H62 (numbered according to the standard database reference sequence NM_000543.4 throughout the review, p.H60 in the publication), whereas that of L-ASM begins at p.G68 (Schissel et al., 1998b) after typical N-terminal processing of lysosomal proteins (Hasilik, 1992). The positions of the N-termini correspond to molecular weights of the remaining deglycosylated full C-terminal portions of 63.6 kDa for S-ASM and 63.0 kDa for L-ASM. The removal of only a few N-terminal amino acids – six in ASM – is reminiscent of the maturation of another lysosomal enzyme, the β-chain of hexosaminidase (Quon et al., 1989). By contrast, p.G85 (61.0 kDa) has been identified as the first amino acid of purified human placental ASM (Lansmann et al., 1996). While the exact molecular weight values differ slightly between publications, the presence of markedly different subunits across human tissues (Jobb and Callahan, 1987) has not been confirmed. ASM may also be delivered from the Golgi directly to the phagosome, bypassing fusion with lysosomal compartments. Sortilin participates in this process via its short cytoplasmic tail (Wähe et al., 2010), which is similar to the cytoplasmic segment of the cation-independent mannose 6-phosphate receptor (Petersen et al., 1997; Nielsen et al., 2001).

In the endolysosomal system, the 72-kDa enzyme is processed at the C-terminus to the 65-kDa L-ASM (55 kDa for the deglycosylated protein). This processing step within the acidic compartment is similar to that for other lysosomal hydrolases (Erickson and Blobel, 1983; Arunachalam et al., 2000) and is essential to generate the catalytically highly active enzyme, most likely by promoting the coordination of lysosomal Zn2+. By contrast, the low activity of the non-lysosomal precursor may prevent enzymatic activity in the Golgi (Jenkins et al., 2011b). The role of the C-terminal region in the proteolytic maturation of L-ASM is indicated by the altered localization and virtual absence of Zn2+-independent activity in the recombinant NPD mutants p.R602H, p.R602P, and p.ΔR610 (Jenkins et al., 2011b). The preliminary detection of p.Q622 as the C-terminus of L-ASM would suggest a loss of only nine amino acids, resulting in a protein core of 62 kDa (Jenkins et al., 2011b). However, another recent study indicates that TNF-mediated stimulation leads to the direct interaction of caspase-7 with the 72-kDa pro-ASM and proteolytic cleavage of the zymogen at the non-canonical cleavage site after p.D253 within TNF receptosomes, generating an active 57-kDa L-ASM (Edelmann et al., 2011). This active L-ASM consists of a protein core of 43 kDa and corresponds to the previously proposed active form after N-terminal cleavage between the second and third glycosylation sites with minimal trimming at the C-terminus (Ferlinz et al., 1994). Unlike its lysosomal counterpart, the C-terminus of S-ASM remains intact (Jenkins et al., 2011b), enabling regulation via its unbound p.C631 (Qiu et al., 2003).

Functional inhibitors of ASM (FIASMAs), e.g., antidepressant drugs such as desipramine and fluoxetine (Kornhuber et al., 2008, 2010, 2011), lead indirectly to the inactivation of L-ASM (Hurwitz et al., 1994a; Kölzer et al., 2004b) by leupeptin-sensitive proteolytic cleavage to a 52-kDa form (Jenkins et al., 2011b). The half-life of L-ASM, as determined by the addition of the protein synthesis inhibitor cycloheximide and pulse chase experiments, of approximately 5–6 h (Hurwitz et al., 1994b; Schissel et al., 1998b; Jenkins et al., 2010) is exceptionally short compared to most lysosomal enzymes, which have half-lives of days to weeks. By contrast, S-ASM is not further processed after the Golgi and is detected as a 75–80-kDa form following its secretion into the extracellular space. In contrast to L-ASM, after secretion, S-ASM is unusually stable and remains at the same activity level over at least 25 h, with profound implications for chronic conditions in which small changes in continued secretion over time compound to high total S-ASM levels (Jenkins et al., 2010).

ASM activity is influenced by further post-translational modifications. An activating effect on L-ASM has been described for the phosphorylation of serine residue p.S510 in the C-terminal domain by the protein kinase Cδ. It has been postulated that this phosphorylation results in the translocation of L-ASM to the plasma membrane (Zeidan and Hannun, 2007a; Zeidan et al., 2008b). The ASM mutant p.S510A, in which this phosphorylation site was altered by site-specific mutagenesis, retains L-ASM activity but is not secreted (Jenkins et al., 2010). This phosphorylation site seems to be essential for constitutive as well as regulated secretion and thus the activity of S-ASM. Furthermore, stimulated exocytosis of ASM is dependent on the t-SNARE protein syntaxin 4 (Perrotta et al., 2010).

L- and S-ASM are activated in vitro via interactions with Zn2+ ions (Schissel et al., 1998b). Zn2+ acquisition itself may represent a form of L-ASM regulation because the partial Zn2+-dependence of L-ASM isolated from lysosomal-rich fractions but not standard cell homogenates suggests an incomplete saturation of this form with Zn2+ (Schissel et al., 1998b), although the isolate might have contained Zn2+-dependent pre-lysosomal forms in addition to the fully mature lysosomal 65-kDa ASM (Jenkins et al., 2011b). Massive overexpression of ASM, which typically results in the saturation of the mannose 6-phosphate receptor shuttling mechanism and secretion of an otherwise lysosomally targeted enzyme (Ioannou et al., 1992), yielded a Zn2+-dependent mannose 6-phosphate receptor-binding species in the supernatant. This finding demonstrates that Zn2+ acquisition is not glycosylation-dependent but occurs during lysosomal trafficking (Schissel et al., 1998b).

Eight disulfide bonds are formed among 17 cysteine residues within mature ASM, which leaves the C-terminal p.C631 unbound (Lansmann et al., 2003). Oxidation, deletion or replacement of this residue activates recombinant human S-ASM by favoring the hydration of Zn2+. Activation is also achieved by copper-promoted dimerization via cysteine residues and by C-terminal truncation by carboxypeptidase Y (Qiu et al., 2003) and likely resembles the cysteine switch mechanism of matrix metalloproteinases (Van Wart and Birkedal-Hansen, 1990) in post-translational regulation. However, this activation is assumed to be limited to S-ASM in which the entire C-terminus is retained; this cysteine is lost in fully mature L-ASM during the final processing step (Jenkins et al., 2011b).

Regulation of S-ASM and L-ASM secretion

The ASM enzyme consists of distinct domains, a proline-rich domain, a metallophosphoesterase domain and the C-terminal domain. Moreover, strong sequence homology (Ponting, 1994) and an analogous arrangement of disulfide bonds (Lansmann et al., 2003) suggest that ASM contains its own intramolecular sphingolipid activator protein (SAP)-type domain as the fourth functional domain and is thus, unlike other lysosomal hydrolases, independent of the presence of SAPs as a cofactor. SAPs are small, enzymatically inactive lysosomal glycoproteins that facilitate sphingolipid degradation at the inner membrane of acidic compartments by promoting substrate binding and lipid extraction (Kolter and Sandhoff, 2005; Remmel et al., 2007). The addition of SAPs can partially rescue inactive ASM with mutations in conserved amino acids within its SAP domain (Kölzer et al., 2004a).

The selective activation of L-ASM by a variety of stress stimuli has been comprehensively investigated (Charruyer et al., 2005; Rotolo et al., 2005; Zeidan et al., 2008a; Milhas et al., 2010). However, stimulating agents can have differential effects on L- and S-ASM activity. The co-regulation of S- and L-ASM activity has been addressed in several cell culture systems. Cultured human platelets exhibit increased S-ASM release but decreased L-ASM activity upon stimulation with thrombin (Romiti et al., 2000). S-ASM activity is increased and L-ASM activity is decreased in MCF7 breast carcinoma cells treated with phorbol myristate acetate or ammonium chloride (Jenkins et al., 2010). S-ASM activity is increased upon stimulation with interleukin (IL) 1β or tumor necrosis factor-alpha (TNFα), with no effect on L-ASM activity (Jenkins et al., 2010). The same pattern was observed following the treatment of human umbilical vein endothelial cells with IL-1β or interferon γ; S-ASM activity was increased by 2–3.5-fold, whereas L-ASM activity was slightly decreased (Marathe et al., 1998). Oxidized low-density lipoprotein (LDL) and oxidized LDL-containing immune complexes differentially regulate L-ASM activity and S-ASM release in macrophages (Truman et al., 2012). A special role of S-ASM in chemokine elaboration was detected following the stimulation of MCF7 cells with TNFα because the chemokine CCL5 appears to be an initial target of the pathway that is selectively induced by S-ASM activity (Jenkins et al., 2011a). Cytokines also affect the ratio of L- and S-ASM activity in a genetic manner (Rhein et al., 2014). Thus, cytokine stimulation of cultured cells may have a short-term effect on S-ASM and appear to increase S-ASM activity and secretion while decreasing L-ASM activity. This effect could evolve by redirecting the ASM precursor protein away from the lysosomal pathway into the secretory pathway. The involvement in this redirection of the cis-Golgi enzyme N-acteylglucosaminyl-1-phosphotransferase, which phosphorylates the mannose residues of the precursor protein for the lysosomal trafficking pathway, has been proposed (Tabas, 1999).

Although originally thought to localize only to the lysosome, under stress conditions, L-ASM may translocate from intracellular compartments to the extracellular leaflet of the cell membrane, where it hydrolyzes SM to ceramide to form signaling platforms (Cifone et al., 1994); however, L-ASM may also contribute to extracellular ASM activity. The designation of the ASM that is translocated to the outer leaflet of the plasma membrane as a response to specific stimuli is not consistent and may, in fact, represent S- or L-ASM or a combination of both species. While some reports refer to the enzyme as ‘secretory’ ASM, other authors suggest it is L-ASM by assuming a lysosomal origin. Although the vesicles that appear to fuse and empty their contents at the extracellular surface may contain the secretory form of ASM, direct data on specific properties or Zn2+-dependence are lacking in most studies. For example, extracellular ASM activity increased following the repair of myoblasts from mechanical injury as a result of vesicle docking and exocytosis (Defour et al., 2014). Cationic cell-penetrating peptides as potential molecular transporters of membrane-impermeable molecules induce the translocation of ASM to the plasma membrane, resulting in an increase in ceramide formation and, in turn, enhancing its own uptake (Verdurmen et al., 2010). Similarly, treatment of Jurkat T cells with ultraviolet light type C also induces the rapid translocation of ASM to the membrane, hydrolysis of extracellular SM and raft clustering (Rotolo et al., 2005). In addition to the ASM-independent functions of CD95, transmission electron microscopy and fluorescence-activated cell sorting analysis have demonstrated that CD95 stimulation also causes ASM to translocate from intracellular, presumably vesicular, compartments to the extracellular leaflet. This process hydrolyzes extracellularly oriented SM, which, in turn, mediates CD95 clustering as an essential step toward CD95-triggered cell death (Grassmé et al., 2001). Together with CD95, TNF-related apoptosis-inducing ligand is involved in the activation of ASM via a redox mechanism that results in the generation of ceramide and the formation of ceramide-enriched membrane platforms (Dumitru and Gulbins, 2006), potentially via the release of ASM of lysosomal origin, as indicated by cell lysate activity in the absence of Zn2+. The application of hydrogen peroxide as the primary form of reactive oxygen species in mammals also induces very rapid calcium-dependent ASM release by lysosomal exocytosis, as detected by the exposure of lysosome-associated protein LAMP1 (Li et al., 2012). The lysosomal origin of released ASM is also supported by the concomitant appearance of LAMP1 on the sarcolemma surface and the release of the lysosomal β-hexosaminidase in a wound repair study (Corrotte et al., 2013).

In fact, ASM has recently emerged as a major regulator facilitating wound repair in eukaryotic cells, such as from pore-forming toxins (Tam et al., 2010; Corrotte et al., 2013). Plasmalemmal injury-triggered Ca2+ influx induces the fusion of lysosomes with the plasma membrane and exocytosis of ASM. Through ceramide generation, ASM in turn triggers lesion removal via endocytosis and intracellular degradation (Tam et al., 2013; Andrews et al., 2014). Transcriptional silencing of ASM abolishes this process, while the addition of exogenous enzyme restores it (Tam et al., 2010). Trypanosoma cruzi subverts the ASM-dependent ceramide-enriched endosomes in the plasma membrane repair pathway to invade host cells (Fernandes et al., 2011). While ceramide production by ASM in the outer leaflet promotes tighter packing and a negative curvature that leads to inward vesiculation (Trajkovic et al., 2008), ceramide generation caused by the activation of the neutral sphingomyelinase species localized to the inner leaflet of the plasma membrane promotes an outward curvature and thus allows the cell to shed membrane-containing toxin pores into the extracellular space (Draeger and Babiychuk, 2013).

Sources of S-ASM

Similar to the ubiquitous expression of L-ASM, which presumably reflects an important housekeeping role, a wide variety of cells have been demonstrated to secrete substantial amounts of Zn2+-dependent S-ASM. In addition to the production of L-ASM, murine brain microglial cells, mouse peritoneal macrophages, J774 macrophages, human skin fibroblasts and several other (untransfected) cultured cells, including Chinese hamster ovary and COS-7 cells, have also been reported to secrete ASM that is activated by physiological levels of Zn2+. This activity is markedly up-regulated during the differentiation of human monocytes to macrophages (Schissel et al., 1996a).

Cultured human coronary artery and umbilical vein endothelial cells secrete massive amounts of ASM both apically and basolaterally (Marathe et al., 1998). However, in contrast to the almost complete Zn2+-dependence of S-ASM originating from macrophages and fibroblasts, endothelium-derived S-ASM exhibits partial activity in the absence of exogenous Zn2+. The up to 20-fold higher levels of basal secretion compared to macrophages are further increased 2- to 3-fold by incubation with inflammatory cytokines such as IL-1β or interferon-gamma (Marathe et al., 1998), demonstrating that human vascular endothelial cells are a rich and regulatable source of S-ASM. The ability of ependymal cells to secrete ASM has not yet been analyzed, although these cells are responsible for most daily cerebrospinal fluid (CSF) production and could be the source of S-ASM detected therein (Mühle et al., 2013). In addition to various endothelial cells, human platelets not only exhibit ASM activity but also release the enzyme in response to stimulation by thrombin (Simon et al., 1998; Romiti et al., 2000), without a significant concomitant change in ceramide levels (Simon and Gear, 1999). As a likely source of ASM in human tear fluid, human corneal cells constitutively secrete ASM and respond with increased secretion to agents that induce stress, including ultraviolet B radiation and hyperosmolarity (Robciuc et al., 2014).

Furthermore, extracellular ASM activity has been detected in muscle cell supernatants after mechanical injury and repair but may represent the lysosomal enzyme that is exocytosed after docking of lysosomes to the plasma membrane (Defour et al., 2014). The release of ASM into the culture medium of human fibroblasts and mouse L-cells has been detected in the presence of the lysosomotropic agent ammonium chloride (Weitz et al., 1983). Similarly, a number of studies have reported that upon stimulation with various agents, vesicles of unspecified identity containing ASM translocate to the plasma membrane and release the enzyme to the outer leaflet or to the extracellular space. Some authors refer to this ASM species as L-ASM (Li et al., 2012; Defour et al., 2014), some as S-ASM (Rotolo et al., 2005) and some do not directly specify the affiliation (Grassmé et al., 2001; Verdurmen et al., 2010), which thus remains to be elucidated. Regardless, the exocytosed ASM localizes to the extracellular space or the outer leaflet of the plasma membrane, in contrast to the orientation of the neutral sphingomyelinase, in which the catalytic site is thought to face the cytosolic inner leaflet (Tani and Hannun, 2007).

While S-ASM is secreted by various cell types, the enzyme can also be internalized and trafficked to the lysosome, where its contact with Zn2+ sufficiently activates the enzyme such that subsequent cell homogenates exhibit sphingomyelinase activity in the absence of added Zn2+, as demonstrated in ASM-negative cells treated with S-ASM (Schissel et al., 1998b). Similar to other lysosomal enzyme defects, this feature renders NPD amenable to enzyme replacement therapy. After intravenous administration of recombinant ASM in ASM knock-out (KO) mice, more than 90% of the injected enzyme was taken up by the liver, of which the majority was delivered to the lysosome. The mannose 6-phosphate receptor system was responsible for 30–50% of the internalization of the enzyme (He et al., 1999), although in ASM KO cells, the mannose receptor plays the major role (Dhami and Schuchman, 2004). As little as four injections resulted in the reversal of symptoms in reticuloendothelial organs, even in older KO mice, but not in neurological improvement because the enzyme cannot traverse the blood-brain-barrier (Miranda et al., 2000).

Estimations of ASM activity in human blood indicate that approximately 17% of the total activity in healthy adults is present as L-ASM in the mixture of cells representing peripheral blood mononuclear cells, while the remaining 83% is detectable as Zn2+-dependent S-ASM in the corresponding plasma (C. Mühle, unpublished data). This proportion, or even absolute ASM levels, may vary between species, as suggested by the observation of elevated S-ASM activity in fetal calf serum, mouse and particularly rat serum (C. Mühle, unpublished data).

Distribution of S-ASM

In contrast to the nearly ubiquitous distribution of the L-ASM enzyme in mammalian tissues (Weinreb et al., 1968) and almost uniform expression across human brain regions (Spence et al., 1979), there is only some basic evidence for the presence of S-ASM in various body fluids other than serum and plasma. Early studies indicated ASM activity in amniotic fluid and urine and confirmed this activity based on phosphorylcholine liberation as well as lower activity levels in patients with NPD (Harzer and Benz, 1973). The enzyme has been purified from human urine and extensively characterized (Weitz et al., 1985; Quintern et al., 1987; Quintern and Sandhoff, 1991). Compared to ASM activity in leukocytes, greater variations in activity were observed in human urine and were best correlated with 24-h creatinine excretion in long-term tests (Seidel et al., 1978). Takahashi et al. (2000) detected slightly weaker S-ASM activity in human urine compared to sera and nearly 2-fold higher activity in synovial fluid aspirated from patients with either rheumatoid arthritis or osteoarthritis. Significant stimulation of SM degradation by Zn2+ ions indicated the presence of S-ASM rather than L-ASM in these samples. Very high Zn2+-dependent S-ASM activity has been observed in salivary fluid and may participate in the digestion of the abundant SM in the normal diet. Similarly, ASM activity detected in human milk efficiently degraded SM as the dominant phospholipid in milk (Nyberg et al., 1998).

Unexpectedly, ASM activity is more than 10-fold higher in tear fluid than in serum. Although the lack of a response to the addition of Zn2+ ions in that study (Takahashi et al., 2000) may indicate that this activity is the result of L-ASM released from cells, a more recent investigation identified both L-ASM and S-ASM in all five tested tear fluid samples (Robciuc et al., 2014). By optimizing the reaction conditions, particularly with respect to the type and concentration of detergent, Zn2+-dependent S-ASM activity has been detected in human CSF and characterized (Mühle et al., 2013). The identity of this enzymatic SM-hydrolyzing activity as S-ASM encoded by SMPD1 was confirmed by its absence in Smpd1 KO mice and overexpression in Smpd1 transgenic mice, which exhibit extraordinarily high levels of S-ASM compared to L-ASM overexpression. ASM activity in human or boar seminal plasma is not influenced by divalent metal ions or chelating agents. Thus, these enzymes most likely originate from the lysosome (Vanha-Perttula, 1988). A SM-degrading enzyme with a slightly acidic pH optimum and a distinct pattern of activation by Co2+ and Mn2+ has been purified from bovine seminal plasma but is assumed to differ from S-ASM (Vanha-Perttula, 1988; Vanha-Perttula et al., 1990).

In a mouse model of NPD type C, lysosomal enlargement in the brain – a feature of these lysosomal storage disorders – correlated with that in circulating B cells (te Vruchte et al., 2014). Thus, measuring the relative acidic compartment volume of these peripheral cells using a fluorescent probe may represent a surrogate marker for the progression of lipid storage in the brain. While these data suggest an association between brain and peripheral L-ASM activities, the higher levels of S-ASM activity in tear fluid compared to serum (Takahashi et al., 2000) and the lack of a correlation between activities in serum and CSF (Mühle et al., 2013) indicate tissue-specific regulation of S-ASM secretion.

