Method for treating a subject suffering from a condition associated with an extracellular zinc sphingomyelinase

The present invention provides for a method for treating a subject suffering from a condition associated with an extracellular zinc sphingomyelinase activity which comprises administering to the subject an amount of a zinc sphingmyelinase inhibitor effective to decrease extracellular zinc sphingomyelinase activity in the subject and thereby treat the subject. The present invention also provides for a method for determining whether a compound inhibits an activity of an extracellular zinc sphingomyelinase involving ceramide formation which comprises: (a) contacting a sample containing the zinc sphingomyelinase under acidic pH conditions known to be associated with the activity of such zinc sphingomyelinase, with: (i) a substrate of the zinc sphingomyelinase enzyme, and (ii) the compound being evaluated; (b) measuring the concentration of ceramide in the sample from (a); (c) determining the amount of zinc sphingomyelinase activity in the sample based upon the concentration of ceramide measured in step (b); and (d) comparing the amount of sphingomyelinase activity determined in step (c) with the amount of sphingomyelinase activity determined in the absence of the compound, so as to determine whether the compound inhibits the activity of zinc sphingomyelinase.

BACKGROUND OF THE INVENTION 
Throughout this application, various publications are referenced by author 
and date. Full citations for these publications may be found listed 
alphabetically at the end of the specification immediately preceding the 
claims. The disclosures of these publications in their entireties are 
hereby incorporated by reference into this application in order to more 
fully describe the state of the art. 
A key early event in atherogenesis is the subendothelial retention of 
atherogenic lipoproteins, including LDL.sup.1 (Schwenke et al., 1989; 
Nievelstein et al., 1991), lipoprotein(a) [Lp(a)] (Kreuzer et al., 1994), 
and triglyceride-rich lipoproteins (Rapp et al., 1994). 
Atherosclerosis-susceptible regions of the arterial tree are distinguished 
by their increased retention of lipoproteins compared with resistant 
regions (Schwenke et al., 1989). The retained lipoproteins are likely to 
trigger a set of biological responses, such as lipoprotein oxidation and 
endothelial changes (Steinberg et al., 1989; Ross, 1995), that are central 
to the atherogenic process (Williams et al., 1995). 
Lipoproteins retained in the subendothelium are often extensively 
aggregated (Nievelstein et al., 1991; Hoff et al., 1985; Guyton et al., 
1996). For example, Hoff and colleagues and others (see Hoff et al., 1985; 
Guyton et al., 1996) have shown that LDL extracted from atherosclerotic 
lesions is aggregated or has an increased tendency to aggregate, whereas 
plasma LDL exposed to the same extraction procedure as a control does not 
aggregate. Furthermore, Frank and colleagues (Nievelstein et al., 1991) 
used freeze-etch electron microscopy to demonstrate aggregated LDL in the 
subendothelium of the rabbit aorta as early as two hours after a large 
bolus injection of human LDL. Subendothelial lipoprotein aggregation is 
likely to be important in atherogenesis for at least two reasons. First, 
processes that promote lipoprotein aggregation before or during retention 
would be expected to increase the amount of material retained (Tabas et 
al., 1993). Second, aggregated LDL, but not unaggregated LDL, is a potent 
inducer of macrophage foam cell formation (Hoff et al., 1990; Khoo et al., 
1988; Suits et al., 1989; Xu et al., 1991). 
SUMMARY OF THE INVENTION 
The present invention provides for a method for treating a subject 
suffering from a condition associated with an extracellular zinc 
sphingomyelinase activity which comprises administering to the subject an 
amount of a zinc sphingmyelinase inhibitor effective to decrease 
extracellular zinc sphingomyelinase activity in the subject and thereby 
treat the subject. The present invention also provides for a method for 
determining whether a compound inhibits an activity of an extracellular 
zinc sphingomyelinase involving ceramide formation which comprises: (a) 
contacting a sample containing the zinc sphingomyelinase under acidic pH 
conditions known to be associated with the activity of such zinc 
sphingomyelinase, with: (i) a substrate of the zinc sphingomyelinase 
enzyme, and (ii) the compound being evaluated; (b) measuring the 
concentration of ceramide in the sample from (a); (c) determining the 
amount of zinc sphingomyelinase activity in the sample based upon the 
concentration of ceramide measured in step (b); and (d) comparing the 
amount of sphingomyelinase activity determined in step (c) with the amount 
of sphingomyelinase activity determined in the absence of the compound, so 
as to determine whether the compound inhibits the activity of zinc 
sphingomyelinase.

DETAILED DESCRIPTION OF THE INVENTION 
The present invention provides for a method for treating a subject 
suffering from a condition associated with an extracellular zinc 
sphingomyelinase activity which comprises administering to the subject an 
amount of a zinc sphingmyelinase inhibitor effective to decrease 
extracellular zinc sphingomyelinase activity in the subject and thereby 
treat the subject. 
In one embodiment of the present invention, the extracellular zinc 
sphingomyelinase is present in the subject at a concentration which is 
higher than that present in the subject prior to the onset of the 
condition. In another embodiment of the present invention, wherein 
extracellular zinc sphingomyelinase is present in the subject at a 
concentration which is lower than that present in the subject prior to the 
onset of the condition. 
As used herein "extracellular zinc sphingomyelinase" is encompassed by 
lysosomal zinc sphingomyelinase or L-SMase; secreted or secretory zinc 
sphingomyelinase or S-SMase; or zinc sphingomyelinase which was originally 
an intracellular zinc sphingomyelinase enzyme and is released into the 
extracellular environment. Extracellular zinc sphingomyelinase is referred 
to herein as the zinc sphingomyelinase. Extracellular zinc 
sphingomyelinase is a metalloenzyme which is capable of binding zinc and 
capable of being activated by zinc. As used herein, extracellular zinc 
sphingomyelinase is encompassed by sphingomyelinase already bound to zinc 
or sphingomyelinase not yet bound to zinc but capable of binding zinc. 
Extracellular zinc sphingomyelinase may be produced from a Golgi secretory 
pathway of a cell or a lysosomal pathway of a cell, or by other processes 
leading to release into an extracellular environment or by a combination 
of these pathways. 
In another embodiment of the present invention, the condition may be an 
atherosclerotic vascular disease, an inflammatory disease, an infectious 
disease, an autoimmune disease, or a demyelinating disease. The 
atherosclerotic vascular disease may be coronary artery disease, cerebral 
vascular disease, peripheral vascular disease, transplantation 
atherosclerosis, vein graft atherosclerosis, or vaculitis-induced 
atherosclerosis. The demyelinating disease may be multiple sclerosis. The 
demyelinating disease may also be progressive multifocal 
leucoencephalopathy, Guillain-Barre syndrome, Retrobulbar neuritis, acute 
rubella encephalitis, chronic relapsing polyneuropathy, intravascular 
lymphomatosis, Krabbe disease, globoid cell leukodystrophy, subacute 
combined degeneration of the spinal cord and brain, allergic encephalitis, 
murine caronavirus, hepatitis virus infection, or Theiler's murine 
encephalomyelitis. The subject may be a mouse which is a Twitcher mouse or 
a rat which is a HAM rat. The subject may be an animal which has 
experimental allergic encephalitis. 
In one embodiment of the present invention, the zinc sphingomyelinase 
inhibitor may comprises a peptide or polypeptide, a peptidomimetic 
compound, an organic compound, a nucleic acid, an inorganic compound, or 
an antibody or fragment thereof. In another embodiment, the inhibitor is 
an antibody capable of binding to and inactivating zinc sphingomyelinase. 
The inhibitor may be an antibody which comprises a monoclonal or a 
polyclonal antibody. 
In another embodiment of the present invention, the inhibitor comprises a 
compound capable of competing with sphingomyelin for binding to the active 
site of naturally occuring sphingomyelinase. In a further embodiment of 
the present invention, the inhibitor may be a pseudoenzyme. 
In one embodiment of the present invention, the administration comprises 
intralesional, intraperitoneal, intramuscular or intravenous injection; 
infusion; liposome-mediated delivery; topical, nasal, oral, anal, 
subcutaneous, vaginal, sublingual, intrathecal, uretheral, transdermal, 
ocular or otic delivery. 
The present invention provides that the zinc sphingomyelinase inhibitor may 
comprise a portion of a naturally occuring zinc sphingomyelinase. The 
portion may consist essentially of a sphingomyelin binding site of the 
sphingomyelinase. In another embodiment, the zinc sphingomyelinase 
inhibitor is a compound having a structure which mimics the structure of a 
substrate of sphingomyelinase or of a product of sphingomyelinase. In 
another embodiment, the substrate of sphingomyelinase is sphingomyelin. In 
another embodiment, the product of sphingomyelinase is ceramide or choline 
phosphate. 
The present invention provides for a method for determining whether a 
compound inhibits an activity of an extracellular zinc sphingomyelinase 
involving ceramide formation which comprises: (a) contacting a sample 
containing the zinc sphingomyelinase under acidic pH conditions known to 
be associated with the activity of such zinc sphingomyelinase, with: (i) a 
substrate of the zinc sphingomyelinase enzyme, and (ii) the compound being 
evaluated; (b) measuring the concentration of ceramide in the sample from 
(a); (c) determining the amount of zinc sphingomyelinase activity in the 
sample based upon the concentration of ceramide measured in step (b); and 
(d) comparing the amount of sphingomyelinase activity determined in step 
(c) with the amount of sphingomyelinase activity determined in the absence 
of the compound, so as to determine whether the compound inhibits the 
activity of zinc sphingomyelinase. The present invention provides for a 
method for determining whether a compound inhibits an activity of an 
extracellular zinc sphingomyelinase involving ceramide formation which 
comprises: (a) contacting a sample containing the zinc sphingomyelinase 
under neutral pH conditions, with: (i) a substrate of the zinc 
sphingomyelinase enzyme, and (ii) the compound being evaluated; (b) 
measuring the concentration of ceramide in the sample from (a); (c) 
determining the amount of zinc sphingomyelinase activity in the sample 
based upon the concentration of ceramide measured in step (b); and (d) 
comparing the amount of sphingomyelinase activity determined in step (c) 
with the amount of sphingomyelinase activity determined in the absence of 
the compound, so as to determine whether the compound inhibits the 
activity of zinc sphingomyelinase. 
As used herein, acidic pH encompasses a pH range from about pH 2.5 to about 
pH 6.9 and neutral pH encompasses a pH range from about pH 7 to about pH 
7.5. The present invention provides for methods which may be carried out 
within a pH range from about 2.5 to about 7.5. 
In one embodiment, the substrate may comprise sphingomyelin or a derivative 
thereof or a lipoprotein. The substrate may be detectably labeled. The 
lipoprotein may be a modified lipoprotein. The detectable label may 
comprise a radioisotope or a fluorophor. The lipoprotein may comprise an 
oxidized lipoprotein, a phospholipase-A-II treated lipoprotein, an 
apolipoprotein-C-III-enriched population of lipoproteins, a lipoprotein 
obtained from an apolipoprotein-E knock-out mouse, or a 
sphingomyelin-enriched population of lipoproteins or emulsions thereof at 
neutral pH. 
In one embodiment, steps (a) through (d) of the above-described method are 
repeated for multiple compounds. 
The present invention also provides for a method for screening a library of 
compounds to identify a compound capable of inhibiting an activity of zinc 
sphingomyelinase involving ceramide formation which comprises: (a) 
contacting a zinc sphingomyelinase under acidic pH conditions known to be 
associated with the activity of such zinc sphingomyelinase, with: (i) a 
substrate of sphingomyelinase, and (ii) a sample from a library of 
compounds being evaluated; (b) measuring the concentration of ceramide in 
the sample from (a); and (c) determining the amount of zinc 
sphingomyelinase activity in the sample based upon the concentration of 
ceramide measured in step (b); and (d) comparing the amount of 
sphingomyelinase activity determined in step (c) with the amount of 
sphingomyelinase activity determined in the absence of the sample, so as 
to determine whether the sample inhibits the activity of zinc 
sphingomyelinase enzyme, and (d) repeating steps (a) through (d) with 
limiting dilutions of the sample so as to identify the compound in the 
sample capable of inhibiting zinc sphingomyelinase. The screen may also be 
carried out at neutral pH. 
The present invention also provides for a method for determining whether a 
subject is at increased risk for becoming afflicted with an increase in 
the concentration of extracellular zinc sphingomyelinase activity in the 
subject which comprises: (a) obtaining a sample of a body fluid from the 
subject; (b) determining the amount of extracellular zinc 
sphyingomyelinase activity in the body fluid sample, and (c) comparing the 
amount of extracellular zinc sphyingomyelinase activity determined in step 
(a) with the amounts of extracellular zinc sphyingomyelinase activity 
determined for the subject at earlier points in time, an increase in the 
amount of such activity indicating that the subject is at increased risk 
for such condition. 
The present invention also provides for a method for determining whether a 
subject is at increased risk for becoming afflicted with an increase in 
the concentration of extracellular zinc sphingomyelinase activity in the 
subject which comprises: (a) obtaining a sample of a body fluid from the 
subject; (b) determining the amount of extracellular zinc 
sphyingomyelinase activity in the body fluid sample, and (c) comparing the 
amount of extracellular zinc sphyingomyelinase activity determined in step 
(a) with the a predetermined standard extracellular zinc sphyingomyelinase 
activity so as to determine whether the subject is at increased risk for 
such condition. 
The predetermined standard extracellular zinc sphingomyelinase activity may 
be determined by identifying the activity present in samples taken from a 
large number of individuals. The activity levels determined could then be 
compared so as to determine a statistical norm for the population of 
individuals. The population may be controlled for age of the individuals, 
sex of the individuals, relative health, relative fitness, etc. From such 
samplings, a standard extracellular sphingomyelinase activity may be 
determined. 
In one embodiment of the present invention, the condition may be an 
atherosclerotic vascular disease, an inflammatory disease, an infectious 
disease, an autoimmune disease, or a demyelinating disease. The 
atherosclerotic vascular disease may be coronary artery disease, cerebral 
vascular disease, peripheral vascular disease, transplantation 
atherosclerosis, vein graft atherosclerosis, or vaculitis-induced 
atherosclerosis. The demyelinating disease may be multiple sclerosis, 
progressive multifocal leucoencephalopathy, Guillain-Barre syndrome, 
retrobulbar neuritis, acute rubella encephalitis, chronic relapsing 
polyneuropathy, intravascular lymphomatosis, Krabbe disease, globoid cell 
leukodystrophy, subacute combined degeneration of the spinal cord and 
brain, allergic encephalitis, murine caronavirus, hepatitis virus 
infection, or Theiler's murine encephalomyelitis. The body fluid may 
comprise plasma, blood, serum, interstitial fluid, cerebrospinal fluid, 
joint fluid, tears, semen, urine, saliva, bile, or amniotic fluid. 
The present invention provides for a method for determining whether a 
subject has lipoproteins susceptible to extracellular zinc 
sphingomyelinase activity and thus is at increased risk for becoming 
afflicted with a condition associated with extracellular zinc 
sphingomyelinase activity which comprises: (a) obtaining a sample of a 
body fluid from the subject; (b) isolating the lipoproteins present in the 
sample; (c) contacting the isolated lipoproteins with zinc 
sphingomyelinase enzyme under acidic pH conditions known to be associated 
with the activity of such zinc sphingomyelinase; (d) measuring the 
concentration of ceramide in step (c); and (e) comparing the amount of 
ceramide detected in step (d) with an amount of ceramide produced from 
lipoproteins isolated from a normal subject, thereby detecting whether the 
subject has lipoproteins susceptible to extracellular zinc 
sphingomyelinase so as to determine whether the subject is at increased 
risk for becoming afflicted with a condition associated with extracellular 
zinc sphingomyelinase activity. The contacting of step (c) may also be 
performed under neutral pH conditions. 
The present invention provides for a pharmaceutical composition comprising 
an amount of an inhibitor of an extracellular zinc sphingomyelinase enzyme 
effective to inhibit the activity of such zinc sphingomyelinase enzyme in 
a subject and a pharmaceutically acceptable carrier. The carrier may 
comprise a diluent. The carrier may further comprise an adjuvant, a 
liposome, a microencapsule, a polymer encapsulated cell, a biodegradable 
plastic or a retroviral vector. The composition may be in a form suitable 
for aerosol, intravenous, oral or topical administration. 
In one embodiment of the invention, the subject may be an animal, an animal 
model or a human. 
In addition to the inhibitor being derived from a naturally-occurring form 
of zinc sphingomyelinase, the present invention also embraces other agents 
such as polypeptide analogs of zinc sphingomyelinase enzyme. Such analogs 
include fragments of zinc sphingomyelinase enzyme. Following the 
procedures of the published application by Alton et al. (WO 83/04053), one 
can readily design and manufacture genes coding for microbial expression 
of polypeptides having primary conformations which differ from the natural 
zinc sphingomyelinase enzyme which would be specified for in terms of the 
identity or location of one or more residues (e.g., substitutions, 
terminal and intermediate additions and deletions). Alternately, 
modifications of cDNA and genomic genes can be readily accomplished by 
well-known site-directed mutagenesis techniques and employed to generate 
analogs and derivatives of zinc sphingomyelinase enzyme. Such agents share 
at least one of the biological properties of zinc sphingomyelinase enzyme 
but may differ in others. As examples, inhibitors of zinc sphingomyelinase 
of the invention include those agents which are foreshortened by e.g., 
deletions; or those which are more stable to hydrolysis (and, therefore, 
may have more pronounced or longerlasting effects than 
naturally-occurring); or which have been altered to delete or to add one 
or more potential sites for O-glycosylation and/or N-glycosylation or 
which have one or more cysteine residues deleted or replaced by e.g., 
alanine or serine residues and are potentially more easily isolated in 
active form from microbial systems; or which have one or more tyrosine 
residues replaced by phenylalanine and bind more or less readily to target 
proteins or to receptors on target cells. Also comprehended are 
polypeptide fragments duplicating only a part of the continuous amino acid 
sequence or secondary conformations within zinc sphingomyelinase, which 
fragments may possess one property of zinc sphingomyelinase (i.e. 
substrate binding) and not others (e.g., enzymatic activity). It is 
noteworthy that activity is not desired for any one or more of the 
products of the invention to have therapeutic utility or utility in other 
contexts, such as in assays of zinc sphingomyelinase activity. Competitive 
antagonists may be quite useful in, for example, cases of overproduction 
of zinc sphingomyelinase or cases of atherosclerosis where the endothelial 
cells associated with lesions overexpress the enzyme. 
One embodiment of the present invention is wherein the agent or the 
inhibitor is a peptidomimetic compound. The peptidomimetic compound may be 
at least partially unnatural. The peptidomimetic compound may be a small 
molecule mimic of the binding site of zinc sphingomyelinase or a mimic of 
the active site of zinc sphingomyelinase. The compound may have increased 
stability, efficacy, potency and bioavailability by virtue of the mimic. 
Further, the compound may have decreased toxicity. The peptidomimetic 
compound may have enhanced mucosal intestinal permeability. The compound 
may be synthetically prepared. The compound of the present invention may 
include L-, D- or unnatural amino acids, alpha, alpha-disubstituted amino 
acids, N-alkyl amino acids, lactic acid (an isoelectronic analog of 
alanine). The peptide backbone of the compound may have at least one bond 
replaced with PSI-[CH.dbd.CH] (Kempf et al. 1991). The compound may 
further include trifluorotyrosine, p-Cl-phenylalanine, p-Br-phenylalanine, 
poly-L-propargylglycine, poly-D,L-allyl glycine, or poly-L-allyl glycine. 
One embodiment of the present invention is a peptidomimetic compound having 
the biological activity of zinc sphingomyelinase wherein the compound has 
a bond, a peptide backbone or an amino acid component replaced with a 
suitable mimic. Examples of unnatural amino acids which may be suitable 
amino acid mimics include .beta.-alanine, L-.alpha.-amino butyric acid, 
L-.gamma.-amino butyric acid, L-.alpha.-amino isobutyric acid, 
L-.epsilon.-amino caproic acid, 7-amino heptanoic acid, L-aspartic acid, 
L-glutamic acid, cysteine (acetamindomethyl), 
N-.epsilon.-Boc-N-.alpha.-CBZ-L-lysine, 
N-.epsilon.-Boc-N-.alpha.-Fmoc-L-lysine, L-methionine sulfone, 
L-norleucine, L-norvaline, N-.alpha.-Boc-N-.delta.CBZ-L-ornithine, 
N-.delta.-Boc-N-.alpha.-CBZ-L-ornithine, Boc-p-nitro-L-phenylalanine, 
Boc-hydroxyproline, Boc-L-thioproline. (Blondelle, et al. 1994; Pinilla, 
et al. 1995). 
