From the Departments of Medicine and ¶ Anatomy & Cell Biology, Columbia University, New York, New York 10032, ** Berlex
Biosciences, Richmond, California 94804, 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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We recently reported that macrophages
and fibroblasts secrete a Zn2+-dependent
sphingomyelinase (S-SMase), which, like lysosomal SMase, is a product
of the acid SMase gene. S-SMase may cause subendothelial retention and
aggregation of lipoproteins during atherogenesis, and the acid SMase
gene has been implicated in ceramide-mediated cell signaling,
especially involving apoptosis of endothelial cells. Because of the
central importance of the endothelium in each of these processes, we
now sought to examine the secretion and regulation of S-SMase by
vascular endothelial cells. Herein we show that cultured human coronary
artery and umbilical vein endothelial cells secrete massive amounts of
S-SMase (up to 20-fold more than macrophages). Moreover, whereas
S-SMase secreted by macrophages and fibroblasts is almost totally
dependent on the addition of exogenous Zn2+,
endothelium-derived S-SMase was partially active even in the absence of
added Zn2+. Secretion of S-SMase by endothelial cells
occurred both apically and basolaterally, suggesting an endothelial
contribution to both serum and arterial wall SMase. When endothelial
cells were incubated with inflammatory cytokines, such as
interleukin-1 and interferon-
, S-SMase secretion by endothelial
cells was increased 2-3-fold above the already high level of basal
secretion, whereas lysosomal SMase activity was decreased. The
mechanism of interleukin-1
-stimulated secretion appears to be
through increased routing of a SMase precursor protein through the
secretory pathway. In summary, endothelial cells are a rich and
regulatable source of enzymatically active S-SMase, suggesting
physiologic and pathophysiologic roles for this enzyme.
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INTRODUCTION |
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The enzyme sphingomyelinase
(SMase)1 (sphingomyelin
phosphodiesterase; EC 3.1.4.12) catalyzes the hydrolysis of
sphingomyelin to ceramide and choline phosphate (1). SMase reactions
have been implicated in specific atherogenic processes (2-5) and in cell signaling events (6-8). For example, partial hydrolysis of
lipoprotein sphingomyelin by bacterial SMase, via the generation of
lipoprotein ceramide, leads to lipoprotein aggregation (2) and
retention onto arterially derived matrix (3). Aggregation and retention
of lipoproteins in the arterial wall are prominent events during
atherogenesis (9-11), and these lipoprotein aggregates are among the
most potent inducers of macrophage cholesteryl ester accumulation
("foam cell" formation) (2, 3, 12-14). Most importantly, recent
findings indicate that subendothelial LDL, including aggregated LDL
isolated from human lesions, is hydrolyzed by an arterial wall SMase
(4). Regarding cell signaling, treatment of certain cell types with
bacterial SMase results in cellular differentiation, cellular
senescence, or apoptosis, which are thought to be triggered by
ceramide-mediated signal transduction pathways (6-8). Moreover,
cell-derived SMase activity and then cellular ceramide content rise
when these cell types are treated with specific inflammatory cytokines,
such as tumor necrosis factor-, interleukin-1
, and interferon-
(6-8).
Experimental evidence suggests that products of the acid SMase (ASM) gene may have a prominent role in both atherogenesis and ceramide-mediated apoptosis. Hydrolysis of lipoprotein SM retained on subendothelial matrix would clearly be expected to be an extracellular event. In this context, our laboratory has reported that the ASM gene in macrophages and fibroblasts gives rise not only to lysosomal SMase (L-SMase) but also, via differential trafficking of the ASM precursor protein,2 to a secretory SMase (S-SMase) (15). Importantly, S-SMase secreted by these cells, which is activated by physiologic levels of Zn2+ (15), can hydrolyze and cause the aggregation of atherogenic lipoproteins, even at neutral pH (16). Several cell culture studies have also implicated a role for acid SMase activity in cytokine-induced, ceramide-mediated apoptosis (17-19), and mice in which the ASM gene has been disabled by homologous recombination show defective radiation- and endotoxin-induced apoptosis in vivo (20, 21). The products of the ASM gene that mediate these cell signaling events have not been identified, but S-SMase might be an excellent candidate, since the outer leaflet of the plasma membrane is rich in SM (22).
