From the Departments of Anatomy & Cell Biology and
Medicine, Columbia University,
New York, New York 10032, ¶ Amgen, Boulder, Colorado 80301, the
Department of Human Genetics, Mount Sinai School
of Medicine, New York, New York 10029, and the ** Dorrance H. Hamilton Research Laboratories, Division of Endocrinology, Diabetes,
and Metabolic Diseases, Thomas Jefferson University,
Philadelphia, Pennsylvania 19107
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ABSTRACT |
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The acid sphingomyelinase (ASM) gene, which has been implicated in ceramide-mediated cell signaling and atherogenesis, gives rise to both lysosomal SMase (L-SMase), which is reportedly cation-independent, and secretory SMase (S-SMase), which is fully or partially dependent on Zn2+ for enzymatic activity. Herein we present evidence for a model to explain how a single mRNA gives rise to two forms of SMase with different cellular trafficking and apparent differences in Zn2+ dependence. First, we show that both S-SMase and L-SMase, which contain several highly conserved zinc-binding motifs, are directly activated by zinc. In addition, SMase assayed from a lysosome-rich fraction of Chinese hamster ovary cells was found to be partially zinc-dependent, suggesting that intact lysosomes from these cells contain subsaturating levels of Zn2+. Analysis of Asn-linked oligosaccharides and of N-terminal amino acid sequence indicated that S-SMase arises by trafficking through the Golgi secretory pathway, not by cellular release of L-SMase during trafficking to lysosomes or after delivery to lysosomes. Most importantly, when Zn2+-dependent S-SMase was incubated with SMase-negative cells, the enzyme was internalized, trafficked to lysosomes, and became zinc-independent. We conclude that L-SMase is exposed to cellular Zn2+ during trafficking to lysosomes, in lysosomes, and/or during cell homogenization. In contrast, the pathway targeting S-SMase to secretion appears to be relatively sequestered from cellular pools of Zn2+; thus S-SMase requires exogeneous Zn2+ for full activity. This model provides important information for understanding the enzymology and regulation of L- and S-SMase and for exploring possible roles of ASM gene products in cell signaling and atherogenesis.
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INTRODUCTION |
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SMases1 (SM phosphodiesterase, EC 3.1.4.12) have been implicated in a wide variety of physiologic and pathophysiologic processes, including lysosomal hydrolysis of endocytosed SM (1, 2), ceramide-mediated cell signaling (3, 4), membrane vesiculation (5, 6), alterations in intracellular cholesterol trafficking (5, 7-9), and atherogenesis (10-13). One type of mammalian SMase is a magnesium-dependent, membrane-bound neutral SMase, and Tomiuk et al. (14) have recently reported the cloning of an enzyme that has several properties in common with this SMase. Two other types of mammalian SMases are lysosomal SMase (L-SMase) and secretory SMase (S-SMase), both of which arise from the "acid SMase" or "ASM" gene (15, 16). Both enzymes are soluble hydrolases that function optimally at acid pH in a standard in vitro micellar assay (16, 17), although we have shown that S-SMase can hydrolyze physiologic SM-containing substrates at neutral pH (Ref. 18 and see below). Both L- and S-SMase are absent from the cells of patients with types A and B Niemann-Pick disease, which is due to mutations in the ASM gene, and from the cells of ASM knock-out mice (16).
S-SMase may have significant physiologic roles, since extracellular SM hydrolysis may be involved in some or all of the non-lysosomal processes listed above. For example, several lines of evidence have implicated extracellular SM hydrolysis in atherogenesis. First, treatment of LDL with SMase in vitro leads to LDL aggregation (10, 11), which is a prominent event during atherogenesis (19-21) and one that leads to massive macrophage foam cell formation (10, 11, 22-24). Second, aggregated LDL from human and animal atherosclerotic lesions shows evidence of hydrolysis by extracellular SMase, and LDL retained in rabbit aortic strips ex vivo is hydrolyzed by an extracellular, cation-dependent SMase (12). Third, S-SMase, a leading candidate for this arterial wall enzyme, is secreted by macrophages (16) and endothelial cells (25), cell types found in atherosclerotic lesions. Fourth, S-SMase is able to hydrolyze the SM in atherogenic lipoproteins at neutral pH (18). Other possible roles for S-SMase may be in ceramide-mediated cell signaling (26-30), perhaps after re-uptake of the secreted enzyme into endosomal vesicles; in extracellular sphingomyelin catabolism after nerve injury and during demyelination (16, 31-33); and in defense against viruses, many of which are enriched in SM (34, 35) and can be inactivated by treatment with SMase in vitro.2
L- and S-SMase are very similar proteins. Previous work from our laboratories has shown that cells transfected with an ASM cDNA overexpress both L-SMase and S-SMase (16), indicating that S-SMase does not arise by alternative processing of the ASM gene. In addition, antibodies made against L-SMase recognize S-SMase, demonstrating that the common mRNA is translated in the same reading frame, and the molecular weights of the enzymes on Western blot are similar (see Ref. 16 and below). Nevertheless, S-SMase requires exogenously added Zn2+ for activation in in vitro assays, whereas L-SMase isolated from cell or tissue homogenates does not (16). In fact, the lack of stimulation of L-SMase by any cations and its lack of inhibition by EDTA has led to a long-standing body of literature labeling L-SMase as a "cation-independent" enzyme (1).