In vitro characterization and function of S-ASM

Coordinated breakdown of SM, the most abundant complex sphingolipid in mammalian cells, is essential for membrane homeostasis. Different structures such as the plasma membrane, axonal myelin sheaths, plasmatic lipoproteins, various particles or vesicles as well as membranes of exogenous origin such as particles could serve as physiological substrates for S-ASM. As a bilayer-destabilizing lipid with a considerably smaller polar head group and thus reduced area within the leaflet compared to SM (López-Montero et al., 2007), ceramide imposes a negative curvature at the bilayer interface (Veiga et al., 1999). Thereby, ceramide generation by ASM in the outer leaflet of the plasma membrane facilitates the formation of endocytic vesicles (Zha et al., 1998). Moreover, the altered fluidity and exclusion of cholesterol promote the formation of large ceramide-enriched platforms, which, in turn, lead to clustering of receptors and signaling molecules (Sot et al., 2008; Staneva et al., 2008, 2009). In addition to SM, ASM may also cleave phosphatidylglycerol and phosphatidylcholine (Quintern et al., 1987; Oninla et al., 2014), although contradictory evidence indicates that neither phosphatidylcholine nor lysophosphatidylcholine or glycerophosphocholine undergo hydrolysis (Spence et al., 1989).

Enzyme kinetics

Experiments using a secretion-incompetent p.S510A mutant enzyme suggest distinct metabolic roles of S-ASM and L-ASM in cellular ceramide formation with respect to specific substrates: while Il-1β induces a time- and dose-dependent up-regulation of S-ASM secretion with selective production of C16-ceramide at the expense of C16-SM, elevations in L-ASM activity are associated with increases in selective, very-long-chain ceramide species, e.g., C26:1-ceramide (Jenkins et al., 2010). Michaelis-Menten kinetic analysis revealed roughly similar values of the Michaelis constant Km for SM affinity despite the use of differently labeled substrates: 77 μm for soluble ASM in the human epidermis (Bowser and Gray, 1978), 5–25 μm for ASM purified from placenta (Jones et al., 1981, 1983; Sakuragawa, 1982), 65 μm for ASM from Bacillus cereus (Fujii et al., 2004), 47 μm for ASM from human fibroblasts (Sato et al., 1988), 25 μm for recombinant ASM from the conditioned medium of infected insect cells (Bartelsen et al., 1998), 20 μm for ASM from human CSF (Mühle et al., 2013), 0.2–0.6 μm for ASM released from human fibroblasts and mouse L-cells, respectively (Weitz et al., 1983), 11 μm with a VMAX of 21 nmol/h/mg protein for HEK 293 whole cell lysates and 2 μm and a VMAX of 4.3 μmol/h/mg protein for ASM immunoprecipitated from Jurkat cells (Gulbins and Kolesnick, 2000). Specific activities ranged from 600 to 2500 μmol/h/mg (Sakuragawa, 1982; Yamanaka and Suzuki, 1982; Jones et al., 1983; Weitz et al., 1985; Lansmann et al., 1996).

Defects in N-glycosylation reduce enzyme stability and activity but do not affect KM values (Newrzella and Stoffel, 1996). Moreover, pH plays an important role in substrate binding but not catalytic velocity (Callahan et al., 1983). The stimulation of cell surface receptors can lead to an increase in VMAX′ of 2–3-fold with little or no change in KM values (Schwandner et al., 1998). In a therapeutically aimed approach, the dipolar aprotic solvent dimethyl sulfoxide caused a slow but marked up to 2-fold increase in sphingomyelinase activity (Vmax) at pH 5.0 and 7.5 over a period of days. This activity was dependent on protein synthesis and did not substantially change the Km value (Sakuragawa et al., 1985; Sato et al., 1988).

Reaction conditions

The influence of detergents on ASM activity was examined by Spence et al. (1989) using fetal bovine serum, with an 8–15-fold increase in response to the addition of Triton X-100 to the reaction. It was also noticeable by the discrepancy in an order of magnitude between optimal Nonidet P-40 concentrations in assays for human serum- vs. CSF-derived S-ASM (Mühle et al., 2013). Detergent also influences the immunoprecipitation of soluble ASM from human urine and placenta (Driessen et al., 1985), but details regarding the different detergent optima remain limited. Storage temperature appears to be an important parameter for comparing absolute S-ASM activities because the activity of endogenous ASM from plasma, CSF and cell lysates (Mühle et al., 2013) as well as that of human recombinant ASM in cell supernatants (Qiu et al., 2003) is higher following long-term storage at -20°C compared to -80°C. This activation is attributed to the loss of the single free cysteine residue p.C631 by chemical modification during the freezing process (Qiu et al., 2003). However, the enzymatic activity remains relatively stable during the storage of CSF (Mühle et al., 2013) as well as undialyzed urine (Seidel et al., 1978) at room temperature for several days.

The dependence on the addition of Zn2+ ions is supported by the stimulation of S-ASM activity by Zn2+, which is increased in case of prior chelation of endogenous Zn2+ by EDTA, and its inhibition by EDTA or the more zinc-specific chelator 1,10-phenanthroline (He et al., 1999). By contrast, typical short-term chelation of L-ASM with EDTA is insufficient to strip the enzyme of its metal ion, consistent with other zinc metalloenzymes (Little and Otnäss, 1975). In contrast to neutral sphingomyelinases, Mg2+ and Mn2+ were ineffective for ASM activity (Spence et al., 1989). The role of other metal cations has not been well investigated, with the exception of a potentially different ASM species that was partially purified from seminal plasma (Vanha-Perttula, 1988). The optimal Zn2+ concentrations are similar to those for matrix metalloproteinases (Sorbi et al., 1993) and lie well within the range present in the extracellular space, e.g., approximately 100 μm in human whole blood or serum (Buxaderas and Farré-Rovira, 1985), although the actual concentration depends on factors such as dietary intake or pregnancy (Donangelo and Chang, 1981). Zn2+ ions move freely through the cerebrospinal and extracellular fluid compartments and are differentially distributed in various regions of the brain (Takeda et al., 1994), exhibiting variations in half-life (Takeda et al., 1995) and age-dependence (Sawashita et al., 1997) that may influence local S-ASM activity. Moreover, Zn2+ levels are elevated in both atherosclerotic and inflammatory lesions (Mendis, 1989; Milanino et al., 1993), and Zn2+ is released during intense neuronal activation (Assaf and Chung, 1984; Frederickson and Moncrieff, 1994). In addition, the regulation and cell-type-dependent variation of the subcellular concentration and localization of Zn2+ ions (Csermely et al., 1987; Brand and Kleineke, 1996) may render S-ASM partially or fully Zn2+-independent under specific conditions. For example, the cysteine-rich protein metallothionein acts as a temporary cellular reservoir of free zinc. Metallothionein may keep cellular concentrations very low or release zinc to intracellular zinc-dependent enzymes in a process that is dynamically controlled by its interactions with the glutathione redox couple (Jiang et al., 1998). Consequently, exogenous Zn2+ ions are not essential for endothelial cell secreted S-ASM activity and cause only 2-fold stimulation (Marathe et al., 1998).

Activity at neutral pH

Given the optimal reaction rate of the enzyme in an acidic range of pH 5.0–5.5 (Spence et al., 1989; Mühle et al., 2013), activity in the neutral pH of blood or CSF of 7.4 appears counterintuitive. Indeed, the ability of S-ASM to hydrolyze SM in a neutral milieu remains controversial and has led some authors to refer to S-ASM as ‘secretory sphingomyelinase’, omitting the specification ‘acidic’ despite its proven origin from the ASM-encoding gene SMPD1. However, this name would only be unambiguous in the absence of any other reported sphingomyelinase species, e.g., neutral or alkaline enzymes that could potentially be detectable as secreted or otherwise released proteins in the extracellular space, which is currently the case (Milhas et al., 2010).

Data concerning the extent of S-ASM activity at neutral pH are inconsistent. Both S-ASM activities in saliva and tear fluid peak at pH 4–5 and are markedly decreased at pH 6, with negligible residual activities at pH 7 (Takahashi et al., 2000). This behavior corresponds well to the pH profiles described for tissue L-ASM (Schneider and Kennedy, 1967; Bowser and Gray, 1978), ASM released from human fibroblasts (optimal pH 4.4) and mouse L-cells (pH 4.8) (Weitz et al., 1983) and our data for S-ASM in human CSF (Mühle et al., 2013), serum, plasma, saliva and urine, which indicate <5% of maximal Zn2+-dependent activity at pH 6.5 and nearly background levels at pH 7.0 (C. Mühle, unpublished data). Purified recombinant human ASM from CHO cells binds tightly to SM at pH 4.0, while binding is not detectable at pH 8.0 (He et al., 1999). By contrast, S-ASM from fetal bovine serum (Spence et al., 1989) or recombinant ASM species (Mintzer et al., 2005) exhibit clearly detectable hydrolysis at a neutral pH of 7.4 of up to 20% of that at the optimal acidic pH. This discrepancy may be attributable to the different characteristics of human endogenous vs. purified (Spence et al., 1989) bacterial or recombinant, sometimes tagged, S-ASM (Mintzer et al., 2005) species utilized in these experiments, different reaction conditions, substrates and purity or glycosylation patterns (Jenkins et al., 2009) of the enzymes. Interestingly, neutral sphingomyelinase C from Staphylococcus aureus selectively catalyzes the hydrolysis of SM in the plasma membrane (Slotte et al., 1990). Its application to hippocampal slices results in the generation of long-chain ceramides and in a positive reversible regulation of neuron excitability via a sphingosine-1-phosphate-mediated mechanism (Norman et al., 2010). Because extracellular ASM catalyzes comparable reactions at the plasma membrane, similar activity can be expected.

Although the biological role of S-ASM is not precisely clear, the enzyme has been implicated in the metabolism of lipoprotein-bound SM to ceramide, the subsequent aggregation and retention of LDL particles and the acceleration of lesion progression as key initial steps in atherogenesis (Marathe et al., 2000a; Tabas et al., 2007; Devlin et al., 2008). After observing that SM-degrading activity of several arterial wall cell types is responsible for subendothelial retention and aggregation of atherogenic lipoproteins in rabbit and human atherosclerotic lesions (Schissel et al., 1996b), Schissel et al. (1998a) identified this activity as a Zn2+-dependent S-ASM capable of hydrolyzing SM associated with atherogenic lipoproteins at a neutral pH. Signs of SM hydrolysis in lesional LDL (Öörni et al., 1998) as well as ASM-induced LDL aggregation (Walters and Wrenn, 2011), depending on the sphingomyelinase-to-LDL molar ratio (Guarino et al., 2006), confirmed the role of ASM and extended its promotion of aggregation to small very-low-density and intermediate-density lipoprotein particles (Öörni et al., 2005). However, another group detected sphingomyelinase activity as an intrinsic property of the multifunctional LDL constituent apolipoprotein B-100 based on the sequence homology of the apoliprotein B-100 α2 domain with bacterial sphingomyelinase or at least as very tightly associated with LDL and absent in oxidized LDL or other lipoproteins (Holopainen et al., 2000; Kinnunen and Holopainen, 2002). A number of atherogenesis-associated modifications of lipoprotein-bound SM appear to increase its susceptibility to hydrolysis: while native LDL is only hydrolyzed and aggregated at acidic pH, oxidation, treatment with phospholipase A2, or enrichment with apolipoprotein CIII promotes prompt hydrolysis at pH 7.4 (Schissel et al., 1998a). LDL particles undergo lipolysis by ASM more readily after prior proteolysis by the extracellular hydrolases chymase and cathepsin S, and this pre-treatment was even essential for the action of the secretory phospholipases A2 (sPLA2-V). This sensitization of LDL by proteolysis may enhance extracellular accumulation of LDL-derived lipids during atherogenesis (Plihtari et al., 2010). Moreover, a high SM-to-phosphatidylcholine ratio appears to promote hydrolysis and aggregation by ASM at neutral pH, as observed for plasmatic lipoproteins from apolipoprotein E KO mice, which are prone to atherosclerosis (Schissel et al., 1998a). Similarly, the high SM content in the slowly cleared remnant lipoproteins of these mice enhances their susceptibility to ASM (Jeong et al., 1998). The most prominent difference was a 10-fold increase in S-ASM activity at pH 7.4 in LDL extracted from atherosclerotic lesions compared to that in plasma LDL from the same human donor (Schissel et al., 1998a).

Moreover, extracellular SM degradation by S-ASM could occur in pockets of acidity that are present in inflammatory lesions of the arterial wall (Menkin, 1934) and in artherosclerotic plaques (Naghavi et al., 2002; Sneck et al., 2012), in proximity to glycosaminoglycans (Maroudas et al., 1988), activated macrophages (Tapper and Sundler, 1992) and osteoclasts (Silver et al., 1988) or close to the cell surface, e.g., in acidic microdomains located predominantly in the distal dendrites of oligodendrocytes (Ro and Carson, 2004). Such acidic microenvironments have been linked to lipid microdomains (Steinert et al., 2008) as the site of optimal ASM action. The translocation of lysosomal V1 H+-ATPase to the cell membrane is a critical contribution to the formation of a local acid microenvironment to facilitate membrane ASM activation and, consequently, redox signalosome formation in coronary arterial endothelial cells (Xu et al., 2012). In addition, substrate binding but not catalytic velocity are predominantly mediated by pH (Callahan et al., 1983). S-ASM may also be active in acidic endosomes following endocytosis of the enzyme by cell surface receptors (Kornfeld, 1987).

Human plasma itself exerts an inhibitory effect on the reaction if sample volumes >2% of the total assay volume are used. This inhibition is only partially explained by the presence of low amounts of EDTA originating from anticoagulation vials. Inhibition has also been observed to a lesser degree for lithium-heparin-plasma as well as serum (C. Mühle, unpublished data). The effect is not observed for CSF but is expected to limit the SM-hydrolyzing activity of S-ASM in the blood in vivo.

Endogenous inhibition of ASM activity

The activity of S-ASM is inhibited by a number of endogenous agents, including adenosine monophosphate and high concentrations of ZnCl2 (6 mm) (Schissel et al., 1996a; Jenkins et al., 2010), as well as in a time- and dose-dependent manner by dithiothreitol and beta-mercaptoethanol (He et al., 1999). Interestingly, S-ASM appears to be less stable than L-ASM at elevated temperatures (Schissel et al., 1996a). Inorganic phosphate, other nucleotides, dolichol phosphate and phosphoinositides are potent noncompetitive inhibitors of ASM (Callahan et al., 1983; Watanabe et al., 1983). Interestingly, while phosphatidylinositol 3,5-bisphosphate (Kölzer et al., 2003) and phosphatidylinositol 3,4,5-trisphosphate (Testai et al., 2004) both inhibit ASM in the low mm range, the enzyme is potently activated by other phosphate-containing lipids such as endocytosed bismonoacylgycerophosphate (Linke et al., 2001) or by plasma membrane-derived phosphatidylglycerol and phosphatidic acid (Oninla et al., 2014). Cholesterol, another endogenous lipid, potently inhibits ASM (Reagan et al., 2000). Removal of lipoproteins from the culture medium of fibroblasts obtained from NPD-C patients who suffer from an additional secondary ASM deficiency restored the activity of ASM (Thomas et al., 1989). The presence of sterols is likely responsible for the inhibitory effect on ASM activity, which may be relevant to inhibitor development because various members of the sterol class, such as 7-ketocholesterol, exhibit inhibitory potential in vitro (Maor et al., 1995).

Role of ASM in microorganism infection

Viral, bacterial and parasitic pathogens are assumed to induce changes in the composition and structure of membranes and microdomains therein to permit the invasion of epithelial cells. The triggered translocation of ASM from endolysosomal compartments to the host cell surface followed by SM hydrolysis to form ceramide-rich domains seems to play a key role in microbial internalization (Riethmüller et al., 2006). ASM has been associated with infection and host defense against Pseudomonas aeruginosa (Grassmé et al., 2003), Listeria monocytogenes (Utermöhlen et al., 2003) and Cryptosporidium parvum (Nelson et al., 2006). ASM mediates the entry of Neisseria gonorrhoeae into nonphagocytic epithelial (Grassmé et al., 1997) and phagocytic (Hauck et al., 2000) cells via the enrichment of CEACAM receptors for the bacterium in ceramide domains, thus functioning as a portal of microbial entry. Infection of macrophages with Salmonella typhimurium induces a dynamic change in the cellular distribution of ASM, with a reduction in L-ASM and a surge in S-ASM activity (McCollister et al., 2007). L-ASM is rapidly activated during the entry of sindbis virus in neuroblastoma cells (Jan et al., 2000). Viruses such as human immunodeficiency virus (Popik et al., 2002) and rhinoviruses (Grassmé et al., 2005) appear to employ sphingomyelin-enriched microdomains to invade epithelial cells. SM functions as a novel receptor for the entry of Helicobacter pylori vacuolating cytotoxin into epithelial cells, and treatment with sphingomyelinase to deplete plasma membrane SM significantly reduces the sensitivity of cells to the pathogen (Gupta et al., 2008). Surface-localized ASM activity and the presence of SM are required for the efficient infection of cells by Ebola virus (Miller et al., 2012). Moreover, there is evidence that pathogens utilize their own sphingolipid repertoire as virulence factors, such as the sphingomyelinase of Streptococcus aureus (Cohen and Barenholz, 1978), or continue to modulate the sphingolipid metabolism of the host after internalization to replicate, survive, and finally infect neighboring cells (reviewed in Zeidan and Hannun, 2007b). An understanding of these interaction processes is a prerequisite for future antimicrobial therapies. In addition, ASM is required for the proper fusion of late phagosomes with lysosomes to efficiently transfer lysosomal antibacterial hydrolases into phagosomes (Schramm et al., 2008).

Involvement of ASM in other cellular processes

In erythrocytes, the activity of sphingomyelinase induces a transition from a discoid to spherical shape, followed by phosphatidylserine exposure and finally a loss of cytoplasmic content (Dinkla et al., 2012). In the context of elevated S-ASM levels in sepsis (Claus et al., 2005; Kott et al., 2014), the resulting enhanced erythrocyte clearance is likely to contribute to anemia. The sensitivity of erythrocytes increases further during blood bank storage and physiological aging (Dinkla et al., 2012).

In contrast to the vast number of experimental conditions that result in increased ASM activity, only one study has reported a condition leading to a reduction of ASM activity. Mechanical injury-triggered ASM secretion is reduced by 70% in C2C12 myoblasts upon knockdown of dysferlin expression, thus mimicking the cells of patients with the progressive muscle wasting disease dysferlinopathy. The defect in cell membrane repair in the immortalized patient myoblasts was fully reversed by treating the cells with extracellular ASM (Defour et al., 2014).

While the ASM/ceramide pathway has been well established as a bona fide system in the chemotherapeutic treatment of cancers and many commonly used chemotherapeutic agents mediate cell death via ASM-induced ceramide formation [reviewed in Henry et al., (2013)], recent evidence (Don et al., 2014) suggests that reducing ASM activity with functional inhibitors such as desipramine may induce lysosomal rupture in cancer cells (Petersen et al., 2013). Although unexpected with respect to reduced SM levels in cancerous compared to normal tissue (Hendrich and Michalak, 2003; Barceló-Coblijn et al., 2011), ASM activity is down-regulated in cancer (Petersen et al., 2013) and may consequently sensitize these cells to further decreases in enzyme activity levels. Lysosomal stability and integrity requires L-ASM activity (Kirkegaard et al., 2010), while the cytoprotective properties of ASM down-regulation in cancer cells are assumed to relate to the secreted form required for vesicle and membrane turnover (Magenau et al., 2011). In the context of reduced SM levels in cancer, the potent antitumor agent 2-hydroxyoleic acid selectively kills cancer cells by stimulating SM synthases, leading to strong augmentation and restoration of the SM concentration (Barceló-Coblijn et al., 2011).

In vivo effects of S-ASM activity on sphingolipid levels

S-ASM activity in plasma or serum is increased under various pathological conditions in humans (Figure 2) and animal models. This increase should result in a reduced concentration of SM and an increased concentration of ceramide. In addition, a negative correlation between S-ASM and SM and a positive correlation between S-ASM and ceramide should be observed. However, such phenomena have been observed in only a few studies. Jenkins et al. (2013) observed increased S-ASM activity in serum with a parallel increase in different ceramide species in patients with lymophohistiocytosis. A particularly large increase in C16-ceramide was observed, and this result was compatible with cell-based studies in which a specific increase in C16-ceramide via S-ASM was noted (Jenkins et al., 2010). Increased S-ASM activity and parallel increases in ceramide levels were observed by Pan et al. (2014) in patients with heart disease and by Hammad et al. (2012) in patients with post-traumatic stress disorder. Irradiation induces increases in S-ASM activity in some but not all tumor patients, and a positive correlation between S-ASM activity and serum ceramide concentration has been observed (Sathishkumar et al., 2005). In rodents, increasing age is associated with increased S-ASM activity and a simultaneous increase in the concentration of individual ceramide species (Kobayashi et al., 2013). Other in vivo studies did not report the expected relationship between S-ASM activity and sphingolipid concentrations. As expected, S-ASM activity is elevated in patients with non-alcoholic fatty liver disease, with a parallel increase in the concentration of ceramide. However, S-ASM activity is increased even further in patients with hepatitis C, although no concomitant change in the concentration of ceramide was observed (Grammatikos et al., 2014). In patients with Alzheimer’s disease, the activity of S-ASM is increased and the concentration of SM is reduced in the plasma, while ceramide levels are unchanged (Lee et al., 2014). S-ASM activity is elevated in the plasma of APP/PS1 mice, while the concentrations of SM and ceramide are unchanged (Lee et al., 2014). Feeding rats grape seed oil vs. butter leads to increased S-ASM activity and, simultaneously, decreased ceramide concentrations (Drachmann et al., 2007). The induction of S-ASM activity by adeno-associated virus in ApoE-/- mice resulted in only minor changes in SM and ceramide: a small reduction of SM and ceramide in plasma at several time points after injection and a reduction of SM and unaltered ceramide concentrations in the liver (Leger et al., 2011). Correlations between S-ASM in plasma SM and ceramide were previously reported only sporadically (Sathishkumar et al., 2005). In the literature, the ratio between SM and ceramide has been used as an indirect measure of S-ASM activity (Drobnik et al., 2003); such an approach is questionable in light of the data described above.