The endothelium is central to these processes. Our previous work on S-SMase focused on macrophages (15), which enter the arterial wall in response to the initial retention of lipoproteins in the subendothelial matrix (5, 23) and therefore may contribute to lipoprotein aggregation after lesion initiation. Lipoprotein aggregation also occurs, however, in prelesional susceptible segments of the subendothelium (10), and thus the endothelium would be a likely candidate for a source of S-SMase in the prelesional arterial wall. Likewise, recent in vivo studies have shown that the endothelium is the key tissue in cytokine-induced, ASM-mediated apoptosis (20, 21). Moreover, cytokines are important regulators of endothelial function (24, 25), atherogenesis, and apoptosis (26, 27), which raises the possibility that cytokines influence endothelial secretion of S-SMase.
In this report, we demonstrate that cultured endothelial cells, including human coronary artery endothelial cells, are an even more abundant source of S-SMase than are macrophages. Furthermore, we show that the secretion of S-SMase by endothelial cells is strongly regulated by cytokines known to be present in atherosclerotic and inflammatory lesions. The mechanism of this regulation is primarily through an alteration in the cellular trafficking of the ASM precursor protein. Thus, the vascular endothelium is likely to be a major, regulatable source of S-SMase that may contribute to atherogenesis and inflammation.
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EXPERIMENTAL PROCEDURES |
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Materials--
The Falcon tissue culture plasticware used in
these studies was purchased from Fisher. Millicell-CM 0.4-µm culture
plate inserts were from Millipore Corp. Tissue culture media and other
tissue culture reagents were obtained from Life Technologies, Inc.
Fetal bovine serum (FBS) was obtained from Hyclone Laboratories (Logan, UT) and was heat-inactivated for 1 h at 65 °C (HI-FBS). Human recombinant cytokines were obtained as follows: interferon- and interferon-
from Biogen (Cambridge, MA), interleukin-4 from
Peprotech (Rocky Hill, NJ), and interleukin-1
from R & D Systems
Inc. (Minneapolis, MN). [9,10-3H]Palmitic acid (56 Ci/mmol) was purchased from DuPont NEN, and [N-palmitoyl-9-10-3H]Sphingomyelin was
synthesized as described previously (15, 28, 29).
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 and Schuell.
FLAG-tagged S-SMase and rabbit anti-S-SMase were kindly provided by Dr.
Henry Lichtenstein (Amgen, Boulder, CO); the FLAG-tagged S-SMase
was purified by anti-FLAG immunoaffinity chromatography from the
conditioned medium of cells transfected with a human ASM-FLAG
cDNA.2 For the immunoprecipitation protocol (see
below), this antibody was further purified on an S-SMase-Affi-Gel
affinity column (Dr. G. Andrew Keesler, Amgen, Boulder, CO). A murine
monoclonal antibody against the FLAG epitope was purchased from Eastman
Kodak Co., and peroxidase-conjugated sheep anti-mouse IgG was from
Amersham Corp. Peroxidase-conjugated goat anti-rabbit IgG and
concanavalin A-Sepharose was purchased from Pierce. DEAE-Sephacel was
from Pharmacia Biotech Inc. Bovine liver 215-kDa mannose 6-phosphate receptor linked to Affi-Gel 15 was made as described by Varki and
Kornfeld (30) and was kindly supplied by Dr. Peter Lobel (Center for
Advanced Biotechnology and Medicine, Piscataway, NJ). All other
chemicals and reagents were from Sigma, and all organic solvents were
from Fisher.