Despite the widespread interest in mammalian SMases in general and in products of the ASM gene in particular, little is known about cellular itineraries of L-SMase and S-SMase or about the mechanism for their apparent difference in zinc dependence. For example, does S-SMase arise by release or exocytosis of L-SMase from lysosomes or by a separate trafficking pathway, and how could two enzymes that are so similar differ in their requirement for zinc? In this report, we show that S-SMase is secreted through a non-lysosomal secretory pathway, and we present evidence that both forms of the enzyme are zinc-activated. According to our model, L-SMase is exposed to cellular Zn2+ during trafficking to lysosomes, in lysosomes, and/or during cell homogenization. Most likely, the Zn2+ dependence of L-SMase has been overlooked because it is already saturated with Zn2+ upon isolation from cell homogenates and thus does not respond to exogenous Zn2+ at the time of assay. Furthermore, as is the case with known zinc metalloenzymes (cf. Ref. 36), the Zn2+ cannot be stripped from L-SMase by simple exposure to EDTA. In contrast, the pathway targeting S-SMase to secretion appears to be relatively sequestered from cellular pools of Zn2+. Thus, this enzyme requires Zn2+ during subsequent in vitro assay. The information in this report should prove useful for future studies that explore the enzymology, regulation, and functions of these important SMases.
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EXPERIMENTAL PROCEDURES |
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Materials--
The Falcon tissue culture plasticware used in
these studies was purchased from Fisher. Tissue culture media and other
tissue culture reagents were obtained from Life Technologies, Inc.
Fetal bovine serum was obtained from HyClone Laboratories (Logan, UT) and was heat-inactivated for 1 h at 65 °C (HI-FBS).
[9,10-3H]Palmitic acid (56 Ci/mmol) was purchased from
NEN Life Science Products, and
[N-palmitoyl-9-10-3H]sphingomyelin
was synthesized as described previously (16, 37, 38).
N,N-Dimethylformamide, 1,3-dicyclohexylcarbodiimide, N-hydroxysuccinimide, and
N,N-diisopropylethylamine were purchased from Aldrich.
Precast 4-20% gradient polyacrylamide gels were purchased from NOVEX
(San Diego, CA). Nitrocellulose was from Schleicher & Schuell. Rabbit
anti-FLAG-tagged S-SMase was kindly provided by Dr. Henri Lichenstein
(Amgen, Boulder, CO). FLAG-tagged S-SMase was purified by anti-FLAG
immunoaffinity chromatography from the conditioned medium of CHO cells
transfected with a human ASM-FLAG cDNA. FLAG-tagged L-SMase was
purified from a 16,000 × g pellet (see below) of
FLAG-ASM-transfected CHO cells using anti-FLAG immunoaffinity
chromatography, Superose-12 gel filtration chromatography, and a second
round of anti-FLAG immunoaffinity chromatography. Peroxidase-conjugated
goat anti-rabbit IgG was purchased from Pierce. Tissue inhibitor of
metalloproteinase-1 (TIMP-1) was a kind gift from Dr. Yasunori Okada
(Kanazawa University, Kanazawa, Japan). The thiol-based
metalloproteinase inhibitors, HS-CH2-R-CH(CH2-CH(CH3)2)-C)-Phe-Ala-NH2
and
HO-NH-CO-CH2-CH(CH2CH(CH3)2)-C)-naphthyl-Ala-NH-CH2-CH2-NH2, were purchased from Peptides International, Inc. (Louisville, KY). -Endo-N-acetylglucosaminidase H (endo H) and
peptide-N-glycanase F were purchased from Boehringer
Mannheim. Bovine liver 215-kDa mannose 6-phosphate receptor linked to
Affi-Gel 15 was made as described by Varki and Kornfeld (39) and was
kindly supplied by Walter Gregory and Dr. Stuart Kornfeld (Washington
University, St. Louis). Sphingosylphosphorylcholine,
1,10-phenanthroline, and all other chemicals and reagents were from
Sigma, and all organic solvents were from Fisher.
