©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Purification and Biochemical Characterization of Membrane-bound Epidermal Ceramidases from Guinea Pig Skin (*)

Yukihiro Yada, Kazuhiko Higuchi, and Genji Imokawa (§)

From the (1) Institute for Fundamental Research, Kao Corporation, 2606 Ichikaimachi, Haga, Tochigi 321-34, Japan

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

Ceramidase (CDase) catalyzes the hydrolysis of ceramides to yield sphingosine and fatty acid. In this paper, two forms of membrane-bound alkaline ceramidase, have been, for the first time, purified from guinea pig epidermis by chromatography on DEAE-cellulose, phenyl-Superose, HCA-hyroxyapatite, isoelectric focusing, Mono Q, and TSK-3000SW column. One species (CDase-I) migrated upon SDS-polyacrylamide gel electrophoresis as a single band with an apparent molecular mass of 60 kDa; the other (CDase-II) was only partially purified with apparent M of about 148,000 estimated by gel filtration. The specific activities of the two species increased by 1.130- (for CDase-I) and 400-fold (for CDase-II) over the original tissue extract. The activity of both enzymes for ceramide species decreased in the order of linoleoyl > oleoyl > palmitoylsphingosine. The optimal pH for enzyme activity was approximately 7.0-9.0 for CDase-I and 7.5-8.5 for CDase-II. Interestingly, both enzymes were inhibited by the reaction product sphingosine with a concentration for half-maximal inhibition (ID) of 100-130 µM, compared to the apparent kinetic parameters with CDase-I (K = 90 µM, V = 0.62 unit) and CDase-II (K = 140 µM, V = 0.50 units). Some lipids, such as phosphatidylcholine and sphingomyelin, are also inhibitory with IC values of 50-250 µM, suggesting well controlled CDase activity by sphingolipid metabolites. These studies begin to elucidate a regulatory mechanism for the balance of the ratio of ceramide/sphingosine which can serve as an intracellular effector molecule in epidermis.


INTRODUCTION

Ceramidase (EC 3.5.1.23) is an enzyme which catalyzes the hydrolysis of ceramide to yield sphingosine and fatty acid and is widely distributed in animal tissues such as brain, kidney, and spleen (1-5). However, the biological role of ceramidase in epidermal sphingolipid metabolism remains unclear because of no success in its purification to homogeneity. The substrate ceramide constitutes the core structure of several sphingolipids such as sphingomyelin, gangliosides, and sulfatides which are reported to play essential roles in cell proliferation and differentiation (6, 7, 8) . Ceramide serves a central role in sphingolipid metabolism and is involved as a modulator in cell growth and vitamin D-induced differentiation (9, 10) . Ceramide is recently implicated as an inducer of programmed cell death (11) . In epidermal tissue, ceramide is an important determinant for the permeability barrier and water reservoir of the upper most layers of the epidermis (12, 13, 14, 15) , the stratum corneum which is directly exposed to outer environmental circumstances. In addition to a physiological role, ceramide is documented as a stimulator for the phosphorylation of epidermal growth factor receptor in epidermoid carcinoma cells (16) . On the other hand, the reaction product sphingosine is an endogenous inhibitor of protein kinase C-mediated biochemical reaction in several cells or tissues (17, 18, 19) . Due to the importance of protein kinase C action in a wide range of cellular events, sphingosine has been implicated as an essential biological mediator in cellular responses to extracellular signals as well as in sustained biological phenomena such as tumorigenesis, cell growth, and differentiation (20, 21, 22, 23) . In mammalian epidermis, where ceramide is a major end product of epidermal differentiation, namely the keratinization process, the balance of sphingosine and ceramide may serve a control mechanism for the regulation of epidermal growth and differentiation through the varied functions of ceramidase (24, 25, 26) . Despite the involvement of ceramidase in the essential hydrolysis of sphingolipids as well as in the control process of epidermal keratinization, little is known about its biochemical properties because of the lack of purification studies. Although Wertz and Downing (27) have recently demonstrated ceramide hydrolytic activity in fractions derived from pig epidermis, the control mechanism of the hydrolysis has remained unclear. Thus, detailed enzymological studies are required for characterization of the physiological function of the enzyme. Based upon the importance of the ceramidases in understanding the regulatory mechanism of sphingolipid metabolism in the epidermis, in this study we describe the purification of two ceramidase species from guinea pig epidermis and their biochemical properties.