Cells--
Primary cultures of HUVEC were obtained from fresh
umbilical cords and maintained in cell culture as described previously (31). Human coronary artery endothelial cells from a 38-year-old female
donor were purchased from Clonetics Corp. (San Diego, CA; catalog
number CC-2585, strain 3033). The cells were grown in endothelial basal
media (Clonetics) supplemented with 10% HI-FBS, 10 µg/ml endothelial
cell growth supplement (Sigma; catalog number E0760), and 50 µg/ml
gentamicin. Primary cultures of bovine aortic endothelial cells were
established and subcultured as described previously (32). For the
experiment in Fig. 2, the cells were plated on Millicell-CM 0.4-µm
culture plate inserts placed in 35-mm wells in -MEM containing 10%
FBS; the inserts had been previously coated with 0.1% gelatin for
1 h and then with 5 µg/ml collagen and 20 µg/ml fibronectin
for 1 h. Monolayer cultures of J774.A1 cells (from the American
Type Culture Collection; see Ref. 33) were grown and maintained in
spinner culture with DMEM/HI-FBS/PSG (penicillin, streptomycin, and
glutamine) as described previously (33, 34). Human peripheral blood
monocytes were isolated from normal subjects as described previously
(35) and induced to differentiate into macrophages by the addition of 1 ng of granulocyte-macrophage colony-stimulating factor/ml of media on
days 1, 4, and 11 of culture as described previously (35); by day 14, the cells were differentiated as assessed by morphological changes
(e.g. increased spreading) and increased expression of
scavenger receptor activity (cf. Ref. 36). Unless indicated
otherwise, the cells were plated in 35-mm (6-well) dishes in media
containing HI-FBS for 48 h. The cells were then washed three times
with PBS and incubated for 24 h in 1 ml of fresh serum-free media
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 medium was 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 dish, respectively. Unless indicated otherwise, the scraped cells were disrupted by sonication on ice using three 5-s bursts (Branson model 450 sonifier), and the cellular homogenates were assayed for total protein by the method of Lowry et al. (22) 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).
SMase Assay-- As described previously (15), 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 was 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 (37); 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.
SMase Immunohistochemistry of Murine Aorta-- Hearts from chow-fed, 4-week-old female ASM knockout mice and wild-type mice of the same genetic background (SV129/C57BL6) were perfused, embedded in optimum-cutting-temperature (OCT) compound (Sakura Finetek, Torrance, CA), and snap-frozen as described previously (38). 8-µm-thick sections of the proximal aorta were cut on a cryostat and fixed in 10% buffered formalin for 5 min at room temperature. The sections were blocked in 2% murine serum in PBS for 2 h at room temperature and then incubated with 20 µg of anti-SMase antibody/ml of PBS containing 0.1% Triton X-100 for 2-4 h at room temperature. Bound antibody was detected with a biotinylated secondary antibody followed by streptavidin peroxidase (Vectastain Elite ABC-peroxidase kit; Vector Laboratories Inc., Burlingame, CA) and 3,3'-diaminobenzidine. The specimens were counterstained with hematoxylin and then viewed with an Olympus IX 70 inverted microscope using a × 100 objective.
Partial Purification of L- and S-SMase from HUVECs-- A modification of previously published procedures (39) was used. Cells were scraped in ice-cold 40 mM Tris-HCl, 0.1 mM ZnCl2, 0.1% Nonidet P-40, pH 7.2 (buffer A), disrupted by sonication on ice using three 5-s bursts (Branson model 450 sonifier), and centrifuged at 100,000 × g for 1 h. The supernatant (<5 ml) was applied to a 5-ml column of DEAE-Sephacel, and the column was capped and incubated end-over-end for 90 min at room temperature. The column was then uncapped, and the flow-through fraction was collected; the column was washed with another 1-2 ml of buffer A, and this flow-through fraction was added to the first. The combined flow-through fractions were loaded onto another DEAE-Sephacel column, and the procedure was repeated. The flow-through fractions from this second column were pooled, concentrated using a Centriprep 30 (see above), assayed for protein concentration by the method of Lowry et al. (40), and subjected to SDS-PAGE and immunoblotting.