In one embodiment of the invention, the substrate comprises sphingomyelin 
or a derivative thereof in a detergent micelle at acidic pH; or a modified 
lipoprotein. In another embodiment of the invention, the substrate is 
detectably labeled. In another embodiment of the invention, the detectable 
label comprises radioisotope or fluorescence. 
The invention also provides for a method for screening a library of 
compounds to identify a compound capable of inhibiting zinc 
sphingomyelinase enzyme activity which comprises: (a) incubating a zinc 
sphingomyelinase enzyme, under acid pH conditions suitable for zinc 
sphingomyelinase enzyme activity, with: (i) a substrate of 
sphingomyelinase enzyme, and (ii) a sample from a library of compounds; 
(b) detecting the amount of sphingomyelinase enzyme activity produced by 
determining whether there exists an increase in ceramide concentration in 
the incubate; and (c) comparing the amount of enzyme activity detected in 
step (b) with an amount of enzyme activity detected in the absence of the 
compound, thereby evaluating the ability of the sample to inhibit activity 
of zinc sphingomyelinase enzyme, and (d) repeating step (a) with limiting 
dilutions of the sample so as to identify the compound in the sample 
capable of inhibiting zinc sphingomyelinase enzyme activity. The 
incubating in step (a) may be carried out at neutral pH. 
In one embodiment of the invention, the carrier comprises a diluent. In 
another embodiment of the invention, the carrier comprises an appropriate 
adjuvant, a liposome, a microencapsule, a polymer encapsulated cell, a 
biodegradable plastic substance or a retroviral vector. In another 
embodiment of the invention, the pharmaceutically acceptable carrier is an 
aerosol, intravenous, oral or topical carrier. 
In one preferred embodiment the pharmaceutical carrier may be a liquid and 
the pharmaceutical composition would be in the form of a solution. In 
another equally preferred embodiment, the pharmaceutically acceptable 
carrier is a solid and the composition is in the form of a powder or 
tablet. In a further embodiment, the pharmaceutical carrier is a gel and 
the composition is in the form of a suppository or cream. In a further 
embodiment the active ingredient may be formulated as a part of a 
pharmaceutically acceptable transdermal patch. 
A solid carrier can include one or more substances which may also act as 
flavoring agents, lubricants, solubilizers, suspending agents, fillers, 
glidants, compression aids, binders or tablet-disintegrating agents; it 
can also be an encapsulating material. In powders, the carrier is a finely 
divided solid which is in admixture with the finely divided active 
ingredient. In tablets, the active ingredient is mixed with a carrier 
having the necessary compression properties in suitable proportions and 
compacted in the shape and size desired. The powders and tablets 
preferably contain up to 99% of the active ingredient. Suitable solid 
carriers include, for example, calcium phosphate, magnesium stearate, 
talc, sugars, lactose, dextrin, starch, gelatin, cellulose, 
polyvinylpyrrolidine, low melting waxes and ion exchange resins. 
Liquid carriers are used in preparing solutions, suspensions, emulsions, 
syrups, elixirs and pressurized compositions. The active ingredient can be 
dissolved or suspended in a pharmaceutically acceptable liquid carrier 
such as water, an organic solvent, a mixture of both or pharmaceutically 
acceptable oils or fats. The liquid carrier can contain other suitable 
pharmaceutical additives such as solubilizers, emulsifiers, buffers, 
preservatives, sweeteners, flavoring agents, suspending agents, thickening 
agents, colors, viscosity regulators, stabilizers or osmo-regulators. 
Suitable examples of liquid carriers for oral and parenteral 
administration include water (partially containing additives as above, 
e.g. cellulose derivatives, preferably sodium carboxymethyl cellulose 
solution), alcohols (including monohydric alcohols and polyhydric 
alcohols, e.g. glycols) and their derivatives, and oils (e.g. fractionated 
coconut oil and arachis oil). For parenteral administration, the carrier 
can also be an oily ester such as ethyl oleate and isopropyl myristate. 
Sterile liquid carriers are useful in sterile liquid form compositions for 
parenteral administration. The liquid carrier for pressurized compositions 
can be halogenated hydrocarbon or other pharmaceutically acceptable 
propellent. 
Liquid pharmaceutical compositions which are sterile solutions or 
suspensions can be utilized by for example, intramuscular, intrathecal, 
epidural, intraperitoneal or subcutaneous injection. Sterile solutions can 
also be administered intravenously. The active ingredient may be prepared 
as a sterile solid composition which may be dissolved or suspended at the 
time of administration using sterile water, saline, or other appropriate 
sterile injectable medium. Carriers are intended to include necessary and 
inert binders, suspending agents, lubricants, flavorants, sweeteners, 
preservatives, dyes, and coatings. 
The active ingredient can be administered orally in the form of a sterile 
solution or suspension containing other solutes or suspending agents, for 
example, enough saline or glucose to make the solution isotonic, bile 
salts, acacia, gelatin, sorbitan monoleate, polysorbate 80 (oleate esters 
of sorbitol and its anhydrides copolymerized with ethylene oxide) and the 
like. 
The active ingredient can also be administered orally either in liquid or 
solid composition form. Compositions suitable for oral administration 
include solid forms, such as pills, capsules, granules, tablets, and 
powders, and liquid forms, such as solutions, syrups, elixirs, and 
suspensions. Forms useful for parenteral administration include sterile 
solutions, emulsions, and suspensions. 
This invention is illustrated in the Experimental Details section which 
follows. These sections are set forth to aid in an understanding of the 
invention but are not intended to, and should not be construed to, limit 
in any way the invention as set forth in the claims which follow 
thereafter. 
EXPERIMENTAL DETAILS 
EXAMPLE 1 
Rabbit Aorta and Human Atherosclerotic Lesions Hydrolyze the Sphingomyelin 
of Retained Low-density Lipoprotein: Role for Arterial-wall 
Sphingomyelinase in Subendothelial Retention and Aggregation of 
Atherogenic Lipoproteins 
Aggregation and retention of low-density lipoprotein (LDL) in the arterial 
wall are key events in atherogenesis, but the mechanisms in vivo are not 
yet fully understood. Exposure of LDL to bacterial sphingomyelinase 
(SMase) in vitro leads to the formation of LDL aggregates that can be 
retained by extracellular matrix and that are able to stimulate macrophage 
foam cell formation. Evidence is provided herein that shows retained LDL 
is hydrolyzed by an arterial-wall SMase activity. First, SMase-induced 
aggregation was demonstrated to be caused by an increase in particle 
ceramide content, even in the presence of excess SM. This finding is 
compatible with data showing that lesional LDL is enriched in SM, though 
its ceramide content has not previously been reported. To address this 
critical compositional issue, the ceramide content of lesional LDL was 
assayed and, remarkably, found to be 10-50-fold enriched compared with 
plasma LDL ceramide. Furthermore, the ceramide was found exclusively in 
lesional LDL that was aggregated; unaggregated lesional LDL, which 
accounted for 20-25% of the lesional material, remained ceramide-poor. 
When [.sup.3 H]SM-LDL was incubated with strips of rabbit aorta ex vivo, a 
portion of the LDL was retained, and the [.sup.3 H]SM of this portion, but 
not that of unretained LDL, was hydrolyzed to [.sup.3 H]ceramide by a 
nonlysosomal arterial hydrolase. In summary, LDL retained in 
atherosclerotic lesions is acted upon by an arterial-wall SMase, which may 
participate in LDL aggregation and possibly other SMase-mediated processes 
during atherogenesis. 
The mechanism of lipoprotein aggregation in lesions is not known. 
Lipoprotein aggregation can be induced in vitro by vortexing (Khoo et al., 
1988), extensive phospholipase C hydrolysis (Suits et al., 1989), 
extensive oxidation (Hoff et al., 1989), and limited hydrolysis with 
bacterial sphingomyelinase (SMase) (Xu et al., 1991). Vortexing and 
extensive hydrolysis by phospholipase C are unlikely to be physiologically 
important. LDL oxidation does occur in arteries (Steinberg et al., 1989), 
but, as mentioned above, subendothelial LDL aggregates have been shown to 
be present in normal rabbit aorta as early as two hours after an 
intravenous bolus injection of LDL (Nievelstein et al., 1991), which may 
be too soon for substantial LDL oxidation to occur in these normal 
vessels. One goal of the current study was to test the possible 
physiological relevance of SMase-induced lipoprotein aggregation. In 
particular, it was demonstrated that the mechanism of SMase-induced 
aggregation of LDL in vitro is entirely consistent with a role for SM 
hydrolysis in LDL aggregation in vivo, that extracellular, retained LDL 
extracted from human atherosclerotic lesions shows evidence of having been 
acted upon by an arterial SMase, and that strips of rabbit aorta ex vivo 
can hydrolyze the SM of retained LDL. The results support a role for 
arterial SMase in LDL aggregation and possibly other SMase-mediated 
processes during atherogenesis. 
Methods 
Materials. sn-1,2-diacylglycerol kinase (from Escherichia coli) was 
purchased from Calbiochem (San Diego, Calif.). Cardiolipin and 
1,2-dioleoyl glycerol were purchased from Avanti Polar Lipids Alabaster, 
Ala.). The Superose 6 HR 10/30 gel filtration column was obtained from 
PHARMACIA.RTM. (Piscataway, N.J.). [9,10-.sup.3 H]palmitic acid, 
[.gamma.-.sup.32 P]ATP, and Na.sup.125 I were obtained from DuPont-New 
England Nuclear (Boston, Mass.). Tissue culture media, tissue culture 
reagents, and human recombinant epidermal growth factor were purchased 
from Life Technologies (Baltimore, Md.), fetal bovine serum was from 
GEMINI.RTM. Bioproducts (Calabasas, Calif.), and glass tissue culture 
plates were from CORNING.RTM. (Corning, N.Y.). SMase (from Bacillus 
cereus) and all other reagents were from SIGMA.RTM. (St. Louis, Mo.). 
Peroxidase-conjugated goat anti-rabbit IgG was purchased from PIERCE.RTM. 
Chemical Co. (Rockford, Ill.). 
Synthesis of [.sup.3 H]SM [N-palmitoyl-9,10-.sup.3 H]SM was synthesized as 
previously described (Sripada et al., 1987; Ahmad et al., 1985). Briefly, 
[9,10-.sup.3 H]palmitic acid (25 mCi, 450 nmol) was stirred for 12 h at 
room temperature with an equimolar equivalent of (N)-hydroxysuccinimide 
and with 3-molar equivalents of 1,3-dicyclohexylcarbodiimide in 
(N,N)-dimethylformamide. The reaction was run under dry argon in the dark. 
Sphingosylphosphorylcholine (300 nmol) and (N,N)-diisopropylethylamine (10 
.mu.l) were then added and the reaction was stirred another 12 h at room 
temperature. The reaction was stopped by evaporating the 
(N,N)-dimethylformamide under a stream of N.sub.2. 
[N-palmitoyl-9,10-.sup.3 H]SM was purified by preparative thin-layer 
chromatography of the reaction products three consecutive times in 
chloroform:methanol (95:5) and then twice in chloroform:methanol:acetic 
acid:water (50:25:8:4). Greater than 95% of the [N-palmitoyl-9,10-.sup.3 
H]SM was converted to [N-palmitoyl-9,10-.sup.3 H]ceramide after treatment 
with 10 mU SMase/ml (Bacillus cereus) for 1 h at 37.degree. C., as assayed 
by TLC, indicating a pure, functional substrate. 
Lipoproteins. LDL (density, 1.020-1.063 g/ml) was isolated from fresh human 
plasma by preparative ultracentrifugation as previously described (Havel 
et al., 1955). Plasma LDL was labeled with [N-palmitoyl-9,10-.sup.3 H]SM 
as follows: .about.3.5 mCi (63 nmol) [N-palmitoyl-9,10-.sup.3 H]SM and 13 
nmol phosphatidylcholine (PC) were mixed in chloroform, and the solvent 
was removed first under a stream of nitrogen and then by lyophilization. 
The dried lipids were resuspended in 1 ml of 150 mM NaCl, 1 mM EDTA, 10 mM 
Tris-HCl, pH 7.5 and, to prepare [.sup.3 H]SM/PC liposomes, sonicated for 
three 50-sec pulses at 4.degree. C. using a tapered microtip on a Branson 
450 sonicator (setting #3). The liposomes were then incubated with 30 mg 
(by protein mass) of LDL, 50 g of partially purified phospholipid transfer 
protein, 100 U penicillin, and 100 g streptomycin for 18 h at 37.degree. 
C. under argon. LDL was then separated from the liposomes after 
phospholipid transfer by centrifuging the mixture at density=1.006 g/ml 
for 8 h at 35,000 rpm in a BECKMAN.RTM. 50.3 rotor; the supernate 
containing the liposomes was removed, and the LDL band at the bottom of 
the tube was harvested. The LDL solution was mixed with PBS and 
centrifuged as before. This wash procedure was performed a total of four 
times, resulting in the removal of 95% of the unreacted [.sup.3 H]SM/PC 
liposomes. All lipoproteins were stored under argon at 4.degree. C. and 
were used within 2 weeks of preparation. 
Control and SM-enriched LDL-lipid emulsions. Two aliquots of LDL (each 12 
mg by protein mass) were extracted by the method of Bligh and Dyer (Bligh 
et al., 1959); 6 mg of SM was added to the lipid extract of one of the 
aliquots. The solvent was then completely evaporated from these samples by 
exposure to a stream of nitrogen, followed by lyophilization. The dried 
lipids were resuspended in 6 ml of PBS and sonicated under a stream of 
argon at 40.degree. C. until translucent (90 min for the control emulsions 
and 130 min for the SM-enriched emulsions). The sonicated material was 
then centrifuged twice at 15,000.times.g, and the supernate was harvested. 
Ceramide and diacylglycerol assays. Ceramide and diacylglycerol were 
measured from aliquots of lipid extracts of LDL and emulsions using the 
method described by Schneider and Kennedy (Schneider et al., 1976) and 
adapted by Preiss et al. (Preiss et al., 1986). In this method, 
diacylglycerol (DAG) kinase phosphorylates ceramide and DAG using 
[.gamma.-.sup.32 P]ATP; the contents of ceramide and DAG in the 
experimental samples are calculated from amount of incorporated .sup.32 P 
label in comparison with standard curves using known quantities of the two 
lipids. For ceramide measurements, the lipids were first incubated with 
0.1 N KOH in methanol for 1 h at 37.degree. C., which hydrolyzes DAG, but 
not ceramide. The extracted lipids were dried under nitrogen and then 
solubilized in 5 mM cardiolipin, 7.5% octyl-.beta.-glucopyranoside, and 1 
mM diethylenetriaminepentaacetic acid by bath sonication. This solution 
was then added to reaction buffer (50 mM imidazole-HCl, pH 6.6, 50 mM 
NaCl, 12.5 mM MgCl.sub.2, 1 mM EGTA) containing sn-1,2-DAG kinase (0.7 
units/ml). The reaction was initiated by the addition of [.gamma.-.sup.32 
P]ATP (final concentration=10 mM). After incubation at room temperature 
for 60 min, the reaction was stopped by lipid extraction with 
chloroform:methanol:HCl (100:100:1, v/v/v) and 10 mM EDTA. 
Ceramide-1-phosphate and phosphatidic acid in the organic phase were 
separated by TLC using chloroform:methanol:acetic acid (65:15:5, v/v/v); 
the lipids were visualized with autoradiography and identified by 
comparing with standards. The spots corresponding to these two lipids were 
scraped and counted, and the masses of the lipids were calculated by 
comparison with standard curves, as described above. 
Sphingomyelin (SM), phosphatidylcholine (PC), and cholesterol assays. Lipid 
extracts (20) of LDL and emulsions were chromatographed by TLC using 
chloroform:methanol:acetic acid:H.sub.2 O (50:25:8:4, v/v/v/v). Individual 
phospholipid subclasses were visualized by iodine vapor staining, and the 
SM and PC spots were identified by comparison with standards. The spots 
were scraped, extracted twice with chloroform:methanol (2:1), and assayed 
for phosphate content by the method of Bartlett (Bartlett, 1959). Total 
cholesterol contents were assayed using an enzymatic colorimetric method 
(Cholesterol C kit, Wako Chemicals U.S.A., Inc., Richmond, Va.); it was 
verified that the values obtained by this method are similar to those 
obtained using gas-liquid chromatography. 
Isolation of LDL from human lesions. LDL was extracted from abdominal 
aortic aneurysm plaque material as previously described (Rapp et al., 
1994). Briefly, aortic plaque was removed from individuals as part of the 
standard reconstructive surgery for abdominal aortic aneurysms at the San 
Francisco Veterans Affairs Medical Center. Plaque material, which ranged 
in weight from 2-12 grams, was obtained in the operating room and 
immediately placed into ice-cold 7-mM citrate buffer, pH 7.4, containing 
15 mM NaCl, 3 mM EDTA, 0.5 mM butylhydroxytoluene, 1 mM 
phenylmethylsulfonylfluoride, 1.5 mg aprotinin/ml, 2 mM benzamidine, and 
0.08 mg gentamycin sulfate/ml. Blood and adherent thrombus were removed by 
blotting with absorbent gauze, scrubbing with a small brush, and sharp 
dissection as necessary. Loosely retained lipoproteins were extracted by 
mincing the plaque into 0.5-1.0 mm.sup.2 pieces and incubating them 
overnight on a LABQUAKE.RTM. shaker at 4.degree. C. in a non-denaturing 
buffer (0.1 M citrate, pH 7.4, with 1 mg EDTA/ml, 0.3 mg benzamidine/ml, 
0.08 mg gentamicin sulfate/ml, 10 .mu.g aprotinin/ml, 10 .mu.g Trolox [an 
anti-oxidant]/ml, and 20 .mu.g phenylmethylsulfonyl fluoride/ml). The 
extracted material was cleared of particulate matter by centrifuging at 
800.times.g for 10 min, and 1.019&lt;d&lt;1.063-g/ml lipoproteins were isolated 
by sequential sodium bromide density ultracentrifugation (Rapp et al., 
1994; Havel et al., 1955). To extract LDL from early human lesions, the 
same procedure was used, except the intima and inner media were peeled 
away from the outer media and adventitia before being minced and processed 
as above. 
Apolipoprotein B-100 slot immunoblot analysis. The samples to be tested 
were applied to a nitrocellulose membrane using a slot blot apparatus. 
Next, the membrane was 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 membrane was then incubated with rabbit anti-human 
apolipoprotein B-100 antiserum (1:1000) in buffer B (buffer A containing 
0.1% Tween-20, 3% nonfat dry milk, and 0.1% bovine serum albumin) for 4 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 membrane was subsequently washed twice with 0.3% 
Tween-20 in buffer A and twice with 0.1% Tween-20 in buffer A. Finally, 
the blot was soaked in the enhanced chemiluminescence reagent for 2 min 
and exposed to X-ray film for 1 min. For FIGS. 5A-D, the relative 
intensities of the slot blot signals were determined by densitometric 
scanning of the X-ray film using a Molecular Dynamics Computing 
Densitometer (model 300A) with IMAGE-QUANT.RTM. software. 
Assay for LDL-SM hydrolysis by strips of rabbit aorta ex vivo. New Zealand 
white male rabbits (2-3 kg) were fed a chow diet containing 0.2% 
cholesterol and 10% soybean oil (w/w) for 18 days. The animals were 
sacrificed, and the thoracic aortae were removed rapidly and placed in a 
dissection dish with physiological Hank's Trizma maleate buffer (133 mM 
NaCl, 3.6 mM KCl, 1.0 mM CaCl.sub.2, 5.0 mM Trizma maleate, 16 mM 
dextrose, pH 7.3). The aortae were then further dissected to remove excess 
fat and any clotted blood and then cut longitudinally and pinned, lumen 
side up, in a tissue culture dish. The aortic strips were washed several 
times with DMEM/0.2% BSA and incubated at 37.degree. C. for 3.5 h in 
DMEM/0.2% BSA containing 2 mg [.sup.3 H]SM-labeled LDL per ml. For certain 
experiments, the strips were preincubated at 37.degree. C. for 30 min in 
medium alone, or supplemented with 15 mM EDTA or 200 .mu.M chloroquine. 
When present, these reagents were also added to the corresponding 
incubation media containing labeled LDL (see legend to FIG. 5). After the 
3.5-h incubation of labeled LDL with aortic strips, media were removed 
(see below), the aortic strips were quickly washed twice in ice-cold PBS, 
and the center of the strip was cut away from the original cut ends. These 
center pieces were then minced and extracted in PBS, pH 7.4, containing 6 
M guanidine-HCl, 5 mM EDTA, 0.02% sodium azide, 35 .mu.M leupeptin, 1.5 mg 
aprotinin/ml, and 5 .mu.M pepstatin A for 36 h at 4.degree. C. The extract 
was centrifuged at 35,000.times.g for 30 min, and lipids were extracted 
from the supernate by the method of Bligh and Dyer (Bligh et al., 1959). 