EXPERIMENTAL PROCEDURES

Materials

DEAE-cellulose was purchased from Whatman BioSystems Ltd. Phenyl-Superose (HR 5/5) and Mono Q (HR 5/5 and HR 10/10) columns were purchased from Pharmacia LKB Biotechnology Inc. TSK-3000SW was obtained from Toyo Soda Industries, LTD. and [1-C]palmitic acid was purchased from Amersham International plc (Bucks, United Kingdom). Sphingosine and phospholipids were obtained from Funakoshi Chemical Company (Tokyo). Preparation of [1-C]Palmitoylsphingosine-Thionyl chloride was refluxed with [1-C]palmitic acid in petroleum to produce [1-C]palmitoylsphingosine. The palmitoyl chloride was condensed with sphingosine benzoate according to the method of Shapiro and Flowers (28) . The condensation product was subjected to mild alkaline hydrolysis with 0.15 ml of sodium acetate (50%) and 120 µl of tetrahydrofuran for 90 min at 25 °C. The reaction was terminated with 2 ml of chloroform/methanol (2:1) and 0.4 ml of water. The mixtures were vortex mixed for 1 min and centrifuged for 5 min at 2,000 g. The upper phase was aspirated, and the lower organic phase was washed once with 1.3 ml of methanol:water (1:1), then evaporated under nitrogen gas. The crude product was isolated by preparative silicic acid column chromatography with a mobile phase of chloroform/methanol (9:1). The extract was dried under nitrogen gas, then 0.5 ml of 0.5 N methanol-saturated NaOH was added, and the mixture was incubated at 37 °C for 2 h. The reaction was terminated by the addition of 1 ml of chloroform, followed by 0.3 ml of water, and centrifugation for 5 min at 2,000 g. The lower phase was washed once with 0.75 ml of methanol/water (1:1). The upper phase was aspirated and the lower phase was evaporated under nitrogen gas. The crude ceramide was applied on a thin layer chromatography Silica Gel G plate and developed with chloroform/methanol/acetic acid (94:1:5). The R value of [1-C]palmitoylsphingosine showed 0.25 in this condition, on the other hand, unreacted [1-C]palmitic acid was R = 0.95 and sphingosine was located in the origin. Furthermore, the preparations of [1-C]linoleoylsphingosine and [1-C]oleoylsphingosine were performed under almost same condition of [1-C]palmitoylsphingosine. The radiochemical purity of the synthetic ceramides checked with a radioscanner was at least 97%, respectively.

Preparation of Membrane Fraction

Epidermal sheets were peeled from the guinea pig skin (Wister species, male) at 4-5 weeks of age after incubation with 1000 units/ml dispase at 4 °C overnight. Briefly, after washing with phosphate-buffered saline, epidermal sheet scissored off chips was resuspended in a 5-fold volume of 60 mM phosphate buffer, pH 7.4, containing 0.25 M sucrose, 1 mM EDTA, 1 mM EGTA, and 0.5 mM phenylmethylsulfonyl fluoride (buffer A), then homogenized on ice for a total of 3 min with 30-s bursts of a Polytron (Kinematica AG, Littau/Luzern, Switzerland). After removing unbroken epidermal tissues by centrifugation at 800 g for 10 min, the supernatant fraction was centrifuged at 10,000 g for 30 min. The pellet was resuspended in buffer A and stored frozen at -135 °C until a sufficient amount was obtained for purification. The collected pellets (about 750 mg/65 ml of protein) were thawed, resuspended in a 3-fold volume of buffer A, and extracted with an equal volume of buffer A containing 1.0% sodium cholate. The effect of sodium cholate on extraction and stability of ceramidase activity from the particle is mostly not affected (see ``Results''). After 2 h at 4 °C, the suspension was sedimented by centrifugation at 18,000 g for 30 min to obtain a cholate extract for subsequent purification.

Purification of Membrane-bound Ceramidase

All procedures were carried out at 4 °C unless otherwise indicated.

DEAE-cellulose Column Chromatography

The cholate extract was applied to a DEAE-cellulose column (3.2 18 cm) equilibrated with 60 mM phosphate buffer, pH 7.4, containing 1 mM EDTA, 1 mM EGTA, 0.5 mM phenylmethylsulfonyl fluoride, and 0.5% sodium cholate (buffer B). The column was washed with 600 ml of the same buffer, then eluted with a linear concentration gradient of NaCl from 0 to 1.0 M in 700 ml of buffer B. The flow-through (Fraction I) and eluted fractions (Fraction II) were collected in 8-ml portions. The active fractions were pooled and concentrated to about 40 ml using a hollow fiber (Mini Modules HC, Asahi Kasei Co.; exclusion limit: 13,000 daltons). The two ceramidase (CDase)() activities of flow-through (Fraction I) and eluted fractions (Fraction II) were designated as CDase-I and CDase-II.

Phenyl-Superose Column Chromatography

Fraction I from the DEAE-cellulose column was supplemented with NaCl to 3 M and applied to a phenyl-Superose packed column (HR 10/10; 1.0 10 cm, Pharmacia) equilibrated with buffer B containing 2.0 M NaCl. The proteins were eluted at a flow rate of 3.0 ml/min with successively decreasing NaCl gradients from 2.0 M to 0 M for 30 min (using total volume of 90 ml) under a high resolution liquid chromatography Bio-Dimension System (Bio-Rad). Fractions of 0.5 ml were collected. The active fractions were pooled and concentrated to about 3.0 ml using a hollow fiber. The concentrated solutions were dialyzed twice for at least 20 h against 2 liters of 50 mM Tris-HCl buffer, pH 7.4 (25 °C), containing 0.5 mM phenylmethylsulfonyl fluoride and 0.5% sodium cholate (buffer C).