The conditioned medium was concentrated, dialyzed against buffer A, and subjected to the same two rounds of DEAE-Sephacel chromatography as above. The combined flow-through fractions from the second DEAE-Sephacel step were then concentrated and dialyzed against 10 mM Tris-HCl, 500 mM NaCl, 1 mM MgCl2, 1 mM MnCl2, 1 mM CaCl2, 0.1% Nonidet P-40, and 0.02% NaN3, pH 7.2 (buffer B). This solution (<1 ml) was added to a 1-ml concanavalin A-Sepharose column and incubated with the column exactly as described for the DEAE-Sephacel step. The column was then washed with two bed volumes of buffer B containing 10 mM methylglucopyranoside, followed by two bed volumes of buffer B containing 1 M methylglucopyranoside. This last eluate was concentrated, assayed for protein concentration by the method of Lowry (40), and subjected to SDS-PAGE and immunoblotting.Northern Blot of SMase mRNA-- Total cellular RNA was isolated from HUVECs using RNAzol B (Tel-Test, Inc., Friendswood, TX). Approximately 10 µg of RNA was separated on a 1% agarose gel and blotted onto a nylon membrane. The membrane was hybridized in a QuikHyb hybridization solution (Stratagene) with a 1.6-kilobase pair EcoRI-XhoI fragment of human ASM cDNA (41) that was labeled with 32P by the random-priming procedure (Life Technologies, Inc.). The amount of RNA loaded for each condition was normalized using a probe to glyceraldehyde-3-phosphate dehydrogenase. The relative intensities of the mRNA bands were determined by densitometric scanning of the autoradiograms using a Molecular Dynamics computing densitometer (model 300A) with ImageQuant software or a Bio-Rad molecular imager (model GS525).
Lactate Dehydrogenase Assay-- Lactate dehydrogenase activity was assayed in medium and cells as reported previously (42) using a kit purchased from Sigma (catalogue number 500); the assay measures the unreacted pyruvic acid using a colorometric assay.
SDS-PAGE 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 milliamps in buffer containing 0.1% SDS. Following electrophoresis, the proteins on the gels were electrotransferred (100 V for 1.5 h) to nitrocellulose for immunoblotting. Next, the nitrocellulose membranes were incubated with 5% Carnation nonfat dry milk in buffer C (24 mM Tris, pH 7.4, containing 0.5 M NaCl) for 3 h at room temperature. The membranes were then incubated with mouse anti-FLAG monoclonal antibody (1:1000) or rabbit anti-L-SMase antibody (1:1000) in buffer D (buffer C 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 C containing 0.1% Tween 20, the blots were incubated with horseradish peroxidase-conjugated sheep anti-mouse IgG (1:20,000) or goat anti-rabbit IgG (1:20,000) for 1 h in buffer D at room temperature. The membranes were subsequently washed twice with 0.3% Tween 20 in buffer C and twice with 0.1% Tween 20 in buffer C. Finally, the blots were soaked in the enhanced chemiluminescence reagent (Pierce "Super Signal" kit) for 2 min and exposed to x-ray film for 1 min.
Immunoprecipitation of S-SMase-- 20 µl of HUVEC conditioned medium, 20 µl of affinity-purified anti-SMase antibody, and 160 µl of PBS were incubated for 1 h at 4 °C. 100 µl of Protein A-Sepharose (50 mg/ml in 50 mM Tris buffer, pH 7.0) was then added to the incubation mixture, and the slurry was mixed end-over-end for 18 h at 4 °C. The suspension was centrifuged for 1 min in a microcentrifuge. The supernatant was harvested and assayed for SMase activity.
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 symbols.
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RESULTS |
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Cultured Human Vascular Endothelial Cells Secrete Abundant Amounts of S-SMase-- Previous work from our laboratories revealed that human and murine macrophages were a relatively abundant source of S-SMase compared with other cell types, such as COS-7 and murine migroglial cells (15). As shown in Fig. 1A, however, both human coronary and umbilical vein endothelial cells (HUVECs) are a much more abundant source of S-SMase; for example, HUVECs secrete almost 20-fold more S-SMase than human macrophages. Interestingly, endothelium-derived S-SMase was partially active in the absence of exogenously added Zn2+ (Fig. 1A), whereas S-SMase secreted by macrophages and other cell types previously examined by us was almost entirely Zn2+-dependent (15).3 This observation may be important in the regulation of endothelium-derived S-SMase activity (see "Discussion"). The data in Fig. 1B show that SMase activity in the cell homogenate, which is not stimulated by exogenous Zn2+ and represents L-SMase activity (15), is also very abundant in endothelial cells (recall that S-SMase and L-SMase originate from the same gene (15), mRNA (15), and protein precursor2). In summary, the data in Fig. 1 demonstrate that human endothelial cells, including those derived from coronary arteries, are an abundant source of S-SMase, much of which is active in the absence of exogenously added Zn2+.