[.sup.3 H]palmitate-SM, .sup.3 [H]palmitate-ceramide, and free [.sup.3 
H]palmitate in the lipid extracts were separated by TLC using 
chloroform:methanol (95:5, v/v) or n-propanol:12.5% ammonium hydroxide 
(80:20, v/v). For controls, [.sup.3 H]SM-LDL that was incubated for 3.5 h 
in media without aortic strips, and [.sup.3 H]SM-LDL that was incubated 
with the strips but not retained (i.e., in the media that was harvested 
after the incubation) were subjected to the same extraction procedures, 
including the 36-h incubation in guanidine buffer. 
Ionizations of EGF and assay for .sup.125 I-EGF degradation. Epidermal 
growth factor (50 .mu.g) was radioiodinated to a specific activity of 5400 
counts/min/ng (IODOBEADS.RTM., Pierce Chemical Co., Rockford, Ill.) and 
was used within 48 h of labeling. After a 3.5-h incubation with aortic 
strips, in the absence or presence of 200 .mu.M chloroquine, degradation 
of .sup.125 I-EGF was assayed by the release of .sup.125 I-tyrosine 
(Golstein et al., 1983). 
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. 
Results 
SMase-induced aggregation of LDL in vitro requires active SMase enzyme and 
is mediated via high particle ceramide content. Determination of whether 
active SMase enzyme is required for LDL aggregation or whether the effect 
is purely structural was first sought; for example, lipoprotein 
lipase-mediated bridging of LDL to cell-surface proteoglycans is a purely 
structural effect of the lipase molecule. In vitro studies utilized 
bacterial SMases, which are a reasonable model for mammalian SMases in 
regard to enzymatic activity but not necessarily in regard to structural 
actions. Careful analysis of the time course of B. cereus SMase-induced 
LDL aggregation and LDL-SM hydrolysis was done. Whereas LDL-SM hydrolysis 
was 70% complete within the first 30 min of exposure to SMase (FIG. 1A, 
closed triangles), substantial LDL aggregation began to occur only after 
30 min of incubation (FIG. 1B, open squares). These kinetics are 
consistent with two possible mechanisms: a slow structural action of 
SMase, or a rapid enzymatic action that alters the particles so that they 
then slowly aggregate. To distinguish between these possibilities, EDTA 
was used to inhibit the enzymatic activity of the bacterial B. cereus 
SMase, which is a Mg.sup.2+ -dependent enzyme (Ikezawa et al., 1986). As 
in FIG. 1A, substantial aggregation of LDL occurred in the absence of EDTA 
after an initial lag period (FIG. 1B, open squares). When EDTA was added 
at the beginning of the reaction, however, there was complete inhibition 
of LDL-SM hydrolysis (inset) and of LDL aggregation (open triangles). To 
show that EDTA is not a direct inhibitor of the aggregation process 
itself, EDTA was added 20 min after the SMase reaction had begun. 68% of 
LDL-SM was hydrolyzed under these conditions (FIG. 1C). Despite the 
presence of EDTA during the period of the aggregation process (i.e., 
following the 20-min incubation with SMase), substantial LDL aggregation 
still occurred (FIG. 1B, closed triangles). The degree of aggregation in 
this setting was consistent with the degree of LDL-SM hydrolysis under 
these conditions. Thus, EDTA can inhibit SMase-induced aggregation of LDL, 
but solely through its inhibition of SMase enzymatic activity, with no 
direct effects on the aggregation process itself. SMase that has been 
enzymatically inactivated by EDTA can no longer provoke LDL aggregation, 
indicating that enzymatic activity is an absolute requirement. 
Next it was determined which of the three consequences of SMase enzymatic 
activity, namely, generation of choline-phosphate, generation of ceramide, 
and depletion of SM, is responsible for LDL aggregation. One of these 
possibilities, SM depletion, cannot be relevant to the pathogenesis of 
atherosclerosis: lesional LDL, which aggregates, is known to be enriched 
in SM compared to plasma LDL. Incubating LDL with a 10-fold molar excess 
of choline-phosphate was performed. This treatment failed to increase the 
turbidity of LDL, indicating that choline-phosphate does not mediate 
SMase-induced aggregation of LDL. 
Work was then done to distinguish whether aggregation results from an 
increase in LDL-ceramide or from a low LDL-SM content. Attempts to 
significantly alter SM and ceramide contents of intact LDL, for example by 
using phospholipid transfer protein or cholesteryl ester transfer protein, 
proved unsuccessful (see below). Therefore, LDL lipids were used to 
prepare protein-free synthetic emulsions, which aggregate like native LDL 
after SMase treatment. Emulsions were prepared with either LDL lipids 
(control emulsions) or LDL lipids to which extra SM was added. Partial 
digestion of these emulsions with SMase was accomplished by adding EDTA to 
stop the reaction after only 15 min at 37.degree. C. (see above). The 
control emulsion showed substantial particle aggregation (compare solid 
bars in first and third triplets of bars in FIG. 2). Partial enzymatic 
digestion of the SM-enriched emulsions generated ceramide-rich particles 
that still had a very high SM content (two-fold greater than in undigested 
control emulsions). These ceramide-rich, SM-rich particles aggregated to 
an even greater degree than SMase-treated control emulsions (compare solid 
bars in third and fourth triplets of bars in FIG. 2), presumably because 
they had more ceramide than the control emulsions (see below). Thus, 
SMase-induced aggregation readily occurs even when the residual SM content 
of the particles is still very high, as it is in lesional LDL (Hoff, 1983; 
Yla-Herttuala et al., 1989). SM depletion cannot explain SMase-induced 
aggregation. These results indicate that the increase in particle ceramide 
is the key factor in this process. To test this relationship directly 
using intact LDL, plasma LDL was treated with SMase for varying times to 
generate particles with varying amounts of ceramide, allowed to aggregate, 
and then assayed for both aggregation and ceramide content. After an 
initial "threshold" level of .about.0.08 nmol ceramide/nmol 
phosphatidylcholine, there was a direct relationship between LDL-ceramide 
content and LDL aggregation (FIG. 3). 
In summary, SMase-induced aggregation of LDL is mediated by enzymatic, not 
structural, actions of the SMase, and the key factor in causing 
aggregation is an increase in particle ceramide content, not a low 
absolute SM content. Both of these findings are consistent with the 
hypothesis that an arterial-wall SMase may contribute to the aggregation 
of lesional LDL. Note also that particle aggregation does not depend on 
apoB. 
Human atheroma-derived LDL is enriched in ceramide. The finding that 
LDL-ceramide content is the key factor in SMase-induced aggregation led to 
an examination of the ceramide content of human lesional LDL. For these 
experiments, material was obtained from two types of human lesions: 
atherectomy material (advanced atherosclerosis) and human aorta with only 
fatty streak involvement (early atherosclerosis). Tissue specimens were 
extracted overnight at 4.degree. C. in a non-denaturing aqueous buffer, 
and 1.019&lt;d&lt;1.063-g/ml lipoproteins were isolated from the extraction 
buffer by sequential ultracentrifugation as previously reported (Rapp et 
al., 1994). 
Three separate experiments, analyzing the ceramide contents of lesional 
material from five different atherectomy samples altogether, are shown in 
FIGS. 4A-C. Same-donor plasma LDL was available for comparison with Lesion 
LDL #1, #3, and #5; Lesion LDL #2 and #4 were compared with two different 
samples of plasma LDL prepared from plasma (labeled Plasma LDL A and 
Plasma LDL B in FIG. 4). Remarkably, the samples of lesional LDL had a 
ceramide content that was 10-50-fold higher than that of plasma LDL, which 
was always ceramide-poor. Further characterization of the lipids in 
lesional LDL by HPLC confirmed the presence of ceramide, with only trace 
amounts of the biosynthetic intermediate, dihydroceramide (see Merrill et 
al., 1990). To verify that the extraction procedure did not cause 
artifactual hydrolysis of LDL-SM, it was shown that [.sup.3 H]SM-labeled 
LDL added to extraction buffer with an unlabeled arterial sample was not 
hydrolyzed during extraction or sequential ultracentrifugation 
(0.006.+-.0.002 percent of the radioactivity in the original preparation 
of [.sup.3 H]SM-LDL co-migrated with ceramide on TLC, and this fraction 
remained exactly the same in [.sup.3 H]SM-LDL subjected to the extraction 
procedure). To show that the ceramide was in LDL, Lesion LDL #1 was 
fractionated using a Superose-6 gel-filtration column (molecular weight 
exclusion=4.times.10.sup.6), a commonly used chromatographic technique for 
separating lipoprotein classes. Note that the column was run in buffer 
containing 6 M guanidine-HCl to partially dissociate large LDL aggregates 
(Hoff et al., 1985; and below), which otherwise might not enter the column 
or elute in the void volume, and thus not be separated from lesional 
debris. As shown in FIG. 4D, cholesterol, ceramide, and apolipoprotein 
B-100 (inset) co-eluted within the included volume of the column. Finally, 
to determine if there had been generalized break-down of phospholipids in 
LDL from advanced lesions, we assayed Lesion and Plasma LDL #3 for DAG, 
which is a product of phosphatidylcholine hydrolysis by phospholipase C. 
Whereas the ceramide content of the lesional LDL was 54 well as of the 
sample of plasma LDL, was below the detection limit of the assay (&lt;2 
pmol/nmol PC). These data indicate specific hydrolysis of SM in lesional 
LDL and no evidence of a significant role for phospholipase C in vivo (cf. 
Suits et al., 1989). 
To examine LDL from early human lesions, the same extraction procedure was 
used to obtain LDL from a segment of aorta from a 43-y/o male donor heart. 
Gross examination of this aortic segment revealed only fatty streaks, 
without any raised lesions. The ceramide content of the lesional LDL was 
0.031.+-.0.002 nmol/nmol PC, whereas same-donor plasma LDL had only 
0.013.+-.0.005 nmol ceramide/nmol PC. Thus, even in relatively early 
atherosclerosis, lesional LDL is 2-3-fold enriched in ceramide compared 
with plasma LDL. 
Only aggregated lesional LDL is enriched in ceramide. Lesion LDL #5 from 
FIG. 4 was characterized for aggregation using centrifugation and 
gel-filtration chromatography, as described previously (Lougheed et al., 
1996). As shown in FIG. 5A, approximately 60% of the lesional 
LDL-cholesterol pelleted during a brief centrifugation (10 min at 
10,000.times.g), indicating the presence of apparently very large 
aggregates. This pelleted material contained ceramide (FIG. 5A), 
consistent with a role for ceramide in LDL aggregation. To determine 
whether the ceramide in the 10,000.times.g supernate (FIG. 5A) was also 
associated with aggregates of LDL, this material was fractionated by 
gel-filtration chromatography using SUPEROSE.RTM. 6 as in FIG. 4, but in 
the absence of guanidine-HCl (FIGS. 5B-D). The column was calibrated with 
plasma LDL, which eluted in a sharp peak centered around fraction #14 
based on cholesterol and apo B-100 immunoreactivity (L in FIG. 5B). The 
data that the cholesterol (FIG. 5B) and apo B-100 (FIG. 5C) of the 
10,000.times.g supernate eluted in two peaks. The second peak, which 
contained approximately 40% of the cholesterol and 50% of the apo B-100, 
eluted in the position of plasma LDL, indicating unaggregated material. 
The first peak eluted four fractions earlier, indicating aggregation. 
Essentially all of the ceramide was associated with the first peak of 
lesional LDL (FIG. 5D). Thus, lesional LDL exists in three forms: 
apparently very large aggregates, smaller aggregates, and particles that 
behave like unaggregated LDL. Importantly, ceramide is present only in the 
aggregated forms of lesional LDL. 
LDL-SM hydrolysis by the arterial wall. Enrichment of lesional LDL with 
ceramide could occur through two processes: hydrolysis of LDL-SM or 
transfer of pre-existing ceramide from cells or cellular debris onto the 
LDL particles. To examine ceramide transfer, enrichment of LDL was 
attempted in vitro by incubation with suspensions of ceramide or with 
sonicated emulsions containing ceramide. Even in the presence of 
phospholipid transfer protein or cholesteryl ester transfer protein, 
however, unable to incorporate pre-existing ceramide into LDL. The only 
method found to enrich LDL with ceramide in vitro was by digestion of the 
lipoprotein with SMase (see FIGS. 1-3). These data indicate that ceramide 
transfers poorly, if at all, and that this mechanism is unlikely to 
explain the lipid composition of lesional LDL. 
To directly examine LDL-SM hydrolysis in arteries, strips of rabbit aorta 
were incubated with [.sup.3 H]palmitate-SM-labeled LDL for 3.5 h at 
37.degree. C. and then extracted the retained LDL in guanidine buffer to 
look for evidence of [.sup.3 H]ceramide generation (cf. Nievelstein-Post 
et al., 1994). Histologic analysis of thin sections of the vessel strips 
after the 3.5-h incubation showed no cellular damage when compared to 
freshly fixed vessels. The data from two separate experiments are shown in 
FIG. 6A. In both experiments, material extracted from the aortic strips 
had markedly increased [.sup.3 H]ceramide compared with unretained [.sup.3 
H]SM-LDL or with [.sup.3 H]SM-LDL incubated for 3.5 h in media without 
aortic strips (compare third pair of bars in FIG. 6A with the first two 
pairs of bars). To determine whether the extraction procedure itself 
caused artifactual hydrolysis of LDL-SM, this procedure was performed 
using [.sup.3 H]SM-LDL added to extraction buffer in the presence of 
minced pieces of aortic strips that had not been incubated previously with 
[.sup.3 H]SM-LDL. Under these conditions, there was no hydrolysis of 
[.sup.3 H]SM-LDL. Thus, the data in FIG. 6A indicate hydrolysis of LDL-SM 
within the arterial wall. 
At least four different SMases have been shown to be present in mammalian 
cells. Two of these are encoded by the acid SMase gene and arise by 
differential post-translational processing (Schissel et al., 1996). One of 
these acid SMase gene products is located in lysosomes (lysosomal SMase), 
does not require added cations, and is not inhibited by EDTA (Spence, 
1993). The other SMase derived from the acid SMase gene is secreted by a 
wide variety of cell types (Schissel et al., 1996) and requires exogenous 
Zn.sup.2+ for activity (Schissel et al., 1996; Spence et al., 1989). In 
addition, cells have a membrane-bound neutral SMase that requires 
Mg.sup.2+ for activity (Chatterjee, 1993) as well as a cytoplasmic neutral 
SMase that is cation-independent (Okazaki et al., 1994). Thus, two of the 
four known mammalian SMases require a divalent cation. To determine if 
aortic hydrolysis of LDL-SM requires divalent cations, portions of the 
aortic strips from the two experiments in FIG. 6A were preincubated with 
15 mM EDTA and then incubated with the [.sup.3 H]SM-LDL plus EDTA. As 
shown in the last doublet of bars in FIG. 6A, EDTA treatment decreased the 
generation of ceramide in the extracted material by approximately 50%. 
These data indicate that at least a portion of the arterial-wall SMase 
activity that hydrolyzes retained [.sup.3 H]SM-LDL requires divalent 
cations. 
The overall hypothesis assumes that LDL-SM is hydrolyzed extracellularly, 
not intracellularly in lysosomes. The fact that a substantial portion of 
the arterial-wall SMase acting on retained LDL is inhibited by EDTA (FIG. 
6A) indicates that much of the LDL-SM is not being hydrolyzed by lysosomal 
SMase, which is EDTA-resistant (see above). Furthermore, the [.sup.3 
H]palmitate-SM-labeled LDL that was retained in the aortic strips showed 
no enrichment in free [.sup.3 H]palmitate, thereby suggesting no contact 
of the particle with lysosomal ceramidase (Bernardo et al., 1995). To 
further assess the contribution of lysosomes, a set of experiments was 
conducted in which 200 .mu.M chloroquine, an inhibitor of lysosomal 
hydrolases (Goldstein et al., 1977), was included in the aortic strip 
assay. Two preliminary studies were done to validate this strategy. First, 
cultured human fibroblasts were incubated in the presence of 50 .mu.M 
chloroquine and found that cellular hydrolysis of [.sup.3 H]SM-LDL was 
inhibited by 73% compared with that seen with untreated fibroblasts (data 
not shown); this finding indicates that chloroquine does indeed block the 
hydrolysis of LDL-SM by lysosomal SMase. Second, the ability of 
chloroquine to work in the aortic strip assay was tested. For this study, 
.sup.125 I-labeled epidermal growth factor (.sup.125 I-EGF) was used, 
which is known to be internalized by receptor-mediated endocytosis and 
degraded in lysosomes in many cell types (Goldstein et al., 1985). Aortic 
strips were pre-incubated at 37.degree. C. for 30 min in medium without 
(control) or with 200 .mu.M chloroquine. .sup.125 I-EGF was added, then 
incubated with these strips for 3.5 h. It was found that chloroquine 
blocked the degradation of .sup.125 I-EGF by 50%, indicating substantial 
inhibition of lysosomal hydrolases in the aortic strips. Next, an 
examination of the effects of 200 .mu.M chloroquine on LDL-SM hydrolysis 
by aortic strips was done. As before, retained material, but not 
unretained [.sup.3 H]SM-LDL, was enriched in [.sup.3 H]ceramide (FIG. 6B: 
compare the first with the second pair of bars). When chloroquine was 
added in the same manner as in the .sup.125 I-EGF experiment, no 
substantial inhibition of [.sup.3 H]ceramide enrichment of the retained 
material was seen (third pair of bars in panel B). In summary, these data 
indicate that SM delivered into the arterial wall on LDL is hydrolyzed by 
a non-lysosomal arterial SMase that is at least partially dependent on 
divalent cations. 
Discussion 
In light of the in-vitro observation that SMase treatment of LDL leads to 
lipoprotein aggregation (Xu et al., 1991), an important atherogenic event 
(Nievelstein et al., 1991; Hoff et al., 1985; Guyton et al., 1996; and 
Tabas et al., 1993), the overall goal of the present study was to seek 
evidence for an arterial-wall SMase acting on LDL retained in vivo. In 
this regard, it has been shown that the mechanism of bacterial 
SMase-induced aggregation of LDL in vitro is consistent with a similar 
process occurring with mammalian SMases in the subendothelium (FIGS. 1-3), 
and direct evidence has been provided that an arterial-wall SMase activity 
hydrolyzes the SM of retained LDL (FIGS. 4-6). Most remarkably, all the 
samples of lesional LDL that were examined were 10-50-fold enriched in 
ceramide compared with plasma LDL (FIG. 4). Although it has not been 
proven that the ceramide in these samples is enough to cause aggregation 
(compare with FIG. 3), only the aggregated forms of lesional LDL are 
enriched in ceramide (FIG. 5). Moreover, one could easily imagine that 
other factors in lesions, such as proteoglycans, collagen, or lipoprotein 
lipase (Williams et al., 1995; Camejo et al., 1993; and Goldberg, 1996), 
could lower the "threshold" for ceramide-induced LDL aggregation in vivo. 
In a similar manner, ceramide enrichment of LDL could lower the 
aggregation threshold for other possible inducers of aggregation, such as 
oxidation (Hoff et al., 1989) or other lipases (Suits et al., 1989). It is 
also possible that the most ceramide-rich and aggregated lipoproteins 
would have already been rapidly ingested and degraded by lesional 
macrophages and thus would not have been present at the time of the 
extractions. 
The mechanism and regulation of ceramide-induced aggregation represents an 
important area requiring further investigation. The mechanism of 
aggregation may be related to the membrane-disruptive and possibly 
"fusogenic" properties of ceramide (van Meer, 1993; Skiba et al., 1996) or 
to hydrogen bonding between ceramide and surface phospholipids of 
neighboring particles (Xu et al., 1991). Clearly, the mechanism does not 
require apolipoproteins, since SMase induces aggregation of protein-free 
emulsions (FIG. 2), though the initial retention to arterial proteoglycans 
(Camejo et al., 1993) or collagen (Bowness et al., 1989) or the 
interaction with lipoprotein lipase (Goldberg, 1996) in vivo presumably 
does require apolipoprotein B. Moreover, other apolipoproteins, such as 
apolipoprotein AI, may prevent lipoprotein aggregation (Khoo et al., 
1990); in fact, it was found that SMase-induced aggregation of LDL is 
completely blocked by free apo AI or apo AI-containing HDL.sub.3 when 
added to LDL at a apo AI:apo B-100 molar ratio of 0.5.sup.3. In this 
regard, the potent anti-atherogenic effect of apo AI and HDL.sub.3 in 
humans (Tall, 1990) and in apo AI-transgenic mice (Rubin et al., 1991) may 
be partially mediated by inhibition of subendothelial lipoprotein 
aggregation. 