Cells-- Monolayer cultures of J774.A1 cells (from the American Type Culture Collection, see Ref. 40) were grown and maintained in spinner culture with DMEM/HI-FBS/PSG as described previously (9, 40). Human skin fibroblasts obtained from a patient with type A Niemann-Pick disease (R496L mutation (41)) were grown in DMEM/HI-FBS/PSG. CHO-K1 cells were grown in Hams' F-12 containing 10% HI-FBS and PSG. CHO cells stably transfected with ASM cDNA3 were maintained in DMEM/HI-FBS/PSG (16). Cells were plated in 35-mm (6-well) or 100-mm dishes in media containing HI-FBS for 48 h. The cells were then washed 3 times with PBS and incubated for 24 h in fresh serum-free media (1 and 6 ml per 35-mm and 100-mm dishes, respectively) containing 0.2% BSA. This 24-h conditioned medium was collected for SMase assays.
Harvesting of Cells and Conditioned Media-- Following the incubations described above and in the figure legends, cells were placed on ice, and the serum-free conditioned media were removed. The cells were washed two times with ice-cold 0.25 M sucrose and scraped into 0.3 and 3.0 ml of this sucrose solution per 35- and 100-mm dishes, respectively. Unless indicated otherwise, the scraped cells were disrupted by sonication on ice using three 5-s bursts (Branson 450 Sonifier), and the cellular homogenates were assayed for total protein by the method of Lowry et al. (42) and for SMase activity as described below. The conditioned media were spun at 800 × g for 5 min to pellet any contaminating cells and concentrated 6-fold using a Centriprep 30 (Amicon; Beverly, MA) concentrator (molecular weight cut-off = 30,000). For the experiment in Fig. 5, CHO-K1 cells were incubated in 100-mm dishes in serum-free media and washed as described above. Cells were then scraped in 5 ml of 0.25 M sucrose and broken open under 500 p.s.i. of nitrogen pressure for 1.5 min using a nitrogen cell disruption bomb (Parr Instruments, Moline, IL). Following disruption, a portion of the cells was subjected to brief sonication as described above; this portion of cells is referred to as the cell homogenate. The remainder of disrupted cells was spun at 1300 × g for 5 min to pellet any remaining intact cells and nuclei. This post-nuclear supernatant was collected, and the volume was increased to 15 ml with 0.25 M sucrose and then spun at 16,000 × g for 30 min. The pellet from this centrifugation was resuspended in 1 ml of 0.25 M sucrose and sonicated as above, and this material, as well as the cell homogenate, was assayed for SMase activity.
SMase Assay-- As described previously (16), the standard 200-µl assay mixture consisted of up to 40 µl of sample (conditioned media or homogenized cells; see above) and a volume of assay buffer (0.1 M sodium acetate, pH 5.0) to bring the volume to 160 µl. The reaction was initiated by the addition of 40 µl of substrate (50 pmol of [3H]sphingomyelin) in 0.25 M sucrose containing 3% Triton X-100 (final concentration of Triton X-100 in the 200-µl assay mix = 0.6%). When added, the final concentrations of EDTA and Zn2+ were 5 and 0.1 mM, respectively, unless indicated otherwise. The assay mixtures were incubated at 37 °C for no longer than 3 h and then extracted by the method of Bligh and Dyer (43); the lower, organic phase was harvested, evaporated under N2, and fractionated by TLC using chloroform/methanol (95:5). The ceramide spots were scraped and directly counted to quantify [3H]ceramide. Typically, our assay reactions contained approximately 20 µg of cellular homogenate protein and a volume of conditioned media derived from a quantity of cells equivalent to approximately 50 µg of cellular protein.