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SMase Is Present in the Endothelium of Murine Aorta-- To complement the cell culture studies described above, we used immunohistochemistry to detect SMase in slices of murine proximal aorta. Because L- and S-SMase are derived from the same protein precursor and differ only by post-translational modifications (15),2 the anti-SMase antibodies currently available to us recognize both forms of the enzyme. To ensure the specificity of the signal for SMase, proximal aorta from ASM knockout mice (44), which lack both L- and S-SMase (15), were used as a negative control (Fig. 3A). Fig. 3B demonstrates substantial SMase staining in the endothelium; much of the stain in this image appears to be intracellular (open arrow), which is most likely L-SMase and possibly some S-SMase in the secretory pathway. By comparison, medial staining was much weaker, although still specific. These data demonstrate that the high levels of intracellular SMase in cultured endothelial cells (Fig. 1B) reflect the situation in an actual vessel wall. In addition, we also noticed areas of dark staining on the lumenal edge of some of the endothelial cells (closed arrow). This staining is specific (i.e., not in panel A) and does not appear lysosomal, and so it is possible that this signal represents S-SMase that has been secreted and perhaps retained on the cell surface.
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Secretion of S-SMase from Human Endothelial Cells Is Regulated by
Cytokines--
Inflammatory cytokines are important constituents of
atherosclerotic lesions and may contribute to various aspects of
atherogenesis (26). A major target of these cytokines is the
endothelium (24, 25). To determine if cytokines known to be present in
atherosclerotic lesions affect endothelium-derived S-SMase, cultured
human endothelial cells were exposed to IL-1, interferon-
,
interferon-
, and IL-4. Each of the first three cytokines
substantially increased the accumulation of S-SMase in the conditioned
media of these cells, although IL-4 had no effect (Fig.
4A). In particular, IL-1
and interferon-
increased S-SMase activity ~3-fold. Interestingly, the stimulatory cytokines led to a decrease in L-SMase activity (Fig.
4B). This pattern is distinct from that observed during monocyte-to-macrophage differentiation, in which both S- and L-SMase activities are increased (15).
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IL-1 Increases S-SMase Secretion by HUVECs via Alteration in the
Trafficking of the ASM Precursor Protein--
We next sought to
determine the mechanism of cytokine-mediated induction of S-SMase
secretion by HUVECs. For these studies, we focused on IL-1
, and we
first determined whether this cytokine increased SMase mRNA levels
in HUVECs. As shown in Fig. 5, there was
an ~40% increase in SMase mRNA in IL-1
-treated HUVECs, when normalized for glyceraldehyde-3-phosphate dehydrogenase mRNA and quantified by either densitometry or molecular imaging. Because differences less than 2-fold in Northern blot assays may not be significant and because the IL-1
-induced increase in S-SMase activity was greater than 3 times control, we conclude that most, if
not all, of the induction by this cytokine was post-transcriptional. This conclusion is consistent with the finding that the
cytokine-mediated increase in S-SMase is accompanied by a decrease in
L-SMase (above).
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DISCUSSION |
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The data in this report demonstrate that endothelial cells are a rich source of active S-SMase, particularly in the presence of inflammatory cytokines. The potential physiologic relevance of these findings is related to the postulated role of the endothelium in extracellular SMase-induced lipoprotein aggregation and retention and to the demonstrated role of this tissue in cytokine-induced, acid SMase-mediated cell signaling (see Introduction and below). In addition, our results reveal several novel mechanisms for the regulation of S-SMase, particularly by alterations in intracellular protein trafficking.
We have previously shown that retained and aggregated lipoproteins in atherosclerotic lesions are hydrolyzed by an extracellular arterial wall SMase (4), and we found that S-SMase was the only SMase secreted by arterial wall cells (15). Mohan Das et al. (49) have reported that a magnesium-dependent SMase is externally oriented on neurons. We have not yet investigated whether this enzyme, which has not yet been cloned, is present on the surface of arterial wall cells. Nonetheless, we imagine that a secreted enzyme would have more access to lipoproteins retained on subendothelial matrix than a cell surface-bound enzyme. These points, together with the findings reported herein, lead us to propose the following model: basal levels of endothelium-derived S-SMase help initiate the atherosclerotic lesion by promoting lipoprotein retention and aggregation (5). Then, as T-cells and macrophages enter lesions (23, 26) and secrete cytokines and (at least in the case of macrophages) additional S-SMase, lipoprotein-SM hydrolysis and lipoprotein retention and aggregation would be amplified.