The data indicate that at least a portion of the arterial-wall SMase 
activity is divalent cation-dependent (FIG. 6A). As described in the 
Results section, two known mammalian SMases have this property: a 
membrane-bound, neutral, Mg.sup.2+ -dependent SMase and a secreted 
Zn.sup.2+ -dependent enzyme. In one report examining neuronal cells, the 
Mg.sup.2+ -dependent SMase was shown to be externally oriented (Mohan Das 
et al., 1984); a similar orientation on arterial-wall cells could result 
in the hydrolysis of extracellularly retained lipoproteins. Homogenates of 
non-lesional rabbit aortic intima have a neutral, Mg.sup.2+ -dependent 
SMase activity (see Spence et al., 1979) that can act on LDL-SM in 
vitro.sup.3, but no direct evidence exists thus far that this enzyme is in 
the proper orientation to hydrolyze LDL retained in the subendothelium in 
vivo. The Zn.sup.2+ -dependent secreted SMase has been found to be 
secreted by several arterial-wall cell types, including macrophages 
(Schissel et al., 1996) and endothelial cells.sup.3. Although Zn-SMase has 
an acidic pH optimum using an SM-micelle substrate (Schissel et al., 1996; 
Spence et al., 1989), this enzyme activity in macrophage-conditioned 
medium can hydrolyze LDL-SM at neutral pH in the presence of lipoprotein 
lipase.sup.3, an arterial-wall enzyme known to be present in 
atherosclerotic lesions (Yla-Herttuala et al., 1991; Jonasson et al., 
1987). The secreted Zn-dependent SMase, acting in co-operation with 
lipoprotein lipase, and perhaps externally oriented Mg.sup.2+ -dependent 
SMase, may be important in subendothelial lipoprotein aggregation. The 
most direct test of these hypotheses will first require further molecular 
characterization of these and possibly other arterial-wall SMases, 
followed by inhibition of these activities in animal models by genetic or 
pharmacologic means. 
Finally, the data herein may have implications beyond the realm of 
subendothelial lipoprotein aggregation. Exposure of cells to SMase has 
been shown to induce inflammatory or apoptotic changes, both of which are 
known to occur in atherosclerosis (Libby et al, 1991; Geng et al., 1995; 
Han et al. 1995), via ceramide-induced signalling pathways (Kolesnick, 
1991; Hannun et al., 1989). Furthermore, treatment of macrophages with 
SMase increases the potency of atherogenic lipoproteins to stimulate the 
cholesterol esterification pathway (Okwu et al., 1994). Thus, the same 
SMase activity that was shown to hydrolyze extracellular, retained LDL-SM 
might also hydrolyze cellular SM, which is known to be concentrated in the 
external leaflet of the plasma membrane (Merrill et al., 1990). 
Furthermore, it is possible that when cells, particularly macrophages, 
ingest large amounts of ceramide-rich lesional LDL, some of the ceramide 
escapes lysosomal hydrolysis and enters the signalling pathway. As with 
the aggregation hypothesis, these ideas will be best tested by using 
experimental systems with altered arterial-wall SMase activity. 
Footnotes 
1. Abbreviations used: apo, apolipoprotein; DMEM, Dulbecco's modified 
Eagle's medium; BSA, bovine serum albumin; EGF, epidermal growth factor; 
LDL, low-density lipoprotein; PBS, phosphate-buffered saline; PC, 
phosphatidylcholine; SM, sphingomyelin; SMase, sphingomyelinase; TLC, 
thin-layer chromatography. 
2. The term "aggregation" is used to designate self-associated LDL due to 
either adherence of individual particles or fusion. 
EXAMPLE 2 
The Sphingomyelin of Atherogenic Lipoproteins is Hydrolyzed by Mammalian 
Secreted Sphingomyelinase A Potential Role for Secreted Sphingomyelinase 
in the Subendothelial Retention and Aggregation of Atherogenic 
Lipoproteins 
The subendothelial aggregation and retention of low-density lipoprotein 
(LDL) are key events in atherogenesis, but the mechanisms in vivo are not 
known. Treatment of LDL with bacterial sphingomyelinase (SMase) in vitro 
leads to the formation of lesion-like LDL aggregates that become retained 
on extracellular matrix and stimulate macrophage foam cell formation. In 
addition, LDL retained in human atherosclerotic lesions shows evidence of 
hydrolysis by an arterial-wall SMase in vivo, and several arterial-wall 
cell types secrete a zinc-activated SMase (S-SMase). S-SMase, however, has 
a sharp acid pH optimum using a standard in vitro SM-micelle assay and its 
ability to act on lipoprotein-SM, particularly at neutral pH, is unknown. 
It is shown herein that S-SMase can hydrolyze and aggregate native plasma 
LDL at pH 5.5, but not at pH 7.4. LDL modified by oxidation, treatment 
with phospholipase A.sub.2, or enrichment with apolipoprotein CIII is 
hydrolyzed readily by S-SMase at pH 7.4. In addition, lipoproteins from 
the plasma of apolipoprotein E knockout mice, which develop extensive 
atherosclerosis, are highly susceptible to hydrolysis and aggregation by 
S-SMase at pH 7.4; a high SM:PC ratio in these lipoproteins appears to be 
an important factor in their susceptibility to S-SMase. Most importantly, 
LDL extracted from human atherosclerotic lesions, which is enriched 
three-fold in sphingomyelin compared to plasma LDL, is hydrolyzed by 
S-SMase at pH 7.4 ten-fold more than same-donor plasma LDL, suggesting 
that LDL is modified in the arterial wall to increase its susceptibility 
to S-SMase. In summary, S-SMase can hydrolyze and aggregate LDL in vitro, 
making it a leading candidate for the arterial-wall SMase that hydrolyzes 
LDL-SM and causes subendothelial LDL aggregation. 
A critical event in early atherogenesis is the subendothelial retention of 
atherogenic lipoproteins, including LDL.sup.1 (Schwenke et al., 1989; 
Nievelstein et al., 1991), lipoprotein(a) [Lp(a)] (Kreuzer et al., 1994), 
and triglyceride-rich lipoproteins (Rapp et al., 1994). Retained 
lipoproteins likely trigger a series of biological responses, such as 
endothelial changes and recruitment of macrophages to the arterial wall, 
that are central to the initiation and progression of atherosclerosis 
(Williams et al., 1995). 
Subendothelial lipoproteins are exposed to several modifying enzymes, 
including lipases (Yla-Herttuala et al., 1991; Schissel et al., 1996; 
Hurt-Camejo et al., 1997), oxidizing enzymes (Leeuwenburgh et al., 1997), 
and proteases (Kaartinen et al., 1994). The actions of these and other 
unknown factors lead to the several prominent lipoprotein modifications 
observed in vivo, including oxidation (Yla-Hettuala et al., 1989), 
enrichment with the phospholipid sphingomyelin (SM) (Yla-Hettuala et al., 
1989; Daugherty et al., 1988; Hoff, 1983), and self-aggregation 
(Neivelstein et al., 1991; Hoff et al., 1985, Guyton et al., 1996). 
Lipoprotein aggregation is likely to be important in atherogenesis for at 
least two reasons. First, processes that promote lipoprotein aggregation 
before or during retention dramatically increase the amount of lipoprotein 
retained (Tabas et al., 1993). Second, aggregated LDL, but not 
unaggregated LDL, is a potent inducer of macrophage foam cell formation 
(Hoff et al, 1990; Khoo et al., 1988; Suits et al., 1989; and Xu et al., 
1991). 
While the mechanism of lipoprotein aggregation in lesions has not yet been 
elucidated, several studies as described herein suggest that the enzyme 
sphingomyelinase (SMase) may be an important mediator of lipoprotein 
aggregation in vivo. First, LDL treated with bacterial SMase forms 
lesion-like self-aggregates (Xu et al., 1991) due to enrichment in 
ceramide (Schissel et al., 1996), the major product of SM hydrolysis; 
furthermore, these aggregates potently induce macrophage foam cell 
formation in vitro (Tabas et al., 1993; Xu et al., 1991). Second, 
aggregated LDL from human atherosclerotic lesions shows evidence of 
hydrolysis by an extracellular SMase, and LDL retained in rabbit aortic 
strips ex vivo is hydrolyzed by an extracellular, cation-dependent SMase 
(Schissel et al., 1996). Third, and most important, several cell types 
present in atherosclerotic lesions, namely endothelial cells and 
macrophages (Schissel et al., 1996), secrete a Zn.sup.2+ -activated SMase 
(S-SMase). 
The cellular origins, secretion, and cation dependency make S-SMase a 
leading candidate for the arterial-wall SMase that acts on retained 
lipoproteins. Nonetheless, two major issues regarding the relevance of 
S-SMase to atherogenesis needed to be addressed. First, mammalian SMases 
are much more selective than bacterial SMases in terms of the milieu in 
which the SM is presented to the enzyme (Spence, 1993). Second, studies on 
the molecular origin of S-SMase have revealed that it is a product of the 
same gene, the acid SMase (ASM) gene, that gives rise to lysosomal SMase 
(L-SMase) (Schissel et al., 1996). Therefore, S-SMase shares with L-SMase 
a sharp acid pH optimum when assayed under standard in-vitro conditions 
using detergent-solubilized SM micelles as a substrate (Schissel et al., 
1996; Spence et al., 1989). While it is possible that acidic enzymes are 
active in advanced atherosclerotic lesions, where local pockets of acidity 
may occur (Menkin, 1934; Smith, 1979; Maroudas et al., 1988, Tapper et 
al., 1992; Silver et al., 1988), a role for such enzymes in pre-lesional 
or early lesional events would require activity at neutral pH with 
physiologic substrates. 
In this context, the goal of the current study was to test whether S-SMase 
can hydrolyze LDL-SM, particularly at neutral pH. Herein, it is shown that 
S-SMase can hydrolyze and aggregate native LDL at acid but not neutral pH. 
LDL modified by several means that have been shown to occur or might occur 
during atherogenesis, however, is an excellent substrate for S-SMase at pH 
7.4. Most importantly, LDL extracted from human atherosclerotic lesions is 
efficiently hydrolyzed by S-SMase at neutral pH, suggesting that LDL is 
modified in the arterial wall to increase its susceptibility to S-SMase. 
The results support a role for S-SMase in the subendothelial hydrolysis of 
LDL-SM, perhaps leading to lipoprotein aggregation and lesion initiation 
and progression. 
Methods 
Materials. sn-1,2-diacylglycerol kinase (from Escherichia coli) was 
purchased from CALBIOCHEM.RTM. (San Diego, Calif.). Cardiolipin and 
1,2-dioleoyl glycerol were purchased from Avanti Polar Lipids Alabaster, 
Ala.). [9,10-.sup.3 H]palmitic acid and [.gamma.-.sup.32 P]ATP, and were 
obtained from DUPONT-NEW ENGLAND NUCLEAR.RTM. (Boston, Mass.). Tissue 
culture media and reagents were purchased from LIFE TECHNOLOGIES.RTM. 
(Baltimore, Md.) and fetal bovine serum was from GEMINI BIOPRODUCTS.RTM. 
(Calabasas, Calif.). Human native apo CIII was prepared as described 
(Clavey et al., 1995). Human recombinant nonpancreatic soluble PLA2 
(sPLA.sub.2) was purified as previously described (Sartipy et al., 1996). 
Partially purified phospholipid transfer protein was prepared as 
previously described (Tollefson et al., 1988). Soybean lipoxygenase and 
all other reagents were from SIGMA.RTM. (St. Louis, Mo.). 
Mice. LDL receptor deficient (LDLr0) (Ishibashi et al., 1993) and 
apolipoprotein E deficient (E0) mice (Plump et al., 1992; Zhang et al., 
1992) were purchased from Jackson Laboratories and crossed into the 
C57BL/6J background. LDL receptor-deficient mice expressing a human apo 
CIII transgene (LDLr0/CIII) were derived as previously described 
(Masucci-Magoulas et al., 1997). 
Lipoprotein Isolation and Modification. Human and murine LDL (density, 
1.020-1.063 g/mL) were isolated from fresh plasma by preparative 
ultracentrifugation as previously described (Havel et al., 1955). LDL (5 
mg/mL) was oxidized by dialysis against 150 mM NaCl, 6 .mu.M FeSO4, 0.04% 
azide for 36 h at room temperature followed by addition of EDTA (1 mM) and 
BHT (150 .mu.M) and then dialysis against 150 mM NaCl, 0.3 mM EDTA (Watson 
et al., 1995). Alternatively, LDL (1 mg) was incubated with 275 U soybean 
lipoxygenase/mL and 60 .mu.g linoleic acid/mL in 50 mM Tris-HCl pH 7.4, 
0.04% azide for 24 h at 37.degree. C. (Dzeletovic et al., 1995); LDL was 
re-isolated using a G-200 gel filtration column and then concentrated 
using a Centricon 30 (molecular weight cut-off=30,000) ultrafiltration 
device. LDL was treated with sPLA.sub.2 as previously described (Sartipy 
et al., 1996). Briefly, LDL (5 mg/mL) was incubated with 15 .mu.g pure 
human recombinant sPLA.sub.2 /mL in 0.12 M Tris-HCl pH 8.0, 12 mM 
CaCl.sub.2, 0.1 mM EDTA, 10 .mu.M BHT for 14 h at 37.degree. C.; LDL (50 
.mu.g protein) was then treated directly with S-SMase as described below. 
Acetyl-5 LDL was prepared by acetylation of LDL with acetic anhydride as 
described previously (Goldstein et al., 1979). 
Isolation of LDL from human lesions. LDL was extracted from abdominal 
aortic aneurysm plaque material as previously described (Rapp et al., 
1994). Briefly, aortic plaque was removed from individuals as part of the 
standard reconstructive surgery for abdominal aortic aneurysms at the San 
Francisco Veterans Affairs Medical Center. Plaque material, which ranged 
in weight from 2-12 grams, was obtained in the operating room and 
immediately placed into ice-cold 7-mM citrate buffer, pH 7.4, containing 
15 mM NaCl, 3 mM EDTA, 0.5 mM butylhydroxytoluene, 1 mM 
phenylmethylsulfonylfluoride, 1.5 mg aprotinin/mL, 2 mM benzamidine, and 
0.08 mg gentamycin sulfate/mL. Blood and adherent thrombus were removed by 
blotting with absorbent gauze, scrubbing with a small brush, and sharp 
dissection as necessary. Loosely retained lipoproteins were extracted by 
mincing the plaque into 0.5-1.0 mm.sup.2 pieces and incubating them 
overnight on a Labquake shaker at 4.degree. C. in a non-denaturing buffer 
(0.1 M citrate, pH 7.4, with 1 mg EDTA/mL, 0.3 mg benzamidine/mL, 0.08 mg 
gentamicin sulfate/mL, 10 .mu.g aprotinin/mL, 10 .mu.g Trolox [an 
anti-oxidant]/mL, and 20 .mu.g phenylmethylsulfonyl fluoride/mL). The 
extracted material was cleared of particulate matter by centrifuging at 
800.times.g for 10 min, and 1.019&lt;d&lt;1.063-g/mL lipoproteins were isolated 
by sequential sodium bromide density ultracentrifugation (Rapp et al., 
1994; Havel et al., 1955). 
Synthesis of [.sup.3 H]SM. [N-palmitoyl-9,10-.sup.3 H]SM was synthesized as 
previously described (Schissel et al., 1996; Sripada et al., 1987; Ahmad 
et al., 1986). [9,10-.sup.3 H]palmitic acid (25 mCi, 450 nmol) was stirred 
for 12 h at room temperature with an equimolar equivalent of 
(N)-hydroxysuccinimide and with 3-molar equivalents of 
1,3-dicyclohexylcarbodiimide in (N,N)-dimethylformamide. The reaction was 
run under dry argon in the dark. Sphingosylphosphorylcholine (300 nmol) 
and (N,N)-diisopropylethylamine (10 .mu.l) were then added and the 
reaction was stirred another 12 h at room temperature. The reaction was 
stopped by evaporating the (N,N)-dimethylformamide under a stream of 
N.sub.2. [N-palmitoyl-9,10-.sup.3 H]SM was purified by preparative 
thin-layer chromatography of the reaction products three consecutive times 
in chloroform:methanol (95:5) and then twice in chloroform:methanol:acetic 
acid:water (50:25:8:4). Greater than 95% of the [N-palmitoyl-9,10-.sup.3 
H]SM was converted to [N-palmitoyl-9,10-.sup.3 H]ceramide after treatment 
with 10 mU SMase/mL (Bacillus cereus) for 1 h at 37.degree. C., as assayed 
by TLC, indicating a pure, functional substrate. 
[.sup.3 H]SM-labeling of LDL. Plasma LDL was labeled with 
[N-palmitoyl-9,10-.sup.3 H]SM as previously described (Schissel et al., 
1996). Briefly, .about.3.5 mCi (63 nmol) [N-palmitoyl-9,10-.sup.3 H]SM and 
13 nmol phosphatidylcholine (PC) were mixed in chloroform, and the solvent 
was removed first under a stream of nitrogen and then by lyophilization. 
The dried lipids were resuspended in 1 mL of 150 mM NaCl, 1 mM EDTA, 10 mM 
Tris-HCl, pH 7.5 and, to prepare [.sup.3 H]SM/PC liposomes, sonicated for 
three 50-sec pulses at 4.degree. C. using a tapered microtip on a 
BRANSON.RTM. 450 sonicator (setting #3). The liposomes were then incubated 
with 30 mg (by protein mass) of LDL, 50 .mu.g of partially purified 
phospholipid transfer protein, 100 U penicillin, and 100 .mu.g 
streptomycin for 18 h at 37.degree. C. under argon. LDL was then separated 
from the liposomes after phospholipid transfer by centrifuging the mixture 
at density=1.006 g/mL for 8 h at 35,000 rpm in a BECKMAN.RTM. 50.3 rotor; 
the supernate containing the liposomes was removed, and the LDL band at 
the bottom of the tube was harvested. The LDL solution was mixed with 
buffer containing 150 mM NaCl, 0.3 mM EDTA pH 7.4 and centrifuged as 
before. This wash procedure was performed a total of four times, resulting 
in the removal of 95% of the unreacted [.sup.3 H]SM/PC liposomes. All 
lipoproteins were stored under argon at 4.degree. C. and were used within 
2 weeks of preparation. 
[.sup.3 H]SM-emulsions. [.sup.3 H]SM-emulsions with a lipid composition 
similar to human LDL were prepared as follows: 5.4 mg cholesteryl oleate, 
0.48 mg triolein, 1.08 mg free cholesterol, 2.04 mg phosphatidylcholine, 
0.96 mg sphingomyelin, and 50 .mu.Ci [N-palmitoyl-9,10-.sup.3 
H]sphingomyelin were all added in chloroform to a sonication vial and the 
solvent completely evaporated by exposure to a stream of nitrogen, 
followed by the high vacuum of a lyophilizer. The dried lipids were 
resuspended in 3 mL of buffer containing 150 mM NaCl, 0.3 mM EDTA pH 7.4 
and sonicated under a stream of argon at 40.degree. C. until translucent 
(approximately 90 min). The sonicated material was then centrifuged twice 
at 15,000.times.g to pellet any titanium shed from the sonication probe. 
SM-rich emulsions were prepared exactly as above except that 1.2 mg of 
phosphatidylcholine and 1.8 mg of SM were used, and sonication time was 
increased to 120 min. [.sup.3 H]SM-emulsions were enriched with apo CIII 
based on the method of Ahmad et al. (Ahmad et al., 1986). Briefly, [.sup.3 
H]SM-emulsions (0.4 mL; 182 nmol SM) were incubated with 200 .mu.g apo 
CIII (22.5 nmol) for 2 h at 40.degree. C. A portion of the emulsions were 
re-isolated from free apo CIII using ultrafiltration (Clavey et al., 1995) 
as follows: the crude emulsion-apo CIII mixture was diluted to 2 mL with 
buffer containing 150 mM NaCl, 0.3 mM EDTA pH 7.4 and ultrafiltered and 
concentrated to 0.2 mL using a CENTRICON.RTM. 30 (molecular weight 
cut-off=30,000); the concentrated emulsions were then diluted to 2 mL and 
the process was repeated 5 times. [.sup.3 H]SM-emulsions run through the 
same enrichment and re-isolation protocols in the absence of apo CIII 
served as the control for the experiments in FIG. 11A. 