HCA®-Hydroxyapatite Column Chromatography

Aliquots (about 4 ml) of CDase-I and -II obtained from the phenyl-Superose and the DEAE-cellulose columns, respectively, were applied to HCA-hydroxyapatite packed columns (P-4001 + A-7610, 4 10 mm, Koken Co., LTD, Japan) equilibrated with buffer C. The proteins were eluted at a flow rate of 1.0 ml/min with successively increasing phosphate buffer, pH 7.4, gradients from 0 to 125 mM for 30 min, from 125 to 250 mM for 5 min (using total volum of 40 ml), then 250 mM for 5 min using the HRLC Bio-Dimension System. Fractions of 0.5 ml were collected. CDase-I and -II were eluted between 70-90 and 200-250 mM, respectively.

Isoelectric Focusing

The solutions obtained from the HCA-hydroxyapatite column were dialyzed for at least 20 h against 10 mM phosphate buffer, pH 7.4, containing 0.5% CHAPS, respectively. Three milliliters of dialyzed solutions were diluted with 52 ml of 1% Bio-Lyte ampholytes, pH 3.0-10.0. All solutions were separately applied to a preparative isoelectric focusing (IEF) chamber (Rotofor Cell, Bio-Rad). The power supply was set to 12 watts constant power and the run proceeded for 4.5 h at 4 °C. The focusing chamber is divided into 20 discrete compartments, each of which was collected simultaneously into 20 tubes at the end of the run. The pH of all fractions was measured and neutralized for measuring the CDase activities. The active fractions were pooled and were dialyzed twice for at least 20 h against 2 liters of buffer B containing 200 mM NaCl. The dialyzed enzyme solutions were concentrated to about 0.4 ml in a Centricon 10 (Amicon, Millipore Corporation).

TSK-3000SW Column Chromatography

Aliquots (about 0.4 ml) of CDase-I and -II obtained from IEF were applied to TSK-3000SW columns (0.6 60 cm) equilibrated with buffer B containing 200 mM NaCl. The enzymes were eluted with the same buffer at a flow rate 1.0 ml/min (using total volum of 50 ml), and 0.2-ml fractions were collected. The active fractions were pooled and concentrated to about 0.15 ml using a Centricon 10 (Amicon) and stored at -135 °C.

Assay of CDase Activity

CDase activity was assayed by measuring the amount of radioactive palmitic acid from [1-C[palmitoylsphingosine as described previously (1, 27) . The standard reaction mixture (final volume, 200 µl) contained 125 mM Tris-HCl buffer, pH 9.0 (at 37 °C), 0.75 µCi of [1-C[palmitoylsphingosine, 100 µg of Tween 20, 250 µg of Triton X-100, and enzyme solution. To examine the effect of pH on purified CDase activity, enzyme solutions were dialyzed twice for at least 20 h against 2 liters of 125 mM acetate (pH 3.0-6.0), phosphate (pH 6.0-8.0), or borate buffers (pH 9.0-11.0), and 0.5% Triton X-100. On the other hand, to examine the effects of various lipids or cations on purified CDase activity, each agent was added to the standard reaction mixture. The reaction mixture was incubated for 60 min at 37 °C and terminated by the addition of 50 µl of carrier palmitic acid, followed by 3.0 ml of Dole's reagent (2-propanol/heptane, 1 N NaOH = 40:10:1) (29) . Heptane, 1.8 ml, and 1.6 ml of water were then added. The mixtures were vortex mixed for 1 min and centrifuged for 5 min at 2,000 g. The upper phase was carefully aspirated, and the under-phase was washed twice with 2 ml of heptane. Thereafter, 1 ml of 1 N HSO and 2.4 ml of heptane was added, and the mixture was vortex-mixed for 1 min, then centrifuged for 10 min at 2,000 g. A 1-ml portion of the upper phase was transferred to a vial and mixed with scintillation fluid. The radioactivity was determined in a liquid scintillation counter. When required, the upper phase was evaporated under nitrogen gas, then residual lipids were dissolved in 20 µl of chloroform/methanol (6:1) and analyzed by thin layer chromatography as described previously (30) . One unit of CDase activity was defined as the amount of enzyme which produced 1 nmol of palmitic acid/min under the described conditions.

Other Methods

Analytical SDS-polyacrylamide gel electrophoresis (PAGE) was performed by the method of Laemmli (31) in a linear polyacrylamide gradient of 8-16%. The protein concentration was determined using a Bio-Rad protein assay kits with bovine serum albumin as the standard.


RESULTS

Purification of Two Ceramidase Activities

Prior to purification experiments, the subcellular distribution of CDase within epidermal cells was examined. About 65% of the total CDase activity in the guinea pig epidermis homogenate was found in the 10,000 g particulate fraction, and the specific activity was the highest in this fraction. Therefore, we purified the CDase from the particulate fraction of guinea pig epidermis. In other studies, Triton X-100 has been shown to enhance the activity of CDase obtained from human spleen (5) , and sodium cholate increased that derived from the rat brain (1) . Therefore, we determined the effect of detergents on extraction and stability of CDase from the particulate fraction by using sodium cholate, Triton X-100, Brij 35, Nonidet P-40, and CHAPS. Sodium cholate, Triton X-100, and Nonidet P-40 at a concentration of 0.5% markedly enhanced the extraction and stimulated, though to a different extent, the palmitoylsphingosine hydrolyzing activity. Sodium cholate enhanced the CDase activity about 4-fold over original 10,000 g particulate fraction. In contrast, Brij 35 and CHAPS were not effective at the same concentration. During solubilization of the particulate fraction by detergents, about 40-60% of CDase activity was lost in the presence of 0.5% Triton X-100 at 4 °C for 48 h, whereas 90-95% of CDase activity retained in the presence of 0.5% sodium cholate (data not shown). Based on this experiment, we used 0.5% sodium cholate upon the extraction and purification of CDase. In the extraction step for 2 h with 0.5% sodium cholate, about 90% of the activity was solubilized with about 50% of the total protein in the particulate fraction.