We have also speculated that endothelium-derived S-SMase may play a
role in ceramide-mediated apoptosis. This speculation is based upon
several pieces of information. First, a product of the acid SMase gene
plays an important role in endothelial apoptosis in vivo
(20, 21). Specifically, the pulmonary endothelium of ASM knockout mice
was the major tissue demonstrating a defect in ceramide generation and
apoptosis in response to total body irradiation; radiation-induced
apoptosis in thymocytes and splenocytes was much less diminished in the
ASM knockout versus wild-type mice (20). Moreover, very
recent studies in mice have shown that the endothelium is the target of
tumor necrosis factor--mediated apoptosis after injection of
lipopolysaccharide. This response is associated with an increase in
endothelial ceramide content and is greatly diminished in ASM knockout
mice, indicating involvement of a SMase arising from the ASM gene (21).
Second, the data in this report show that endothelial cells are a rich
and cytokine-regulatable source of S-SMase, a product of the ASM gene.
Third, S-SMase may have more access to the most abundant pools of
cellular SM (22). In contrast, since lysosomal membranes have little SM
(50), signaling by L-SMase would have to occur during trafficking of the nascent enzyme to lysosomes or would require delivery of SM into
lysosomes. Thus, we propose that apoptotic stimuli, such as radiation
and cytokines, increase the amount of endothelium-derived S-SMase from
a subthreshold basal level to a level capable of generating enough
cellular ceramide to trigger or enhance cell-signaling events. It is
important to note, however, that this hypothesis is based upon several
controversial assumptions. For example, despite the compelling data
obtained using ASM knockout mice (20, 21), there are some cell culture
systems that have shown a role for the neutral,
magnesium-dependent SMase in ceramide-mediated apoptosis
(6). In addition, there are conflicting data regarding the role of cell
surface versus intracellular pools of SM in
ceramide-mediated signaling (cf. Ref. 51, and see
"Discussion" therein). Nonetheless, the data presented in this
report give impetus for further in vitro and in
vivo studies on the role of endothelium-derived S-SMase in
atherogenesis and ceramide-mediated apoptosis.
Regarding the regulation of S-SMase, the data in this report provide evidence to support at least three separate mechanisms for control of S-SMase. Two of these mechanisms, alterations in protein trafficking and accessibility to cellular zinc, are based upon the following model of how the ASM gene gives rise to both L- and S-SMase2; when a common precursor protein derived from the ASM gene is mannose-phosphorylated and thus is targeted to lysosomes, it becomes L-SMase. During this targeting, the enzyme acquires cellular Zn2+ and so does not require exogenous Zn2+ for enzymatic activity. In contrast, when the common precursor is not mannose-phosphorylated, and thus is targeted to the Golgi-secretory pathway, it gives rise to S-SMase. In the secretory pathway, the enzyme does not acquire cellular Zn2+, so S-SMase secreted by macrophages requires exogenous Zn2+ for enzymatic activity.
Based on this model, we predicted that one level of control of S-SMase
secretion might be the proportion of the common ASM precursor
trafficked into the lysosomal versus secretory pathways. The
current data indicate that this mechanism is indeed primarily responsible for the increase in S-SMase in response to IL-1. How
might a cytokine influence protein trafficking? The key regulatory step
in the trafficking of the SMase precursor is mannose phosphorylation of
the precursor by N-acetylglucosaminyl-1-phosphotransferase (52, 53),2 and we speculate that cytokines may affect
(e.g. by protein phosphorylation) either the activity of
this phosphotransferase or the suitability of the SMase precursor as
its substrate. Further work will be needed to test this and other
possible mechanisms.