SDS-Polyacrylamide Gel Electrophoresis and Immunoblotting-- Protein samples were boiled in buffer containing 1% SDS and 10 mM dithiothreitol for 10 min, loaded onto 4-20% gradient polyacrylamide gels, and electrophoresed for 50 min at 35 mA in buffer containing 0.1% SDS (SDS-PAGE). Following electrophoresis, some gels were fixed in methanol/glacial acetic acid/water (5:2:3, v/v) and then silver-stained using reagents from Bio-Rad. Other gels were electrotransferred (100 V for 1.5 h) to nitrocellulose for immunoblotting. For immunoblotting, the nitrocellulose membranes were incubated with 5% Carnation nonfat dry milk in buffer A (24 mM Tris, pH 7.4, containing 0.5 M NaCl) for 3 h at room temperature. The membranes were then incubated with rabbit anti-FLAG-tagged S-SMase polyclonal antiserum (1:2000) in buffer B (buffer A containing 0.1% Tween 20, 3% nonfat dry milk, and 0.1% bovine serum albumin) for 1 h at room temperature. After washing four times with buffer A containing 0.1% Tween 20, the blots were incubated with horseradish peroxidase-conjugated goat anti-rabbit IgG (1:2000) for 1 h in buffer B at room temperature. The membranes were subsequently washed twice with 0.3% Tween 20 in buffer A and twice with 0.1% Tween 20 in buffer A. Finally, the blots were soaked in the enhanced chemiluminescence reagent (NEN Life Science Products) for 2 min and exposed to x-ray film for 1 min.
Glycosidase Treatments--
We followed the procedure described
by Hurwitz et al. (44). CHO-K1 cells were incubated
overnight with serum-free medium (CHO-S-SFM II from Life Technologies,
Inc.). Fifty µg of 30-fold-concentrated conditioned medium and 50 µg of cell homogenate were diluted 1:1 (v/v) with 50 mM
sodium acetate buffer, pH 5.0, containing 2% SDS and 20 mM
-mercaptoethanol (44). One set of aliquots of the diluted
conditioned medium and cell homogenate was treated for 16 h at
37 °C with 4 milliunits of endo H. Another set of aliquots was
diluted another 15-fold with 50 mM sodium phosphate buffer,
pH 7.2, containing 1% Nonidet P-40 and treated for 16 h at
37 °C with 100 milliunits of peptide-N-glycanase F. The
endo H digest and a trichloroacetic acid pellet of the
peptide-N-glycanase F digest (44) were boiled in
SDS/dithiothreitol buffer and then electrophoresed and immunoblotted as
described above.
Zinc-Chelate Chromatography-- We used a modification of the method of Hortin and Gibson (45). Packed 1-ml columns of chelating Sepharose 6B (iminodiacetic acid coupled to agarose gel via a hydrophilic spacer; from Amersham Pharmacia Biotech) were washed with 10 mM sodium acetate buffer, pH 6.0, containing 10 mM EDTA, to leave the column uncharged, or containing 60 mM ZnCl2, to charge the column with Zn2+. The columns were then equilibrated with 50 mM Hepes buffer, pH 7.4, containing 50 mM NaCl. One-ml samples of a 1:1 (v/v) mixture of this equilibration buffer and unconcentrated conditioned medium from ASM-transfected CHO cells (above) were loaded onto the columns and incubated for 15 min at room temperature. The columns were then washed with 7.5 ml of 50 mM Hepes, pH 7.4, containing either 100 mM NaCl or 1 M NaCl, which was collected as 10 0.75-ml fractions. The columns were eluted with 3.75 ml of 50 mM Hepes, pH 7.4, containing 50 mM EDTA plus 1 mM 1,10-phenanthroline, which was collected as 5 0.75-ml fractions. Aliquots of each of the fractions were spotted on nitrocellulose using a slot-blot apparatus and then immunoblotted using goat anti-human L-SMase polyclonal antiserum as described above.
Statistics-- Unless otherwise indicated, results are given as means ± S.D. (n = 3); absent error bars in the figures signify S.D. values smaller than the graphic symbols.