Relative S-ASM activities in plasma or serum across different studies in humans. In clinical studies, the activity values in the control population were set as 100%; in genetic studies, activities of homocygotes for the major allele were set as 100%. The activities range from approximately 0.2% in NPD-B to nearly 2000% in lymphohistiocytosis, thus representing four orders of magnitude. Severe reductions as well as elevations of peripheral S-ASM activities appear to be associated with human diseases and occur via different mechanisms. Decreased S-ASM activities are caused by genetic mechanisms: NPD-B involves mutations in the SMPD1 gene. However, heterozygous gene carriers also exhibit reduced S-ASM activity. Common or rare polymorphisms in the SMPD1 gene (rs1050239, rs141641266) and a shorter repeat length in the signal peptide also lead to reduced peripheral S-ASM activity. Because of the association of the rs1050239 polymorphism with allergy, slightly decreased values of peripheral S-ASM are found in allergy sufferers. Elevated levels of S-ASM are found in a variety of diseases, such as Alzheimer’s disease, heart disease, diabetes mellitus II, alcohol dependence and severe inflammatory diseases such as hepatitis C, sepsis, systemic inflammatory response syndrome, inflammatory renal disease, systemic vasculitis and lymphohistiocytosis. The mechanisms underlying these elevations of S-ASM include enhanced release of the enzyme and activation of the enzyme by oxidative stress and cytokine activation. Reduced S-ASM activity is mediated by genetic mechanisms and must therefore be regarded as a trait marker, while increased activity is mediated by cytokines, LPS or oxidative stress and must therefore be regarded as a state marker of a condition. Please note that the values obtained in different studies are not directly comparable because S-ASM activities were determined using different methods and different reference values. SIRS, systemic inflammatory response syndrome; NPD, Niemann-Pick disease. Literature (in alphabetical order of the description in the Figure): acute myocardial infarction (Pan et al., 2014), alcohol detoxification (Reichel et al., 2011), allergy (Reichel et al., 2014), Alzheimer’s disease (Lee et al., 2014), chronic heart failure (Pan et al., 2014), diabetes mellitus II (Górska et al., 2003), hepatitis C (Grammatikos et al., 2014), inflammatory renal disease (Kiprianos et al., 2012), lymphohistiocytosis (Takahashi et al., 2002; Jenkins et al., 2013), non-alcoholic fatty liver (Grammatikos et al., 2014), NPD-B (Abe et al., 1999; He et al., 2003), NPD-B carrier (He et al., 2003), post-operative (Kott et al., 2014), post-traumatic stress disorder (Hammad et al., 2012), rs1050239 (Reichel et al., 2014), rs141641266 (Rhein et al., 2013), sepsis (Claus et al., 2005), signal peptide repeat 4/4 (Rhein et al., 2014), SIRS (Kott et al., 2014), stable angina pectoris (Pan et al., 2014), systemic vasculitis (Kiprianos et al., 2012), unstable angina pectoris (Pan et al., 2014), and Wilson disease (Lang et al., 2007).
Figure 2:

Relative S-ASM activities in plasma or serum across different studies in humans.

In clinical studies, the activity values in the control population were set as 100%; in genetic studies, activities of homocygotes for the major allele were set as 100%. The activities range from approximately 0.2% in NPD-B to nearly 2000% in lymphohistiocytosis, thus representing four orders of magnitude. Severe reductions as well as elevations of peripheral S-ASM activities appear to be associated with human diseases and occur via different mechanisms. Decreased S-ASM activities are caused by genetic mechanisms: NPD-B involves mutations in the SMPD1 gene. However, heterozygous gene carriers also exhibit reduced S-ASM activity. Common or rare polymorphisms in the SMPD1 gene (rs1050239, rs141641266) and a shorter repeat length in the signal peptide also lead to reduced peripheral S-ASM activity. Because of the association of the rs1050239 polymorphism with allergy, slightly decreased values of peripheral S-ASM are found in allergy sufferers. Elevated levels of S-ASM are found in a variety of diseases, such as Alzheimer’s disease, heart disease, diabetes mellitus II, alcohol dependence and severe inflammatory diseases such as hepatitis C, sepsis, systemic inflammatory response syndrome, inflammatory renal disease, systemic vasculitis and lymphohistiocytosis. The mechanisms underlying these elevations of S-ASM include enhanced release of the enzyme and activation of the enzyme by oxidative stress and cytokine activation. Reduced S-ASM activity is mediated by genetic mechanisms and must therefore be regarded as a trait marker, while increased activity is mediated by cytokines, LPS or oxidative stress and must therefore be regarded as a state marker of a condition. Please note that the values obtained in different studies are not directly comparable because S-ASM activities were determined using different methods and different reference values. SIRS, systemic inflammatory response syndrome; NPD, Niemann-Pick disease. Literature (in alphabetical order of the description in the Figure): acute myocardial infarction (Pan et al., 2014), alcohol detoxification (Reichel et al., 2011), allergy (Reichel et al., 2014), Alzheimer’s disease (Lee et al., 2014), chronic heart failure (Pan et al., 2014), diabetes mellitus II (Górska et al., 2003), hepatitis C (Grammatikos et al., 2014), inflammatory renal disease (Kiprianos et al., 2012), lymphohistiocytosis (Takahashi et al., 2002; Jenkins et al., 2013), non-alcoholic fatty liver (Grammatikos et al., 2014), NPD-B (Abe et al., 1999; He et al., 2003), NPD-B carrier (He et al., 2003), post-operative (Kott et al., 2014), post-traumatic stress disorder (Hammad et al., 2012), rs1050239 (Reichel et al., 2014), rs141641266 (Rhein et al., 2013), sepsis (Claus et al., 2005), signal peptide repeat 4/4 (Rhein et al., 2014), SIRS (Kott et al., 2014), stable angina pectoris (Pan et al., 2014), systemic vasculitis (Kiprianos et al., 2012), unstable angina pectoris (Pan et al., 2014), and Wilson disease (Lang et al., 2007).

It is unclear why the available in vivo data on the relationship between S-ASM activity and sphingolipid concentrations are so heterogeneous and predominantly negative. Various explanations may be considered. (1) The measurement of total sphingolipids easily overlooks significant and relevant changes in subspecies. This is exemplified in the work of Jenkins et al. (2013). While the total ceramide concentration is moderately increased in patients with lymphohistiocytosis, the analysis of single ceramide species reveals a more differentiated picture, with highly elevated concentrations of C16-ceramide but decreased concentrations of C24-ceramide. (2) If relevant changes in sphingolipid concentrations occur only in small subcompartments, they may be missed in the overall analysis. (3) Ceramides may rapidly enter different metabolic pathways after formation (the enzymes involved include sphingomyelin synthase, ceramidase, glucosylceramide synthase, and ceramide kinase). The measurement of static concentrations provides little information about turnover. Therefore, future studies should analyze not only total sphingolipids but sphingolipid species at greater spatial and temporal resolution and assess the activities of the enzymes involved.


The enzymes involved in sphingolipid metabolism are considered important therapeutic targets (Billich and Baumruker, 2008; Arenz, 2010; Kornhuber et al., 2010). A pronounced increase in the activity of S-ASM would have obvious negative clinical consequences, spurring interest in the development of inhibitors of S-ASM. In addition to classifications based on endogenous and exogenous inhibitors (Canals et al., 2011), we can also distinguish direct, competitive inhibitors from FIASMAs. An indirect mechanism of irreversible inactivation or degradation of cellular ASM and lipidosis by cationic amphiphilic drugs was assumed in the early 1980s (Albouz et al., 1981, 1983; Yoshida et al., 1985) but was not further pursued directly. FIASMAs such as desipramine or fluoxetine (Kornhuber et al., 2008, 2010, 2011) accumulate in acidic intracellular environments (Trapp et al., 2008) and induce a detachment of L-ASM from the inner lysosomal membrane by shielding the abundant negatively charged bis(monoacylglycero)phosphate, resulting in subsequent proteolytic degradation of L-ASM (Hurwitz et al., 1994a; Kölzer et al., 2004b). Interestingly, treatment with the conserved chaperone heat shock protein 70, which binds to endolysosomal bis(monoacylglycero)phosphate to provide lysosomal stability, effectively corrects the NPD phenotype in vitro (Kirkegaard et al., 2010). In agreement with this model, the prototypical FIASMA desipramine does not affect purified ASM in in vitro micellar assays (Yoshida et al., 1985) or S-ASM in a cell culture system (Jenkins et al., 2011b). However, another report suggested an inhibition of endogenous as well as TNFα-stimulated secretion of recombinant S-ASM by desipramine in stably transfected MCF7 cells (Jenkins et al., 2011a). Thus, it is conceivable that FIASMAs not only act within acidic intracellular compartments but also extracellularly by another mechanism. Although they possess some degree of selectivity (Kornhuber et al., 2010), these FIASMA drugs are not completely specific for ASM and exhibit inhibitory effects against other lysosomal enzymes, including acid ceramidase (Zeidan et al., 2006).

Potent and selective competitive inhibitors of ASM are rare. In a cell-free drug screen of a wide range of compounds, no drug-like competitive inhibitors of S-ASM were identified (Mintzer et al., 2005). More recently, new competitive inhibitors of ASM have been identified and/or developed (Roth et al., 2009a,b, 2010; Arenz, 2010) that should function against both L-ASM and S-ASM. Available compounds include substrate analogues such as L-carnitine, phosphate analogues such as phosphatidylinositol-3,5-bisphosphate and natural compounds such as α-mangostin (Arenz, 2010). The systemic availability of these molecules remains unclear, and most of these substances are not yet available for clinical use. Competitive inhibitors of ASM that cannot pass through the cell membrane and therefore have no intracellular effects may help clarify the physiological effects of S-ASM relative to L-ASM.

However, there are applications for increasing ASM activity. In addition to obvious therapeutic applications for patients with NPD-A and -B, including enzyme replacement and gene therapy, NPD-C patients may benefit from ASM treatment despite their intact SMPD1 gene. Mutations in NPC1 or NPC2 result in impaired cholesterol trafficking in the endolysosome (Madra and Sturley, 2010) and in a secondary deficiency in ASM activity (Tamura et al., 2006). Restoration of ASM to normal levels in NPC1-deficient cells improves several defects, including a reduction in lysosomal cholesterol (Devlin et al., 2010), underlining the important role of ASM in this disease. Another field of application of ASM is cancer therapy, with the aim of pharmacologically modifying the deregulated sphingolipid metabolism that occurs in many cancers, as evident, for example, in the reduced levels of ASM mRNA. A number of studies have revealed the potential of ASM overexpression or supplementation to either directly kill tumor cells or sensitize them to chemotherapy (Smith and Schuchman, 2008; He and Schuchman, 2012; Savic and Schuchman, 2013).

Animal models

S-ASM in brain function and behavior

The development of ASM KO mice has permitted the investigation of the role of ASM in behavior. Homozygous ASM KO mice develop normally and do not exhibit impairments until they are 8 weeks of age. Mild tremor and locomotor ataxia then emerge. Tottering with zigzag movements becomes evident and is characteristic of cerebellar dysfunction. Between 12 and 16 weeks, ASM KO mice become lethargic and unresponsive to stimuli. They increasingly experience problems feeding and much reduced weight compared to WT mice. Beyond 4 months of age, these animals exhibit severe ataxia and die between 6 and 8 months of age. ASM KO animals exhibit a significant reduction in brain volume. A striking finding was the significant loss of Purkinje cells in the cerebellum and atrophy of the mesencephalon beginning at 10 weeks of age (Horinouchi et al., 1995; Otterbach and Stoffel, 1995). Both regions appear to be involved in movement control and motivated behavior. Homozygous ASM KO mice exhibited reduced ceramide levels in the hippocampus and displayed some signs of reduced anxiety and a reduction of depression-related behaviors. Young ASM KO mice did not exhibit altered synaptic structure or functioning at the level of the hippocampus (Gulbins et al., 2013). Interestingly, heterozygous ASM KO mice, which possess approximately 50% of normal ASM activity, did not exhibit any motor problems, reduced feeding, or enhanced mortality. This finding may suggest that 50% enzyme activity is sufficient for normal brain function and a normal behavioral phenotype (Horinouchi et al., 1995), consistent with the rare phenotype observed in NPD carriers (Lee et al., 2003; Schuchman, 2007). While these findings suggest an important role of ASM in brain function and locomotor behavior, it was unclear if these effects were mediated by L-ASM or S-ASM because the KO affects both subtypes. In a subsequent study, this question was addressed by fusing the Smpd1 gene with the lysosomal Lamp1 gene and thus targeting all ASM to the lysosome. The mutant mice still exhibited low levels of L-ASM activity (11.5–18.2% of WT) but a complete absence of S-ASM activity. These mice exhibited neither brain pathology nor signs of locomotor impairment. This finding suggests that severe deficits in locomotor function in ASM KO mice are mediated predominantly by a lack of L-ASM, while S-ASM may not be required (Marathe et al., 2000b).

The role of ASM in depression/anxiety has been investigated in animal studies utilizing a transgene for ASM (tgASM). These mice exhibited higher ASM activity and ceramide production in the hippocampus than WT mice (Gulbins et al., 2013). The enhanced ceramide levels in the hippocampus were paralleled by a decline in neurogenesis, neuronal maturation, and neuronal survival (Gulbins et al., 2013), which are depression-related neuronal markers (Santarelli et al., 2003; Krishnan and Nestler, 2008). S-ASM activity was increased more than 10-fold in the CSF of tgASM mice compared to the WT controls (Mühle et al., 2013). At the behavioral level, tgASM mice exhibited a depression/anxiety-like phenotype in several tests (Gulbins et al., 2013); for a review see Kornhuber et al. (2014). This ASM effect is most likely mediated in the brain because the chronic application of C16-ceramide in the dorsal hippocampus of mice induced the same depression-like behavioral phenotype observed in tgASM mice (Gulbins et al., 2013). This finding is consistent with the effects of chronic unpredictable stress, which not only double hippocampal ceramide levels and reduced neurogenesis and neuronal maturation but also induce depression-like behavior in mice (Gulbins et al., 2013). Further support for the role of ceramide is provided by the elevated plasma ceramide levels detected in patients with major depression (Gracia-Garcia et al., 2011). Whether these effects are mediated by S-ASM or L-ASM remains to be determined.

Many antidepressant drugs are functional inhibitors of ASM (Albouz et al., 1983; Kornhuber et al., 2008, 2010, 2011). These drugs can normalize the effects of chronic unpredictable stress in WT and tgASM but not ASM KO mice. This observation suggests that ASM functional inhibition is a major pathway for the pharmacological effects of antidepressants (Gulbins et al., 2013; Kornhuber et al., 2014). This effect, however, predominantly affects L-ASM. Interestingly, evidence suggests that serum zinc levels are significantly lower in patients with major depression (Levenson, 2006; Swardfager et al., 2013a). Low dietary zinc intake has been associated with a greater incidence of depression (Vashum et al., 2014), and an antidepressant effect of zinc supplementation has been observed in laboratory rodents (reviewed in Levenson, 2006, and Swardfager et al., 2013b) and in human trials (Lai et al., 2012; Swardfager et al., 2013b). While the underlying mechanism remains a subject of speculation, a possible connection to the stimulatory effects of zinc on S-ASM activity has not been explored.

S-ASM and atherosclerosis

Inflammation plays an important role in the development of atherosclerosis. In mice, inflammation induced by lipopolysaccharide (LPS) increases S-ASM activity in the serum 3 h after injection. This increase was dependent on IL-1 activity. Serum S-ASM was also increased in mice that were treated with IL-1β or TNFα. These findings raise the possibility that an increase in S-ASM activity contributes to inflammatory cytokine activity in atherosclerosis (Wong et al., 2000). The ASM activity present in atherosclerotic lesions (Schissel et al., 1996b) is dependent on S-ASM and is associated with the extracellular arterial wall. In this bound state, S-ASM retains enzymatic activity that stimulates subendothelial retention and aggregation of atherogenic lipoproteins (Marathe et al., 1999). When mice are fed an atherogenic diet containing saturated fats and cholesterol, serum S-ASM activity is up-regulated, which facilitates the conversion of SM to ceramide (Deevska et al., 2012). However, another study found increased activity of S-ASM in rats fed for 14 weeks with a diet rich in n-3 polyunsaturated fatty acids (PUFA) vs. butter, and a strong negative correlation was observed between the serum concentration of n-3-PUFA and S-ASM activity (Drachmann et al., 2007). The association of atherosclerosis with increased age is mediated, at least in part, by S-ASM. S-ASM is increased in mice at 65 weeks of age. Apolipoprotein E-/- (ApoE-/-) mice develop atherosclerosis at 15 weeks of age. Plasma ceramide levels are generally higher in ApoE-/- than WT mice but decrease with age in parallel with an increase in S-ASM activity. When ApoE-/- mice develop atherosclerosis, several ceramide species increase in the aorta compared to WT mice: C18:0, C22:0 and C24:0. At 65 weeks of age, C16:0 and C24:1 species are increased compared to WT mice. These findings indicate that an age-related increase in S-ASM activity, which results in an elevation of certain ceramide species, may contribute to age-related atherosclerosis (Kobayashi et al., 2013). When ApoE-/- mice are treated with a recombinant adeno-associated virus (AAV) that constitutively expresses high levels of human ASM in the liver and plasma, S-ASM levels are persistently elevated. However, this phenomenon leads to only a short reduction in plasma SM and no changes in serum lipoprotein levels. Plaque formation in the aortic sinus after 17 weeks did not differ in mice treated with a control AAV. These findings suggest that S-ASM has no accelerating or exacerbating role in atherosclerotic lesion formation in the ApoE-/- model of atherosclerosis (Leger et al., 2011).

S-ASM in diabetes

Diabetes mellitus, a metabolic disorder characterized by hyperglycemia, is associated with changes in ceramide metabolism. The induction of a diabetic state with streptozotocin causes a significant increase in blood glucose in rats. Eight weeks after induction, plasma and renal ceramide levels appear to increase. While neutral sphingomyelinase levels are unchanged in the liver and kidney, plasma S-ASM activity increases. These findings suggest that the induction of diabetes increases plasma ceramide levels by enhancing S-ASM activity (Kobayashi et al., 2013).

S-ASM in sepsis

Early studies show that ASM KO mice are protected against effects of LPS (Haimovitz-Friedman et al., 1997). However, the differential role of L- and S-ASM cannot be derived from these studies. Claus et al. (2005) find increased S-ASM activity 24 h after endotoxin application in mice; this increase did not occur after pre-treatment of mice with the FIASMA NB6. In a subsequent study, the group demonstrates an increase of the S-ASM activity after induction of sepsis in WT mice, but not in ASM KO mice (Jbeily et al., 2013). ASM KO mice exhibit enhanced basal ceramide levels in the blood but a significantly reduced ceramide response when sepsis is experimentally induced by peritoneal contamination and infection. There is an increase in bacterial burden, increased phagocytic activity and an enhanced cytokine storm in the blood after sepsis induction in ASM KO mice. Thus, in murine models of sepsis, S-ASM activity is increased, which may contribute to the effective defense of septic pathogens. S-ASM triggered hydrolysis of membrane-bound SM may play an important role in the primary defense against invading microorganisms (Jbeily et al., 2013).

S-ASM in pulmonary edema

In the context of non-cardiogenic pulmonary edema, treatment of mice with platelet-activating factor (PAF) leads to a 50% increase in lung tissue concentrations of ceramides, and this increase is thought to contribute to the development of severe edema (Göggel et al., 2004). However, in contrast to the PAF-stimulated transient accumulation of ceramide in P388D1 macrophages by de novo synthesis (Balsinde et al., 1997), this increase in ceramide is attributed to the PAF-induced release of ASM from the lungs, which results in a 28% increase in strictly Zn2+-dependent serum ASM activity without altering the activities of neutral sphingomyelinase or ceramide synthase. The application of ceramide-specific antisera, pharmacological disruption of ceramide formation or functional inhibition of ASM all specifically prevent PAF-triggered pulmonary edema. These data corroborate the hypothesis that PAF-induced pulmonary edema is mediated by ASM and ceramide as the second major pathway in addition to the activation of EP-3-receptors by PAF (Göggel et al., 2004).