The cholate extract was directly applied to a DEAE-cellulose column (Fig. 1). At this stage, the elution profile exhibited two peaks of CDase activities. One appeared as the initial pass-through fraction (Fraction I), while the other activity lied between 0 and 0.2 M NaCl gradient (Fraction II). More than 85% of the total CDase activity was recovered in these two fractions. Fraction I contained about 75% of the total activity. These two highly active fraction were designated as CDase-I and -II, respectively. This DEAE-cellulose column chromatography was efficient in separating and purifying these CDase species, demonstrating 141- and 34-fold increase in the specific activity for CDase-I and -II, respectively (). Each active fraction was subsequently eluted through a phenyl-Superose column after supplementing with NaCl to 3 M (data not shown). The CDase was bound to the hydrophobic column and eluted by decreasing NaCl gradient from 2 to 0 M, demonstrating 97-fold increase in the specific activity, whereas CDase-II was not bound to this hydrophobic column without any resulting increase in the specific activity (). This led us to omit this procedure for the purification of CDase-II.


Figure 1: Elution profiles on DEAE-cellulose column of CDase-I and -II. The cholate extract was applied to a DEAE-cellulose column, and the column was washed and eluted as described under ``Experimental Procedures.'' CDase activities were measured in the presence of 0.5% sodium cholate. CDase-I and -II activities, indicated by a bar, were pooled. -, absorbance at 280 nm; , , CDase activities.



These CDase isozymes were separately purified by successive chromatography upon HCA-hydroxyapatite, Mono Q, IEF, and TSK-3000SW under the HRLC Bio-Dimension System. Upon HCA-hydroxyapatite, the activity of CDase-I was detected in the fractions through 80 to 100 mM phosphate buffer, pH 7.4, while CDase-II was eluted at 250 mM of the same phosphate buffer (Fig. 2). The elution pattern for the two enzyme species was the same for both pH 4.5 and 9.0 at which the activity was assayed, suggesting that both represent alkaline ceramidases. This chromatographic process provided 447- and 17-fold purification index for CDase-I and -II, respectively (). On IEF, the pI value of CDase-I and -II was 6.8 and 7.2, respectively (Fig. 3). Both enzyme species were not inhibited by ampholytes that were used in the IEF. The specific activity following this chromatographic process showed 820- and 162-fold increases for CDase-I and -II as compared to the original fraction (). During the above successive chromatographic process, a split or a reduction in the molecular weight of CDases was not detected. Activity of CDase-II was relatively unstable with lower recovery, whereas CDase-I was stable during repeated chromatography. The extent of purification from the original membrane estimated from the final specific activity following TSK-3000SW gel chromatography was 1,130-fold for CDase-I and 400-fold for CDase-II (). The apparent molecular weight of these two CDase species as estimated on the final gel chromatography was 150,000 (CDase-I) and 62,000 (CDase-II) (Fig. 4A). These fractions were separated on SDS-PAGE, demonstrating that CDase-I migrated as a single band (Fig. 4B). On the other hand, CDase-II still migrated as five bands on SDS-PAGE (data not shown), indicating that it was partially purified. The results of the purification of CDase species are summarized in .


Figure 2: Elution profiles on HCA-hydroxyapatite columns of CDase-I and -II. The dialysate from phenyl-Superose (CDase-I) or DEAE-cellulose (CDase-II) were separately applied to HCA-hydroxyapatite columns and eluted as described under ``Experimental Procedures.'' Samples were as follows: A, the concentrated solutions which were eluted fraction from phenyl-Superose, followed by dialysis against buffer C. B, the NaCl-eluted fraction (Fraction II) of DEAE-cellulose. -, absorbance at 280 nm; , , CDase activities.




Figure 3: Isoelectric focusing of CDase species. The CDase species were isoelectrically focused with a power supply set to 12 watts constant power and run for 4.5 h at 4 °C. The 20 compartments were collected simultaneously in 20 tubes at the end of the run, neutralized, and assayed as described under ``Experimental Procedures.'' , pH; , CDase activities.




Figure 4: Molecular mass determination of the two CDase species at the final step of gel filtration and SDS-polyacrylamide gel electrophoresis of CDase-I. The apparent molecular masses of both CDase species were determined at the final step of purification by gel filtration through TSK-3000SW. The protein standards were 440,000 (ferritin), 232,000 (catalase), 158,000 (aldolase), 67,000 (bovin serum albumin), and 45,000 (ovalbumin). Protein bands were visualized by means of Coomassie Blue staining. Details are described under ``Experimental Procedures.'' CDase-I was obtained by chromatography on a TSK-3000SW column (7 µg of protein). Molecular mass markers; 200,000, myosin; 93,000, phosphorylase b; 66,000, bovine serum albumin; 45,000, ovalbumin; 30,000 carbonic anhydrase; 20,000, soybean trypsin inhibitor.