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RESULTS |
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Zn2+ Requirement for S-SMase Does Not Involve a Zn2+-dependent Cofactor-- We sought to address how L- and S-SMase acquire their apparent differences in zinc dependence. One explanation would be that the secreted form requires a Zn2+-dependent cofactor. Because many lysosomal enzymes undergo proteolytic activation (46), an obvious candidate for a Zn2+-dependent cofactor would be a zinc metalloproteinase. Five sets of results, however, ruled out this possibility. First, Zn2+-activated S-SMase can be subsequently inactivated by Zn2+ chelation (see below); reversibility of Zn2+-induced activation is not consistent with proteolytic activation. Second, inhibitors of zinc metalloproteinases, such as tissue inhibitor of metalloproteinase-1 (TIMP-1) (47) and two different thiol-based peptide inhibitors, HS-CH2-R-CH(CH2-CH(CH3)2)-C)-Phe-Ala-NH2 and HO-NH-CO-CH2-CH(CH2CH(CH3)2)-C)-naphthyl-Ala-NH-CH2-CH2-NH2 (48), did not affect the ability of Zn2+ to activate S-SMase (data not shown). Third, mammalian zinc metalloproteinases require Ca2+ as well as Zn2+ for activity (49), whereas Ca2+ is not a requirement for the activation of S-SMase (16). Fourth, comparison of Zn2+-activated S-SMase from CHO cells with that of the intracellular (lysosomal) enzyme by immunoblot analysis showed that the activated secreted form had a somewhat higher, not lower, apparent molecular weight (see control data in Fig. 3, below); in addition, S-SMase not activated with Zn2+ had the same apparent molecular weight as Zn2+-activated S-SMase (data not shown). Fifth, we found that highly purified S-SMase, obtained by either anti-FLAG immunoaffinity purification of a FLAG-tagged S-SMase or by concanavalin A chromatography followed by anti-SMase immunoaffinity purification of S-SMase from ASM-transfected CHO cells (16), was ~95% Zn2+-dependent. Thus, neither a zinc metalloproteinase nor any other Zn2+-dependent cofactor appears to be involved in the activation of S-SMase, suggesting direct activation of S-SMase by Zn2+.
To support this conclusion further, we sought to demonstrate that S-SMase directly binds Zn2+ by subjecting conditioned media from ASM-transfected CHO cells (16) to zinc-chelate chromatography (cf. Ref. 45 and "Experimental Procedures"). None of the S-SMase from the conditioned medium bound to an uncharged column, whereas >95% of the S-SMase bound to a Zn2+-charged column, even when washed with buffer containing 1 M NaCl; all of the bound material was eluted by EDTA plus 1,10-phenanthroline. These data and the previous data are consistent with the conclusion that S-SMase binds and is directly activated by Zn2+.Evidence for Direct Activation of L-SMase by Zn2+-- Despite the long-standing tenet that L-SMase is a cation-independent enzyme (2), several lines of evidence initially suggested to us that L-SMase was a zinc-activated enzyme. First, L-SMase and S-SMase come from the same gene and same mRNA in the same reading frame (16), and S-SMase binds and is directly activated by Zn2+ (above). Second, there are seven aminoacyl sequences in the enzyme that are homologous to Zn2+-binding sequences in known zinc metalloenzymes (50), including one sequence that is very similar to that in another phosphodiesterase enzyme (Table I). Histidine residues in two of these sequences (His-425 and His-575) are highly conserved and are sites of mutations in Niemann-Pick disease.3 Moreover, His-421 is conserved in mouse ASM and in a homologue of ASM in Caenorhabditis elegans that is zinc-dependent but not in an ASM homologue in C. elegans that is zinc-independent (51). Third, L-SMase shares two other properties of known zinc metalloenzymes, namely inhibition by phosphate ions (1, 2), which are thought to block the Zn2+-binding pocket(s) in zinc metalloenzymes (52) and inhibition by high concentrations (e.g. 6 mM) of ZnCl2 (49, 53).
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The Difference in Requirement for Zn2+ in the in Vitro Assays of L- and S-SMase: Differential Zn2+ Affinity Versus Differential Exposure to Cellular Zn2+ Prior to the Assay-- One possible explanation for the difference in Zn2+ requirement in the in vitro assays of L- and S-SMase is that the two enzymes would both be exposed to the same, although limiting, concentration of intracellular Zn2+ but that the lysosomal enzyme would have a higher affinity for the cation, perhaps owing to a difference in post-translation modification. Thus, L-SMase would already have bound Zn2+ at the time of the assay. The secreted enzyme would have lower affinity for Zn2+, and thus excess exogenous Zn2+ would have to be added for activation in vitro.
We sought to estimate the relative Zn2+ affinities of these two enzymes by assaying their inactivation as a function of increasing exposure to metal chelators (36). Therefore, we incubated a cellular homogenate of J774 macrophages and the conditioned medium from these cells with EDTA plus the 1,10-phenanthroline for increasing times at 4 °C and then assayed these two fractions for SMase activity at each time point. As expected (above), both enzymes lost activity with increasing duration of chelation (Fig. 2), whereas incubation in the absence of the chelators for 8 h at 4 °C resulted in no loss of either secreted or cellular SMase activity (not shown). The data show that cellular SMase activity decreased at a greater rate and to a greater extent than secreted SMase activity, which is not consistent with the hypothesis that L-SMase has a higher affinity for Zn2+ than S-SMase.