A slow, modest but significant increase in S-ASM but not L-ASM or neutral sphingomyelinase was also noted in a pulmonary emphysema mouse model after VEGF receptor blockade in which the prompt and up to 3-fold activation of ceramide synthase was the main contributor to increased lung ceramide levels associated with alveolar septal destruction (Petrache et al., 2005). The authors speculate that, due to the high abundance of S-ASM from endothelial cells, injured lung capillary endothelial cells aggravate cellular damage in a vicious cycle, resulting in the sustained lung damage observed in patients despite smoking cessation. In a neonatal pigled model of pulmonary inflammation, S-ASM activity and ceramide concentration are reduced in serum after treatment of lungs with inhaled surfactant factor combined with the FIASMA imipramine (von Bismarck et al., 2008; Preuß et al., 2012).

Wilson disease, an autosomal recessive disorder, is characterized by an accumulation of Cu2+, which leads to severe consequences including progressive liver cirrhosis, neurological and psychiatric symptoms and, occasionally, anemia. Patients with Wilson disease exhibit a constitutive increase in ASM activity in the blood plasma even during treatment with D-penicillamine or trientine, which decrease total but not free Cu2+ concentrations (Lang et al., 2007). In isolated murine hepatocytes or cell lines, Cu2+ activates ASM but not neutral sphingomyelinase and triggeres ceramide release within 10–20 min after application. Both effects were absent or reduced in ASM KO cells and cells preincubated with the antioxidants N-acetylcysteine and Tiron, suggesting that Cu2+ may directly or indirectly stimulate ASM via oxygen radicals. In whole blood and in purified mouse leukocytes, Cu2+ treatment results in the stimulation and secretion of ASM into the supernatant or plasma, followed by apoptosis-like changes including shrinkage of erythrocytes and phosphatidylserine exposure. In patients, increased plasma ASM activity also correlates with a constitutively increased number of erythrocytes exposing ceramide or phosphatidylserine on the cell surface, which may mediate the rapid clearance of erythrocytes and thus result in anemia. The causal connection between Wilson disease symptoms and ASM is supported by the observation of a delayed onset of disease and protection against the development of fibrosis and liver failure in a rat model of Wilson disease treated with amitriptyline as a pharmacological inhibitor of ASM (Lang et al., 2007).

Clinical studies

Cross-sectional clinical studies

Several studies have investigated peripheral S-ASM activities in humans under various conditions. Most of these studies analyzed serum or plasma. Some studies have also examined urine, salivary fluid, tear fluid, synovial fluid (Takahashi et al., 2000) and CSF (Mühle et al., 2013). Figure 2 summarizes all available studies using plasma or serum that also included a control group. The consideration of relative changes allows comparisons between different types of disorders. However, the studies used different methods, and S-ASM activity was related to a certain volume in most studies but to the amount of protein in others. Many of these studies suffer from the limitation of a small number of investigated cases. Furthermore, the study designs are heterogeneous; some studies used a retrospective design, while other studies used a prospective design. However, Figure 2 illustrates the variation of S-ASM in different clinical situations. Reduced activity values arise because of genetic variations in the SMPD1 gene. Very low activity values (<5%) are measured in NPD-B patients (Abe et al., 1999; He et al., 2003). Heterozygous NPD-B gene carriers exhibit intermediate S-ASM activity levels (≈20%) (He et al., 2003) (the NPD patients reported in the publications by Takahashi et al. (2000, 2002) are likely identical to those reported in Abe et al. (1999) and were therefore omitted in Figure 2). Only slightly reduced activity values (40–90%) are observed for the polymorphisms rs1050239 (Reichel et al., 2014) and rs141641266 (Rhein et al., 2013) and short repeat lengths in the signal peptide (Rhein et al., 2014). Because the minor A allele of polymorphism rs1050239 is associated with allergy, slightly reduced values of peripheral S-ASM are also observed in allergy patients (Reichel et al., 2014). Based on current information, decreased S-ASM activity is genetically determined and must therefore be regarded as a trait marker.

Until now, increased S-ASM activity has not been associated with genetic changes in the SMPD1 gene but has been related to the enhanced release of the enzyme and enhanced activation via oxidative stress, inflammation and cytokines (Marathe et al., 1998; Wong et al., 2000; Jenkins et al., 2010). Increased activity of S-ASM is induced by LPS, cytokines or oxidative stress and must therefore be regarded as a state marker of these conditions. The data in Figure 2 show no clear boundaries, and thus up to 200% activation of S-ASM is considered a mild increase. Mildly elevated activities are observed in Alzheimer’s disease (Lee et al., 2014), type II diabetes mellitus (Górska et al., 2003), stable angina pectoris, acute myocardial infarction (Pan et al., 2014), chronic heart failure (Doehner et al., 2007), post-traumatic stress disorder (Hammad et al., 2012) and in response to ionizing radiation treatment in cancer patients (Sathishkumar et al., 2005). Marked increases in activity (>200%) are observed in unstable angina pectoris (Pan et al., 2014), Wilson disease (Lang et al., 2007), sepsis (Claus et al., 2005), systemic inflammatory response syndrome (Kott et al., 2014), lymphohistiocytosis (Takahashi et al., 2002; Jenkins et al., 2013), non-alcoholic fatty liver, hepatitis C (Grammatikos et al., 2014), systemic vasculitis, inflammatory renal disease (Kiprianos et al., 2012) and in patients with alcohol dependency undergoing withdrawal (Reichel et al., 2011). In sepsis, lymphohistiocytosis, systemic inflammatory response syndrome and chronic heart failure, high S-ASM activity is associated with higher mortality (Claus et al., 2005; Doehner et al., 2007; Jenkins et al., 2013; Kott et al., 2014). In accordance with this, the plasma ceramide content is an independent risk factor for mortality in patients with chronic heart failure (Yu et al., 2015). High S-ASM activity has been related to reduced skeletal muscle strength (Doehner et al., 2007). In several studies, a negative correlation between S-ASM activity and body mass index (BMI) has been observed (Doehner et al., 2007; Reichel et al., 2011; Mühle et al., 2014), while this has not been detected in other studies (Grammatikos et al., 2014). In patients with chronic heart failure, S-ASM activity increases with age (Doehner et al., 2007), in agreement with studies conducted in mice (Kobayashi et al., 2013). By contrast, Grammatikos et al. (2014) did not observe a correlation between age and S-ASM activity. A diet rich in n-3 PUFAs reduces S-ASM activity in rats (Drachmann et al., 2007) but not in humans (measured in samples from the study by Rees et al. (2006); Lars I. Hellgren, personal communication). Indirect evidence for increased S-ASM activity based on an elevated ceramide to SM ratio in septic patients has been reported (Drobnik et al., 2003). In the plasma or CSF of patients with Alzheimer’s disease, increases in this ratio have been associated with greater cognitive progression (Mielke et al., 2011). Although increased ceramide levels could originate from an inhibition of ceramide-degrading enzymes (Satoi et al., 2005), aberrantly high concentrations of total and individual ceramide species correlate with an up-regulation of ASM transcription in patients with neurodegeneration (Filippov et al., 2012).

L-ASM activity is reduced in the lysosomes of cells from patients with I-cell disease (mucolipidosis II) who are deficient in N-acetylglucosaminyl-1-phosphotransferase activity and thus unable to add mannose 6-phosphate residues to enzymes targeted to the lysosome, including L-ASM (Wenger et al., 1976; Weitz et al., 1983). Although either increased S-ASM activity in the plasma of these patients or intracellular retention would be expected, the fate and localization of the assumed excessive enzymes remain unknown.

Longitudinal clinical studies

Several clinical studies have longitudinally investigated the activity of S-ASM, yielding interesting conclusions regarding the value of S-ASM as a biomarker. In contrast to a pronounced decrease in C-reactive protein (CRP), S-ASM activity only normalizes to a minor extent within 2 weeks in patients with systemic vasculitis undergoing immunosuppressive therapy, resulting in a dissociation of S-ASM activity and CRP-levels during the treatment of these patients (Kiprianos et al., 2012). Increased S-ASM activity was measured at baseline in male alcohol-dependent patients entering an alcohol withdrawal program. S-ASM activity steadily declined from day 0 (baseline) to days 1 and 2 and nearly reached control levels on days 7–10 (Reichel et al., 2011). Similar results were reported in an independent sample of alcohol-dependent subjects, with males exhibiting higher initial S-ASM activities compared to females (Mühle et al., 2014). In a single patient with lymphohistiocytosis, S-ASM activities slowly declined over several weeks in response to active treatment with dexamethasone (Takahashi et al., 2002). In patients with unstable angina pectoris, high S-ASM activity was detected on the day of symptom onset; only slightly normalized values were observed on day 1 after onset, and slightly increased S-ASM values were observed on the day of percutaneous coronary intervention. In patients with acute myocardial infarction who underwent conventional treatment, high S-ASM activity was observed up to at least day 7 after onset (Pan et al., 2014). In a group of septic patients, S-ASM activity was increased compared to controls at baseline. S-ASM activity decreased in survivors, while S-ASM activity further increased in non-survivors (Claus et al., 2005). Similarly, in patients with severe systemic inflammation, S-ASM activity predicted mortality in patients with low procalcitonin values (Kott et al., 2014). Serial measurements of S-ASM during the days after abdominal surgery revealed increased S-ASM values that remained high until the end of the observation period at day 5 (Kott et al., 2014). Serial measurements indicated increased ceramide/SM ratios as an indirect measure of S-ASM activity in septic patients at day 11 in non-survivors compared to survivors (Drobnik et al., 2003). These studies all suggest a relatively slow decrease in S-ASM activity in response to treatment for diseases in which S-ASM activity is initially increased. This effect may be related to the slow turnover of S-ASM. In cell culture experiments, S-ASM activity remains high over 24 h despite the inhibition of protein synthesis, whereas L-ASM activity rapidly declines (Jenkins et al., 2010). The slow turnover of S-ASM indicates that small persistent changes in the release of S-ASM result in profound and long-lasting changes in peripheral S-ASM activity. If the pathological condition is removed, such as during alcohol withdrawal treatment (Reichel et al., 2011; Mühle et al., 2014), S-ASM activity slowly decreases to control levels. This underscores the notion of increased ASM activity as a state marker of disease.

S-ASM in clinical routine

Should S-ASM activity be used as a chemistry marker in the clinical routine? A moderate reduction of S-ASM activities per se is not clearly related to the pathophysiology of disease. The association of slightly reduced S-ASM activity with allergy appears to be indirectly mediated via genetic mechanisms; however, it is unlikely that reduced S-ASM activity results in an increased risk of allergy. A severe reduction of S-ASM activity is observed in patients with NPD-B. However, all diseases that are associated with increased S-ASM activity are also associated with mechanisms that lead to activation of S-ASM, namely oxidative stress, increased cytokine production and/or endothelial damage. While ASM and its reaction product ceramide have been implicated in all of these diseases, the specific role of S-ASM in relation to L-ASM remains to be clarified. Diseases with hepatic and endothelial involvement preferentially appear to present a marked elevation of peripheral S-ASM activity. This elevation may be related to the characteristics of the endothelium as a rich source of S-ASM (Marathe et al., 1998). The mechanisms leading to marked elevations of S-ASM activity must be investigated in future studies. Because of the slow turnover of S-ASM, its measured activity may represent not only the current status but also past events.

A determination of peripheral S-ASM activity appears to be useful in suspected NPD patients and carriers and in severe acute inflammatory diseases. Increased S-ASM activity is observed in many different disease states and thus can be used as a general marker that reflects inflammation, cytokine release, oxidative stress and endothelial damage as a pathological state. Although it is not applicable as a biomarker of a single disease, S-ASM levels may be helpful for monitoring disease progression and severity and therapeutic efficiency or for evaluating the prognosis as well as the differentiation between subtypes of disorders with variable involvement of inflammatory processes. In severely diseased patients, S-ASM activity is associated with mortality and low BMI.

Open questions

S-ASM is a potential biomarker and therapeutic target, but important questions remain to be resolved before its inclusion in the clinical routine:

  • The precise origin of S-ASM activity in the blood should be identified to further elucidate the relevance of altered (mainly increased) S-ASM activity under various clinical conditions. Indeed, there seems to be no clear consensus on the subcellular origin of ASM that is translocated to the outer leaflet of the plasma membrane in response to different stimulatory signals: the lysosome, other vesicles or compartments at or near the cell surface or even from cytosolic pools. Similarly, the fate of endocytosed S-ASM has not yet been determined.

  • Many details of regulation and processing, including modifications in the Golgi complex, trafficking, differentiation between L- and S-ASM and activation, remain elusive. Moreover, we have observed higher S-ASM activity in bovine and rodent serum compared to human serum. However, the activity of ASM and related enzymes of sphingolipid metabolism from different species in relation to their evolution and regulation has not been systematically investigated, despite the use of various animal species in models of human diseases.

  • With respect to enzymatic analysis, there are insufficient data regarding reaction requirements for S- (and L-) ASM originating from different sources, including cofactors, inhibitors and pH profiles as well as the substrate specificity such as fatty acid chain length and explanations for so far observed differences are lacking. The reported activation of S-ASM from the cell supernatant, serum and CSF following storage at -20°C is relevant for patient material and clinical studies, but specific mechanisms and temperature profiles are not yet available.

  • A detailed understanding of the mechanisms of pharmacological inhibition of both ASM enzymes, or specifically L- or S-ASM, would facilitate the development of common and selective inhibitors for in vitro experimental work as well as for clinical applications.

  • To date, only a few in vitro and in vivo models are able to separate the effects of L-and S-ASM (Marathe et al., 2000b; Jenkins et al., 2010). The development of further models that permit distinction between S- and L-ASM is urgently needed to study the differential effects and responses of these enzyme species in a variety of normal and pathological conditions. Inducible and tissue-specific knockdown or overexpression of S- or L-ASM will aid in dissecting the biological function of these enzyme species.

  • The methods used to determine S-ASM activities should be standardized to enable comparisons of values across studies and to establish reference values; in the recent literature, different radioactive and fluorescent methods with various conditions, sub-steps and detection techniques have been applied. Other biological materials including saliva, urine, tear fluid as well as synovial or peritoneal fluid could be studied for the correlation of S-ASM activities with that in blood. The potential of these materials as sources of S-ASM as a biomarker remains largely unexplored.

  • The relationships between S-ASM activities and specific clinical features remain to be elucidated for both very low as well as high S-ASM activities. Cut-off values and sensitivity and specificity data for diagnostic (such as NPD) and prognostic purposes (such as mortality) should be established when using S-ASM activity either alone or in combination with other biomarkers.

  • In future clinical studies, known confounders such as alcohol consumption, BMI, and genetic variants should be controlled for. To obtain a complete understanding of sphingolipid turnover, the activities of other enzymes such as L-ASM, neutral sphingomyelinase, ceramidases, sphingosine kinase, ceramide synthases and lipids such as ceramide and sphingosine-1-phosphate should be quantified in parallel to the Zn2+-dependent and Zn2+-independent fractions of S-ASM activity. The levels of inhibitors and activators of S-ASM, such as Zn2+ ions, TNFα, oxidative stress, and inflammatory markers should be determined concomitantly. Further confounders such as fasting, chronobiology or epigenetic influences, as well as the influence of sex and developmental stage, should be investigated.


We have summarized the current state of research on S-ASM. This enzyme is encoded by the acid sphingomyelinase gene (SMPD1), is subject to complex intracellular glycosylation and is secreted into the extracellular space. S-ASM requires the addition of Zn2+ ions to achieve full activity. LPS, cytokines and oxidative stress cause an increase in the release and activation of S-ASM. S-ASM hydrolyzes SM to ceramide and phophorylcholine at the outer cell membranes and in the blood. Ceramide in turn integrates and mediates different cellular stress signals. Therefore, S-ASM is considered a proinflammatory enzyme. Many questions about the specific effects of S-ASM vs. L-ASM remain unanswered because both enzymes arise from the same gene. The cellular and animal models required to address these questions are only partially available. Genetic alterations such as mutations, polymorphisms and changes in the signal peptide result in reduced activity of serum/plasma S-ASM in humans, suggesting a trait marker. Conversely, the peripheral activity of S-ASM is elevated in diseases that are associated with acute and/or chronic inflammation, suggesting a state marker of a condition. The role of S-ASM as a potential clinical chemical marker and therapeutic target requires further exploration.


This work was supported by funding from the Interdisciplinary Center for Clinical Research Erlangen, Project E13, the Forschungsstiftung Medizin at the University Hospital Erlangen and by the Scholarship Programme ‘Equality for Women in Research and Teaching’, University Erlangen-Nuremberg (to C.R.).


  • Abe, T., Takahashi, T., Takahashi, I., Shoji, Y., and Takada, G. (1999). Serum Zn2+-stimulated sphingomyelinase deficiency in type B Niemann-Pick disease. Eur. J. Pediatr. 158, 953.Google Scholar

  • Albouz, S., Hauw, J.J., Berwald-Netter, Y., Boutry, J.M., Bourdon, R., and Baumann, N. (1981). Tricyclic antidepressants induce sphingomyelinase deficiency in fibroblast and neuroblastoma cell cultures. Biomedicine 35, 218–220.PubMedGoogle Scholar

  • Albouz, S., Vanier, M.T., Hauw, J.J., Le Saux, F., Boutry, J.M., and Baumann, N. (1983). Effect of tricyclic antidepressants on sphingomyelinase and other sphingolipid hydrolases in C6 cultured glioma cells. Neurosci. Lett. 36, 311–315.Google Scholar

  • Andrews, N.W., Almeida, P.E., and Corrotte, M. (2014). Damage control: cellular mechanisms of plasma membrane repair. Trends Cell Biol. 24, 734–742.CrossrefPubMedGoogle Scholar

  • Arenz, C. (2010). Small molecule inhibitors of acid sphingomyelinase. Cell. Physiol. Biochem. 26, 1–8.CrossrefPubMedGoogle Scholar

  • Arunachalam, B., Phan, U.T., Geuze, H.J., and Cresswell, P. (2000). Enzymatic reduction of disulfide bonds in lysosomes: characterization of a gamma-interferon-inducible lysosomal thiol reductase (GILT). Proc. Natl. Acad. Sci. USA 97, 745–750.Google Scholar

  • Assaf, S.Y. and Chung, S.H. (1984). Release of endogenous Zn2+ from brain tissue during activity. Nature 308, 734–736.Google Scholar

  • Balsinde, J., Balboa, M.A., and Dennis, E.A. (1997). Inflammatory activation of arachidonic acid signaling in murine P388D1 macrophages via sphingomyelin synthesis. J. Biol. Chem. 272, 20373–20377.Google Scholar

  • Barceló-Coblijn, G., Martin, M.L., de Almeida, R.F.M., Noguera-Salvà, M.A., Marcilla-Etxenike, A., Guardiola-Serrano, F., Lüth, A., Kleuser, B., Halver, J.E., and Escribá, P.V. (2011). Sphingomyelin and sphingomyelin synthase (SMS) in the malignant transformation of glioma cells and in 2-hydroxyoleic acid therapy. Proc. Natl. Acad. Sci. USA 108, 19569–19574.CrossrefGoogle Scholar

  • Bartelsen, O., Lansmann, S., Nettersheim, M., Lemm, T., Ferlinz, K., and Sandhoff, K. (1998). Expression of recombinant human acid sphingomyelinase in insect Sf21 cells: purification, processing and enzymatic characterization. J. Biotechnol. 63, 29–40.Google Scholar

  • Billich, A. and Baumruker, T. (2008). Sphingolipid metabolizing enzymes as novel therapeutic targets. In: Lipids in Health and Disease, Vol. 49, P.J. Quinn and X. Wang, eds. (Springer), pp. 487–522.Google Scholar

  • Bowser, P.A. and Gray, G.M. (1978). Sphingomyelinase in pig and human epidermis. J. Invest. Dermatol. 70, 331–335.CrossrefGoogle Scholar

  • Brady, R.O., Kanfer, J.N., Mock, M.B., and Fredrickson, D.S. (1966). The metabolism of sphingomyelin. II. Evidence of an enzymatic deficiency in Niemann-Pick disease. Proc. Natl. Acad. Sci. USA 55, 366–369.Google Scholar

  • Brand, I.A. and Kleineke, J. (1996). Intracellular zinc movement and its effect on the carbohydrate metabolism of isolated rat hepatocytes. J. Biol. Chem. 271, 1941–1949.Google Scholar

  • Buxaderas, S.C. and Farré-Rovira, R. (1985). Whole blood and serum zinc levels in relation to sex and age. Rev. Esp. Fisiol. 41, 463–470.PubMedGoogle Scholar