Substrate Specificity of CDase Species

The CDase activities were compared at pH 4.5 and 9.0 among palmitoylsphingosine, linoleoylsphingosine, and oleoylsphingosine as the substrate in the presence of 0.5% sodium cholate. The activity of both enzymes for ceramide species was in the order of linoleoyl oleoyl palmitoylsphingosines at pH 4.5 or 9.0 ().

Effects of Phospholipids

The CDase activities were markedly suppressed by either phosphatidylcholine (PC) (porcine brain or egg origin) or sphingomyelin (SM) (porcine brain or egg origin) in the presence of 0.5% sodium cholate at pH 7.4 (Fig. 5). When the final purified enzymes were dialyzed for 24 h against 50 mM Tris-HCl buffer, pH 7.4, containing 0.5% CHAPS, the similar inhibitory effect of SM or PC on these isozyme activities remained (data not shown). Half-maximal inhibition (IC) by PC (egg yolk) was about 50 µM (CDase-I; Fig. 5A) and 125 µM (CDase-II; Fig. 5C). Furthermore, the IC for SM (egg yolk) was about 350 µM (CDase-I; Fig. 5B) and 700 µM (CDase-II; Fig. 5D). The activities of CDase-I and -II were reduced by only 5-15% in the presence of 1 mM PE, PI, or PG (Fig. 5, A and C)). Thus, phosphatidylglycerol (PG) (brain), phosphatidylinositol (PI) (soybean), or phophosphatidylethanolamine (PE) (egg) were not effective (Fig. 5, A and C). Neither caldiolipin (brain) nor 1,2-diacylglycerol (brain) had any significant inhibitory effect. (Fig. 5, B and D).


Figure 5: Effect of lipids on CDase activities. The purified species (each approximately 0.09 µg of protein) were assayed in the presence or absence of lipids, which were sonicated separately and added to the reaction mixture before the assay as described under ``Experimental Procedures.'' The control values are expressed as 100% for specific activity of 1.20 nmol/min (CDase-I) and 0.97 nmol/min (CDase-II). Results are expressed as means of three separate experiments. PI, ; PE, ; PG, ; PS, ; PA, ; PC-b, ; PC-e, ; diacylglycerol (DG), ; caldiolipin (CAL), ▾; GC, ; BRAIN EXT, +; dimyristorylphosphatidylcholine (DMPC), ; SM-b, ; SM-e, or .



Effects of Lysophospholipids

Fig. 6 shows the effect of five lysophospholipids on the activities of both isozymes. Lysophosphatidic acid (lysoPA) was inhibitory to CDase-I but not CDase-II. Lysophosphatidylethanolamine (lysoPE), lysophosphtidylcholines (lysoPC), or lysophosphatidylserine (lysoPS) did not affect the enzyme activity.


Figure 6: Effects of lysophospholipids on CDase activities. The CDase species (approximately 0.10 µg of protein each) were assayed in the presence or absence of lysolipids, which were sonicated separately and added to the reaction mixture before the assay. The control values are expressed as 100% for specific activity of 1.32 nmol/min (CDase-I) and 1.08 nmol/min (CDase-II). The reaction proceeded as described under ``Experimental Procedures.'' Results are expressed as means of three separate experiments. LysoPC-e, ; lysoPC-b, ; lysoPE, ; lysoPS, ; lysoPA, .



Effects of Sphingosine

The effects of sphingosine upon the CDases were examined (Fig. 7). The addition of 0.05-1 mM of sphingosine (bovine brain) significantly suppressed the catalytic activities of both CDase-I and -II. The two species had almost the same sensitivity to sphingosine with an IC of about 100 µM (CDase-I) and 130 µM (CDase-II), respectively. The mode of product inhibition by sphingosine in palmitoylsphingosine hydrolysis by these two CDase species was competitive (Fig. 8, A and B). When palmitoylsphingosine was used as the substrate, the apparent Kinetic inhibitory parameters (K ) determined were about 260 µM (CDase-I; K = 90 µM, V = 0.62 unit) and 225 µM (CDase-II; K = 140 µM, V = 0.50 unit).


Figure 7: Effects of sphingosine on CDase activities. The CDase species were assayed in the presence of varying concentrations of sphingosine (bovine brain), which was sonicated separately and added to the reaction mixture before the assay. The reaction proceeded as described under ``Experimental Procedures.'' , CDase-I activity. , CDase-II activity. Results are expressed as means from three separate experiments.




Figure 8: Effects of sphingosine on CDase activities according to double-reciprocal Lineweaver-Burk plot analysis. The ceramide hydrolysis activities of CDase species were measured in the presence () or absence () of varying concentrations of sphingosine (bovine brain), which was sonicated separately and added to the reaction mixture before the assay. The reaction proceeded as described under ``Experimental Procedures.'' A, Lineweaver-Burk plots of CDase-I. B, Lineweaver-Burk plots of CDase-II. Results are expressed as means of three separate experiments.