Ceramide assay. Ceramide was measured from LDL lipid extracts using the 
method described by Schneider and Kennedy (Schneider et al., 1976) and 
adapted by Preiss et al. (Preiss et al., 1986). In this method, 
diacylglycerol (DAG) kinase phosphorylates ceramide and DAG using 
[.gamma.-.sup.32 P]ATP. For ceramide measurement, the lipids were first 
incubated with 0.1 N KOH in methanol for 1 h at 37.degree. C., which 
hydrolyzes DAG, but not ceramide. The extracted lipids were dried under 
nitrogen and then solubilized in 5 mM cardiolipin, 7.5% 
octyl-.beta.-glucopyranoside, and 1 mM diethylenetriaminepentaacetic acid 
by bath sonication. This solution was then added to reaction buffer (50 mM 
imidazole-HCl, pH 6.6, 50 mM NaCl, 12.5 mM MgCl.sub.2, 1 mM EGTA) 
containing sn-1,2-DAG kinase (0.7 units/mL). The reaction was initiated by 
the addition of [.gamma.-.sup.32 P]ATP (final concentration=10 mM). After 
incubation at room temperature for 60 min, the reaction was stopped by 
lipid extraction with chloroform:methanol:HCl (100:100:1, v/v/v) and 10 mM 
EDTA. Ceramide-1-[.sup.32 P] phosphate in the organic phase was separated 
by TLC using chloroform:methanol:acetic acid (65:15:5, v/v/v) and 
visualized with autoradiography and identified by comparing with 
standards. The spots corresponding to ceramide-1-[.sup.32 P] phosphate 
were scraped and counted, and the mass calculated by comparison with a 
ceramide standard curve. 
Sphingomyelin (SM) and phosphatidylcholine (PC) assays. Lipid extracts 
(Bligh et al., 1959) of lipoproteins were chromatographed by TLC using 
chloroform:methanol:acetic acid:H.sub.2 O (50:25:8:4, v/v/v/v). Individual 
phospholipid subclasses were visualized by iodine vapor staining, and the 
SM and PC spots were identified by comparison with standards. The spots 
were scraped, extracted twice with chloroform:methanol (2:1), and assayed 
for phosphate content by the method of Bartlett (Bartlett, 1959). 
LDL Oxidation Assays. LDL lipid peroxides were measured using the method of 
El-Saadani et al. (El-Saadani et al., 1989). LDL (50-150 .mu.g protein), 
in a volume of no more than 100 .mu.L, was added to 1 mL color reagent 
[0.2 M KH.sub.2 PO.sub.4, 0.12 M KI, 0.15 mM NaN.sub.3, 2 g 
Triton-X-100/L, 0.1 g benzalkonium chloride/L, 10 .mu.M ammonium 
molybdate, 20 .mu.M BHT, 25 .mu.M EDTA, pH 6.2] and incubated in the dark 
for 30 min at room temperature; light absorbance at 365 nm was then 
measured and lipid peroxides were quantified by comparison with a H.sub.2 
O.sub.2 standard curve. Thiobarbituric acid-reactive substances (TBARS) 
were measured using a standard method (Puhl et al., 1994). Briefly, LDL 
(100 .mu.L, 100-200 .mu.g protein), was mixed with 1 mL 20% 
trichloroacetic acid and incubated on ice for 30 min. Following 
precipitation, 1 mL 1% thiobarbituric acid was added and the samples 
heated at 95.degree. C. for 45 min. After cooling, the samples were 
centrifuged at 1000.times.g for 20 min. and the light absorbance at 532 nm 
was measured. TBARS were quantified by comparison with a malonaldehyde 
standard curve prepared using tetramethoxypropane. LDL electrophoretic 
mobility was assayed by loading 30 .mu.g of LDL protein onto a 
polyacrylamide 0.75%-27% gradient gel (Lipogel; Zaxis, Hudson Ohio) and 
electrophoresing in 0.1 M Tris-base, 0.1 M boric acid, 20 mM EDTA (upper 
chamber=pH 8.7; lower chamber=pH 8.3) for 12 h at 100 V. The gel was then 
stained with sudan black and the bands visualized by counter-staining with 
methanol:acetic acid:water (10:7:83 v/v/v). 
S-SMase. The source of S-SMase was serum-free conditioned medium from DG44 
CHO cells stably transfected with the human acid SMase cDNA (Schissel et 
al., 1996; Schuchman et al., 1991). It was demonstrated that S-SMase is 
the only detectable SMase secreted into the culture medium (Schissel et 
al., 1996). Cells were plated in 100-mm dishes and cultured for 48 h in 
DMEM/FBS/PSG. The cells were then changed to low protein serum-free media 
for 12 h, washed 3 times with PBS, and finally incubated for 18 h in fresh 
serum-free media (6 mL per 100-mm dish). The "18-h conditioned medium" was 
then collected, centrifuged at 1000.times.g to pellet any cells and, 
except where indicated, ZnCl.sub.2 (final concentration=100 .mu.M) was 
added to fully activate and stabilize S-SMase; this S-SMase-containing 
conditioned media was then used fresh to treat LDL and lipid emulsions. 
S-SMase Treatment of LDL and Emulsions. The standard incubation mixture 
consisted of up to 50 .mu.l of sample (LDL or emulsions), 25 .mu.l of 
S-SMase-containing conditioned media (see above) and a volume of assay 
buffer (0.1 M Tris-HCl, pH 7.4, 0.04% azide; or, where indicated, 0.1 M 
sodium acetate, pH 5.5, 0.04% azide) to bring the final volume to 200 
.mu.l. The reactions were incubated at 37.degree. C. for no longer than 16 
h and then extracted by the method of Bligh and Dyer (Bligh et al., 1959). 
For the samples containing [N-palmitoyl-9-10.sup.3 H]SM-LDL or 
[N-palmitoyl-9-10-.sup.3 H]SM-emulsions, the lower, organic phase was 
harvested, evaporated under N.sub.2, and fractionated by TLC using 
chloroform:methanol (95:5). The ceramide spots were scraped and directly 
counted to quantify [.sup.3 H]ceramide. In all other samples ceramide was 
determined 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. 
Results 
Native LDL can be hydrolyzed and aggregated by S-SMase. Previous LDL 
aggregation studies with SMase utilized bacterial enzymes (Tabas et al., 
1993; Xu et al., 1991). S-SMase, like moss mammalian SMases, displays 
significant substrate specificity in terms of the milieu in which SM is 
presented to the enzyme (Spence, 1993); the most commonly used in-vitro 
assays for mammalian SMases use detergent-solubilized SM micelles as the 
substrate (Spence, 1993). The first question was whether S-SMase can act 
on intact LDL and, if so, whether it causes LDL aggregation. Although 
treatment of native LDL with S-SMase at pH 7.4 caused very little SM 
hydrolysis, S-SMase hydrolyzed significant amounts (nearly 90%) of LDL-SM 
at pH 5.5 (FIG. 7A), leading to LDL aggregation (FIG. 7B). These data are 
important for two reasons. First, they demonstrate that S-SMase can 
hydrolyze the SM in intact LDL without the need for detergent. Second, 
these data formally demonstrate that a mammalian SMase can cause LDL 
aggregation. 
While S-SMase-induced LDL aggregation at acid pH may be important in more 
advanced atherosclerotic lesions, early subendothelial lipoprotein 
aggregation likely occurs at a more neutral, physiologic pH. If S-SMase 
mediates early aggregation, therefore, then lipoprotein-SM should be a 
substrate for S-SMase at neutral pH. Interestingly, studies by Callahan et 
al. (Callahan et al., 1983) demonstrate that only the affinity of L-SMase 
for SM-micelles (i.e. K.sub.m) is highly sensitive to changes in pH, 
whereas the maximal velocity (V.sub.max) for SM hydrolysis is 
pH-independent. Since the kinetic properties of S-SMase and L-SMase should 
be similar (Schissel et al., 1996), it was reasoned that LDL-SM would be 
hydrolyzed at neutral pH if it could access the active site of S-SMase. 
Moreover, several physiologically relevant modifications of LDL, including 
oxidation, hydrolysis with phospholipase A.sub.2, and sphingomyelin 
enrichment, alter the structure of the lipoprotein surface, perhaps 
allowing S-SMase to bind LDL-SM at neutral pH. These ideas prompted us to 
test whether modified forms of LDL are better substrates than native LDL 
for S-SMase at neutral pH. 
Oxidized LDL and sPLA-treated LDL are hydrolyzed by S-SMase at neutral pH. 
S-SMase hydrolysis of [.sup.3 H]SM-labeled LDL oxidized by two independent 
methods, FeSO.sub.4 and lipoxygenase was analyzed. Whereas oxidation alone 
caused no artifactual LDL-[.sup.3 H]SM hydrolysis (FIG. 8A, open bars), 
oxidized [.sup.3 H]SM-LDL was hydrolyzed 5-6 fold more than native [.sup.3 
H]SM-LDL by S-SMase at pH 7.4 (FIG. 8A, solid bars). Similar results were 
obtained when LDL-SM hydrolysis was assayed by measuring generation of 
ceramide mass. Note that FeSO.sub.4 and lipoxygenase caused similar 
degrees of LDL oxidation as indicated by the levels of thiobarbituric 
acid-reactive substances [TBARS] (FIG. 8B) and lipid peroxides [LPO] (FIG. 
8C) in the LDL. To more precisely define the relationship between LDL 
oxidation and its susceptibility to S-SMase hydrolysis, [.sup.3 H]SM-LDL 
was incubated with FeSO.sub.4 for increasing amounts of time and compared 
its extent of oxidation to its hydrolysis by S-SMase at pH 7.4. [.sup.3 
H]SM-LDL oxidation, assayed by measuring lipid peroxides (FIG. 9, open 
circles) increased almost linearly with increasing time of incubation with 
FeSO.sub.4 ; electrophoretic mobility of LDL as assessed by native 
polyacrylamide gradient gel electrophoresis also increased with time of 
oxidation. Most importantly, the susceptibility of [.sup.3 H]SM-LDL to 
S-SMase hydrolysis at neutral pH was, after an initial threshold level of 
oxidation, closely correlated with the extent of [.sup.3 H]SM-LDL 
oxidation (FIG. 9, closed squares). 
Oxidation likely increases S-SMase hydrolysis of LDL-SM through one of two 
general mechanisms: either oxidized SM itself is a better substrate for 
S-SMase than non-oxidized SM, or another consequence of LDL oxidation 
enhances the interaction between LDL-SM and S-SMase. To address the first 
possible mechanism, [.sup.3 H]SM-vesicles, which contained no other 
lipids, were incubated in the absence or presence of FeSO.sub.4 under the 
exact conditions used to oxidize [.sup.3 H]SM-LDL for the experiment in 
FIG. 7, followed by incubation with S-SMase at pH 7.4. [.sup.3 
H]SM-vesicles exposed to FeSO.sub.4 were hydrolyzed less than two-fold 
greater than untreated control vesicles, suggesting that another 
consequence of LDL oxidation is important for stimulating S-SMase 
hydrolysis of LDL. 
The biological actions of oxidized LDL (e.g. scavenger receptor binding, 
cytotoxicity, and monocyte chemotaxis) are largely attributed to its 
overall negative charge and enrichment in lysophosphatidylcholine (Krieger 
et al., 1994; Quinn et al., 1988; Weltzien, 1979; Niewoehner et al., 
1989). Thus, these modifications were studied individually to define the 
critical component(s) in oxidized LDL that stimulates S-SMase. To examine 
negative charge, the hydrolysis of acetylated-LDL, a highly negatively 
charged form of LDL (Goldstein et al., 1979), was compared to the 
hydrolysis of native LDL by S-SMase at pH 7.4 and no significant 
difference in LDL-sphingomyelin hydrolysis were found. Thus, adding 
negative charges to LDL, at least via acetylation, does not increase its 
hydrolysis by S-SMase. Lysophosphatidylcholine enrichment of LDL occurs 
during oxidation due to activation of an apparently latent phospholipase 
A.sub.2 activity (Parthasarathy et al., 1985). Importantly, Camejo and 
colleagues (Hurt-Cameho et al., 1997; Sartipy et al., 1996) have shown 
that the nonpancreatic secretory phospholipase A.sub.2 type II 
(sPLA.sub.2) is abundant in both normal and lesional human arteries and 
can act on LDL in vitro, suggesting that LDL may be enriched with 
lysophosphatidylcholine by sPLA.sub.2 in vivo. Moreover, Lusis and 
colleagues (Ivandic et al., 1996) have reported that transgenic mice 
overexpressing sPLA.sub.2 have more extensive atherosclerotic lesions than 
wild-type mice. With this background, the next question was whether LDL 
treated with sPLA.sub.2 was more susceptible than native LDL to S-SMase 
hydrolysis at neutral pH. Whereas sPLA.sub.2 treatment alone caused no 
[.sup.3 H]SM-LDL hydrolysis, S-SMase hydrolysis of [.sup.3 H]SM-LDL at pH 
7.4 was markedly enhanced by treating the LDL with sPLA.sub.2 (FIG. 10). 
Furthermore, exogenously added lysophosphatidylcholine stimulated S-SMase 
hydrolysis of [.sup.3 H]SM-LDL 2-3 fold at pH 7.4. In summary, LDL 
oxidation markedly increases S-SMase hydrolysis of LDL-SM at neutral pH. 
While it is possible that several oxidation products are acting together, 
lysophosphatidylcholine appears to be particularly effective in 
stimulating S-SMase hydrolysis of LDL. 
Apolipoprotein CIII stimulates S-SMase hydrolysis of lipoprotein-SM. 
Apolipoprotein CIII (apo CIII) is associated with severe 
hypertriglyceridemia (Ito et al., 1990; Dammerman et al., 1993) and, 
recently, has been identified as a possible contributing factor in 
familial combined hyperlipidemia (Masucci-Magoulas et al., 1997; 
Dallinga-Thie et al., 1997). In addition, apo CIII has been linked to 
certain forms of coronary heart disease (Rigoli et al., 1995), and its 
overexpression in mice significantly increases atherosclerosis 
(Masucci-Magoulas et al., 1997). While apo CIII is known to inhibit 
lipoprotein lipolysis (Ito et al., 1990), the atherogenic mechanisms of 
apo CIII are not known. Interestingly, Ahmad et al. (Ahmad et al., 1986) 
have shown that Apo CIII, but not other apolipoproteins, stimulates 
L-SMase hydrolysis of SM-liposomes. Given that L-SMase and S-SMase are 
such similar enzymes (Schissel et al., 1996), the next question was 
whether Apo CIII could stimulate S-SMase hydrolysis of lipoprotein-SM at 
neutral pH. Because enriching native LDL with apo CIII in vitro proved 
extremely difficult, the effect of apo CIII on S-SMase hydrolysis of 
[.sup.3 H]SM-labeled emulsions containing lipids in the same proportions 
as in LDL was examined first. For this experiment, three different types 
of emulsions were treated with S-SMase at pH 7.4: control emulsions not 
enriched with Apo CIII (control), emulsions mixed with apo CIII (+CIII, 
emulsion-bound+free CIII), and emulsions mixed with apo CIII followed by 
re-isolation from free apo CIII (+CIII, emulsion-bound only). While 
S-SMase hydrolysis of the emulsions with bound plus free CIII was two-fold 
greater than hydrolysis of the control emulsions, similar to Ahmad et al. 
(Ahmad et al., 1989), it was found that emulsions with only bound CIII 
were hydrolyzed approximately ten-fold more than control emulsions (FIG. 
11A). Thus, apo CIII stimulates S-SMase hydrolysis of LDL-like emulsions, 
particularly in the absence of free apo CIII. 
To determine whether apo CIII stimulates S-SMase hydrolysis of 
lipoprotein-SM, the hydrolysis of LDL from LDLr0 mice expressing a human 
apo CIII transgene (LDLr0/CIII) (Masucci-Magoulas et al., 1997) was 
compared with the hydrolysis of LDL from LDLr0 mice (Ishibashi et al., 
1993); LDL from the LDLr0/CIII mouse is enriched in apo CIII, whereas LDL 
from the LDLr0 mouse is not. The data in FIG. 11B clearly show that LDL 
from LDLr0/CIII mice is significantly hydrolyzed by S-SMase at pH 7.4, 
whereas LDL from LDLr0 mice is hydrolyzed very little. Importantly, the 
turbidity at A.sub.430 nm of LDLr0/CIII LDL, but not for the LDLr0 LDL, 
increased 1.5 fold following treatment with S-SMase, indicating particle 
aggregation. Lipoprotein enrichment with apo CIII, therefore, increases 
its susceptibility to hydrolysis and aggregation induced by S-SMase. 
Plasma LDL from E0 mice is efficiently hydrolyzed by S-SMase at neutral pH. 
The E0 mouse develops widely distributed and complex atherosclerotic 
lesions like those seen in humans and thus has become a widely used model 
of atherosclerosis (Plump et al., 1993; Zhang et al., 1992). A likely 
factor contributing to the extensive atherosclerosis in these mice is the 
atherogenicity of E0 lipoproteins, but the mechanisms whereby apo 
E-deficient lipoproteins lead to foam cell formation and other lesional 
events is not known. Another question was whether plasma LDL from these 
mice was susceptible to S-SMase at neutral pH; as a comparison, plasma LDL 
from LDLr0 mice was used, which mice also develop atherosclerosis, though 
to a lesser degree than E0 mice (Ishibashi et al., 1993; Plump et al., 
1992; Zhang et al., 1992). While the ceramide contents of native plasma 
LDL from these two strains of mice are equivalent (FIG. 12A, hatched 
bars), treatment with S-SMase at pH 7.4 resulted in generation of 4-5-fold 
more ceramide in LDL from E0 mice than in LDL from LDLr0 mice (FIG. 12A, 
solid bars), which resulted in aggregation of E0 LDL. E0 LDL has a high 
SM:PC molar ratio (0.36) compared with LDLr0 LDL, which has a SM:PC ratio 
of 0.19; the mechanism appears to be a combination of increased SM 
synthesis and decreased plasma SM clearance in the E0 mice. SM-enrichment 
of LDL may increase substrate availability to S-SMase and thereby promote 
ceramide generation, which is required for LDL aggregation (Schissel et 
al., 1996); note that particles enriched in both SM and ceramide aggregate 
readily (Schissel et al., 1996). Thus, one possible factor contributing to 
the increased susceptibility of E0 LDL to S-SMase is its relatively high 
SM:PC ratio. To test the importance of the membrane SM:PC ratio on S-SMase 
activity in a more controlled system, [.sup.3 H]SM-labeled emulsions were 
prepared containing lipids similar to those in human LDL with a SM:PC 
ratio of either 0.5 (control emulsions) or 1.5 (SM-rich emulsions) and 
then treated with S-SMase at pH 7.4. Although the control emulsions, 
unlike native plasma LDL, were significantly hydrolyzed by S-SMase, 
SM-enrichment resulted in a two-fold stimulation of [.sup.3 H]SM-emulsion 
hydrolysis by S-SMase (FIG. 12B). Together, these findings demonstrate 
that SM enrichment of lipoproteins may markedly enhance their hydrolysis 
by S-SMase at neutral pH. Note, however, that other factors must be 
involved. For example, human native plasma LDL has a higher SM:PC ratio 
than E0 LDL and the same ratio as the "control" emulsions in FIG. 12B, and 
yet it is a worse substrate for S-SMase at neutral pH than either of these 
two particles (see FIG. 12). 
Human atheroma-derived LDL is hydrolyzed by S-SMase at neutral pH. The data 
presented thus far clearly demonstrate that LDL modified in vitro to forms 
reported to be present in vivo is a significantly better substrate than 
native LDL for S-SMase at neutral pH. A critical issue regarding the 
physiologic relevance of these findings, however, is whether LDL modified 
in the arterial wall shows greater susceptibility to hydrolysis by S-SMase 
than native LDL. To address this important point, we compared the S-SMase 
hydrolysis of lesional LDL, extracted from two different atherectomy 
samples, to the hydrolysis of same-donor plasma LDL. First, the data in 
FIG. 13 confirm the report that lesion LDL is markedly enriched in 
ceramide compared to plasma LDL (compare first pair of bars to second pair 
of bars), indicating that LDL is hydrolyzed by a SMase in the arterial 
wall. Most importantly, however, treatment of lesional LDL with S-SMase at 
pH 7.4 generates nearly ten-fold more ceramide than treatment of plasma 
LDL (FIG. 13, compare third pair of bars to fourth pair of bars). LDL 
retained in the arterial wall, therefore, is an excellent substrate for 
S-SMase at neutral pH. 