  • Callahan, J.W., Jones, C.S., Davidson, D.J., and Shankaran, P. (1983). The active site of lysosomal sphingomyelinase: evidence for the involvement of hydrophobic and ionic groups. J. Neurosci. Res. 10, 151–163.CrossrefGoogle Scholar

  • Canals, D., Perry, D.M., Jenkins, R.W., and Hannun, Y.A. (2011). Drug targeting of sphingolipid metabolism: sphingomyelinases and ceramidases. Br. J. Pharmacol. 163, 694–712.Google Scholar

  • Charruyer, A., Grazide, S., Bezombes, C., Müller, S., Laurent, G., and Jaffrézou, J.P. (2005). UV-C light induces raft-associated acid sphingomyelinase and JNK activation and translocation independently on a nuclear signal. J. Biol. Chem. 280, 19196–19204.Google Scholar

  • Cifone, M.G., De Maria, R., Roncaioli, P., Rippo, M.R., Azuma, M., Lanier, L.L., Santoni, A., and Testi, R. (1994). Apoptotic signaling through CD95 (Fas/Apo-1). activates an acidic sphingomyelinase. J. Exp. Med. 180, 1547–1552.Google Scholar

  • Claus, R.A., Bunck, A.C., Bockmeyer, C.L., Brunkhorst, F.M., Lösche, W., Kinscherf, R., and Deigner, H.-P. (2005). Role of increased sphingomyelinase activity in apoptosis and organ failure of patients with severe sepsis. FASEB J. 19, 1719–1721.PubMedGoogle Scholar

  • Cohen, R. and Barenholz, Y. (1978). Correlation between the thermotropic behavior of sphingomyelin liposomes and sphingomyelin hydrolysis by sphingomyelinase of Staphylococcus aureus. Biochim. Biophys. Acta 509, 181–187.Google Scholar

  • Corrotte, M., Almeida, P.E., Tam, C., Castro-Gomes, T., Fernandes, M.C., Millis, B.A., Cortez, M., Miller, H., Song, W., Maugel, T.K., et al. (2013). Caveolae internalization repairs wounded cells and muscle fibers. eLife 2, e00926.Google Scholar

  • Csermely, P., Fodor, P., and Somogyi, J. (1987). The tumor promoter tetradecanoylphorbol-13-acetate elicits the redistribution of heavy metals in subcellular fractions of rabbit thymocytes as measured by plasma emission spectroscopy. Carcinogenesis 8, 1663–1666.CrossrefGoogle Scholar

  • Deevska, G.M., Sunkara, M., Morris, A.J., and Nikolova-Karakashian, M.N. (2012). Characterization of secretory sphingomyelinase activity, lipoprotein sphingolipid content and LDL aggregation in ldlr-/- mice fed on a high-fat diet. Biosci. Rep. 32, 479–490.CrossrefGoogle Scholar

  • Defour, A., Van der Meulen, J.H., Bhat, R., Bigot, A., Bashir, R., Nagaraju, K., and Jaiswal, J.K. (2014). Dysferlin regulates cell membrane repair by facilitating injury-triggered acid sphingomyelinase secretion. Cell Death Dis. 5, e1306.Google Scholar

  • Deng, H., Xiu, X., and Jankovic, J. (2014). Genetic convergence of Parkinson’s disease and lysosomal storage disorders. Mol. Neurobiol., in press.Google Scholar

  • Desnick, J.P., Kim, J., He, X., Wasserstein, M.P., Simonaro, C.M., and Schuchman, E.H. (2010). Identification and characterization of eight novel SMPD1 mutations causing types A and B Niemann-Pick disease. Mol. Med. 16, 316–321.Google Scholar

  • Devlin, C.M., Leventhal, A.R., Kuriakose, G., Schuchman, E.H., Williams, K.J., and Tabas, I. (2008). Acid sphingomyelinase promotes lipoprotein retention within early atheromata and accelerates lesion progression. Arterioscler. Thromb. Vasc. Biol. 28, 1723–1730.PubMedCrossrefGoogle Scholar

  • Devlin, C., Pipalia, N.H., Liao, X., Schuchman, E.H., Maxfield, F.R., and Tabas, I. (2010). Improvement in lipid and protein trafficking in Niemann-Pick C1 cells by correction of a secondary enzyme defect. Traffic 11, 601–615.CrossrefGoogle Scholar

  • Dhami, R. and Schuchman, E.H. (2004). Mannose 6-phosphate receptor-mediated uptake is defective in acid sphingomyelinase-deficient macrophages: implications for Niemann-Pick disease enzyme replacement therapy. J. Biol. Chem. 279, 1526–1532.Google Scholar

  • Dinkla, S., Wessels, K., Verdurmen, W.P.R., Tomelleri, C., Cluitmans, J.C.A., Fransen, J., Fuchs, B., Schiller, J., Joosten, I., Brock, R., et al. (2012). Functional consequences of sphingomyelinase-induced changes in erythrocyte membrane structure. Cell Death Dis. 3, e410.Google Scholar

  • Doehner, W., Bunck, A.C., Rauchhaus, M., von Haehling, S., Brunkhorst, F.M., Cicoira, M., Tschope, C., Ponikowski, P., Claus, R.A., and Anker, S.D. (2007). Secretory sphingomyelinase is upregulated in chronic heart failure: a second messenger system of immune activation relates to body composition, muscular functional capacity, and peripheral blood flow. Eur. Heart J. 28, 821–828.CrossrefGoogle Scholar

  • Don, A.S., Lim, X.Y., and Couttas, T.A. (2014). Re-configuration of sphingolipid metabolism by oncogenic transformation. Biomolecules 4, 315–353.CrossrefPubMedGoogle Scholar

  • Donangelo, C.M. and Chang, G.W. (1981). An enzymatic assay for available zinc in plasma and serum. Clin. Chim. Acta 113, 201–206.Google Scholar

  • Drachmann, T., Mathiessen, J.H., Pedersen, M.H., and Hellgren, L.I. (2007). The source of dietary fatty acids alters the activity of secretory sphingomyelinase in the rat. Eur. J. Lipid Sci. Technol. 109, 1003–1009.CrossrefGoogle Scholar

  • Draeger, A. and Babiychuk, E.B. (2013). Ceramide in plasma membrane repair. Handb. Exp. Pharmacol. 216, 341–353.Google Scholar

  • Driessen, M., Weitz, G., Brouwer-Kelder, E.M., Donker-Koopman, W.E., Bastiaannet, J., Sandhoff, K., Barranger, J.A., Tager, J.M., and Schram, A.W. (1985). The effect of detergents on immunoprecipitability of lysosomal sphingomyelinase. Biochim. Biophys. Acta 841, 97–102.Google Scholar

  • Drobnik, W., Liebisch, G., Audebert, F.X., Fröhlich, D., Glück, T., Vogel, P., Rothe, G., and Schmitz, G. (2003). Plasma ceramide and lysophosphatidylcholine inversely correlate with mortality in sepsis patients. J. Lipid Res. 44, 754–761.CrossrefGoogle Scholar

  • Dumitru, C.A. and Gulbins, E. (2006). TRAIL activates acid sphingomyelinase via a redox mechanism and releases ceramide to trigger apoptosis. Oncogene 25, 5612–5625.CrossrefGoogle Scholar

  • Eckhardt, M. (2010). Pathology and current treatment of neurodegenerative sphingolipidoses. Neuromolecular. Med. 12, 362–382.CrossrefPubMedGoogle Scholar

  • Edelmann, B., Bertsch, U., Tchikov, V., Winoto-Morbach, S., Perrotta, C., Jakob, M., Adam-Klages, S., Kabelitz, D., and Schütze, S. (2011). Caspase-8 and caspase-7 sequentially mediate proteolytic activation of acid sphingomyelinase in TNF-R1 receptosomes. EMBO J. 30, 379–394.Google Scholar

  • Erickson, A.H. and Blobel, G. (1983). Carboxyl-terminal proteolytic processing during biosynthesis of the lysosomal enzymes β-glucuronidase and cathepsin D. Biochemistry 22, 5201–5205.PubMedCrossrefGoogle Scholar

  • Ferlinz, K., Hurwitz, R., Vielhaber, G., Suzuki, K., and Sandhoff, K. (1994). Occurrence of two molecular forms of human acid sphingomyelinase. Biochem. J. 301, 855–862.Google Scholar

  • Ferlinz, K., Hurwitz, R., Moczall, H., Lansmann, S., Schuchman, E.H., and Sandhoff, K. (1997). Functional characterization of the N-glycosylation sites of human acid sphingomyelinase by site-directed mutagenesis. Eur. J. Biochem. 243, 511–517.Google Scholar

  • Fernandes, M.C., Cortez, M., Flannery, A.R., Tam, C., Mortara, R.A., and Andrews, N.W. (2011). Trypanosoma cruzi subverts the sphingomyelinase-mediated plasma membrane repair pathway for cell invasion. J. Exp. Med. 208, 909–921.Google Scholar

  • Filippov, V., Song, M.A., Zhang, K., Vinters, H.V., Tung, S., Kirsch, W.M., Yang, J., and Duerksen-Hughes, P.J. (2012). Increased ceramide in brains with Alzheimer’s and other neurodegenerative diseases. J. Alzheimers Dis. 29, 537–547.Google Scholar

  • Foo, J.N., Liany, H., Bei, J.X., Yu, X.Q., Liu, J., Au, W.L., Prakash, K.M., Tan, L.C., and Tan, E.K. (2013). Rare lysosomal enzyme gene SMPD1 variant (p. R591C). associates with Parkinson’s disease. Neurobiol. Aging 34, 2890–2895.Google Scholar

  • Frederickson, C.J. and Moncrieff, D.W. (1994). Zinc-containing neurons. Biol. Signals 3, 127–139.CrossrefPubMedGoogle Scholar

  • Fujii, S., Yoshida, A., Sakurai, S., Morita, M., Tsukamoto, K., Ikezawa, H., and Ikeda, K. (2004). Chromogenic assay for the activity of sphingomyelinase from Bacillus cereus and its application to the enzymatic hydrolysis of lysophospholipids. Biol. Pharm. Bull. 27, 1725–1729.CrossrefGoogle Scholar

  • Gan-Or, Z., Ozelius, L.J., Bar-Shira, A., Saunders-Pullman, R., Mirelman, A., Kornreich, R., Gana-Weisz, M., Raymond, D., Rozenkrantz, L., Deik, A., et al. (2013). The p. L302P mutation in the lysosomal enzyme gene SMPD1 is a risk factor for Parkinson disease. Neurology 80, 1606–1610.Google Scholar

  • Gault, C.R., Obeid, L.M., and Hannun, Y.A. (2010). An overview of sphingolipid metabolism: from synthesis to breakdown. Adv. Exp. Med. Biol. 688, 1–23.Google Scholar

  • Göggel, R., Winoto-Morbach, S., Vielhaber, G., Imai, Y., Lindner, K., Brade, L., Brade, H., Ehlers, S., Slutsky, A.S., Schütze, S., et al. (2004). PAF-mediated pulmonary edema: a new role for acid sphingomyelinase and ceramide. Nat. Med. 10, 155–160.PubMedCrossrefGoogle Scholar

  • Goni, F.M. and Alonso, A. (2002). Sphingomyelinases: enzymology and membrane activity. FEBS Lett. 531, 38–46.Google Scholar

  • Górska, M., Baranczuk, E., and Dobrzyn, A. (2003). Secretory Zn2+-dependent sphingomyelinase activity in the serum of patients with type 2 diabetes is elevated. Horm. Metab. Res. 35, 506–507.Google Scholar

  • Gracia-Garcia, P., Rao, V., Haughey, N.J., Ratnam Banduru, V.V., Smith, G., Rosenberg, P.B., Lobo, A., Lyketsos, C.G., and Mielke, M.M. (2011). Elevated plasma ceramides in depression. J. Neuropsychiatry Clin. Neurosci. 23, 215–218.CrossrefGoogle Scholar

  • Grammatikos, G., Mühle, C., Ferreiros, N., Schroeter, S., Bogdanou, D., Schwalm, S., Hintereder, G., Kornhuber, J., Zeuzem, S., Sarrazin, C., et al. (2014). Serum acid sphingomyelinase is upregulated in chronic hepatitis C infection and non alcoholic fatty liver disease. Biochim. Biophys. Acta 1841, 1012–1020.Google Scholar

  • Grassmé, H., Gulbins, E., Brenner, B., Ferlinz, K., Sandhoff, K., Harzer, K., Lang, F., and Meyer, T.F. (1997). Acidic sphingomyelinase mediates entry of N. gonorrhoeae into nonphagocytic cells. Cell 91, 605–615.Google Scholar

  • Grassmé, H., Schwarz, H., and Gulbins, E. (2001). Molecular mechanisms of ceramide-mediated CD95 clustering. Biochem. Biophys. Res. Commun. 284, 1016–1030.Google Scholar

  • Grassmé, H., Jendrossek, V., Riehle, A., von Kürthy, G., Berger, J., Schwarz, H., Weller, M., Kolesnick, R., and Gulbins, E. (2003). Host defense against Pseudomonas aeruginosa requires ceramide-rich membrane rafts. Nat. Med. 9, 322–330.CrossrefGoogle Scholar

  • Grassmé, H., Riehle, A., Wilker, B., and Gulbins, E. (2005). Rhinoviruses infect human epithelial cells via ceramide-enriched membrane platforms. J. Biol. Chem. 280, 26256–26262.Google Scholar

  • Guarino, A.J., Tulenko, T.N., and Wrenn, S.P. (2006). Sphingomyelinase-to-LDL molar ratio determines low density lipoprotein aggregation size: biological significance. Chem. Phys. Lipids 142, 33–42.Google Scholar

  • Gulbins, E. and Kolesnick, R. (2000). Measurement of sphingomyelinase activity. Methods Enzymol. 322, 382–388.Google Scholar

  • Gulbins, E., Palmada, M., Reichel, M., Lüth, A., Böhmer, C., Amato, D., Müller, C.P., Tischbirek, C.H., Groemer, T.W., Tabatabai, G., et al. (2013). Acid sphingomyelinase/ceramide system mediates effects of antidepressant drugs. Nat. Med. 19, 934–938.PubMedCrossrefGoogle Scholar

  • Gupta, V.R., Patel, H.K., Kostolansky, S.S., Ballivian, R.A., Eichberg, J., and Blanke, S.R. (2008). Sphingomyelin functions as a novel receptor for Helicobacter pylori VacA. PLoS Pathog. 4, e1000073.CrossrefGoogle Scholar

  • Haimovitz-Friedman, A., Cordon-Cardo, C., Bayoumy, S., Garzotto, M., McLoughlin, M., Gallily, R., Edwards, C.K., III, Schuchman, E.H., Fuks, Z., and Kolesnick, R. (1997). Lipopolysaccharide induces disseminated endothelial apoptosis requiring ceramide generation. J. Exp. Med. 186, 1831–1841.Google Scholar

  • Hammad, S.M., Truman, J.P., Al Gadban, M.M., Smith, K.J., Twal, W.O., and Hamner, M.B. (2012). Altered blood sphingolipidomics and elevated plasma inflammatory cytokines in combat veterans with post-traumatic stress disorder. Neurobiol. Lipids 10, 2.Google Scholar

  • Harzer, K. and Benz, H.U. (1973). Letters: a simple sphingomyelinase determination for Niemann-Pick disease: differential diagnosis of types A, B and C. J. Neurochem. 21, 999–1001.CrossrefGoogle Scholar

  • Hasilik, A. (1992). The early and late processing of lysosomal enzymes: proteolysis and compartmentation. Experientia 48, 130–151.PubMedCrossrefGoogle Scholar

  • Hauck, C.R., Grassmé, H., Bock, J., Jendrossek, V., Ferlinz, K., Meyer, T.F., and Gulbins, E. (2000). Acid sphingomyelinase is involved in CEACAM receptor-mediated phagocytosis of Neisseria gonorrhoeae. FEBS Lett. 478, 260–266.Google Scholar

  • He, X. and Schuchman, E.H. (2012). Potential role of acid sphingomyelinase in environmental health. Zhong. Nan. Da. Xue. Xue. Bao. Yi. Xue. Ban. 37, 109–125.Google Scholar

  • He, X., Miranda, S.R., Xiong, X., Dagan, A., Gatt, S., and Schuchman, E.H. (1999). Characterization of human acid sphingomyelinase purified from the media of overexpressing Chinese hamster ovary cells. Biochim. Biophys. Acta 1432, 251–264.Google Scholar

  • He, X., Chen, F., Dagan, A., Gatt, S., and Schuchman, E.H. (2003). A fluorescence-based, high-performance liquid chromatographic assay to determine acid sphingomyelinase activity and diagnose types A and B Niemann-Pick disease. Anal. Biochem. 314, 116–120.Google Scholar

  • Hendrich, A.B. and Michalak, K. (2003). Lipids as a target for drugs modulating multidrug resistance of cancer cells. Curr. Drug Targets 4, 23–30.PubMedCrossrefGoogle Scholar

  • Henry, B., Möller, C., Dimanche-Boitrel, M.T., Gulbins, E., and Becker, K.A. (2013). Targeting the ceramide system in cancer. Cancer Lett. 332, 286–294.PubMedCrossrefGoogle Scholar

  • Holopainen, J.M., Medina, O.P., Metso, A.J., and Kinnunen, P.K.J. (2000). Sphingomyelinase activity associated with human plasma low density lipoprotein. J. Biol. Chem. 275, 16484–16489.Google Scholar

  • Horinouchi, K., Erlich, S., Perl, D.P., Ferlinz, K., Bisgaier, C.L., Sandhoff, K., Desnick, R.J., Stewart, C.L., and Schuchman, E.H. (1995). Acid sphingomyelinase deficient mice: a model of types A and B Niemann-Pick disease. Nat. Genet. 10, 288–293.CrossrefGoogle Scholar

  • Hurwitz, R., Ferlinz, K., and Sandhoff, K. (1994a). The tricyclic antidepressants desipramine causes proteolytic degradation of lysosomal sphingomyelinase in human fibroblasts. Biol. Chem. Hoppe Seyler 375, 447–450.Google Scholar

  • Hurwitz, R., Ferlinz, K., Vielhaber, G., Moczall, H., and Sandhoff, K. (1994b). Processing of human acid sphingomyelinase in normal and I-cell fibroblasts. J. Biol. Chem. 269, 5440–5445.Google Scholar

  • Ioannou, Y.A., Bishop, D.F., and Desnick, R.J. (1992). Overexpression of human α-galactosidase A results in its intracellular aggregation, crystallization in lysosomes, and selective secretion. J. Cell Biol. 119, 1137–1150.Google Scholar

  • Jan, J.T., Chatterjee, S., and Griffin, D.E. (2000). Sindbis virus entry into cells triggers apoptosis by activating sphingomyelinase, leading to the release of ceramide. J. Virol. 74, 6425–6432.CrossrefGoogle Scholar

  • Jbeily, N., Suckert, I., Gonnert, F.A., Acht, B., Bockmeyer, C.L., Grossmann, S.D., Blaess, M.F., Lueth, A., Deigner, H.P., Bauer, M., et al. (2013). Hyperresponsiveness of mice deficient in plasma-secreted sphingomyelinase reveals its pivotal role in early phase of host response. J. Lipid Res. 54, 410–424.CrossrefGoogle Scholar

  • Jenkins, R.W., Canals, D., and Hannun, Y.A. (2009). Roles and regulation of secretory and lysosomal acid sphingomyelinase. Cell. Signal. 21, 836–846.PubMedCrossrefGoogle Scholar

  • Jenkins, R.W., Canals, D., Idkowiak-Baldys, J., Simbari, F., Roddy, P., Perry, D.M., Kitatani, K., Luberto, C., and Hannun, Y.A. (2010). Regulated secretion of acid sphingomyelinase: implications for selectivity of ceramide formation. J. Biol. Chem. 285, 35706–35718.Google Scholar

  • Jenkins, R.W., Clarke, C.J., Canals, D., Snider, A.J., Gault, C.R., Heffernan-Stroud, L., Wu, B.X., Simbari, F., Roddy, P., Kitatani, K., et al. (2011a). Regulation of CC ligand 5/RANTES by acid sphingomyelinase and acid ceramidase. J. Biol. Chem. 286, 13292–13303.Google Scholar

  • Jenkins, R.W., Idkowiak-Baldys, J., Simbari, F., Canals, D., Roddy, P., Riner, C.D., Clarke, C.J., and Hannun, Y.A. (2011b). A novel mechanism of lysosomal acid sphingomyelinase maturation – requirement for carboxy-terminal proteolytic processing. J. Biol. Chem. 286, 3777–3788.Google Scholar