Effects of pH

When palmitoylsphingosine was used as the substrate, the highest activities were observed at pH 7.0-10.0 and 8.0-9.0, respectively, in the presence of 0.5% sodium cholate (Fig. 9). Even when the each finally purified enzyme solution was dialyzed for 24 h against 50 mM Tris-HCl buffer, pH 7.4, containing 0.5% CHAPS, the pH optima of each isozyme was not affected (data not shown).


Figure 9: Effects of pH on CDase activities. Assays contained 125 mM acetate (, pH 3.0-6.0), or phosphate (, pH 6.0-8.0), Tris-HCl (, pH 8.0-10.0), or borate buffers (, pH 9.0-11.0). The reaction proceeded as described under ``Experimental Procedures.'' Results are expressed as means of two separate experiments.




DISCUSSION

Sphingolipid metabolism is a key cellular event involved in the regulation of cell proliferation and differentiation in many tissues (6-8). Ceramide is involved as a structural and functional component in the sphingolipid metabolism. As evidence is accumulating that ceramide plays an important role as an intracellular effector molecule (32, 33), enzymes that regulate metabolism of ceramide stand as potential regulators of ceramide levels and consequently ceramide-mediated function. Among them, the process of ceramide breakdown mediated by CDase seems important in controlling the cellular level of a series of sphingolipid metabolites. Recent evidence suggests that cellular levels of ceramides are deeply associated with stimulation of the phosphorylation of several kinases, the activity of protein kinase, the levels of the c-myc protooncogene, the activity of phospholipase A, and prostagalandin release (17-19), resulting in a significant modification of several cellular functions. Thus, ceramide analogs exert specific and potent antiproliferative effects in HL-60 cells at concentrations as low as 1-10 µM, mimicking the action of tumor necrosis factor , interleukin-1, 25-dihydroxyvitamin D, and -interferon on HL-60 cells (32) . They are also active against other leukemia cells, malignant cells in tissue culture, and normal fibroblasts in logarithmic phase of growth (for review, see Ref. 34). In myeloid, lymphoid cells, and fibroblasts, ceramide analogs caused early, potent, and specific internucleosomal DNA fragmentation, a hallmark of apoptosis (35) , suggesting that ceramide mediates the effects of tumor necrosis factor on programmed cell death and participate in other events of apoptosis, as evidenced by the fact that ceramide levels are significantly increased in HIV-infected T lymphocytes undergoing apoptosis. Other studies raise the possibility that ceramide may function as a regulator of protein trafficking in which C-ceramide inhibits secretion of vesicular stomatitis virus glycoprotein from infected Chinese hamster ovary cells (36). Studies in human fibroblasts suggest that ceramide may play a role in modulation of immune function and inflammatory response by modulating secretion of prostaglandin E in response to the action of interleukin-1 (37) . On the other hand, the hydrolysis product, sphingosine is also known to affect a wide variety of biological activities (38) . Sphingosine may exert many of its biological effects through its ability to inhibit protein kinase C activity. In addition, a role for sphingosine in PI turnover has been suggested in which it stimulates PI hydrolysis by stimulating phospholipase C activity and may represent a pathway by which it may exert some of its many effects upon cell function. Based upon evidence that sphingosine and ceramide play an important role in such fundamental biological processes as cell proliferation, differentiation, receptor function, and oncogenesis, the biological analysis and the enzymatic regulation mechanisms of ceramide hydrolysis and sphingosine producing enzyme, CDases should provide insight into the role of sphingolipid metabolites as the ultimate endogenous control molecules for maintaining tissue homeostasis.

Our present studies are the first to describe the extensive purification of an enzyme utilizing ceramide in mammalian tissues. We demonstrated that at least two CDase species with different molecular masses and enzymatic properties are present in guinea pig skin. For the first time, of the two, one designated herein as CDase-I, was purified to apparent homogeneity in mammalian tissues (Fig. 4B). CDase activity was first described by Gatt (1) , who characterized and partially purified the enzyme from rat brain. Although there are a few reports of the partial purification of membrane-bound CDase isozymes from human spleen (5, 39, 40) , the enzymatic and regulatory properties of CDase remain largely unknown. Al et al. have reported an acid CDase activity partially isolated from human spleen, which has an molecular mass of about 100 kDa. On the other hand, Nilsson (3) described a neutral CDase in human small intestine. Sugita et al. (4) reported an alkaline CDase activity in the human cerebellum. However, the molecular mass of these CDases was unknown, and then biochemical properties of these CDase proteins did not coincide with those of epidermal isozymes. Therefore, the two CDase species purified from guinea pig epidermis in the present study appear to be unique for enzymological and biological aspects of ceramide metabolism.