What are the critical modifications to lesional LDL that stimulate its 
hydrolysis by S-SMase? To begin addressing this question, lesional LDL was 
examined for the presence of the LDL modifications that have been 
identified above as increasing susceptibility to hydrolysis by S-SMase at 
neutral pH in vitro. To determine whether lesional LDL shows evidence of 
oxidation, the levels of TBARS was compared and lipid peroxides in the 
lesional LDL samples from FIG. 13 to those in the same-donor plasma LDL. 
The levels of TBARS and lipid peroxides, however, in both plasma and 
lesional LDL were below the detection limit of the assay (&lt;0.5 nmol/mg 
protein and &lt;4 nmol/mg protein for TBARS and lipid peroxides, 
respectively). Although one interpretation of these findings is that the 
lesional LDL was not oxidized, other studies have suggested that these 
markers of oxidation can be lost during the isolation of LDL (Esterbauer 
et al., 1987; Steinbrecher et al., 1987) or that other markers of 
oxidation may be more relevant to in-vivo events (Leeuwenburgh et al., 
1997). Thus, pending further study, oxidation may still prove to be an 
important modification to lesional LDL that stimulates S-SMase. Similarly, 
detection of apo CIII in the two samples of lesional LDL was not 
accomplised, but it is possible that less dense lesional lipoproteins, or 
lipoproteins isolated from other individuals or from earlier lesions might 
have shown evidence of apo CIII. 
The next question was whether the ratio of SM to PC was different for 
lesional and plasma LDL. Note that other investigators have shown 
previously that LDL extracted from both animal and human atherosclerotic 
lesions is enriched in sphingomyelin compared with plasma LDL 
(Yla-Herttuala et al., 1989; Daugherty et al., 1988; Hoff, 1983). The data 
in Table 1 show the SM and PC contents of the lesional and same-donor 
plasma LDL samples from FIG. 13. Whereas the PC level in lesional LDL was 
modestly reduced compared with plasma LDL, suggesting limited 
arterial-wall PC hydrolysis, the SM content of lesional LDL was nearly 
three-fold higher than the level of SM in plasma LDL. As a result, the 
SM:PC ratio for lesional LDL was 3-4-fold higher than the ratio for plasma 
LDL (Table 1). Thus, lesional LDL is highly enriched in SM, a modification 
that stimulates S-SMase hydrolysis of LDL at neutral pH in vitro (see FIG. 
12). 
Discussion 
LDL hydrolysis by bacterial SMase causes LDL self-aggregation, an important 
process in atherogenesis (see Introduction), and that aggregated LDL 
retained in vivo shows evidence of hydrolysis by a SMase in the 
subendothelium (Schissel et al., 1996). Although the mammalian SMase 
responsible for subendothelial lipoprotein-SM hydrolysis has yet to be 
identified, several cell types, including macrophages, secrete a 
zinc-activated SMase (S-SMase) (Schissel et al., 1996). Cultured human 
coronary artery endothelial cells secrete abundant amounts of S-SMase 
basolaterally, making the endothelium a potential source of the enzyme in 
the pre-lesional artery. Furthermore, preliminary immunohistochemistry 
studies indicate that S-SMase is present in both the pre-lesional artery 
and in atherosclerotic lesions. Thus, S-SMase is a leading candidate for 
the arterial-wall SMase that hydrolyzes retained lipoproteins. S-SMase, 
however, has an acid pH optimum using a standard in-vitro, detergent-based 
SM-micelle assay (Schissel et al., 1996; Spence et al., 1989). Thus, the 
goal of this study was to determine whether LDL-SM could be hydrolyzed by 
S-SMase under more physiologic conditions. S-SMase can hydrolyze and 
aggregate native plasma LDL, but only at acid pH (FIG. 7). Although 
S-SMase-induced LDL aggregation under these conditions may be relevant in 
more advanced lesions, where an acidic environment may exist in the 
vicinity of lesion macrophages (Tapper et al., 1992; Silver et al., 1988) 
or as a result of hypoxia-induced metabolic acidosis (Tsukamoto et al., 
1996), early lipoprotein aggregation likely occurs at a more neutral, 
physiologic pH. Remarkably, several modified forms of LDL, known to be 
atherogenic and occur in vivo, are much better substrates for S-SMase at 
neutral pH than native LDL. 
Oxidized LDL, for example, is significantly hydrolyzed by S-SMase at 
neutral pH (FIGS. 2 and 3). Although this hydrolysis was not sufficient to 
cause visible aggregation, the level of ceramide generated (.about.0.5 
nmol ceramide/.mu.g protein) was very close to the threshold level 
(.about.0.8 nmol ceramide/.mu.g protein) required for LDL aggregation in 
vitro (Schissel et al., 1996). Furthermore, it is possible that other 
arterial-wall factors, such as proteoglycans, collagen, and lipoprotein 
lipase (5, Camejo et al., 1993; Goldber et al., 1996), lower the threshold 
for ceramide-mediated LDL aggregation in vivo. Interestingly, oxidized LDL 
treated with S-SMase at neutral pH contained similar levels of ceramide as 
aggregated lesional LDL (Schissel et al., 1996) (also, compare FIG. 8A 
with FIG. 13), consistent with the idea that the threshold for 
ceramide-mediated aggregation may be lowered in vivo. Although several 
consequences of LDL oxidation may contribute to stimulation of S-SMase, 
one of these, namely, hydrolysis of LDL by PLA.sub.2 (Parthasarathy et 
al., 1985; Ivandic et al., 1996), may be particularly important. 
Specifically, LDL hydrolyzed by sPLA.sub.2, an enzyme known to be present 
in the arterial wall (Hurt-Camejo et al., 1997), was hydrolyzed 
significantly more than native LDL by S-SMase at neutral pH (FIG. 10). 
Other atherogenic particles are susceptible to S-SMase at neutral pH 
include apo CIII-enriched lipoproteins (FIG. 11) and plasma LDL from E0 
mice (FIG. 12A); the E0 particles aggregate after treatment with S-SMase 
at neutral pH (see text). Since susceptibility of SM and apo 
CIII-containing emulsions to hydrolysis by S-SMase is greater in the 
absence of free apo CIII (FIG. 11A), apo CIII may act as a bridge between 
S-SMase and membrane SM, thereby stimulating SM hydrolysis. A property of 
E0 LDL that likely plays a role in its susceptibility to S-SMase is the 
high SM:PC ratio in these particles (FIG. 12B), although other factors are 
probably also involved. In particular, human native plasma LDL has a SM:PC 
ratio that is greater than E0 LDL and equal to the "control" emulsions in 
FIG. 12B and yet is substantially less susceptible to hydrolysis by 
S-SMase at neutral pH than these particles. Since E0 LDL contains mostly 
apo B48 (Plump et al., 1992; Zhang et al., 1992) and the emulsions contain 
no protein, it is possible that apo B100, the protein in human native 
plasma LDL, confers relative resistance to hydrolysis by S-SMase. In this 
context, it is possible that oxidative degradation of apo B100 contributes 
to the enhanced susceptibility of oxidized LDL to hydrolysis by S-SMase. 
Most importantly, LDL extracted from atherosclerotic lesions was hydrolyzed 
ten-fold more than same-donor plasma LDL by S-SMase at neutral pH (FIG. 
13), indicating that lesional LDL is modified in vivo, either before or 
after retention in lesions (see below), to a form that renders it more 
susceptible to hydrolysis by S-SMase. Although it is difficult to 
determine the specific characteristics of lesional LDL that increase its 
susceptibility to S-SMase, lesional LDL is enriched in SM (Table 1), which 
is shown here to be a potent stimulator of S-SMase hydrolysis of LDL (FIG. 
12B). Evidence of oxidation was not found. Detecting oxidation of lesional 
LDL may require more sensitive and/or specific techniques, such as 
immunostaining for oxidized epitopes (Yla-Herttuala et al., 1989) or 
evidence of myeloperoxidase-induced oxidation (Leeuwenburgh et al., 1997). 
Thus, oxidation may, in fact, be an important modification of LDL in vivo 
that increases LDL-SM hydrolysis by S-SMase. Nonetheless, SM-enrichment of 
LDL in the arterial wall may prove to be an important process regulating 
hydrolysis and aggregation of LDL in the subendothelium. The mechanism of 
SM enrichment of lesional LDL is not known. The possibilities include SM 
enrichment of LDL retained in the subendothelium (e.g., by increased 
arterial-wall synthesis of SM followed by transfer of the SM to retained 
LDL) or selective uptake of SM-rich LDL from the plasma (see, for example, 
Portman et al., 1970). 
In summary, lipoprotein-SM hydrolysis, which occurs in the subendothelium 
in vivo (Schissel et al., 1996), causes LDL aggregation and enhances 
lipoprotein retention on matrix (Tabas et al., 1993; Xu et al., 1991). 
S-SMase, which is secreted by arterial-wall and lesional cells in culture, 
can efficiently hydrolyze and aggregate modified LDL at neutral pH. Most 
importantly, LDL extracted from atherosclerotic lesions, but not plasma 
LDL, is significantly hydrolyzed by S-SMase at neutral pH, indicating that 
modifications present in vivo can stimulate S-SMase hydrolysis of LDL. 
S-SMase, therefore, is a prime candidate for the arterial-wall SMase that 
acts on retained lipoproteins, causing lipoprotein aggregation and leading 
to lesion initiation and progression. 
Footnotes 
1. Abbreviations used: apo, apolipoprotein; DMEM, Dulbecco's modified 
Eagle's medium; LDL, low-density lipoprotein; L-SMase, lysosomal 
sphingomyelinase; PBS, phosphate-buffered saline; PC, phosphatidylcholine; 
SM, sphingomyelin; SMase, sphingomyelinase; sPLA.sub.2, nonpancreatic 
secretory phospholipase A.sub.2 ; S-SMase; secretory sphingomyelinase; 
TBARS, thiobarbituric acid-reactive substances; TLC, thin-layer 
chromatography. 
TABLE I 
______________________________________ 
SPHINGOMYELIN AND PHOSPHATIDYLCHOLINE CONTENTS 
OF PLASMA AND LESION LDL 
Sphingomyelin 
Phosphatidylcholine 
Content (nmol/mg 
Content SM:PC 
Sample protein) (nmol/mg protein) 
Molar Ratio 
______________________________________ 
Plasma LDL 
376 .+-. 20 785 .+-. 13 0.478 .+-. 0.022 
#1 
Plasma LDL 
352 .+-. 10 727 .+-. 6 0.473 .+-. 0.025 
#2 
Lesion LDL 
858 .+-. 19 710 .+-. 14 1.203 .+-. 0.040 
#1 
Lesion LDL 
902 .+-. 50 516 .+-. 22 1.740 .+-. 0.170 
#2 
______________________________________ 
Plasma and lesional LDL from FIG. 13 (25 .mu.g protein) were 
lipid-extracted, and the sphingomyelin and phosphatidylcholine contents in 
the lipid extracts were assayed as described under "Experimental 
Procedures". 
EXAMPLE 3 
Evidence that Both Secreted and Lysosomal Sphingomyelinase are Directly 
Activated by Zinc but Differ in Their Exposure to Intracellular Zinc* 
Many cell types secrete a Zn.sup.2+ -dependent SMase (S-SMase). This enzyme 
arises from the same gene and the same mRNA that gives rise to lysosomal 
SMase (L-SMase), which has been reported to be a cation-independent 
enzyme. Herein, evidence is presented for a model to explain how a single 
mRNA gives rise to two forms of SMase with apparent differences in 
Zn.sup.2+ -dependency. First,.sup.2+ Zn-induced activation of S-SMase 
does not involve a Zn.sup.2+ -dependent cofactor, indicating direct 
activation by Zn.sup.2+, and the enzyme binds Zn.sup.2+ as assessed by 
zinc-chelate chromatography. Second, L-SMase activity in sonicated cell 
homogenates was inhibited by vigorous chelation of Zn.sup.2+ and 
partially reactivated by addition of exogenous Zn.sup.2+. Thus, both L- 
and S-SMase are zinc-activated enzymes, and, indeed, the amino acid 
sequence includes several Zn.sup.2+ -binding motifs. Interestingly, SMase 
activity in a lysosome-rich 16,000.times.-x-g pellet was .about.50% 
Zn.sup.2+ -dependent, suggesting sub-saturating levels of Zn.sup.2+ in 
intact lysosomes and raising the possibility of regulation by Zn.sup.2+ 
availability. Third, when S-SMase was incubated with SMase-negative cells, 
the enzyme was internalized and trafficked to lysosomes; when subsequently 
assayed in the cell homogenate, the enzyme became active in the absence of 
exogenously added Zn.sup.2+ and was inactivated by subsequent chelation 
of Zn.sup.2+. These data suggest a model in which L-SMase is exposed to 
cellular Zn.sup.2+ during trafficking to lysosomes or in lysosomes, and 
some additional Zn.sup.2+ exposure may occur during preparation of the 
sonicated cell homogenate. In contrast, the pathway targeting S-SMase to 
secretion, which we showed is clearly distinct from the lysosomal 
targeting pathway, appears to be sequestered from cellular pools of 
Zn.sup.2+ ; thus S-SMase requires exogenous Zn.sup.2+ for activity. This 
model provides important information for understanding the enzymology and 
regulation of L- and S-SMase and for the design of animal models to 
explore their functions in vivo. 
SMases.sup.1 (SM phosphodiesterase, E.C. 3.1.4.12) have been implicated in 
a wide variety of physiological and pathophysiological processes, 
including lysosomal hydrolysis of endocytosed SM (Levade et al., 1986; 
Brady, 1983), ceramide-mediated cell signalling (Kolesnick, 1991; Hannun 
et al., 1989), membrane vesiculation (Skiba et al., 1996), alterations in 
intracellular cholesterol trafficking (Skiba et al., 1996; Slotte et al., 
1988; Porn et al., 1995; Okwu et al., 1994), and atherogenesis (Xu et al., 
1991; Tabas et al., 1993; Schissel et al., 1996; Williams et al., 1995). 
So far, only one mammalian SMase gene has been cloned, the "acid SMase" or 
"ASM" gene, which gives rise to lysosomal SMase (L-SMase) (Schuchman et 
al., 1991). This gene can also give rise to a secreted SMase (S-SMase) 
(Schissel et al., 1996). 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 ASM knock-out mice (Schissel et al., 
1996). The secreted form of the enzyme, which requires exogenously added 
Zn.sup.2+ for activity (Schissel et al., 1996), may also be involved in 
some of the processes mentioned above. 
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 (Xu et al., 1991; Tabas et al., 
1993), which is a prominent event during atherogenesis (Hoff et al., 1985; 
Nievelstein et al., 1991; Guyton et al., 1996) and one that leads to 
massive macrophage foam cell formation (Xu et al., 1991; Tabas et al., 
1993; Hoff et al., 1990; Khoo et al., 1988; suits et al., 1989). 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 (Schissel et al., 1996). Third, S-SMase, a leading candidate for 
this arterial-wall enzyme, is secreted by macrophages (Schissel et al., 
1996) and endothelial cells.sup.2, cell types found in atherosclerotic 
lesions. Fourth, S-SMase is able to hydrolyze the SM in atherogenic 
lipoproteins at neutral pH..sup.3 Other possible roles for S-SMase may be 
in ceramide-mediated cell signalling, perhaps after reuptake of the 
secreted enzyme into endosomal vesicles (cf. Wiegmann et al., 1994; Cifone 
et al., 1995), in extracellular sphingomyelin catabolism after nerve 
injury and during demyelination (Schissel et al., 1996; Svensson et al., 
1993; Bauer et al., 1994; Hartung et al., 1992), and in defense against 
viruses, many of which are enriched in SM (van Genderen et al., 1994; 
Aloia et al., 1988) and can be inactivated by treatment with SMase in 
vitro. 
L- and S-SMase are very similar proteins. Cells transfected with an ASM 
cDNA overexpress both L-SMase and S-SMase (Schissel et al., 1996), 
indicating that S-SMase does not arise by alternative processing of the 
ASM gene. In addition, antibodies made against L-SMase recognize S-SMase, 
indicating that the common mRNA is translated in the same reading frame, 
and the molecular weights of the enzymes on Western blot are very close to 
one another (see Schissel et al., 1996). Nevertheless, S-SMase requires 
exogenously added Zn.sup.2+ for activation in in-vitro assays whereas 
L-SMase does not (Schissel et al., 1996). 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 (Lebade et al., 1986). 
What is the basis for this apparent difference in Zn.sup.2+ dependency 
between L- and S-SMase? Data is presented herein to support the hypothesis 
that, in fact, both forms of the enzyme are zinc-activated enzymes. The 
Zn.sup.2+ -dependency of L-SMase has been overlooked because it is already 
saturated with Zn.sup.2+ upon isolation from cell homogenates and thus 
does not respond to exogenous Zn.sup.2+ at the time of assay. 
Furthermore, as is the case with known zinc-metalloenzymes (cf. Little et 
al., 1975), the Zn.sup.2+ cannot be stripped from L-SMase by simple 
exposure to EDTA. S-SMase escapes lysosomal targeting and travels through 
a secretory pathway that does not come into contact with intracellular 
stores of Zn.sup.2+. Thus, this enzyme requires Zn.sup.2+ during 
subsequent in-vitro assay. The information herein presented should prove 
useful for future studies that explore the enzymology and regulation of 
these important SMases and for the design of animal models to explore 
their functions in vivo. 
Experimental Procedures 
Materials--The FALCON.RTM. tissue culture plasticware used in these studies 
was purchased from FISHER.RTM. Scientific Co. (Springfield, N.J.). Tissue 
culture media and other tissue culture reagents were obtained from LIFE 
TECHNOLOGIES.RTM. (Baltimore, Md.). Fetal bovine serum (FBS) was obtained 
from HYCLONE LABORATORIES.RTM. (Logan, Utah) and was heat-inactivated for 
1 h at 65.degree. C. (HI-FBS). [9,10-.sup.3 -H]Palmitic acid (56 Ci/mmol) 
was purchased from DUPONT NEN.RTM. and [N-palmitoyl-9-10-.sup.3 
H]Sphingomyelin was synthesized as previously described (Schissel et al., 
1996; Sripada et al., 1987; Ahmad et al., 1985). (N,N)-Dimethylformamide; 
1,3-dicyclohexylcarbodiimide; (N)-hydroxysuccinimide; and 
(N,N)-diisopropylethylamine were purchased from ALDRICH.RTM. Chemical Inc. 
Precast 4-20% gradient polyacrylamide gels were purchased from NOVEX.RTM. 
(San Diego, Calif.). Nitrocellulose was from Schleicher and Schuell 
(Keene, N.H.). FLAG-tagged S-SMase and rabbit anti-FLAG-tagged S-SMase was 
from (AMGEN.RTM., Boulder, Colo.); 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. Peroxidase-conjugated goat 
anti-rabbit IgG was purchased from PIERCE.RTM. Chemical Co. (Rockford, 
Ill.). The thiol-based metalloproteinase inhibitors, HS--CH.sub.2 
--R--CH(CH.sub.2 --CH(CH.sub.3).sub.2)--C)-Phe-Ala-NH.sub.2 and 
HO--NH--CO--CH.sub.2 --CH(CH.sub.2 
CH(CH.sub.3).sub.2)--C)-Nal-Ala-NH--CH.sub.2 --CH.sub.2 --NH.sub.2, were 
purchased from Peptides International, Inc. (Louisville, Ky.). 
.beta.-endo-N-acetylglucosaminidase H (endo H) and peptide-N-glycanase F 
were purchased from Boehringer Mannheim. Bovine liver 215-kD 
mannose-6-phosphate receptor linked to Affigel 15 was made as described by 
Varki and Kornfeld (Varki et al., 1983). Sphingosylphosphorylcholine, 
1,10-phenanthroline, and all other chemicals and reagents were from 
SIGMA.RTM. Chemical Co. (St. Louis, Mo.), and all organic solvents were 
from FISHER.RTM. Scientific Co. 
Cells--Monolayer cultures of J774.A1 cells (from the American Type Culture 
Collection--see Khoo et al., 1989) were grown and maintained in spinner 
culture with DMEM/HI-FBS/PSG as described previously (Okwu et al., 1994; 
Khoo et al., 1989). Human skin fibroblasts obtained from a patient with 
type A Niemann-Pick disease (R496L mutation Levran et al., 1991) were 
grown in DMEM/HI-FBS/PSG. CHO-K1 cells were grown in Ham's F-12 containing 
10% HI-FBS and PSG. CHO cells stably transfected with ASM cDNA were 
maintained in DMEM/HI-FBS/PSG (Schissel et al., 1996). 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 ml 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 was removed. The cells were washed two 
times with ice-cold 0.25 M sucrose and scraped into 0.3 ml and 3.0 ml of 
this sucrose solution per 35-mm and 100-mm dishes, respectively. Unless 
indicated otherwise, the scraped cells were disrupted by sonication on ice 
using three 5-second bursts (Branson 450 Sonifier), and the cellular 
homogenates were assayed for total protein by the method of Lowry et al. 