  • Jenkins, R.W., Clarke, C.J., Lucas, J.T., Jr., Shabbir, M., Wu, B.X., Simbari, F., Mueller, J., Hannun, Y.A., Lazarchick, J., and Shirai, K. (2013). Evaluation of the role of secretory sphingomyelinase and bioactive sphingolipids as biomarkers in hemophagocytic lymphohistiocytosis. Am. J. Hematol. 88, E265–E272.Google Scholar

  • Jeong, T., Schissel, S.L., Tabas, I., Pownall, H.J., Tall, A.R., and Jiang, X. (1998). Increased sphingomyelin content of plasma lipoproteins in apolipoprotein E knockout mice reflects combined production and catabolic defects and enhances reactivity with mammalian sphingomyelinase. J. Clin. Invest. 101, 905–912.CrossrefGoogle Scholar

  • Jiang, L.J., Maret, W., and Vallee, B.L. (1998). The glutathione redox couple modulates zinc transfer from metallothionein to zinc-depleted sorbitol dehydrogenase. Proc. Natl. Acad. Sci. USA 95, 3483–3488.CrossrefGoogle Scholar

  • Jobb, E. and Callahan, J.W. (1987). The subunit of human sphingomyelinase is not the same size in all tissues: studies with a polyclonal rabbit serum. J. Inher. Metab. Dis. 10 (Suppl. 2), 326–328.Google Scholar

  • Jones, C.S., Shankaran, P., and Callahan, J.W. (1981). Purification of sphingomyelinase to apparent homogeneity by using hydrophobic chromatography. Biochem. J. 195, 373–382.Google Scholar

  • Jones, C.S., Shankaran, P., Davidson, D.J., Poulos, A., and Callahan, J.W. (1983). Studies on the structure of sphingomyelinase. Amino acid composition and heterogeneity on isoelectric focusing. Biochem. J. 209, 291–297.Google Scholar

  • Kim, Y. and Sun, H. (2012). ASM-3 Acid sphingomyelinase functions as a positive regulator of the DAF-2/AGE-1 signaling pathway and serves as a novel anti-aging target. PLoS One 7, e45890.Google Scholar

  • Kinnunen, P.K. and Holopainen, J.M. (2002). Sphingomyelinase activity of LDL: a link between atherosclerosis, ceramide, and apoptosis? Trends Cardiovasc. Med. 12, 37–42.Google Scholar

  • Kiprianos, A.P., Morgan, M.D., Little, M.A., Harper, L., Bacon, P.A., and Young, S.P. (2012). Elevated active secretory sphingomyelinase in antineutrophil cytoplasmic antibody-associated primary systemic vasculitis. Ann. Rheum. Dis. 71, 1100–1102.CrossrefPubMedGoogle Scholar

  • Kirkegaard, T., Roth, A.G., Petersen, N.H.T., Mahalka, A.K., Olsen, O.D., Moilanen, I., Zylicz, A., Knudsen, J., Sandhoff, K., Arenz, C., et al. (2010). Hsp70 stabilizes lysosomes and reverts Niemann-Pick disease-associated lysosomal pathology. Nature 463, 549–553.Google Scholar

  • Kobayashi, K., Nagata, E., Sasaki, K., Harada-Shiba, M., Kojo, S., and Kikuzaki, H. (2013). Increase in secretory sphingomyelinase activity and specific ceramides in the aorta of apolipoprotein E knockout mice during aging. Biol. Pharm. Bull. 36, 1192–1196.PubMedCrossrefGoogle Scholar

  • Kolter, T. and Sandhoff, K. (2005). Principles of lysosomal membrane digestion: stimulation of sphingolipid degradation by sphingolipid activator proteins and anionic lysosomal lipids. Annu. Rev. Cell Dev. Biol. 21, 81–103.PubMedCrossrefGoogle Scholar

  • Kölzer, M., Arenz, C., Ferlinz, K., Werth, N., Schulze, H., Klingenstein, R., and Sandhoff, K. (2003). Phosphatidylinositol-3,5-bisphosphate is a potent and selective inhibitor of acid sphingomyelinase. Biol. Chem. 384, 1293–1298.Google Scholar

  • Kölzer, M., Ferlinz, K., Bartelsen, O., Hoops, S.L., Lang, F., and Sandhoff, K. (2004a). Functional characterization of the postulated intramolecular sphingolipid activator protein domain of human acid sphingomyelinase. Biol. Chem. 385, 1193–1195.Google Scholar

  • Kölzer, M., Werth, N., and Sandhoff, K. (2004b). Interactions of acid sphingomyelinase and lipid bilayers in the presence of the tricyclic antidepressant desipramine. FEBS Lett. 559, 96–98.Google Scholar

  • Kornfeld, S. (1987). Trafficking of lysosomal enzymes. FASEB J. 1, 462–468.PubMedGoogle Scholar

  • Kornhuber, J., Tripal, P., Reichel, M., Terfloth, L., Bleich, S., Wiltfang, J., and Gulbins, E. (2008). Identification of new functional inhibitors of acid sphingomyelinase using a structure-property-activity relation model. J. Med. Chem. 51, 219–237.CrossrefGoogle Scholar

  • Kornhuber, J., Tripal, P., Reichel, M., Mühle, C., Rhein, C., Muehlbacher, M., Groemer, T.W., and Gulbins, E. (2010). Functional inhibitors of acid sphingomyelinase (FIASMAs).: a novel pharmacological group of drugs with broad clinical applications. Cell. Physiol. Biochem. 26, 9–20.PubMedCrossrefGoogle Scholar

  • Kornhuber, J., Muehlbacher, M., Trapp, S., Pechmann, S., Friedl, A., Reichel, M., Mühle, C., Terfloth, L., Groemer, T.W., Spitzer, G.M., et al. (2011). Identification of novel functional inhibitors of acid sphingomyelinase. PLoS One 6, e23852.Google Scholar

  • Kornhuber, J., Müller, C.P., Becker, K.A., Reichel, M., and Gulbins, E. (2014). The ceramide system as a novel antidepressant target. Trends Pharmacol. Sci. 35, 293–304.PubMedCrossrefGoogle Scholar

  • Kott, M., Elke, G., Reinicke, M., Winoto-Morbach, S., Schädler, D., Zick, G., Frerichs, U., Weiler, N., and Schütze, S. (2014). Acid sphingomyelinase serum activity predicts mortality in intensive care unit patients after systemic inflammation: a prospective cohort study. PLoS One 9, e112323.Google Scholar

  • Krishnan, V. and Nestler, E.J. (2008). The molecular neurobiology of depression. Nature 455, 894–902.Google Scholar

  • Lai, J., Moxey, A., Nowak, G., Vashum, K., Bailey, K., and McEvoy, M. (2012). The efficacy of zinc supplementation in depression: systematic review of randomised controlled trials. J. Affect. Disord. 136, e31–e39.Google Scholar

  • Lang, P.A., Schenck, M., Nicolay, J.P., Becker, J.U., Kempe, D.S., Lupescu, A., Koka, S., Eisele, K., Klarl, B.A., Rübben, H., et al. (2007). Liver cell death and anemia in Wilson disease involve acid sphingomyelinase and ceramide. Nat. Med. 13, 164–170.CrossrefPubMedGoogle Scholar

  • Langmann, T., Buechler, C., Ries, S., Schaeffler, A., Aslanidis, C., Schuierer, M., Weiler, M., Sandhoff, K., de Jong, P.J., and Schmitz, G. (1999). Transcription factors Sp1 and AP-2 mediate induction of acid sphingomyelinase during monocytic differentiation. J. Lipid Res. 40, 870–880.Google Scholar

  • Lansmann, S., Ferlinz, K., Hurwitz, R., Bartelsen, O., Glombitza, G., and Sandhoff, K. (1996). Purification of acid sphingomyelinase from human placenta: characterization and N-terminal sequence. FEBS Lett. 399, 227–231.Google Scholar

  • Lansmann, S., Schuette, C.G., Bartelsen, O., Hoernschemeyer, J., Linke, T., Weisgerber, J., and Sandhoff, K. (2003). Human acid sphingomyelinase. Eur. J. Biochem. 270, 1076–1088.PubMedCrossrefGoogle Scholar

  • Lee, C.Y., Krimbou, L., Vincent, J., Bernard, C., Larramée, P., Genest, J., Jr., and Marcil, M. (2003). Compound heterozygosity at the sphingomyelin phosphodiesterase-1 (SMPD1). gene is associated with low HDL cholesterol. Hum. Genet. 112, 552–562.Google Scholar

  • Lee, J.K., Jin, H.K., Park, M.H., Kim, B.R., Lee, P.H., Nakauchi, H., Carter, J.E., He, X., Schuchman, E.H., and Bae, J.S. (2014). Acid sphingomyelinase modulates the autophagic process by controlling lysosomal biogenesis in Alzheimer’s disease. J. Exp. Med. 211, 1551–1570.Google Scholar

  • Leger, A.J., Mosquea, L.M., Li, L., Chuang, W., Pacheco, J., Taylor, K., Luo, Z., Piepenhagen, P., Ziegler, R., Moreland, R., et al. (2011). Adeno-associated virus-mediated expression of acid sphingomyelinase decreases atherosclerotic lesion formation in apolipoprotein E-/- mice. J. Gene Med. 13, 324–332.CrossrefGoogle Scholar

  • Levenson, C.W. (2006). Zinc: the new antidepressant? Nutr. Rev. 64, 39–42.PubMedCrossrefGoogle Scholar

  • Li, X., Gulbins, E., and Zhang, Y. (2012). Oxidative stress triggers Ca-dependent lysosome trafficking and activation of acid sphingomyelinase. Cell. Physiol. Biochem. 30, 815–826.PubMedCrossrefGoogle Scholar

  • Lin, X., Hengartner, M.O., and Kolesnick, R. (1998). Caenorhabditis elegans contains two distinct acid sphingomyelinases. J. Biol. Chem. 273, 14374–14379.Google Scholar

  • Linke, T., Wilkening, G., Lansmann, S., Moczall, H., Bartelsen, O., Weisgerber, J., and Sandhoff, K. (2001). Stimulation of acid sphingomyelinase activity by lysosomal lipids and sphingolipid activator proteins. Biol. Chem. 382, 283–290.Google Scholar

  • Little, C. and Otnäss, A.B. (1975). The metal ion dependence of phospholipase C from Bacillus cereus. Biochim. Biophys. Acta 391, 326–333.Google Scholar

  • López-Montero, I., Vélez, M., and Devaux, P.F. (2007). Surface tension induced by sphingomyelin to ceramide conversion in lipid membranes. Biochim. Biophys. Acta 1768, 553–561.Google Scholar

  • Madra, M. and Sturley, S.L. (2010). Niemann-Pick type C pathogenesis and treatment: from statins to sugars. Clin. Lipidol. 5, 387–395.CrossrefPubMedGoogle Scholar

  • Magenau, A., Benzing, C., Proschogo, N., Don, A.S., Hejazi, L., Karunakaran, D., Jessup, W., and Gaus, K. (2011). Phagocytosis of IgG-coated polystyrene beads by macrophages induces and requires high membrane order. Traffic 12, 1730–1743.CrossrefPubMedGoogle Scholar

  • Maor, I., Mandel, H., and Aviram, M. (1995). Macrophage uptake of oxidized LDL inhibits lysosomal sphingomyelinase, thus causing the accumulation of unesterified cholesterol-sphingomyelin-rich particles in the lysosomes. A possible role for 7-ketocholesterol. Arterioscler. Thromb. Vasc. Biol. 15, 1378–1387.CrossrefPubMedGoogle Scholar

  • Marathe, S., Schissel, S.L., Yellin, M.J., Beatini, N., Mintzer, R., Williams, K.J., and Tabas, I. (1998). Human vascular endothelial cells are a rich and regulatable source of secretory sphingomyelinase. Implications for early atherogenesis and ceramide-mediated cell signaling. J. Biol. Chem. 273, 4081–4088.Google Scholar

  • Marathe, S., Kuriakose, G., Williams, K.J., and Tabas, I. (1999). Sphingomyelinase, an enzyme implicated in atherogenesis, is present in atherosclerotic lesions and binds to specific components of the subendothelial extracellular matrix. Arterioscler. Thromb. Vasc. Biol. 19, 2648–2658.CrossrefPubMedGoogle Scholar

  • Marathe, S., Choi, Y., Leventhal, A.R., and Tabas, I. (2000a). Sphingomyelinase converts lipoproteins from apolipoprotein E knockout mice into potent inducers of macrophage foam cell formation. Arterioscler. Thromb. Vasc. Biol. 20, 2607–2613.PubMedCrossrefGoogle Scholar

  • Marathe, S., Miranda, S.R.P., Devlin, C., Johns, A., Kuriakose, G., Williams, K.J., Schuchman, E.H., and Tabas, I. (2000b). Creation of a mouse model for non-neurological (type B). Niemann-Pick disease by stable, low level expression of lysosomal sphingomyelinase in the absence of secretory sphingomyelinase: relationship between brain intra-lysosomal enzyme activity and central nervous system function. Hum. Mol. Genet. 9, 1967–1976.PubMedGoogle Scholar

  • Maroudas, A., Weinberg, P.D., Parker, K.H., and Winlove, C.P. (1988). The distributions and diffusivities of small ions in chondroitin sulphate, hyaluronate and some proteoglycan solutions. Biophys. Chem. 32, 257–270.PubMedCrossrefGoogle Scholar

  • McCollister, B.D., Myers, J.T., Jones-Carson, J., Voelker, D.R., and Vázquez-Torres, A. (2007). Constitutive acid sphingomyelinase enhances early and late macrophage killing of Salmonella enterica serovar Typhimurium. Infect. Immun. 75, 5346–5352.CrossrefGoogle Scholar

  • Mendis, S. (1989). Magnesium, zinc, and manganese in atherosclerosis of the aorta. Biol. Trace Elem. Res. 22, 251–256.PubMedCrossrefGoogle Scholar

  • Menkin, V. (1934). Studies on inflammation: X. The cytological picture of an inflammatory exudate in relation to its hydrogen ion concentration. Am. J. Pathol. 10, 193–210.PubMedGoogle Scholar

  • Mielke, M.M., Haughey, N.J., Bandaru, V.V., Weinberg, D.D., Darby, E., Zaidi, N., Pavlik, V., Doody, R.S., and Lyketsos, C.G. (2011). Plasma sphingomyelins are associated with cognitive progression in Alzheimer’s disease. J. Alzheimers Dis. 27, 259–269.Google Scholar

  • Mielke, M.M., Maetzler, W., Haughey, N.J., Bandaru, V.V.R., Savica, R., Deuschle, C., Gasser, T., Hauser, A.K., Gräber-Sultan, S., Schleicher, E., et al. (2013). Plasma ceramide and glucosylceramide metabolism is altered in sporadic Parkinson’s disease and associated with cognitive impairment: a pilot study. PLoS One 8, e73094.Google Scholar

  • Milanino, R., Marrella, M., Gasperini, R., Pasqualicchio, M., and Velo, G. (1993). Copper and zinc body levels in inflammation: an overview of the data obtained from animal and human studies. Agents Actions 39, 195–209.PubMedCrossrefGoogle Scholar

  • Milhas, D., Clarke, C.J., and Hannun, Y.A. (2010). Sphingomyelin metabolism at the plasma membrane: implications for bioactive sphingolipids. FEBS Lett. 584, 1887–1894.Google Scholar

  • Miller, M.E., Adhikary, S., Kolokoltsov, A.A., and Davey, R.A. (2012). Ebolavirus requires acid sphingomyelinase activity and plasma membrane sphingomyelin for infection. J. Virol. 86, 7473–7483.CrossrefGoogle Scholar

  • Mintzer, R.J., Appell, K.C., Cole, A., Johns, A., Pagila, R., Polokoff, M.A., Tabas, I., Snider, R.M., and Meurer-Ogden, J.A. (2005). A novel high-throughput screening format to identify inhibitors of secreted acid sphingomyelinase. J. Biomol. Screen. 10, 225–234.CrossrefGoogle Scholar

  • Miranda, S.R.P., He, X., Simonaro, C.M., Gatt, S., Dagan, A., Desnick, R.J., and Schuchman, E.H. (2000). Infusion of recombinant human acid sphingomyelinase into Niemann-Pick disease mice leads to visceral, but not neurological, correction of the pathophysiology. FASEB J. 14, 1988–1995.PubMedCrossrefGoogle Scholar

  • Mühle, C., Huttner, H.B., Walter, S., Reichel, M., Canneva, F., Lewczuk, P., Gulbins, E., and Kornhuber, J. (2013). Characterization of acid sphingomyelinase activity in human cerebrospinal fluid. PLoS One 8, e62912.Google Scholar

  • Mühle, C., Amova, V., Biermann, T., Bayerlein, K., Richter-Schmidinger, T., Kraus, T., Reichel, M., Gulbins, E., and Kornhuber, J. (2014). Sex-dependent decrease of sphingomyelinase activity during alcohol withdrawal treatment. Cell. Physiol. Biochem. 34, 71–81.PubMedCrossrefGoogle Scholar

  • Murate, T., Suzuki, M., Hattori, M., Takagi, A., Kojima, T., Tanizawa, T., Asano, H., Hotta, T., Saito, H., Yoshida, S., et al. (2002). Up-regulation of acid sphingomyelinase during retinoic acid-induced myeloid differentiation of NB4, a human acute promyelocytic leukemia cell line. J. Biol. Chem. 277, 9936–9943.Google Scholar

  • Naghavi, M., John, R., Naguib, S., Siadaty, M.S., Grasu, R., Kurian, K.C., van Winkle, W.B., Soller, B., Litovsky, S., Madjid, M., et al. (2002). pH Heterogeneity of human and rabbit atherosclerotic plaques; a new insight into detection of vulnerable plaque. Atherosclerosis 164, 27–35.Google Scholar

  • Nelson, J.B., O’Hara, S.P., Small, A.J., Tietz, P.S., Choudhury, A.K., Pagano, R.E., Chen, X.M., and LaRusso, N.F. (2006). Cryptosporidium parvum infects human cholangiocytes via sphingolipid-enriched membrane microdomains. Cell. Microbiol. 8, 1932–1945.CrossrefGoogle Scholar

  • Newrzella, D. and Stoffel, W. (1996). Functional analysis of the glycosylation of murine acid sphingomyelinase. J. Biol. Chem. 271, 32089–32095.Google Scholar

  • Ni, X. and Morales, C.R. (2006). The lysosomal trafficking of acid sphingomyelinase is mediated by sortilin and mannose 6-phosphate receptor. Traffic 7, 889–902.PubMedGoogle Scholar

  • Nielsen, M.S., Madsen, P., Christensen, E.I., Nykjaer, A., Gliemann, J., Kasper, D., Pohlmann, R., and Petersen, C.M. (2001). The sortilin cytoplasmic tail conveys Golgi-endosome transport and binds the VHS domain of the GGA2 sorting protein. EMBO J. 20, 2180–2190.PubMedCrossrefGoogle Scholar

  • Nishikawa, A., Gregory, W., Frenz, J., Cacia, J., and Kornfeld, S. (1997). The phosphorylation of bovine DNase I Asn-linked oligosaccharides is dependent on specific lysine and arginine residues. J. Biol. Chem. 272, 19408–19412.CrossrefGoogle Scholar

  • Norman, E., Cutler, R.G., Flannery, R., Wang, Y., and Mattson, M.P. (2010). Plasma membrane sphingomyelin hydrolysis increases hippocampal neuron excitability by sphingosine-1-phosphate mediated mechanisms. J. Neurochem. 114, 430–439.CrossrefGoogle Scholar

  • Nyberg, L., Farooqi, A., Bläckberg, L., Duan, R.D., Nilsson, A., and Hernell, O. (1998). Digestion of ceramide by human milk bile salt-stimulated lipase. J. Pediatr. Gastroenterol. Nutr. 27, 560–567.CrossrefGoogle Scholar

  • Oninla, V.O., Breiden, B., Babalola, J.O., and Sandhoff, K. (2014). Acid sphingomyelinase activity is regulated by membrane lipids and facilitates cholesterol transfer by NPC2. J. Lipid Res. 55, 2606–2619.CrossrefGoogle Scholar

  • Öörni, K., Hakala, J.K., Annila, A., Ala-Korpela, M., and Kovanen, P.T. (1998). Sphingomyelinase induces aggregation and fusion, but phospholipase A2 only aggregation, of low density lipoprotein (LDL). particles. Two distinct mechanisms leading to increased binding strength of LDL to human aortic proteoglycans. J. Biol. Chem. 273, 29127–29134.Google Scholar

  • Öörni, K., Posio, P., Ala-Korpela, M., Jauhiainen, M., and Kovanen, P.T. (2005). Sphingomyelinase induces aggregation and fusion of small very low-density lipoprotein and intermediate-density lipoprotein particles and increases their retention to human arterial proteoglycans. Arterioscler. Thromb. Vasc. Biol. 25, 1678–1683.PubMedCrossrefGoogle Scholar