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L-SMase Activity from a Lysosome-rich 16,000 × g Pellet of CHO Cells Demonstrates Some Zinc Dependence-- The Zn2+ dependence of L-SMase and previous work demonstrating discrete intracellular Zn2+ pools that can change under certain metabolic conditions (cf. Refs. 70 and 71) led us to consider the hypothesis that Zn2+ availability to lysosomes and to L-SMase might be involved in the regulation of this enzyme. A prediction of our hypothesis is that L-SMase may not always be maximally stimulated by intracellular Zn2+. In the standard L-SMase assay, cells or tissues are completely homogenized, and the cell homogenate is assayed. As shown in Fig. 5A for CHO-K1 cells disrupted by sonication, the intracellular enzyme is maximally activated, and exogenous Zn2+ has no effect (Fig. 5A). To obtain a less damaged lysosomal preparation, a separate aliquot of these CHO cells was disrupted under 500 p.s.i. of nitrogen pressure for 1.5 min, and a 16,000 × g pellet was isolated, which consists of intact lysosomes, as well as mitochondria and peroxisomes (cf. Ref. 59).2 This 16,000 × g pellet was then sonicated and assayed for SMase activity. Remarkably, under these conditions, the enzyme was only ~50% activated and was substantially stimulated by exogenous Zn2+ (Fig. 5A).
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DISCUSSION |
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Our model to explain the cellular trafficking and apparent difference in Zn2+ dependence of L-SMase versus S-SMase is shown in Fig. 6. Based upon our previous work, the ASM gene gives rise to a common precursor protein (16), which is then modified by typical high mannose oligosaccharide residues (44, 62, 63). We propose that this mannosylated precursor then traffics into either the lysosomal or the secretory pathway. In the lysosomal pathway, the SMase undergoes modification and trafficking that is typical for lysosomal enzymes: acquisition of mannose-phosphate residues by the sequential action of N-acetylglucosamine-1-phosphotransferase and N-acetylglucosamine phosphodiesterase on the mannose residues of the precursor (Fig. 3 and Refs. 44 and 63). Vesicles containing mannose-phosphate receptors then shuttle this modified SMase to early endosomes or late endosomes/prelysosomes (63, 72, 73), and we propose that at some point along this pathway the enzyme encounters cellular Zn2+ and thus becomes at least partially activated. As mentioned under "Results," L-SMase, at least in CHO cells, appears to be exposed to subsaturating concentrations of Zn2+ in lysosomes and thus is potentially subject to regulation by changes in Zn2+ availability.
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L-SMase has been studied for many years, particularly in the context of its absence in a human disease, namely types A and B Niemann-Pick disease (1, 2). Throughout this period of study, the enzyme has been reported to be cation-independent (1, 2). Although plasma emission spectrometry and x-ray crystallography of large amounts of homogeneously purified L-SMase will be needed to define the stoichiometry and location of zinc interaction with L-SMase, the data in this report strongly support the conclusion that this enzyme is, indeed, a zinc-activated enzyme. The most compelling data are those in Fig. 1B, Fig. 5, and Table I, especially footnotes d and e. In fact, the information in Table I raises the possibility that some cases of Niemann-Pick disease may be due to mutations in the zinc-binding domain, possibly resulting in defective binding of Zn2+ and thus loss of enzymatic activity. Along these lines, He et al.3 have shown that chelation of Zn2+ from SMase by 1,10-phenanthroline results in defective SM binding to the enzyme. We believe the reason why this fundamental property of this widely studied enzyme has been overlooked is because the enzyme at the time of isolation from whole cell homogenates, which has been the source of L-SMase for the previous studies (54-59), is already tightly bound to Zn2+. In view of the data in Fig. 5, some of this Zn2+ may come from pools of zinc that are released during the homogenization of cells or tissues. Thus, exogenous Zn2+ is not needed for the in vitro assay, and typical short-term EDTA chelation incubations will not strip the enzyme of its metal, similar to findings with other known zinc metalloenzymes (36).