In epidermal tissue, Wertz and Downing (27) reported only the existence of enzymatic ceramide hydrolytic activity in porcine epidermis, but the examination of enzymatic properties was not sufficient to clarify the biochemical properties because the enzymatic characterization, including the pH profile or the K for ceramide, revealed the possible additive or mixed effects of two or more CDase species. In this study, we biochemically and kinetically characterized two membrane-bound CDase species as the terminal enzyme of sphingolipid metabolism. The specific activity of ceramide hydrolysis in guinea pig epidermis homogenate (5.05 units/mg protein) was higher than that of brain (3.24 units/mg protein), suggesting that CDase activity has an important role in epidermal metabolism, because in addition to the cellular action as biochemical effector molecules, ceramides themselves are the principal differentiation products in epidermis and the major constituents of the stratum corneum lipids which serve the epidermal barrier against water loss (12) . Furthermore, the long chain base sphingosine is a potent endogenous inhibitor of protein kinase C in vitro with the ability to inhibit the proliferation of germitive epidermal cells (17, 18, 19) . The injured stratum corneum may facilitate the mutal contact between its main lipid constituent, ceramides, and living epidermal cells, probably leading to the stimulation of reproduction of keratinocytes, as evidenced by our preliminary study in that in contrast to the inhibitory effect of short chain base alkyl acyl sphingosines (41) , authentic ceramides elicit a marked stimulation of the proliferation of human keratinocytes (data not shown). Because our study demonstrated that a high specific activity of CDase is also localized in the stratum corneum, the uppermost layer of the epidermis, it is conceivable that under such injured conditions, the function of CDase in the stratum corneum becomes crucial in reducing the biochemical activity of the ceramide by hydrolyzing it and subsequently in abrogating the activated cellular events by liberating sphingosine. The fact that two CDase species in guinea pig epidermis were suppressed by the sphingosine (Fig. 7) suggests that ceramide hydrolysis liberating free sphingosine could also provide a feedback mechanism for regulating ceramide breakdown, supporting the notion that CDase activities are controlled by the balance of substrate degradation and production. Mass contents of ceramides exist at least 50-100-fold greater than those of sphingosine bases in mammalian epidermis, and Wertz et al.(42, 43, 44) have recently reported that ceramides serve about 40% of the total lipid in the stratum corneum, while sphingosine bases account for about 0.44% in same portion (42, 43, 44) , indicating that control metabolism underlying the well regulated balance of ceramide/sphingosine may be the important physiological phenomena in epidermal keratinaization process. Our study also demonstrated that several lipids, in particular SM and PC, markedly inhibited the purified enzyme activities in the presence of sodium cholate (Fig. 5, A-D). SM is metabolized by SMase to yield ceramides and PC in the upper subcorneal layers where ceramides are essential components to subsequently form bilayer of lipids in the intercellular spaces between stratum corneum cells for serving barrier and water-holding function. Therefore, the inhibitory effects by SM and PC can be viewed as precursor suppression for preventing ceramides from prior degradation, although we could not conclude whether or not these lipids affected the enzymes or the ceramides.

Our study revealed that major CDases present in the guinea pig epidermis are alkaline CDases with the optimal pH of 7.0-9.0, while this is the case for the stratum corneum, the upper layers of the skin. On the other hand, the surface pH of mammalian skin is maintained in acidic side with human skin being at pH 4.2-5.6 (45) and that of guinea pig skin shows pH 5.0-6.5. It is, therefore, likely that the epidermal alkaline CDase present in the stratum corneum does not function effectively in normal physiological conditions. However, under some barrier-disturbed conditions where living layer (having alkaline side of pH) can get in contact with the stratum corneum, CDase may perform the function to degrade ceramides. On the other hand, Anderson (46) has reported that the surface pH of some dermatitis, for example atopic dermatitis or ichthyosis, is observed on alkaline side (46) . Although the alteration of surface pH may relate with the depression of barrier function or the abnormality of lipid metabolism in these disorders, they would provide a physiological basis for activating CDase, leading to the deficiency in the mass of ceramides and the abnormalities in keratinization process due to the loss of balance between ceramides and sphingosine.

In conclusion, our present study elucidated the biochemical properties of the epidermal CDases which regulate the quantity of sphingolipid metabolites involved as regulatory molecules in cell growth and differentiation during epidermal keratinization process. The regulatory features of epidermal CDases through the enzyme-associated sphingolipid metabolites are suggestive of the notion that sphingolipid metabolism undergoes the strictly controlled regulations in epidermal tissue through the product feedback and precursor suppression mechanisms. Based upon the presence of CDase species in epidermal tissue, it would be of importance to define the participation of each CDase species in the regulation of cell growth or keratinization in the epidermis, and further characterization of each is required for clarifying their roles in regulating sphingolipid metabolism.

  
Table: Purification of CDase isozymes from guinea pig epidermis

The ceramidase activities at each step were determined using [1-C]palmitoylsphingosine as a substrate as described under ``Experimental Procedures.''


  
Table: Substrate specificity of CDase species

The CDase activities were compared at pH 4.5 and 9.0 among palmitoylsphingosine, linoleoylsphingosine, and oleoylsphingosine as substrates. The activity was expressed as disintegrations/min (dpm) of each released radioactive free fatty acid by subtracting background disintegrations/min (dpm) (approximately 10,000 disintegrations/min (dpm)). The percent value represents relative activity for each substrate when the hydrolysis value for palmitoylsphingosine is taken as 100%. Results are expressed as means of two separate experiments.



FOOTNOTES

*
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Institute for Fundamental Research, Kao Corporation, 2606 Ichikaimachi, Akabane, Haga, Tochigi 321-34, Japan. Tel.: 81-285-68-7467; Fax: 81-285-68-7469.