  • Otterbach, B. and Stoffel, W. (1995). Acid sphingomyelinase-deficient mice mimic the neurovisceral form of human lysosomal storage disease (Niemann-Pick disease). Cell 81, 1053–1061.PubMedCrossrefGoogle Scholar

  • Pan, W., Yu, J., Shi, R., Yan, L., Yang, T., Li, Y., Zhang, Z., Yu, G., Bai, Y., Schuchman, E.H., et al. (2014). Elevation of ceramide and activation of secretory acid sphingomyelinase in patients with acute coronary syndromes. Coron. Artery Dis. 25, 230–235.PubMedGoogle Scholar

  • Pavlu-Pereira, H., Asfaw, B., Poupctová, H., Ledvinová, J., Sikora, J., Vanier, M.T., Sandhoff, K., Zeman, J., Novotná, Z., Chudoba, D., et al. (2005). Acid sphingomyelinase deficiency. Phenotype variability with prevalence of intermediate phenotype in a series of twenty-five Czech and Slovak patients. A multi-approach study. J. Inherit. Metab. Dis. 28, 203–227.CrossrefGoogle Scholar

  • Pavoine, C. and Pecker, F. (2009). Sphingomyelinases: their regulation and roles in cardiovascular pathophysiology. Cardiovasc. Res. 82, 175–183.PubMedGoogle Scholar

  • Perrotta, C., Bizzozero, L., Cazzato, D., Morlacchi, S., Assi, E., Simbari, F., Zhang, Y., Gulbins, E., Bassi, M.T., Rosa, P., et al. (2010). Syntaxin 4 is required for acid sphingomyelinase activity and apoptotic function. J. Biol. Chem. 285, 40240–40251.Google Scholar

  • Petersen, C.M., Nielsen, M.S., Nykjaer, A., Jacobsen, L., Tommerup, N., Rasmussen, H.H., Roigaard, H., Gliemann, J., Madsen, P., and Moestrup, S.K. (1997). Molecular identification of a novel candidate sorting receptor purified from human brain by receptor-associated protein affinity chromatography. J. Biol. Chem. 272, 3599–3605.CrossrefGoogle Scholar

  • Petersen, N.H.T., Olsen, O.D., Groth-Pedersen, L., Ellegaard, A.M., Bilgin, M., Redmer, S., Ostenfeld, M.S., Ulanet, D., Dovmark, T.H., Lønborg, A., et al. (2013). Transformation-associated changes in sphingolipid metabolism sensitize cells to lysosomal cell death induced by inhibitors of acid sphingomyelinase. Cancer Cell 24, 379–393.PubMedCrossrefGoogle Scholar

  • Petrache, I., Natarajan, V., Zhen, L., Medler, T.R., Richter, A.T., Cho, C., Hubbard, W.C., Berdyshev, E.V., and Tuder, R.M. (2005). Ceramide upregulation causes pulmonary cell apoptosis and emphysema-like disease in mice. Nat. Med. 11, 491–498.CrossrefPubMedGoogle Scholar

  • Pittis, M.G., Ricci, V., Guerci, V.I., Marcais, C., Ciana, G., Dardis, A., Gerin, F., Stroppiano, M., Vanier, M.T., Filocamo, M., et al. (2004). Acid sphingomyelinase: identification of nine novel mutations among Italian Niemann Pick type B patients and characterization of in vivo functional in-frame start codon. Hum. Mutat. 24, 186–187.CrossrefPubMedGoogle Scholar

  • Plihtari, R., Hurt-Camejo, E., Öörni, K., and Kovanen, P.T. (2010). Proteolysis sensitizes LDL particles to phospholipolysis by secretory phospholipase A2 group V and secretory sphingomyelinase. J. Lipid Res. 51, 1801–1809.CrossrefGoogle Scholar

  • Ponting, C.P. (1994). Acid sphingomyelinase possesses a domain homologous to its activator proteins: saposins B and D. Protein Sci. 3, 359–361.Google Scholar

  • Popik, W., Alce, T.M., and Au, W.C. (2002). Human immunodeficiency virus type 1 uses lipid raft-colocalized CD4 and chemokine receptors for productive entry into CD4+ T cells. J. Virol. 76, 4709–4722.Google Scholar

  • Preuß, S., Omam, F.D., Scheiermann, J., Stadelmann, S., Winoto-Morbach, S., von Bismarck, P., Adam-Klages, S., Knerlich-Lukoschus, F., Lex, D., Wesch, D., et al. (2012). Topical application of phosphatidyl-inositol-3,5-bisphosphate for acute lung injury in neonatal swine. J. Cell Mol. Med. 16, 2813–2826.Google Scholar

  • Qiu, H., Edmunds, T., Baker-Malcolm, J., Karey, K.P., Estes, S., Schwarz, C., Hughes, H., and Van Patten, S.M. (2003). Activation of human acid sphingomyelinase through modification or deletion of C-terminal cysteine. J. Biol. Chem. 278, 32744–32752.Google Scholar

  • Quintern, L.E. and Sandhoff, K. (1991). Human acid sphingomyelinase from human urine. Methods Enzymol. 197, 536–540.Google Scholar

  • Quintern, L.E., Weitz, G., Nehrkorn, H., Tager, J.M., Schram, A.W., and Sandhoff, K. (1987). Acid sphingomyelinase from human urine: purification and characterization. Biochim. Biophys. Acta 922, 323–336.Google Scholar

  • Quon, D.V.K., Proia, R.L., Fowler, A.V., Bleibaum, J., and Neufeld, E.F. (1989). Proteolytic processing of the β-subunit of the lysosomal enzyme, β-hexosaminidase, in normal human fibroblasts. J. Biol. Chem. 264, 3380–3384.Google Scholar

  • Reagan, J.W., Jr., Hubbert, M.L., and Shelness, G.S. (2000). Posttranslational regulation of acid sphingomyelinase in niemann-pick type C1 fibroblasts and free cholesterol-enriched chinese hamster ovary cells. J. Biol. Chem. 275, 38104–38110.Google Scholar

  • Rees, D., Miles, E.A., Banerjee, T., Wells, S.J., Roynette, C.E., Wahle, K.W.J., and Calder, P.C. (2006). Dose-related effects of eicosapentaenoic acid on innate immune function in healthy humans: a comparison of young and older men. Am. J. Clin. Nutr. 83, 331–342.Google Scholar

  • Reichel, M., Beck, J., Mühle, C., Rotter, A., Bleich, S., Gulbins, E., and Kornhuber, J. (2011). Activity of secretory sphingomyelinase is increased in plasma of alcohol-dependent patients. Alcohol. Clin. Exp. Res. 35, 1852–1859.PubMedCrossrefGoogle Scholar

  • Reichel, M., Richter-Schmidinger, T., Mühle, C., Rhein, C., Alexopoulos, P., Schwab, S.G., Gulbins, E., and Kornhuber, J. (2014). The common acid sphingomyelinase polymorphism p. G508R is associated with self-reported allergy. Cell. Physiol. Biochem. 34, 82–91.CrossrefGoogle Scholar

  • Reitman, M.L. and Kornfeld, S. (1981). UDP-N-acetylglucosamine:glycoprotein N-acetylglucosamine-1-phosphotransferase. Proposed enzyme for the phosphorylation of the high mannose oligosaccharide units of lysosomal enzymes. J. Biol. Chem. 256, 4275–4281.Google Scholar

  • Remmel, N., Locatelli-Hoops, S., Breiden, B., Schwarzmann, G., and Sandhoff, K. (2007). Saposin B mobilizes lipids from cholesterol-poor and bis(monoacylglycero)phosphate-rich membranes at acidic pH. Unglycosylated patient variant saposin B lacks lipid-extraction capacity. FEBS J. 274, 3405–3420.Google Scholar

  • Rhein, C., Tripal, P., Seebahn, A., Konrad, A., Kramer, M., Nagel, C., Kemper, J., Bode, J., Mühle, C., Gulbins, E., et al. (2012). Functional implications of novel human acid sphingomyelinase splice variants. PLoS One 7, e35467.Google Scholar

  • Rhein, C., Naumann, J., Mühle, C., Zill, P., Adli, M., Hegerl, U., Hiemke, C., Mergl, R., Möller, H. -J., Reichel, M., et al. (2013). The acid sphingomyelinase sequence variation p. A487V is not associated with decreased levels of enzymatic activity. JIMD Rep 8, 1–6.Google Scholar

  • Rhein, C., Reichel, M., Mühle, C., Rotter, A., Schwab, S.G., and Kornhuber, J. (2014). Secretion of acid sphingomyelinase is affected by its polymorphic signal peptide. Cell. Physiol. Biochem. 34, 1385–1401.CrossrefPubMedGoogle Scholar

  • Riethmüller, J., Riehle, A., Grassmé, H., and Gulbins, E. (2006). Membrane rafts in host-pathogen interactions. Biochim. Biophys. Acta 1758, 2139–2147.Google Scholar

  • Ro, H.A. and Carson, J.H. (2004). pH microdomains in oligodendrocytes. J. Biol. Chem. 279, 37115–37123.Google Scholar

  • Robciuc, A., Rantamäki, A.H., Jauhiainen, M., and Holopainen, J.M. (2014). Lipid-modifying enzymes in human tear fluid and corneal epithelial stress response. Invest. Ophthalmol. Vis. Sci. 55, 16–24.CrossrefPubMedGoogle Scholar

  • Romiti, E., Vasta, V., Meacci, E., Farnararo, M., Linke, T., Ferlinz, K., Sandhoff, K., and Bruni, P. (2000). Characterization of sphingomyelinase activity released by thrombin-stimulated platelets. Mol. Cell Biochem. 205, 75–81.Google Scholar

  • Roth, A.G., Drescher, D., Yang, Y., Redmer, S., Uhlig, S., and Arenz, C. (2009a). Potent and selective inhibition of acid sphingomyelinase by bisphosphonates. Angew. Chem. Int. Ed. Engl. 48, 7560–7563.PubMedCrossrefGoogle Scholar

  • Roth, A.G., Redmer, S., and Arenz, C. (2009b). Potent inhibition of acid sphingomyelinase by phosphoinositide analogues. ChemBioChem 10, 2367–2374.CrossrefPubMedGoogle Scholar

  • Roth, A.G., Redmer, S., and Arenz, C. (2010). Development of carbohydrate-derived inhibitors of acid sphingomyelinase. Bioorg. Med. Chem. 18, 939–944.PubMedCrossrefGoogle Scholar

  • Rotolo, J.A., Zhang, J., Donepudi, M., Lee, H., Fuks, Z., and Kolesnick, R. (2005). Caspase-dependent and -independent activation of acid sphingomyelinase signaling. J. Biol. Chem. 280, 26425–26434.CrossrefGoogle Scholar

  • Sakuragawa, N. (1982). Acid sphingomyelinase of human placenta: purification, properties, and 125iodine labeling. J. Biochem. 92, 637–646.Google Scholar

  • Sakuragawa, N., Sato, M., Yoshida, Y., Kamo, I., Arima, M., and Satoyoshi, E. (1985). Effects of dimethylsulfoxide on sphingomyelinase in cultured human fibroblasts and correction of sphingomyelinase deficiency in fibroblasts from Niemann-Pick patients. Biochem. Biophys. Res. Commun. 126, 756–762.CrossrefPubMedGoogle Scholar

  • Santarelli, L., Saxe, M., Gross, C., Surget, A., Battaglia, F., Dulawa, S., Weisstaub, N., Lee, J., Duman, R., Arancio, O. et al. (2003). Requirement of hippocampal neurogenesis for the behavioral effects of antidepressants. Science 301, 805–809.Google Scholar

  • Sathishkumar, S., Boyanovsky, B., Karakashian, A.A., Rozenova, K., Giltiay, N.V., Kudrimoti, M., Mohiuddin, M., Ahmed, M.M., and Nikolova-Karakashian, M. (2005). Elevated sphingomyelinase activity and ceramide concentration in serum of patients undergoing high dose spatially fractionated radiation treatment: implications for endothelial apoptosis. Cancer Biol. Ther. 4, 979–986.CrossrefPubMedGoogle Scholar

  • Sato, M., Yoshida, Y., Sakuragawa, N., and Arima, M. (1988). Effects of dimethylsulfoxide on sphingomyelinase activities in normal and Niemann-Pick type A, B and C fibroblasts. Biochim. Biophys. Acta 962, 59–65.Google Scholar

  • Satoi, H., Tomimoto, H., Ohtani, R., Kitano, T., Kondo, T., Watanabe, M., Oka, N., Akiguchi, I., Furuya, S., Hirabayashi, Y., et al. (2005). Astroglial expression of ceramide in Alzheimer’s disease brains: a role during neuronal apoptosis. Neuroscience 130, 657–666.Google Scholar

  • Savic, R. and Schuchman, E.H. (2013). Use of acid sphingomyelinase for cancer therapy. In: The Role of Sphingolipids in Cancer Development and Therapy, J.S. Norris, ed. (Academic Press), pp. 91–115.Google Scholar

  • Sawashita, J., Takeda, A., and Okada, S. (1997). Change of zinc distribution in rat brain with increasing age. Brain Res. Dev. Brain Res. 102, 295–298.CrossrefPubMedGoogle Scholar

  • Schissel, S.L., Schuchman, E.H., Williams, K.J., and Tabas, I. (1996a). Zn2+-stimulated sphingomyelinase is secreted by many cell types and is a product of the acid sphingomyelinase gene. J. Biol. Chem. 271, 18431–18436.Google Scholar

  • Schissel, S.L., Tweedie-Hardman, J., Rapp, J.H., Graham, G., Williams, K.J., and Tabas, I. (1996b). Rabbit aorta and human atherosclerotic lesions hydrolyze the sphingomyelin of retained low-density lipoprotein. Proposed role for arterial-wall sphingomyelinase in subendothelial retention and aggregation of atherogenic lipoproteins. J. Clin. Invest. 98, 1455–1464.CrossrefGoogle Scholar

  • Schissel, S.L., Jiang, X., Tweedie-Hardman, J., Jeong, T., Camejo, E.H., Najib, J., Rapp, J.H., Williams, K.J., and Tabas, I. (1998a). Secretory sphingomyelinase, a product of the acid sphingomyelinase gene, can hydrolyze atherogenic lipoproteins at neutral pH. Implications for atherosclerotic lesion development. J. Biol. Chem. 273, 2738–2746.Google Scholar

  • Schissel, S.L., Keesler, G.A., Schuchman, E.H., Williams, K.J., and Tabas, I. (1998b). The cellular trafficking and zinc dependence of secretory and lysosomal sphingomyelinase, two products of the acid sphingomyelinase gene. J. Biol. Chem. 273, 18250–18259.Google Scholar

  • Schneider, P.B. and Kennedy, E.P. (1967). Sphingomyelinase in normal human spleens and in spleens from subjects with Niemann-Pick disease. J. Lipid Res. 8, 202–209.Google Scholar

  • Schramm, M., Herz, J., Haas, A., Krönke, M., and Utermöhlen, O. (2008). Acid sphingomyelinase is required for efficient phago-lysosomal fusion. Cell Microbiol. 10, 1839–1853.PubMedCrossrefGoogle Scholar

  • Schuchman, E.H. (2007). The pathogenesis and treatment of acid sphingomyelinase-deficient Niemann-Pick disease. J. Inherit. Metab. Dis. 30, 654–663.CrossrefGoogle Scholar

  • Schuchman, E.H. (2010). Acid sphingomyelinase, cell membranes and human disease: lessons from Niemann-Pick disease. FEBS Lett. 584, 1895–1900.Google Scholar

  • Schuchman, E.H. and Miranda, S.R.P. (1997). Niemann-Pick disease: mutation update, genotype/phenotype correlations, and prospects for genetic testing. Genet. Test. 1, 13–19.Google Scholar

  • Schuchman, E.H., Suchi, M., Takahashi, T., Sandhoff, K., and Desnick, R.J. (1991). Human acid sphingomyelinase. Isolation, nucleotide sequence and expression of the full-length and alternatively spliced cDNAs. J. Biol. Chem. 266, 8531–8539.Google Scholar

  • Schwandner, R., Wiegmann, K., Bernardo, K., Kreder, D., and Krönke, M. (1998). TNF receptor death domain-associated proteins TRADD and FADD signal activation of acid sphingomyelinase. J. Biol. Chem. 273, 5916–5922.Google Scholar

  • Seidel, D., Klenke, J., Fischer, G., and Pilz, H. (1978). An improved and simple micro-method of sphingomyelinase assay in leukocytes and urine. J. Clin. Chem. Clin. Biochem. 16, 407–411.Google Scholar

  • Sharom, F.J. (2011). Flipping and flopping–lipids on the move. IUBMB Life 63, 736–746.PubMedGoogle Scholar

  • Silver, I.A., Murrills, R.J., and Etherington, D.J. (1988). Microelectrode studies on the acid microenvironment beneath adherent macrophages and osteoclasts. Exp. Cell Res. 175, 266–276.Google Scholar

  • Simon, C.G., Jr. and Gear, A.R.L. (1999). Sphingolipid metabolism during human platelet activation. Thromb. Res. 94, 13–23.CrossrefPubMedGoogle Scholar

  • Simon, C.G., Jr., Chatterjee, S., and Gear, A.R.L. (1998). Sphingomyelinase activity in human platelets. Thromb. Res. 90, 155–161.CrossrefPubMedGoogle Scholar

  • Simonaro, C.M., Desnick, R.J., McGovern, M.M., Wasserstein, M.P., and Schuchman, E.H. (2002). The demographics and distribution of type B Niemann-Pick disease: novel mutations lead to new genotype/phenotype correlations. Am. J. Hum. Genet. 71, 1413–1419.CrossrefGoogle Scholar

  • Slotte, J.P., Härmälä, A.S., Jansson, C., and Pörn, M.I. (1990). Rapid turn-over of plasma membrane sphingomyelin and cholesterol in baby hamster kidney cells after exposure to sphingomyelinase. Biochim. Biophys. Acta 1030, 251–257.Google Scholar

  • Smith, E.L. and Schuchman, E.H. (2008). The unexpected role of acid sphingomyelinase in cell death and the pathophysiology of common diseases. FASEB J. 22, 3419–3431.PubMedCrossrefGoogle Scholar

  • Sneck, M., Nguyen, S.D., Pihlajamaa, T., Yohannes, G., Riekkola, M.L., Milne, R., Kovanen, P.T., and Öörni, K. (2012). Conformational changes of apoB-100 in SMase-modified LDL mediate formation of large aggregates at acidic pH. J. Lipid Res. 53, 1832–1839.CrossrefGoogle Scholar

  • Sorbi, D., Fadly, M., Hicks, R., Alexander, S., and Arbeit, L. (1993). Captopril inhibits the 72 kDa and 92 kDa matrix metalloproteinases. Kidney Int. 44, 1266–1272.Google Scholar

  • Sot, J., Ibarguren, M., Busto, J.V., Montes, L.R., Goñi, F.M., and Alonso, A. (2008). Cholesterol displacement by ceramide in sphingomyelin-containing liquid-ordered domains, and generation of gel regions in giant lipidic vesicles. FEBS Lett. 582, 3230–3236.Google Scholar

  • Spatola, M. and Wider, C. (2014). Genetics of Parkinson’s disease: the yield. Parkinsonism Relat. Disord. 20 (Suppl 1), S35–S38.Google Scholar

  • Spence, M.W., Burgess, J.K., and Sperker, E.R. (1979). Neutral and acid sphingomyelinases: somatotopographical distribution in human brain and distribution in rat organs. A possible relationship with the dopamine system. Brain Res. 168, 543–551.Google Scholar

  • Spence, M.W., Byers, D.M., Palmer, F.B.S.C., and Cook, H.W. (1989). A new Zn2+-stimulated sphingomyelinase in fetal bovine serum. J. Biol. Chem. 264, 5358–5363.Google Scholar

  • Staneva, G., Chachaty, C., Wolf, C., Koumanov, K., and Quinn, P.J. (2008). The role of sphingomyelin in regulating phase coexistence in complex lipid model membranes: competition between ceramide and cholesterol. Biochim. Biophys. Acta 1778, 2727–2739.Google Scholar

  • Staneva, G., Momchilova, A., Wolf, C., Quinn, P.J., and Koumanov, K. (2009). Membrane microdomains: role of ceramides in the maintenance of their structure and functions. Biochim. Biophys. Acta 1788, 666–675.Google Scholar

  • Steinert, S., Lee, E., Tresset, G., Zhang, D., Hortsch, R., Wetzel, R., Hebbar, S., Sundram, J.R., Kesavapany, S., Boschke, E., et al. (2008). A fluorescent glycolipid-binding peptide probe traces cholesterol dependent microdomain-derived trafficking pathways. PLOS One 3, e2933.Google Scholar