To explain the origin of S-SMase, we propose that a portion of the common precursor, via a potentially regulated process (see below), bypasses N-acetylglucosamine-1-phosphotransferase and thus is directed into the secretory pathway, not the lysosomal targeting pathway (63) (Fig. 6). The difference in susceptibility of S- and L-SMase to endo H (Fig. 3) and the differences in N-terminal proteolytic processing (see "Results") provide direct support for this component of the model. Importantly, our data suggest that SMase in the secretory pathway is not exposed to pools of cellular Zn2+, thus explaining the requirement for exogenously added Zn2+ when the secreted enzyme is assayed in vitro. As mentioned under "Results," however, the subcellular location of Zn2+ may be subject to cell-type variation or regulation (60, 70, 71). For example, recent data from Vallee and colleagues (74) suggest that the redox state of the cell may be an important factor in the transfer of zinc from metallothionein, a cellular zinc reservoir, to intracellular zinc-dependent enzymes. Therefore, it is possible that S-SMase may, under certain circumstances or in certain cell types, be fully or partially Zn2+-independent. In fact, we have observed that SMase secreted by endothelial cells, unlike that secreted by macrophages (16), is active in the absence of Zn2+ and stimulated only 2-fold by exposure to exogenous Zn2+ (25).
According to this model and work by other researchers (46, 63), the key step that would determine the fate of SMase is catalysis of the common mannosylated precursor by N-acetylglucosamine-1-phosphotransferase. Extensive work by Kornfeld and colleagues (75-77) has shown that N-acetylglucosamine-1-phosphotransferase recognizes a particular three-dimensional structure of lysosomal enzyme precursors, and induced modifications that alter this structure can have profound effects on lysosomal enzyme modification and targeting. Moreover, these workers have found that at least one enzyme, bovine DNase I, is a suboptimal substrate for the phosphotransferase, thus presumably giving rise to both intralysosomal and secretory forms (78)8 If the enzymes that undergo secretion by this mechanism can function at neutral pH (see below) or if the cells are in an acidic environment, this process may enable cells to acquire two groups of functions from a single enzyme, namely functions in lysosomes and functions in the extracellular milieu. In the case of S-SMase, there is an additional requirement for extracellular Zn2+, which is known to exist in sufficient extracellular concentrations in vivo to activate the enzyme (cf. Refs. 16 and 53). Interestingly, we found that certain cytokines increase the secretion of SMase from endothelial cells without affecting L-SMase activity, suggesting that the phosphotransferase reaction or perhaps another critical step responsible for determining the fate of SMase may be subject to specific regulation (25). Finally, C. elegans has two separate genes that encode SMases that are highly homologous to mammalian S- and L-SMase; one of these SMases is almost entirely secreted and the other is mostly intracellular (51). Thus, organisms evidently need both intracellular and extracellular SMases; C. elegans has two genes to meet these needs, and it appears as if higher species (i.e. mammals) have evolved the mechanism described above to meet these needs using one gene.
Our current data and previous work by others (79) indicate difficulties with the prior nomenclature of these SMases. First, we now know that both forms of the enzymes are zinc-activated enzymes, and so our previous designation of the secreted form as "Zn-SMase" (16) is obsolete. Second, the "acid SMase" nomenclature reflects the acid pH optima of the lysosomal and secreted forms of the enzyme in standard in vitro detergent-based micellar assays and the ability of the lysosomal form to function in the acid environment of lysosomes (1). Kinetic studies, however, have shown that acid pH is needed only for proper interaction of the enzyme with the SM in these micelles (i.e. Km) and that Vmax for hydrolysis is relatively pH-independent (79). Furthermore, we have recently demonstrated that S-SMase can hydrolyze the SM of certain lipoproteins quite well at neutral pH (18). Thus, the SM in certain physiological substrates may be in an orientation that allows ready interaction with the enzyme at neutral pH, which, based upon the above-mentioned kinetic data, would then result in neutral SM hydrolysis. For these reasons, and since lysosomes and conditioned media of cells contain no other known SMase activity (1, 2, 16, 80, 81), we suggest that the nomenclature in this paper (L-SMase and S-SMase) is preferred. To maintain consistency with prior literature, however, we still refer to their common gene of origin as the ASM gene.
The original impetus for the current mechanistic study was evidence gathered by our laboratories supporting a role for an extracellular arterial wall SMase in atherogenesis (10-13). As outlined in the Introduction, S-SMase is a leading candidate for this arterial wall activity. Furthermore, there is evidence that one or more products of the ASM gene triggers ceramide-mediated cell-signaling processes (14, 26-30), and S-SMase is a candidate since it would have access to the extracellular leaflet of the plasma membrane, which is where most cellular sphingomyelin is located (42). It is also possible that S-SMase plays roles in extracellular SM catabolism in the central nervous system and in anti-viral host defense mechanisms (see Introduction). In this context, the information reported herein on the cellular trafficking and zinc dependence of the S-SMase and L-SMase should prove useful in further regulatory studies on these enzymes and in designing strategies to test their possible roles in atherogenesis, ceramide-mediated cell signaling, and possibly other physiologic and pathophysiologic processes.