(Merrill et al., 1990) and for SMase activity as described below. The 
conditioned media were spun at 800.times.g for 5 min to pellet any 
contaminating cells and concentrated six-fold using a Centriprep 30 
(Amicon; Beverly, Mass.) concentrator (molecular weight cut off=30,000). 
For the experiment in FIG. 18, 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 psi of 
nitrogen pressure for 1.5 minutes using a nitrogen cell disruption bomb 
(Parr Instrument Company, Moline, Ill.). 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.times.g for 5 min to pellet any remaining 
intact cells and nuclei. This post-nuclear supernate (PNS) was collected, 
and the volume was increased to 15 ml with 0.25 M sucrose and then spun at 
16,000.times.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, were assayed for SMase activity. 
SMase Assay--As previously described (Schissel et al., 1996), the standard 
200-.mu.l assay mixture consisted of up to 40 .mu.l 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 .mu.l. The reaction was 
initiated by the addition of 40 .mu.l substrate (50 pmol [.sup.3 
H]sphingomyelin) in 0.25 M sucrose containing 3% Triton X-100 (final 
concentration of Triton X-100 in the 200-.mu.l assay mix=0.6%). When 
added, the final concentrations of EDTA and Zn.sup.2+ were 5 mM and 0.1 
mM, respectively, unless indicated otherwise. The assay mixtures were 
incubated at 37.degree. C. for no longer than 3 h and then extracted by 
the method of Bligh and Dyer (Bligh et al., 1959); the lower, organic 
phase was harvested, evaporated under N.sub.2, and fractionated by TLC 
using chloroform:methanol (95:5). The ceramide spots were scraped and 
directly counted to quantify [.sup.3 H]ceramide. Typically, the assay 
reactions contained approximately 20 .mu.g of cellular homogenate protein 
and a volume of conditioned media derived from a quantity of cells 
equivalent to approximately 50 .mu.g 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 milliamps 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/v) and then silver-stained using reagents from 
BioRad (Hercules, Calif.). Other gels were electrotransferred (100V 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 a 
t 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 (DuPont 
NEN.RTM.) for 2 min and exposed to X-ray film for 1 min. 
Glycosidase Treatments--The procedure described by Hurwitz et al. (Hurwitz 
et al., 1994) was followed. CHO-K1 cells were incubated overnight with 
serum-free medium (CHO-S-SFM II from GIBCO). Fifty .mu.g of 
30-fold-concentrated conditioned medium and 50 .mu.g of cell homogenate 
were diluted 1:1 (v/v) with 50 mM sodium acetate buffer, pH 5.0, 
containing 2% SDS and 20 mM .beta.-mercaptoethanol (Hurwitz et al., 1994). 
One set of aliquots of the diluted conditioned medium and cell homogenate 
were treated for 16 h at 37.degree. C. with 4 mU endo H. Another set of 
aliquots w as 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.degree. C. 
with 100 mU peptide-N-glycanase F. The endo H digest and a 
trichloroacetic-acid pellet of the peptide-N-glycanase F digest (Hurwitz 
et al., 1994) 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 (Hortin et al., 1989). Packed one-ml columns of chelating 
Sepharose 6B (iminodiacetic acid coupled to agarose gel via a hydrophilic 
spacer; from Pharmacia) were washed with 10 mM sodium acetate buffer, pH 
6.0, containing 10 mM EDTA, to leave the column uncharged, or containing 
60 mM ZnCl.sub.2, to charge the column with Zn.sup.2+. The columns were 
then equilibrated with 50 mM Hepes 
(N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid) 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 ten 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 five 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. 
Results 
Zn.sup.2+ requirement for S-SMase does not involve a Zn.sup.2+ -dependent 
cofactor. One question was how L- and S-SMase acquire their apparent 
differences in zinc-dependency. One explanation would be that the secreted 
form requires a Zn.sup.2+ -dependent co-factor. Because many lysosomal 
enzymes undergo proteolytic activation (Hasilik, 1992), an obvious 
candidate for a Zn.sup.2+ -dependent co-factor would be a 
zinc-metalloproteinase. The results of five experiments, however, ruled 
out this possibility. First, Zn.sup.2+ -activated S-SMase can be 
subsequently inactivated by Zn.sup.2+ -chelation (see below); 
reversibility of Zn.sup.2+ -induced activation is not consistent with 
proteolytic activation. Second, inhibitors of zinc-metalloproteinases, 
such as tissue inhibitor of metalloproteinase-1 (TIMP-1) (Nagase et al., 
1996) and two different thiol-based peptide inhibitors, HS--CH.sub.2 
--R--CH(CH.sub.2 --CH(CH.sub.3).sub.2)--C)-Phe-Ala-NH.sub.2 and 
HO--NH--CO--CH.sub.2 --CH(CH.sub.2 
CH(CH.sub.3).sub.2)--C)-Nal-Ala-NH--CH.sub.2 --CH.sub.2 --NH.sub.2 
(Panchenko et al., 1996), did not affect the ability of Zn.sup.2+ to 
activate S-SMase. Third, mammalian zinc-metalloproteinases require 
Ca.sup.2+ as well as Zn.sup.2+ for activity (Reynolds, 1996), whereas 
Ca.sup.2+ is not a requirement for the activation of S-SMase (Schissel et 
al., 1996). Fourth, comparison of Zn.sup.2+ -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 M.sub.r (see control data in FIG. 16, below); in 
addition, S-SMase not activated with Zn.sup.2+ had the same apparent 
M.sub.r as Zn.sup.2+ -activated S-SMase. 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 (Schissel et al., 1996), was .about.95% Zn.sup.2+ -dependent. Thus, 
neither a zinc-metalloproteinase nor any other Zn.sup.2+ -dependent 
cofactor appears to be involved in the activation of S-SMase, suggesting 
direct activation of S-SMase by Zn.sup.2+. 
To further support this conclusion, a demonstration that S-SMase directly 
binds Zn.sup.2+ by subjecting conditioned media from ASM-transfected CHO 
cells (Schissel et al., 1996) to zinc-chelate chromatography (cf. Hortin 
et al., 1989) was undertaken. None of the S-SMase from the conditioned 
medium bound to an uncharged column, whereas &gt;95% of the S-SMase bound to 
a Zn.sup.2+ -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 Zn.sup.2+. 
Evidence for direct activation of L-SMase by Zn.sup.2+ --Despite the 
long-standing tenet that L-SMase is a cation-independent enzyme (Brady, 
1983), it was hypothesized 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 (Schissel et al., 1996), and S-SMase binds and is 
directly activated by Zn.sup.2+ (above). Second, there are seven amino 
acyl sequences in the enzyme that are homologous to Zn.sup.2+ -binding 
sequences in known zinc-metalloenzymes.sup.4 (Valle et al., 1990), 
including one sequence that is very similar to that in another 
phosphodiesterase enzyme (Table 2). Third, L-SMase shares two other 
properties of known zinc-metalloenzymes, namely, inhibition by phosphate 
ions (Levade et al., 1986; Brady, 1983), which are thought to block the 
Zn.sup.2+ -binding pocket(s) in zinc-metalloenzymes (Hough et al., 1989), 
and inhibition by high concentrations (e.g., 6 mM) of ZnCl.sub.2 
(Reynolds, 1996; Spence et al., 1989). 
To directly test whether L-SMase is a zinc-activated enzyme, chelation of 
Zn.sup.2+ away from each enzyme in vitro was tried to determine the 
effect on catalytic activity. In fact, the conclusion by others that 
L-SMase is a cation-independent enzyme is based partly on the observation 
that EDTA does not inhibit activity (Schneider et al., 1967; Rao et al., 
1976; Yamaguchi et al., 1977; Callahan t al., 1978; Bowser et al., 1978; 
Watanabe et al., 1983). Many zinc-metalloenzymes, however, bind the metal 
very tightly and thus require more potent chelation, such as by long-term 
incubation with the Zn.sup.2+ chelator 1,10-phenanthroline (Little et 
al., 1975). To begin, studies were conducted with S-SMase, which was shown 
herein to bind and be directly activated by Zn.sup.2+. In FIG. 14A, 
conditioned medium from J774 macrophages was incubated with either EDTA or 
Zn.sup.2+ (first two bars) and then assayed for SMase activity. The 
S-SMase is markedly activated by Zn.sup.2+ (Schissel et al., 1996). An 
aliquot of Zn.sup.2+ -activated S-SMase was incubated for 18 h with EDTA 
plus 1,10-phenanthroline in an attempt to chelate the enzyme-bound 
Zn.sup.2+. As shown in the third bar in FIG. 14A, this treatment resulted 
in an approximately 50% loss of activity; treatment with EDTA alone did 
not affect enzyme activity. Finally, the partially inactivated S-SMase was 
dialyzed against Zn.sup.2+ -containing buffer (fourth bar), which restored 
activity to the original level observed when Zn.sup.2+ was added 
initially to the conditioned medium (compare fourth and second bars in 
FIG. 14A). 
Next, cellular (i.e., lysosomal) SMase.sup.5 (FIG. 14B) was examined. As 
reported (Levade et al., 1986; Brady, 1983; Schissel et al., 1996), and in 
contrast to the situation with the secreted enzyme, L-SMase in cell 
homogenates shows maximal activity without added Zn.sup.2+ and is not 
inhibited by EDTA (first three bars of FIG. 14B). The original, active 
cellular enzyme (i.e., not exposed to exogenous Zn.sup.2+ in vitro) was 
incubated with EDTA plus 1,10-phenanthroline for 8 h (fourth bar). This 
treatment resulted in almost total inactivation of the enzyme, and 
substantial activity was restored by dialyzing against Zn.sup.2+ (fifth 
bar) or by directly adding back excess Zn.sup.2+..sup.6 The actual degree 
of inhibition by chelation and re-activation by Zn.sup.2+ -dialysis 
differed somewhat between the secreted and cellular SMases, perhaps due to 
a lesser stability of the cellular enzyme under the incubation conditions 
employed. Nonetheless, the overall patterns of inhibition and reactivation 
shown in FIG. 14, together with the lines of evidence mentioned earlier, 
provide strong evidence that L-SMase, like S-SMase, is a zinc-activated 
enzyme. 
The difference in requirement for Zn.sup.2+ in the in-vitro assays of L- 
and S-SMase: differential Zn.sup.2+ -affinity versus differential exposure 
to cellular Zn.sup.2+ prior to the assay--One possible explanation for 
the difference in Zn.sup.2+ -requirement in the in-vitro assays of L- and 
S-SMase is that the two enzymes would both be exposed to the same, though 
limiting, concentration of intracellular Zn.sup.2+, 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 Zn.sup.2+ at the time of the assay. The secreted enzyme would 
have lower affinity for Zn.sup.2+, and thus excess exogenous Zn.sup.2+ 
would have to be added for activation in vitro. 
The relative Zn.sup.2+ -affinities of these two enzymes was estimated by 
assaying their inactivation as a function of increasing exposure to metal 
chelators (Little et al., 1975). Therefore, a cellular homogenate of J774 
macrophages and the conditioned medium from these cells was incubated with 
EDTA plus the 1,10-phenanthroline for increasing times at 4.degree. C. and 
then assayed these two fractions for SMase activity at each time point. 
Both enzymes lost activity with increasing duration of chelation (FIG. 
15), whereas incubation in the absence of the chelators for 8 h at 
4.degree. C. resulted in no loss of either secreted or cellular SMase 
activity. Surprisingly, 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 
Zn.sup.2+ than S-SMase. 
The other possibility is that both enzymes bind Zn.sup.2+ with similar 
affinities, but only the lysosomal enzyme would be exposed to pools of 
intracellular Zn.sup.2+ prior to the assay; this exposure to Zn.sup.2+ 
could occur during transit to or residence in lysosomes and/or during 
preparation of the cell homogenate. Indeed, studies in many different cell 
types have shown that Zn.sup.2+ is distributed in various intracellular 
organelles, including lysosomes (Bettger et al., 1981) and cytoplasmic 
vesicles (Sanscher et al., 1985). This model makes several assumptions and 
predictions that we tested experimentally. First, the idea that exposure 
of L-SMase to Zn.sup.2+ could occur during transit to or residence in 
lysosomes assumes that S-SMase does not simply arise by exocytosis of 
lysosomal vesicles (cf. Ref. Hasilik, 1992). To directly test this 
important point, data was obtained on the carbohydrates of L- and S-SMase. 
The lysosomal targeting of L-SMase is typical for most lysosomal enzymes: 
acquisition of Asn-linked high-mannose oligosaccharides (Hurwitz et al., 
1994; Newrzella et al., 1996) followed by phosphorylation of some of the 
mannose residues and shuttling from the trans-Golgi to late 
endosomes/prelysosomes via mannose-phosphate receptor-containing vesicles 
(Hasilik, 1992; Kornfeld, 1987). In the typical (i.e., non-lysosomal) 
secretory pathway, however, the original high-mannose oligosaccharides on 
the SMase would be expected to undergo processing to complex 
oligosaccharides during transit through the Golgi (Hurwitz et al., 1994; 
Hasilik, 1992; Newrzella et al., 1996; Kornfeld, 1987). Therefore, 
aliquots of conditioned medium and homogenates from untransfected CHO 
cells were incubated with endo H, which is specific for high mannose-type 
Asn-linked oligosaccharides (Yamamoto, 1994); other aliquots were 
incubated with peptide-N-glycanase F, which cleaves both high mannose and 
complex Asn-linked oligosaccharides (Yamamoto, 1994). These incubations 
were then analyzed by anti-SMase immunoblots. As shown in FIG. 16, S-SMase 
was completely resistant to endo H but susceptible to peptide-N-glycanase 
F, indicating the presence of complex-type Asn-linked oligosaccharides. In 
contrast, L-SMase was susceptible to both glycosidases, which confirms 
that this form of the enzyme has high mannose-type oligosaccharides. These 
data indicate that S-SMase does not arise via exocytosis of lysosomes or 
vesicles in transit to lysosomes but rather through the typical secretory 
pathway. These distinctly divergent pathways provide the opportunity for 
one of the SMase to be exposed to different levels of cellular Zn.sup.2+ 
than the other form of the enzyme. 
Second, the model implies that it is the sequestration of S-SMase away from 
Zn.sup.2+ in the lysosomal pathway, not the oligosaccharide processing of 
S-SMase per se, that confers Zn.sup.2+ -dependency on S-SMase. To test 
this idea, a system in which cells secreted S-SMase that was 
mannose-phosphorylated but not exposed to the lysosomal pathway was 
investigated. In transfected cells that massively overexpress a lysosomal 
enzyme, a substantial portion of the mannose-phosphorylated form of this 
enzyme saturates the mannose-6-phosphate receptor shuttling mechanism 
(Ioannou et al., 1992). Therefore, these cells secrete lysosomal enzymes 
that are mannose-phosphorylated but that have not been exposed to the 
lysosomal targeting pathway or to lysosomes (Ioannu et al., 1992). Indeed, 
.about.80% of S-SMase from ASM-transfected CHO cells bound to a 
mannose-6-phosphate receptor affinity column and could be eluted with 
mannose-6-phosphate (cf. Faust et al., 1987). This secreted SMase was 98% 
Zn.sup.2+ -dependent, while the lysosomal SMase in these cells required no 
added Zn.sup.2+ when assayed in cellular homogenates. Thus, even though 
the S-SMase from these overexpressing cells underwent the typical 
carbohydrate modifications of a lysosomal, not a secretory, enzyme, it had 
the same degree of Zn.sup.2+ -dependency seen with S-SMase from 
non-transfected cells. This finding is consistent with the model herein, 
since the S-SMase from these cells bypassed lysosomal targeting and thus 
would not be exposed to cellular pools of Zn.sup.2+. 
Third, the model predicts that secreted, Zn.sup.2+ -dependent S-SMase, when 
endocytosed by cells and delivered to lysosomes, would be exposed to 
cellular Zn.sup.2+ and thus no longer require exogenously added 
Zn.sup.2+. To test this prediction, highly purified secreted FLAG-tagged 
SMase, which is 99.6% Zn.sup.2+ -dependent, and fibroblasts from a patient 
with type A Niemann-Pick disease, which completely lack both L- and 
S-SMase were used (Schissel et al., 1996). To introduce the FLAG-S-SMase 
into intact Niemann-Pick fibroblasts, the enzyme was added to media on 
living cells and then incubated for 16 h at 37.degree. C. It was found 
that these cells can endocytose S-SMase and target it to lysosomes in a 
catalytically active form, as evidenced by a substantial reduction in 
lysosomal SM mass, which otherwise accumulates in these mutant fibroblasts 
(cf. Hurwitz et al., 1994; Neufeld, 1980). After the incubation, media and 
cells were harvested, cells were homogenized, and the media and sonicated 
cell homogenates were assayed for SMase activity (FIG. 17). As expected, 
the SMase activity that remained in the media (i.e., the portion of the 
enzyme that was not internalized) was almost entirely Zn.sup.2+ 
-dependent. In contrast, the SMase activity in the sonicated cell 
homogenates, which originated entirely via internalization of the 
exogenously added secretory enzyme, was maximally activated in the absence 
of Zn.sup.2+ (FIG. 17). Addition of Zn.sup.2+ not only failed to 
increase the cellular enzymatic activity, but for unclear reasons produced 
a somewhat lower activity. To demonstrate that the Zn.sup.2+ -independent 
SMase activity in the cell homogenates was due to exposure of the enzyme 
to cellular pools of Zn.sup.2+, homogenates were subsequently incubated 
for 24 h with EDTA plus 1,10-phenanthroline, to chelate Zn.sup.2+ (see 
FIGS. 1 and 2), or with buffer alone as a control. As shown in FIG. 17, 
the cellular SMase activity was specifically inhibited by Zn.sup.2+ 
-chelation. Similar results were obtained when the Niemann-Pick cells were 
incubated with native (i.e., non-FLAG-tagged) S-SMase. These data 
demonstrate that Zn.sup.2+ -dependent secreted SMase can become maximally 
activated by exposure to intracellular pools of Zn.sup.2+ during 
internalization, intracellular sorting, and/or cellular sonication. Once 
sonic disruption of the cells is completed, however, these pools of 
Zn.sup.2+ are too diluted to activate the enzyme, since simply adding 
S-SMase to sonicated cell homogenates does not reduce the Zn.sup.2+ 
-dependency of the enzyme. 
The Zn.sup.2+ -dependency of L-SMase and previous work demonstrating 
discrete intracellular Zn.sup.2+ pools that can change under certain 
metabolic conditions (cf. Csermely et al., 1987; Brand et al., 1996) led 
us to consider the hypothesis that Zn.sup.2+ 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 Zn.sup.2+. In the standard L-SMase assay, 
transfected CHO cells are completely homogenized (by sonication), and the 
cell homogenate is assayed. Under these conditions, the enzyme is 
maximally activated, and exogenous Zn.sup.2+ has no effect (FIG. 18A). To 
obtain a less damaged lysosomal preparation, a separate aliquot of these 
CHO cells was disrupted under 500 psi of nitrogen pressure for 1.5 min, 
and a 16,000-x-g pellet was isolated, which consists of intact lysosomes, 
as well as mitochondria and peroxisomes (cf. Watanabe et al., 1983). This 
16,000-x-g pellet was then sonicated and assayed for SMase activity. 
Remarkably, under these conditions, the enzyme was only .about.50% 
activated and was substantially stimulated by exogenous Zn.sup.2+ (FIG. 
18B). Thus, L-SMase encounters sub-saturating levels of Zn.sup.2+ during 
transit to lysosomes and/or after subsequent storage there. In contrast, 
when the cells are disrupted by sonication, nearby intracellular pools of 
Zn.sup.2+ are released, which leads to saturation of the enzyme with 
Zn.sup.2+ prior to dissipation of these pools throughout the entire 
homogenate. Thus, in the standard in-vitro assay, which uses sonicated 
whole-cell homogenates, L-SMase is fully saturated with Zn.sup.2+. These 
findings raise the intriguing possibility that the activity of L-SMase in 
intact cells may be subject to regulation by changes in the concentration 
or availability of Zn.sup.2+ in lysosomes. 
Discussion 
A model to explain the apparent difference in Zn.sup.2+ -dependency of 
L-SMase versus S-SMase is shown in FIG. 19. The ASM gene gives rise to a 
common precursor protein (Schissel et al., 1996), which is then modified 
by typical high-mannose oligosaccharide residues (Hurwitz et al., 1994; 
Newrzella et al., 1996; Kornfeld, 1987). 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. 16 and 
Hurwitz et al., 1994; Kornfeld, 1987). Vesicles containing 
mannose-phosphate receptors then shuttle this modified SMase to late 
endosomes/prelysosomes (Kornfeld, 1987), and at some point along this 
pathway the enzyme encounters cellular Zn.sup.2+ and thus becomes at 
least partially activated. As mentioned in the Results section, L-SMase 
appears to be exposed to subsaturating concentrations of Zn.sup.2+ in 
lysosomes and thus potentially subject to regulation by changes in 
Zn.sup.2+ availability. 