  • Swardfager, W., Herrmann, N., Mazereeuw, G., Goldberger, K., Harimoto, T., and Lanctôt, K.L. (2013a). Zinc in depression: a meta-analysis. Biol. Psychiatry 74, 872–878.Google Scholar

  • Swardfager, W., Herrmann, N., McIntyre, R.S., Mazereeuw, G., Goldberger, K., Cha, D.S., Schwartz, Y., and Lanctôt, K.L. (2013b). Potential roles of zinc in the pathophysiology and treatment of major depressive disorder. Neurosci. Biobehav. Rev. 37, 911–929.CrossrefPubMedGoogle Scholar

  • Tabas, I. (1999). Secretory sphingomyelinase. Chem. Phys. Lipids 102, 123–130.Google Scholar

  • Tabas, I., Williams, K.J., and Borén, J. (2007). Subendothelial lipoprotein retention as the initiating process in atherosclerosis: update and therapeutic implications. Circulation 116, 1832–1844.Google Scholar

  • Takahashi, I., Takahashi, T., Abe, T., Watanabe, W., and Takada, G. (2000). Distribution of acid sphingomyelinase in human various body fluids. Tohoku J. Exp. Med. 192, 61–66.Google Scholar

  • Takahashi, T., Abe, T., Sato, T., Miura, K., Takahashi, I., Yano, M., Watanabe, A., Imashuku, S., and Takada, G. (2002). Elevated sphingomyelinase and hypercytokinemia in hemophagocytic lymphohistiocytosis. J. Pediatr. Hematol. Oncol. 24, 401–404.CrossrefGoogle Scholar

  • Takahashi, I., Takahashi, T., Mikami, T., Komatsu, M., Ohura, T., Schuchman, E.H., and Takada, G. (2005). Acid sphingomyelinase: relation of 93lysine residue on the ratio of intracellular to secreted enzyme activity. Tohoku J. Exp. Med. 206, 333–340.Google Scholar

  • Takatsu, H., Tanaka, G., Segawa, K., Suzuki, J., Nagata, S., Nakayama, K. and Shin, H.W. (2014). Phospholipid flippase activities and substrate specificities of human type IV P-type ATPases localized to the plasma membrane. J. Biol. Chem. 289, 33543–33556.Google Scholar

  • Takeda, A., Sawashita, J., and Okada, S. (1994). Localization in rat brain of the trace metals, zinc and manganese, after intracerebroventricular injection. Brain Res. 658, 252–254.Google Scholar

  • Takeda, A., Sawashita, J., and Okada, S. (1995). Biological half-lives of zinc and manganese in rat brain. Brain Res. 695, 53–58.Google Scholar

  • Tam, C., Idone, V., Devlin, C., Fernandes, M.C., Flannery, A., He, X., Schuchman, E., Tabas, I., and Andrews, N.W. (2010). Exocytosis of acid sphingomyelinase by wounded cells promotes endocytosis and plasma membrane repair. J. Cell Biol. 189, 1027–1038.Google Scholar

  • Tam, C., Flannery, A.R., and Andrews, N. (2013). Live imaging assay for assessing the roles of Ca2+ and sphingomyelinase in the repair of pore-forming toxin wounds. J. Vis. Exp. e50531.Google Scholar

  • Tamura, H., Takahashi, T., Ban, N., Torisu, H., Ninomiya, H., Takada, G., and Inagaki, N. (2006). Niemann-Pick type C disease: novel NPC1 mutations and characterization of the concomitant acid sphingomyelinase deficiency. Mol. Genet. Metab. 87, 113–121.Google Scholar

  • Tani, M. and Hannun, Y.A. (2007). Neutral sphingomyelinase 2 is palmitoylated on multiple cysteine residues. Role of palmitoylation in subcellular localization. J. Biol. Chem. 282, 10047–10056.Google Scholar

  • Tapper, H. and Sundler, R. (1992). Cytosolic pH regulation in mouse macrophages. Proton extrusion by plasma-membrane-localized H+-ATPase. Biochem. J. 281 (Pt 1), 245–250.Google Scholar

  • te Vruchte, D., Speak, A.O., Wallom, K.L., Al Eisa N., Smith, D.A., Hendriksz, C.J., Simmons, L., Lachmann, R.H., Cousins, A., Hartung, R., et al. (2014). Relative acidic compartment volume as a lysosomal storage disorder-associated biomarker. J. Clin. Invest. 124, 1320–1328.CrossrefGoogle Scholar

  • Testai, F.D., Landek, M.A., Goswami, R., Ahmed, M., and Dawson, G. (2004). Acid sphingomyelinase and inhibition by phosphate ion: role of inhibition by phosphatidyl-myo-inositol 3,4,5-triphosphate in oligodendrocyte cell signaling. J. Neurochem. 89, 636–644.Google Scholar

  • Thomas, G.H., Tuck-Muller, C.M., Miller, C.S., and Reynolds, L.W. (1989). Correction of sphingomyelinase deficiency in Niemann-Pick type C fibroblasts by removal of lipoprotein fraction from culture media. J. Inherit. Metab. Dis. 12, 139–151.CrossrefGoogle Scholar

  • Trajkovic, K., Hsu, C., Chiantia, S., Rajendran, L., Wenzel, D., Wieland, F., Schwille, P., Brügger, B., and Simons, M. (2008). Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science 319, 1244–1247.Google Scholar

  • Trapp, S., Rosania, G.R., Horobin, R.W., and Kornhuber, J. (2008). Quantitative modeling of selective lysosomal targeting for drug design. Eur. Biophys. J. 37, 1317–1328.PubMedCrossrefGoogle Scholar

  • Truman, J.P., Al Gadban, M.M., Smith, K.J., Jenkins, R.W., Mayroo, N., Virella, G., Lopes-Virella, M.F., Bielawska, A., Hannun, Y.A., and Hammad, S.M. (2012). Differential regulation of acid sphingomyelinase in macrophages stimulated with oxidized low-density lipoprotein (LDL) and oxidized LDL immune complexes: role in phagocytosis and cytokine release. Immunology 136, 30–45.Google Scholar

  • Utermöhlen, O., Karow, U., Löhler, J., and Krönke, M. (2003). Severe impairment in early host defense against Listeria monocytogenes in mice deficient in acid sphingomyelinase. J. Immunol. 170, 2621–2628.Google Scholar

  • van Meer, G., Voelker, D.R., and Feigenson, G.W. (2008). Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 9, 112–124.CrossrefGoogle Scholar

  • Van Wart, H.E. and Birkedal-Hansen, H. (1990). The cysteine switch: a principle of regulation of metalloproteinase activity with potential applicability to the entire matrix metalloproteinase gene family. Proc. Natl. Acad. Sci. USA 87, 5578–5582.Google Scholar

  • Vanha-Perttula, T. (1988). Sphingomyelinases in human, bovine and porcine seminal plasma. FEBS Lett. 233, 263–267.Google Scholar

  • Vanha-Perttula, T., Rönkkö, S., and Lahtinen, R. (1990). Hydrolases from bovine seminal vesicle, prostate and Cowper’s gland. Andrologia 22 (Suppl 1), 10–24.Google Scholar

  • Vanier, M.T., Revol, A., and Fichet, M. (1980). Sphingomyelinase activities of various human tissues in control subjects and in Niemann-Pick disease – development and evaluation of a microprocedure. Clin. Chim. Acta 106, 257–267.Google Scholar

  • Vashum, K.P., McEvoy, M., Milton, A.H., McElduff, P., Hure, A., Byles, J., and Attia, J. (2014). Dietary zinc is associated with a lower incidence of depression: findings from two Australian cohorts. J. Affect. Disord. 166, 249–257.Google Scholar

  • Veiga, M.P., Arrondo, J.L. R., Goñi, F.M., and Alonso, A. (1999). Ceramides in phospholipid membranes: effects on bilayer stability and transition to nonlamellar phases. Biophys. J. 76, 342–350.PubMedCrossrefGoogle Scholar

  • Verdurmen, W.P. R., Thanos, M., Ruttekolk, I.R., Gulbins, E., and Brock, R. (2010). Cationic cell-penetrating peptides induce ceramide formation via acid sphingomyelinase: implications for uptake. J. Control. Release 147, 171–179.Google Scholar

  • von Bismarck, P., Wistädt, C.F., Klemm, K., Winoto-Morbach, S., Uhlig, U., Schütze, S., Adam, D., Lachmann, B., Uhlig, S., and Krause, M.F. (2008). Improved pulmonary function by acid sphingomyelinase inhibition in a newborn piglet lavage model. Am. J. Respir. Crit. Care Med. 177, 1233–1241.Google Scholar

  • Wähe, A., Kasmapour, B., Schmaderer, C., Liebl, D., Sandhoff, K., Nykjaer, A., Griffiths, G., and Gutierrez, M.G. (2010). Golgi-to-phagosome transport of acid sphingomyelinase and prosaposin is mediated by sortilin. J. Cell Sci. 123, 2502–2511.Google Scholar

  • Walters, M.J. and Wrenn, S.P. (2011). Mechanistic roles of lipoprotein lipase and sphingomyelinase in low density lipoprotein aggregation. J. Colloid Interface Sci. 363, 268–274.Google Scholar

  • Wan, Q. and Schuchman, E.H. (1995). A novel polymorphism in the human acid sphingomyelinase gene due to size variation of the signal peptide region. Biochim. Biophys. Acta 1270, 207–210.Google Scholar

  • Wasserstein, M.P., Aron, A., Brodie, S.E., Simonaro, C., Desnick, R.J., and McGovern, M.M. (2006). Acid sphingomyelinase deficiency: prevalence and characterization of an intermediate phenotype of Niemann-Pick disease. J. Pediatr. 149, 554–559.Google Scholar

  • Watanabe, K., Sakuragawa, N., Arima, M., and Satoyoshi, E. (1983). Partial purification and properties of acid sphingomyelinase from rat liver. J. Lipid Res. 24, 596–603.Google Scholar

  • Weinreb, N.J., Brady, R.O., and Tappel, A.L. (1968). The lysosomal localization of sphingolipid hydrolases. Biochim. Biophys. Acta 159, 141–146.Google Scholar

  • Weitz, G., Lindl, T., Hinrichs, U., and Sandhoff, K. (1983). Release of sphingomyelin phosphodiesterase (acid sphingomyelinase). by ammonium chloride from CL 1D mouse L-cells and human fibroblasts. Partial purification and characterization of the exported enzymes. Hoppe-Seylers Z. Physiol. Chem. 364, 863–871.Google Scholar

  • Weitz, G., Driessen, M., Brouwer-Kelder, E.M., Sandhoff, K., Barranger, J.A., Tager, J.M., and Schram, A.W. (1985). Soluble sphingomyelinase from human urine as antigen for obtaining anti-sphingomyelinase antibodies. Biochim. Biophys. Acta 838, 92–97.Google Scholar

  • Wenger, D.A., Sattler, M., Clark, C., and Wharton, C. (1976). I-cell disease: activities of lysosomal enzymes toward natural and synthetic substrates. Life. Sci. 19, 413–420.CrossrefPubMedGoogle Scholar

  • Wong, M.L., Xie, B., Beatini, N., Phu, P., Marathe, S., Johns, A., Gold, P.W., Hirsch, E., Williams, K.J., Licinio, J., et al. (2000). Acute systemic inflammation up-regulates secretory sphingomyelinase in vivo: a possible link between inflammatory cytokines and atherogenesis. Proc. Natl. Acad. Sci. USA 97, 8681–8686.CrossrefGoogle Scholar

  • Wu, R.M., Lin, C.H., and Lin, H.I. (2014). The p. L302P mutation in the lysosomal enzyme gene SMPD1 is a risk factor for Parkinson disease. Neurology 82, 283.Google Scholar

  • Xu, M., Xia, M., Li, X.X., Han, W.Q., Boini, K.M., Zhang, F., Zhang, Y., Ritter, J.K., and Li, P.L. (2012). Requirement of translocated lysosomal V1 H+-ATPase for activation of membrane acid sphingomyelinase and raft clustering in coronary endothelial cells. Mol. Biol. Cell 23, 1546–1557.CrossrefGoogle Scholar

  • Yamamoto, K. (1994). Microbial endoglycosidases for analyses of oligosaccharide chains in glycoproteins. J. Biochem. 116, 229–235.Google Scholar

  • Yamanaka, T. and Suzuki, K. (1982). Acid sphingomyelinase of human brain: purification to homogeneity. J. Neurochem. 38, 1753–1764.CrossrefGoogle Scholar

  • Yook, K., Harris, T.W., Bieri, T., Cabunoc, A., Chan, J., Chen, W.J., Davis, P., de la Cruz, N., Duong, A., Fang, R., et al. (2012). WormBase 2012: more genomes, more data, new website. Nucleic Acids Res. 40, D735–D741.Google Scholar

  • Yoshida, Y., Arimoto, K., Sato, M., Sakuragawa, N., Arima, M., and Satoyoshi, E. (1985). Reduction of acid sphingomyelinase activity in human fibroblasts induced by AY-9944 and other cationic amphiphilic drugs. J. Biochem. (Tokyo) 98, 1669–1679.Google Scholar

  • Yu, J., Pan, W., Shi, R., Yang, T., Li, Y., Yu, G., Bai, Y., Schuchman, E.H., He, X., and Zhang, G. (2015). Ceramide is upregulated and associated with mortality in patients with chronic heart failure. Can. J. Cardiol., in press.Google Scholar

  • Zeidan, Y.H. and Hannun, Y.A. (2007a). Activation of acid sphingomyelinase by protein kinase Cδ-mediated phosphorylation. J. Biol. Chem. 282, 11549–11561.Google Scholar

  • Zeidan, Y.H. and Hannun, Y.A. (2007b). Translational aspects of sphingolipid metabolism. Trends Mol. Med. 13, 327–336.CrossrefPubMedGoogle Scholar

  • Zeidan, Y.H., Pettus, B.J., Elojeimy, S., Taha, T., Obeid, L.M., Kawamori, T., Norris, J.S., and Hannun, Y.A. (2006). Acid ceramidase but not acid sphingomyelinase is required for tumor necrosis factor-α-induced PGE2 production. J. Biol. Chem. 281, 24695–24703.Google Scholar

  • Zeidan, Y.H., Jenkins, R.W., and Hannun, Y.A. (2008a). Remodeling of cellular cytoskeleton by the acid sphingomyelinase/ceramide pathway. J. Cell Biol. 181, 335–350.Google Scholar

  • Zeidan, Y.H., Wu, B.X., Jenkins, R.W., Obeid, L.M., and Hannun, Y.A. (2008b). A novel role for protein kinase Cδ-mediated phosphorylation of acid sphingomyelinase in UV light-induced mitochondrial injury. FASEB J. 22, 183–193.PubMedGoogle Scholar

  • Zha, X., Pierini, L.M., Leopold, P.L., Skiba, P.J., Tabas, I., and Maxfield, F.R. (1998). Sphingomyelinase treatment induces ATP-independent endocytosis. J. Cell Biol. 140, 39–47.CrossrefGoogle Scholar

About the article

Corresponding author: Johannes Kornhuber, Department of Psychiatry and Psychotherapy, Friedrich Alexander University of Erlangen-Nürnberg (FAU), Schwabachanlage 6, D-91054 Erlangen, Germany, e-mail:

Received: 2015-01-20

Accepted: 2015-02-16

Published Online: 2015-03-24

Published in Print: 2015-06-01

Citation Information: Biological Chemistry, Volume 396, Issue 6-7, Pages 707–736, ISSN (Online) 1437-4315, ISSN (Print) 1431-6730, DOI: https://doi.org/10.1515/hsz-2015-0109.

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©2015, Johannes Kornhuber et al., published by De Gruyter. This work is licensed under the Creative Commons Attribution-NonCommercial-NoDerivatives 3.0 License. BY-NC-ND 3.0

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Scientific Reports, 2018, Volume 8, Number 1
Liubov S. Kalinichenko, Erich Gulbins, Johannes Kornhuber, and Christian P. Müller
Journal of Neural Transmission, 2018
Shuichiro Yoshida, Atsuko Noguchi, Wataru Kikuchi, Hiroshi Fukaya, Kiyoshi Igarashi, and Tsutomu Takahashi
The Tohoku Journal of Experimental Medicine, 2017, Volume 243, Number 4, Page 275
Ha-Yeun Chung, C. Julius Witt, Nayla Jbeily, Jorge Hurtado-Oliveros, Benjamin Giszas, Amelie Lupp, Markus H. Gräler, Tony Bruns, Andreas Stallmach, Falk A. Gonnert, and Ralf A. Claus
Scientific Reports, 2017, Volume 7, Number 1
Jingyuan Li, Ying Liu, Wei Li, Zhe Wang, Pan Guo, Lin Li, and Naijing Li
Behavioural Brain Research, 2017
Wing-Kee Lee and Richard N. Kolesnick
Cellular Signalling, 2017, Volume 38, Page 134
Alexei Gorelik, Katalin Illes, Giulio Superti-Furga, and Bhushan Nagar
Journal of Biological Chemistry, 2016, Volume 291, Number 12, Page 6376
Alexei Gorelik, Leonhard X. Heinz, Katalin Illes, Giulio Superti-Furga, and Bhushan Nagar
Journal of Biological Chemistry, 2016, Volume 291, Number 46, Page 24054
Alexei Gorelik, Fangyu Liu, Katalin Illes, and Bhushan Nagar
Journal of Biological Chemistry, 2017, Volume 292, Number 17, Page 7087
Thomas Pinkert, David Furkert, Thomas Korte, Andreas Herrmann, and Christoph Arenz
Angewandte Chemie, 2017, Volume 129, Number 10, Page 2834
Thomas Pinkert, David Furkert, Thomas Korte, Andreas Herrmann, and Christoph Arenz
Angewandte Chemie International Edition, 2017, Volume 56, Number 10, Page 2790
Joaquin Bobillo Lobato, Maria Jiménez Hidalgo, and Luis Jiménez Jiménez
Diseases, 2016, Volume 4, Number 4, Page 40
Richard S. Hoehn, Peter L. Jernigan, Lukasz Japtok, Alex L. Chang, Emily F. Midura, Charles C. Caldwell, Burkhard Kleuser, Alex B. Lentsch, Michael J. Edwards, Erich Gulbins, and Timothy A. Pritts
Annals of Surgery, 2017, Volume 265, Number 1, Page 218
Christiane Mühle and Johannes Kornhuber
Journal of Chromatography A, 2017, Volume 1481, Page 137
Maria Rachele Ceccarini, Michela Codini, Samuela Cataldi, Samuele Vannini, Andrea Lazzarini, Alessandro Floridi, Massimo Moretti, Milena Villarini, Bernard Fioretti, Tommaso Beccari, and Elisabetta Albi
Lipids in Health and Disease, 2016, Volume 15, Number 1
Cosima Rhein, Martin Reichel, Marcel Kramer, Andrea Rotter, Bernd Lenz, Christiane Mühle, Erich Gulbins, and Johannes Kornhuber
Journal of Affective Disorders, 2017, Volume 209, Page 10
Phillips-Farfán Bryan, Carvajal Karla, Medina-Torres Edgar Alejandro, Espinosa-Padilla Sara Elva, Fabrias Gemma, and Camacho Luz
Mediators of Inflammation, 2016, Volume 2016, Page 1
Alexei Gorelik, Katalin Illes, Leonhard X. Heinz, Giulio Superti-Furga, and Bhushan Nagar
Nature Communications, 2016, Volume 7, Page 12196
Zi-Jian Xiong, Jingjing Huang, Gennady Poda, Régis Pomès, and Gilbert G. Privé
Journal of Molecular Biology, 2016, Volume 428, Number 15, Page 3026
Joseph P. Huston, Johannes Kornhuber, Christiane Mühle, Lukasz Japtok, Mara Komorowski, Claudia Mattern, Martin Reichel, Erich Gulbins, Burkhard Kleuser, Bianca Topic, Maria A. De Souza Silva, and Christian P. Müller
Journal of Neurochemistry, 2016, Volume 137, Number 4, Page 589
Clara Di Vito, Loubna Abdel Hadi, Stefania Elena Navone, Giovanni Marfia, Rolando Campanella, Maria Elisa Mancuso, and Laura Riboni
Platelets, 2016, Volume 27, Number 5, Page 393
Yuuki Konno, Ikuko Takahashi, Ayuko Narita, Osamu Takeda, Hiromi Koizumi, Masamichi Tamura, Wataru Kikuchi, Akira Komatsu, Hiroaki Tamura, Satoko Tsuchida, Atsuko Noguchi, and Tsutomu Takahashi
The Tohoku Journal of Experimental Medicine, 2015, Volume 237, Number 2, Page 133
Motohide Murate and Toshihide Kobayashi
Chemistry and Physics of Lipids, 2016, Volume 194, Page 58
Gemma Fabrias and Richard M. Epand
Chemistry and Physics of Lipids, 2016, Volume 197, Page 1

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