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ACKNOWLEDGEMENTS |
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We thank Xingxuan He for preparing the CHO cells overexpressing L- and S-SMase and for the purified preparations of S-SMase from the conditioned medium of these cells; Dr. Henri Lichenstein (Amgen, Boulder, CO) for the anti-FLAG-tagged S-SMase antiserum; Dr. Yasunori Okada (Kanazawa University, Kanazawa, Japan) for the TIMP-1; Dr. Stuart Kornfeld and Walter Gregory (Washington University, St. Louis, MO) for the mannose 6-phosphate receptor resin; and Dr. Kornfeld for helpful discussions related to lysosomal enzyme targeting.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grants HL39703 (to I. T.), HL 21006 (to I. T.), HD28607 (to E. H. S.), and HL38956 (to K. J. W.); March of Dimes Birth Defects Foundation Basic Research Grant 1-1224 (to E. H. S.); and an Established Investigator award from the American Heart Association and Genentech (to K. J. W.). This work was also supported in part by a research grant from Berlex Biosciences.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ Supported by National Institutes of Health Medical Scientist Training Grant 5T32GM07367.
§§ To whom correspondence and reprint requests should be addressed: Dept. of Medicine, Columbia University, 630 West 168th St., New York, NY 10032. Tel.: 212-305-9430; Fax: 212-305-5052; E-mail: iat1{at}columbia.edu.
1
The abbreviations used are: SMase,
sphingomyelinase; S-SMase, secreted sphingomyelinase; ASM, acid
sphingomyelinase; CHO, Chinese hamster ovary; DMEM, Dulbecco's
modified Eagle's medium; endo H,
-endo-N-acetylglucosaminidase H; HI-FBS, heat-inactivated fetal bovine serum; LDL, low density lipoprotein; L-SMase, lysosomal sphingomyelinase; PBS, phosphate-buffered saline; PSG, penicillin, streptomycin, and glutamine; PAGE, polyacrylamide gel electrophoresis; SM, sphingomyelin; TIMP-1, tissue inhibitor of metalloproteinase-1; BSA, bovine serum albumin.
2 S. L. Schissel and I. Tabas, unpublished data.
3 X. He, S. R. P. Miranda, A. Dagan, S. Gatt, and E. H. Schuchman, submitted for publication.
4 Consistent with prior literature (cf. Ref. 2), SMase activity in whole cell homogenates using the standard acidic micellar assay, particularly when EDTA is added, has been equated with "lysosomal" SMase activity. Other types of cellular SMase are not active at acidic pH in this assay, and one of these other SMases also requires Mg2+ for activity (2).
5 In pilot experiments, we found that 1,10-phenanthroline alone was not as effective as EDTA plus phenanthroline in inhibiting the activity of S- and L-SMase. One possible explanation is that the enzymes bind another divalent cation in addition to Zn2+, and removal of this cation by EDTA facilitates the removal of Zn2+ by 1,10-phenanthroline (cf. Refs. 36 and 53). Whatever the mechanism, the fact that 1,10-phenanthroline alone does not inhibit S- or L-SMase argues against an unlikely alternative interpretation of the data in Fig. 1, namely that 1,10-phenanthroline is a direct SMase inhibitor that becomes inactive as an inhibitor when the compound binds Zn2+.
6 Note that treatment with chelators results in almost total inhibition of the internalized, activated S-SMase. This near total effect of chelators is similar to that observed with L-SMase but not with zinc-activated S-SMase from conditioned medium, which is only partially inhibited by chelators (Figs. 1 and 2). Therefore, when S-SMase is delivered to lysosomes, it appears to be converted into a form that allows more complete chelation of its Zn2+. It is possible that this conversion is related to the N-terminal proteolytic processing of SMase in lysosomes (cf. Ref. 46 and data in next section).
7 Simply adding S-SMase to cell homogenates after sonication is complete does not reduce the Zn2+ dependence of the enzyme (data not shown), suggesting that cellular Zn2+ under these conditions is too dilute. To explain this result, we propose that the sequestered pools of Zn2+ released during sonication are in close proximity to the lysosomes, which allows exposure of L-SMase to the released Zn2+ prior to dissipation of the Zn2+ throughout the entire homogenate.
8 S. Kornfeld, personal communication.
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