A second level of regulation of S-SMase is via Zn2+-induced activation (15). The zinc requirement of S-SMase is similar to that of matrix metalloproteinases (15, 43, 54, 55), and so under conditions in which these proteinases are active, such as in atherosclerotic lesions (56), one would expect S-SMase to be active as well. In addition, Zn2+ levels have been reported to be elevated in atherosclerotic (57) and inflammatory (58) lesions. Nonetheless, the accessibility of extracellular zinc, perhaps modulated by zinc-binding proteins such as metallothionein (59), may represent a regulatory mechanism. Our current results indicate that accessibility of cell-derived zinc is also relevant; S-SMase from endothelial cells was partially activated in the absence of exogenously added Zn2+ (Fig. 1). In contrast, S-SMase from macrophages, fibroblasts, and Chinese hamster ovary cells is almost entirely inactive in the absence of added Zn2+ (Fig. 1 and Ref. 15). Based upon our model (above), we propose that SMase in the secretory pathway of endothelial cells, unlike SMase in the secretory pathway of the other cells examined, has partial access to cellular pools of Zn2+.3 The findings that endothelial cells secrete abundant amounts of S-SMase and that this S-SMase is partially active in the absence of added Zn2+ suggest that endothelium-derived S-SMase has unique physiologic roles.
A third point of S-SMase regulation is extracellular pH. S-SMase, like L-SMase, has an acidic pH optimum when assayed in vitro using sphingomyelin in detergent micelles as substrate (15). Thus, S-SMase may be particularly active in environments in which the pH is relatively low, such as in advanced atherosclerotic lesions (60-63), in certain types of inflammatory processes (62, 64), and possibly after reuptake into acidic endosomes. Calahan (65) noted, however, that pH affects only the Km, not the Vmax, of L-SMase. This finding suggests that access to SM is the issue, and we showed recently that S-SMase can extensively hydrolyze the SM of atherogenic lipoproteins (e.g., oxidized LDL) and lesional LDL at neutral pH (16). This point is of particular importance regarding our hypothesis that endothelium-derived S-SMase plays a role in early lesional events, where the arterial wall pH would be expected to be neutral. These observations may also be relevant to the hydrolysis of cellular sphingomyelin by endothelium-derived S-SMase in neutral pH environments.
In summary, endothelial cells, which we have postulated are important in subendothelial, extracellular lipoprotein SM hydrolysis and which others have shown are important in cytokine-induced, ASM-mediated cell signaling, are a rich and regulatable source of the ASM gene product, S-SMase. These findings have formed the basis of ongoing work directed at further testing the physiologic and pathophysiologic roles of endothelium-derived S-SMase.
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ACKNOWLEDGEMENTS |
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We thank Dr. Henry Lichtenstein (Amgen, Boulder, CO) for the secreted FLAG-tagged S-SMase and the anti-FLAG-tagged S-SMase antiserum, Dr. G. Andrew Keesler (Amgen, Boulder, CO) for the affinity-purified anti-S-SMase antibody, Dr. Louis A. Peña (Memorial Sloan-Kettering Cancer Center, New York, NY) for advice on the immunoprecipitation of S-SMase, and Dr. Peter Lobel (Center for Advanced Biotechnology and Medicine, Piscataway, NJ) for the mannose 6-phosphate receptor resin.
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FOOTNOTES |
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* This study was supported by National Institutes of Health Grants HL39703 (to I. T.), HL56984 (to I. T. and K. J. W.), HL38956 (to K. J. W.), and AR44198 (to M. J. Y.); an Established Investigator award from the American Heart Association and Genentech (to K. J. W.); and a grant-in-aid from the American Heart Association (to M. J. Y.). 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.
§ These authors contributed equally to this work.
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 W. 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; ASM, acid sphingomyelinase; BSA, bovine serum
albumin; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine
serum; HI-FBS, heat-inactivated FBS; HUVEC, human umbilical vein
endothelial cell; IL, interleukin; LDL, low density lipoprotein;
L-SMase, lysosomal sphingomyelinase; PBS, phosphate-buffered saline;
PAGE, polyacrylamide gel electrophoresis; SM, sphingomyelin; S-SMase,
secretory sphingomyelinase; -MEM,
-minimum essential
medium.
2 S. L. Schissel, E. H. Schuchman, K. J. Williams, and I. Tabas, submitted for publication.
3 95% of S-SMase activity from HUVECs was immunoprecipitated by an affinity-purified anti-S-SMase antibody, and 97% of HUVEC S-SMase activity was inactivated by chelation of Zn2+ with 10 mM EDTA plus 10 mM 1,10-phenanthroline (see Footnote 2). These data indicate that a single enzyme accounts for SMase activity in the conditioned medium of HUVECs.
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