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 
(Levade et al., 1986; Brady, 1983). Throughout this period of study, the 
enzyme has been reported to be "cation-independent" (Levade et al., 1986; 
Brady, 1983). The data in this report strongly support the conclusion that 
this enzyme is, indeed, a zinc-activated enzyme. 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 (Schneider 
et al., 1967; Rao et al., 1976; Yamaguchi et al., 1977; Callahan et al., 
1978; Bowser et al., 1978; Watanabe et al., 1983), is already tightly 
bound to Zn.sup.2+. Thus, exogenous Zn.sup.2+ is not needed for the 
in-vitro assay, and typical short-term EDTA chelation incubations will not 
strip the enzyme of its metal, which has been reported for known 
zinc-metalloenzymes (Little et al., 1975). 
To explain the origin of S-SMase, a portion of the common precursor, via a 
potentially regulated process, bypasses 
N-acetylglucosamine-1-phosphotransferase and thus is directed into the 
secretory pathway, not the lysosomal targeting pathway (Kornfeld, 1987) 
(FIG. 19). The difference in susceptibility of S- and L-SMase to endo H 
(FIG. 16) provide direct support for this component of the model. 
Importantly, the data suggest that SMase in the secretory pathway is not 
exposed to pools of cellular Zn.sup.2+, thus explaining the requirement 
for exogenously added Zn.sup.2+ when the secreted enzyme is assayed in 
vitro. As mentioned in the Results section, however, the subcellular 
location of Zn.sup.2+ may be subject to cell-type variation or regulation 
(Bettger et al., 1981; Csermely et al., 1987; Brand et al., 1996). 
Therefore, it is possible that S-SMase may, under certain circumstances or 
in certain cell types, be fully or partially Zn.sup.2+ -independent. In 
fact, it was observed that SMase secreted by endothelial cells, unlike 
that secreted by macrophages (Schissel et al., 1996), is active in the 
absence of Zn.sup.2+ and stimulated only two-fold by exposure to 
exogenous Zn.sup.2+..sup.2 
According to this model, 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 (Baranski et al., 1992; Cantor et al., 1992; Dustin et al., 
1995) 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..sup.6 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 Zn.sup.2+, 
which is known to exist in sufficient extracellular concentrations in vivo 
to activate the enzyme (cf. Schissel et al., 1996; Spence et al., 1989). 
Interestingly, certain cytokines increase the secretion of SMase from 
endothelial cells without affecting L-SMase activity.sup.2, suggesting 
that the phosphotransferase reaction or perhaps another critical step 
responsible for determining the fate of SMase may be subject to specific 
regulation. 
The current data and previous work by others (Callahan et al., 1983) 
indicate difficulties with the prior nomenclature of these SMases. First, 
both forms of the enzymes are zinc-activated enzymes, and so previous 
designation of the secreted form as "Zn-SMase" (Schissel et al., 1996) 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 (Levade et al., 
1986). 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., 
K.sub.m) and that V.sub.max for hydrolysis is relatively pH-independent 
(Callahan et al., 1983). Furthermore, it was recently demonstrated that 
S-SMase can hydrolyze the SM of certain lipoproteins quite well at neutral 
pH..sup.3 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 (Levade 
et al., 1986; Brady, 1983; Schissel et al., 1996; Horinouchi et al., 1995; 
Otterbach et al., 1995), the nomenclature herein (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 
supporting a role for an extracellular arterial-wall SMase in 
atherogenesis (u et al., 1991; Tabas et al., 1993; Schissel et al., 1996; 
Schuchman et al., 1991). S-SMase is a leading candidate for this 
arterial-wall activity. Furthermore, it is theoretically possible that 
S-SMase plays roles in cell-signalling processes, in extracellular SM 
catabolism in the central nervous system, and in anti-viral host defense 
mechanisms. In this context, the information reported herein on the 
molecular and cellular origin of S-SMase should prove useful in further 
regulatory studies on this enzyme and in designing strategies to test the 
role of this enzyme in atherogenesis and possibly other physiologic and 
pathophysiologic processes. 
Footnotes 
1. The abbreviations used are: ASM, acid sphingomyelinase; CHO, Chinese 
hamster ovary; DMEM, Dulbecco's modified Eagle's medium; endo H, 
.beta.-endo-N-acetylglucosaminidase H; FBS, fetal bovine serum; HI-FBS, 
heat-inactivated FBS; LDL, low-density lipoprotein; L-SMase, lysosomal 
sphingomyelinase; PBS, phosphate-buffered saline; PSG, penicillin, 
streptomycin, & glutamine; SDS-PAGGE, sodium dodecylsulfate polyacrylamide 
gradient-gel electrophoresis; SM, sphingomyelin; SMase, sphingomyelinase; 
S-SMase, secreted sphingomyelinase; TIMP-1, tissue inhibitor of 
metalloproteinase-1; TLC, thin-layer chromatography. 
4. The term "zinc-metalloenzyme", while often used to describe enzymes 
directly activated by Zn.sup.2+, should be formally reserved for enzymes 
that have been purified to homogeneity and shown by atomic absorption 
spectroscopy or X-ray crystallography to contain one or more moles of zinc 
per mole of enzyme (Vallee et al., 1990). In this report, the term 
"zinc-activated" was used to describe current knowledge of S- and L-SMase, 
namely, Zn.sup.2+ -dependence of catalytic activity. The presence of 
zinc-binding motifs in the amino acyl sequence of these enzymes, together 
with the S-SMase data showing binding to the zinc-chelate column and 
absence of co-factor involvement, suggest that these enzymes are, indeed, 
"zinc-metalloenzymes", but this designation must await detailed structural 
studies. 
5. Consistent with prior literature (cf. Brady, 1983), 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 Mg.sup.2+ for 
activity (Brady, 1983). 
6. In experiments, it was 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 Zn.sup.2+, and removal of this cation by 
EDTA facilitates the removal of Zn.sup.2+ by 1,10-phenanthroline (cf. 
Little et al., 1975; Spence et al., 1989). Whatever the mechanism, the 
fact that 1,10-phenanthroline alone does not inhibit S- or L-SMase argues 
against an unlikely, though formally possible, alternative interpretation 
of the data in FIG. 14, namely, that 1,10-phenanthroline is a direct SMase 
inhibitor that becomes inactive as an inhibitor when the compound binds 
Zn.sup.2+. 
TABLE 2 
______________________________________ 
Comparison of amino acid sequences of portions of L/S- 
SMase with known Zn.sup.2+ -binding consensus sequences of zinc- 
metalloenzymes.sup.a 
Enzyme Amino acid sequence 
______________________________________ 
L/S-SMase (282).sup.b: 
H-(X).sub.3 -H-(X).sub.33 -E.sup.c 
L/S-SMase (421) H-(X).sub.3 -H-(X).sub.21 -E 
Astracin zinc-protease gene 
H-(X).sub.3 -H-(X).sub.n -E (n = 21, 28, 35) 
family 
cGMP-specific H-(X).sub.3 -H-(X).sub.24 -E 
phosphodiesterase 
Thermolysins & B. Cereus 
H-(X).sub.3 -H-(X).sub.19 -E 
neutral protease 
L/S-SMase (221) C-(X)4-C-(X).sub.23 -C-(X).sub.2 -P 
Aspartate transcarbamoylase 
C-(X).sub.4 -C-(X).sub.22 -C-(X).sub.2 -C 
L/S-SMase (136) H-(X).sub.2 -E-(X).sub.142 -H 
L/S-SMase (459) H-(X).sub.2 -E-(X).sub.115 -H 
Carboxypeptidase A & B 
H-(X).sub.2 -E-(X).sub.123 -H 
L/S-SMase (575) H-(X).sub.2 -H-(X).sub.30 -H 
DD carboxypeptidase 
H-(X).sub.2 -H-(X).sub.40 -H 
L/S-SMase (457) H-(X)-H-(X).sub.118 -H 
.beta.-lactamase H-(X)-H-(X).sub.121 -H 
______________________________________ 
.sup.a L/SSMase refers to the common amino acid sequence of L and SSMase, 
derived from the ASM cDNA (Schuchman et al., 1991); the sequences of the 
other enzymes are from Ref. (Vallee et al., 1990), except those of the 
astracin gene family, which is from Ref. (Hung et al., 1997), and of 
cGMPspecific phosphodiesterase which is from Ref. (Francis et al., 1994). 
.sup.b The number in parentheses represents the position of the first 
amino acid (H or C) of the SMase sequences displayed. 
.sup.c X refers to any amino acid. 
EXAMPLE 4 
Role of Macrophage-Secreted Zn.sup.2+ -Stimulated Sphingomyelinase in 
Atherogenesis 
Focal retention of lipoproteins in the subendothelium is a key step in both 
the initiation and propagation of atherosclerotic lesions (reviewed in 
Williams and Tabas, 1995)). Retained lipoproteins undergo marked 
aggregation (Nievelstein et al., 1991; Hoff and Morton, 1985), which is 
important for two major reasons: (1) aggregation of lipoproteins greatly 
increases the amount of lipoprotein retained (Tabas et al., 1993) and thus 
increases the atherogenic responses to this retained material; and (2) 
aggregated lipoproteins are potent stimulators of macrophage foam cell 
formation (Hoff et al., 1990; Khoo et al, 1988; Suits et al., 1989; Xu and 
Tabas, 1991), greatly exceeding the potency of oxidized LDL in inducing 
macrophage cholesteryl ester loading (Hoppe et al., 1994). Macrophage foam 
cells themselves have been shown to be very important in early lesion 
development (Smith et al., 1995) and probably also play a major role in 
late lesion complications, such as plaque rupture (Libby and Clinton, 
1993). Thus, interventions directed at blocking subendothelial lipoprotein 
aggregation may be anti-atherogenic at both early and late stages of 
lesion development. Furthermore, their mechanism of action would be 
strongly complementary to the currently available statins, which lower 
serum LDL levels. Along these lines, it should be noted that although the 
use of statins alone has led to decreased risk of both primary and 
secondary coronary artery disease, the risk reduction is far from 
complete. Therefore, the addition of another agent whose mechanism of 
action is complementary to that of lipid-lowering agents would be expected 
to have a major beneficial impact on the large number of patients with 
"statin-resistant" ischemic heart disease (Crouse, 1984). 
Sphingomyelinase-Induced LDL Aggregation 
Hydrolysis of a relatively small portion of the sphingomyelin content of 
LDL, using a commercially available bacterial sphingomyelinase (SMase), 
leads to aggregation of LDL which morphologically resembled aggregates 
seen in atheromata. These SMase-induced LDL aggregates were potent 
stimulators of macrophage foam cell formation. Furthermore, SMase was able 
to enhance LDL retention by 100-fold in a tissue culture model that 
utilized components thought to be involved in LDL retention in lesions, 
namely, smooth muscle cell-derived extracellular matrix and the 
lipoprotein lipase. When macrophages were added to this tissue culture 
model, massive foam cell formation was seen (Tabas et al., 1993). Thus, in 
vitro, SMase-induced LDL aggregation resulted in the two consequences 
described above--foam cell formation and enhanced LDL retention. 
The role of SMase in LDL aggregation is enzymatic, not structural (Schissel 
et al, 1994). Thus, although initial experiments used bacterial SMase, 
similar results can be expected for mammalian SMases. Second, 
SMase-induced LDL aggregation was due to an increase in LDL ceramide 
content, not to low LDL sphingomyelin content or to the generation of 
choline-phosphate (Schissel et al., 1994). In fact, sphingomyelin 
enrichment of particles was shown to enhance subsequent SMase-induced 
aggregation. This was an important finding, since aggregation that depends 
on a low LDL SM content could not be physiologically significant: lesional 
LDL is enriched in SM. Overall, the mechanistic studies support a role for 
arterial-wall SMase in LDL trapping and aggregation. 
The next question was whether LDL extracted from human and rabbit 
atherosclerotic lesions is enriched in ceramide. Using both early and late 
human and rabbit atheromatous material, lesion LDL is markedly enriched in 
ceramide, whereas plasma LDL has very little ceramide. [.sup.3 
H]sphingomyelin-labeled LDL was incubated with strips of rabbit aorta ex 
vivo. A portion of the labeled LDL was retained by the aorta, and this 
material was extracted and found to have a substantial amount of [.sup.3 
H]ceramide. The hydrolysis of the LDL-sphingomyelin was enhanced in 
lesional vs. normal aorta, was not inhibited by chloroquine (and thus did 
not involve lysosomal SMase), and was inhibited by EDTA. Thus, in both 
humans and rabbits, retained LDL is acted upon by an arterial wall 
non-lysosomal, cation-dependent SMase activity. 
To look for such an activity, a determination of whether cell types 
prominent in lesions could secrete a Smase was done. In a remarkable 
series of experiments, it was demonstrated that both murine and human 
macrophages secrete an abundant Zn.sup.2+ -stimulated sphingomyelinase 
activity. This macrophage-secreted Zn-SMase has properties identical to a 
SMase described in 1989 by Spence et al. These workers found the enzyme in 
fetal calf serum, and there have been no follow-up studies since this 
original report; the cellular source of their enzyme remained a mystery 
until this recent discovery. Follow-up studies of the discovery revealed 
the following: (1) Zn-SMase is markedly up-regulated during 
monocyte-to-macrophage differentiation; (2) Zn-SMase has an acidic pH 
optimum using sphingomyelin-detergent micelles as substrate; (3) Zn-SMase 
can hydrolyze LDL-SM at physiological pH, in the presence of 
subendothelial factors such as extracellular matrix and lipoprotein lipase 
(see above); and (4) Zn-SMase is secreted by other cells, including 
brain-derived microglial cells. 
The acidic pH optimum of Zn-SMase when sphingomyelin-detergent micelles 
were used as substrate prompted us to question whether this enzyme 
originated form the same gene as acidic lysosomal SMase (lys-SMase). 
Zn-SMase activity was measured in peritoneal macrophages from wild-type 
mice and mice that were made to lack one or both acid SMase genes by gene 
targeting. Remarkably, macrophages from the heterozygous knockout mice had 
half the normal amount of secreted Zn-SMase activity, and those from the 
homozygous knockout mouse secreted no Zn-SMase. Further studies revealed 
that CHO cells stably transfected with lys-SMase cDNA massively 
overexpressed both intracellular lysosomal SMase activity and secreted 
Zn-SMase activity. Lastly, when conditioned media cell extracts were 
mixed, neither the mounts nor the properties of the two SMases were 
altered, suggesting that cellular or secreted co-factor interaction with 
the enzymes cannot explain their differences. Thus, secreted Zn-SMase 
arises from the same gene as lys-SMase and arises by post-translational 
modification of the lysosomal enzyme. Studies are currently in progress to 
determine the exact molecular differences between the two SMases and to 
elucidate the intracellular pathway that converts lys-SMase into Zn-SMase. 
Screening of Inhibitors for Zn-SMase and Testing in Vitro and in Vivo 
One embodiment of the present invention is to identify an inhibitor of 
Zn-SMase and determine whether it is anti-atherogenic in animal models of 
atherosclerosis. First, a high throughput assay will need to be developed. 
The source of Zn-SMase activity will be conditioned media from macrophages 
or transfected CHO cells (see above). There are colorometric assays for 
acid SMases available and these will need to be tested using secreted 
Zn-SMase. If necessary, to avoid contaminating activities, a partially 
purified preparation of Zn-SMase, such as by passing the conditioned media 
over a sphingosylphosphorylcholine column, can be used as a source of 
enzyme for these inhibition assays. Once a rapid assay is identified and 
verified, screening can begin using compounds derived from libraries or 
other sources. Positive hits would be tested in a more detailed assay that 
would assess potency, specificity (e.g., Zn-SMase vs. lys-SMase vs. other 
Zn-requiring enzymes), and mechanism of action. Even compounds that can 
not distinguish between Zn-SMase and lys-SMase may be useful if they are 
unable to enter lysosomes or become inactive at acid pH. 
Specific and potent Zn-SMase inhibitors will then be tested in a 
cell-culture system. In this system, smooth muscle cells, lipoprotein 
lipase, LDL, and macrophages will be coincubated in an attempt to 
reproduce the synergistic interactions described above for exogenously 
added bacterial SMase and macrophage conditioned media. The goal will be 
to establish a "self-sufficient" model of subendothelial LDL retention, 
aggregation, and foam cell formation. Once established, the Zn-SMase 
inhibitors will be tested for their ability to block these events. Other 
responses to retention and aggregation, such as LDL oxidation and monocyte 
chemotaxis & adhesion, could be modeled by adding an endothelial cell 
monolayer via a Transwell insert. Inhibition of these responses by 
Zn-SMase inhibitors would then be tested as above. 
The effect of these agents may be tested in animal models of 
atherosclerosis. The two models that will be used are fat-fed apo B 
transgenic mice (Purcell-Huynh et al., 1995) and Watanabe heritable 
hyperlipidemic (WHHL) rabbits (Buja et al., 1983), both of which develop 
extensive LDL-induced atherosclerotic lesions. The most potent and 
specific Zn-SMase inhibitors will be fed to the animals at various doses 
to first determine toxicity and then effectiveness in inhibiting aortic 
Zn-SMase activity and atherogenesis. Toxicity could arise from both 
"non-specific" and specific effects of the compounds. The specific effects 
would be related to physiologically important functions of Zn-SMase. 
Regarding this point, it is possible that microglial cell-secreted 
Zn-SMase (above) plays an important role in scavenging myelin in response 
to nerve injury. Thus, an inhibitor that does not cross the blood-brain 
barrier might be most useful for the atherosclerosis studies. 
Protocols testing both primary prevention and secondary regression will be 
established, and the extent and nature of atherosclerosis will be 
determined by both quantitative (Paigen et al, 1987) and morphological 
(Nakashima et al., 1994) assays. Careful documentation of plasma 
lipoprotein levels and properties will also be carried out. Evidence of 
anti-atherogenic effects would be followed by more detailed mechanistic 
studies that will specifically assess LDL retention (Kreuzer et al, 1994) 
and aggregation in vivo. 
Creation and Examination of Mice that Selectively Lack Zn-SMase 
Another embodiment of the present invention is to determine whether mice 
that specifically lack Zn-SMase, when bred into an atherogenic background 
(i.e., apo B transgenic mice), will be protected from atherosclerosis. 
From the recent discovery, it is now known the currently available acid 
SMase knockout mice lack two distinct enzymes. Since the absence of 
lys-SMase might complicate the evaluation of atherogenesis, it will be 
necessary to create a Zn-SMase "knockout" mouse. The overall strategy will 
be as follows: (1) based upon specific strategies, create a genetic 
construct that can encode only lysosomal SMase; (2) create transgenic mice 
based upon this selective lys-SMase construct. Natural flanking regions 
will be utilized in the transgene to avoid overexpression of the enzyme; 
and (3) cross these mice with acid SMase knockout mice to create a mouse 
that lacks only Zn-SMase. 
Once created and verified, the Zn-SMase knockout mice will be crossed with 
apo B transgenic mice. The Zn-SMase knockout x apo B transgenic mice will 
be compared with apo B transgenic mice for the development of 
atherosclerosis exactly as described in Section I above, including the 
in-vivo retention and aggregation mechanistic studies. The creation of a 
mouse that selectively overexpresses Zn-SMase, using a genetic construct 
based on the results of our structural studies is also possible. The 
construct would be ligated to a macrophage-specific promoter, such as the 
scavenger receptor promoter (Horvai et al, 1995), so that the mice would 
oversecrete the enzyme specifically in macrophages. These mice would 
develop accelerated atherosclerosis that could be ameliorated by the 
Zn-SMase inhibitors developed. 
Arterial wall factors undoubtedly account for a large portion of 
atherosclerotic disease in humans and probably explain the large number of 
patients that develop atherosclerosis despite lipid-lowering therapy. In 
this light, arterial wall molecules that promote subendothelial 
lipoprotein retention and aggregation are probably very important. A 
macrophage-secreted molecule--Zn-SMase--may play a key role in these 
processes. By using both pharmacological inhibitors and genetic 
manipulations in animal models of atherosclerosis, it is possible to 
critically test the role of macrophage-secreted Zn-SMase in lipoprotein 
retention & aggregation and atherogenesis. The usefulness of the 
pharmacological inhibitors will be evaluated, leading to the development 
of important statin-complementary anti-atherogenic therapy in humans. 
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