The abbreviations used are: CDase, ceramidase; Fraction I, pass-through fraction; Fraction II, eluted fraction; PAGE, polyacrylamide gel electrophoresis; PC-e, phosphatidylcholine (egg origin); PC-b, phosphatidylcholine (brain origin); SM-e, sphingomyelin (egg origin); SM-b, sphingomyelin (brain origin); PG, phosphatidylglycerol; PI, phosphatidylinositol; PE, phosphatidylethanolamine; lysoPC-e, lysophosphatidylcholine (egg origin); lysoPC-b, lyso-phosphatidylcholine (brain origin); IEF, isoelectric focusing; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid.


REFERENCES
  1. Gatt, S.(1966) J. Biol. Chem. 241, 3724-3730 [Abstract/Free Full Text]
  2. Yavin, E., and Gatt, S.(1969) Biochemistry 8, 1692-1698 [Medline] [Order article via Infotrieve]
  3. Nilsson, A.(1969) Biochim. Biophys. Acta 176, 339-347 [Medline] [Order article via Infotrieve]
  4. Sugita, M., Williams, M., Dulaney, J. T., and Moser, H. W.(1975) Biochim. Biophys. Acta 398, 125-131 [Medline] [Order article via Infotrieve]
  5. Al, B. J. M., Tiffany, C. W., Gomes de Mesquita, D. S., Moser H. W., Tager, J. M., and Schram, A. W.(1989) Biochim. Biophys. Acta 1004, 245-251 [Medline] [Order article via Infotrieve]
  6. Bremer, E. G., Schlessinger, J., and Hakomori, S.-I.(1986) J. Biol. Chem. 261, 2434-2440 [Abstract/Free Full Text]
  7. Kolesnick, R. N., Hemer, M.(1990) J. Biol. Chem. 265, 18803-18808 [Abstract/Free Full Text]
  8. Yada, Y., Okano, Y., and Nozawa, Y.(1991) Biochem. J. 279, 665-670 [Medline] [Order article via Infotrieve]
  9. Okazaki, T., Bell, R. M., and Hannun, Y. A.(1989) J. Biol. Chem. 264, 19076-19080 [Abstract/Free Full Text]
  10. Okazaki, T., Bielawaska, A., Bell, R. M., and Hannun, Y. A.(1990) 265, 15823-15831
  11. Obeid, L. M., Linardic, C. M., Karolak, L. A., and Hannun, Y. A.(1993) Science 259, 1769-1771 [Medline] [Order article via Infotrieve]
  12. Elias, P. M., and Friend, D. S.(1975) J. Cell Biol. 65, 180-191 [Abstract]
  13. Melton, J. L., Wertz, P. W., Swartzendruber, D. C., and Downing, D. T. (1987) Biochim. Biophys. Acta 921, 191-197 [Medline] [Order article via Infotrieve]
  14. Imokawa, G., and Hattori, M.(1985) J. Invest. Dermatol. 83, 282-284
  15. Imokawa, G., Akasaki, S., Hattori, M., and Yoshizuka, N.(1986) J. Invest. Dermatol. 87, 758-761 [Abstract]
  16. Goldkorn, T., Dressler, K. A., Muindi, J., Radin, N. S., Mendelsohn, J., Menaldino, D., Liotta, D., and Kolesnick, R. N.(1991) J. Biol. Chem. 266, 16092-16097 [Abstract/Free Full Text]
  17. Wilson, E., Olcott, M. C., Bell, R. M., Merrill, A. H., Jr., and Lambeth, J. D(1986) J. Biol. Chem. 261, 12616-12623 [Abstract/Free Full Text]
  18. Oishi, K., Raynor, R. L., Charp, P. A., and Kuo, J. F.(1988) J. Biol. Chem. 263, 6865-6871 [Abstract/Free Full Text]
  19. El Touny, S., Khan, W., and Hannun, Y.(1990) J. Biol. Chem. 265, 16437-16443 [Abstract/Free Full Text]
  20. Stevens, V. L., Winton, E. F., Smith, E. E., Qwens, N. E., Kinkade, J. M., Jr., and Merrill, A. H., Jr.(1989) Cancer Res. 49, 3229-3234 [Abstract]
  21. Hannun, Y., Loomis, C. R., Merrill, A., and Bell, R. M.(1986) J. Biol. Chem. 261, 12604-12609 [Abstract/Free Full Text]
  22. Nutter, L. M., Grill, S. P., Li, J. S., Tan, R. S., and Cheng, Y. C. (1987) Cancer Res. 47, 4407-4412 [Abstract]
  23. Merrill, A. H., Jr., Sereni, A. M., Stevens, V. L., Hannun, Y. A., Bell, R. M., and Kinkade, J. M., Jr.(1986) J. Biol. Chem. 261, 12610-12615 [Abstract/Free Full Text]
  24. Koval, M., and Pagano, R. E.(1991) Biochim. Biophys. Acta 1082, 113-125 [Medline] [Order article via Infotrieve]
  25. Merrill, A. H., Jr.(1991) J. Bioener. Biomem. 23, 83-104 [Medline] [Order article via Infotrieve]
  26. Kolesnick, R. N.(1991) Prog. Lipid Res. 30, 1-38 [CrossRef][Medline] [Order article via Infotrieve]
  27. Wertz, P. W., and Downing, D. T.(1990) FEBS Lett. 268, 110-112 [CrossRef][Medline] [Order article via Infotrieve]
  28. Shapiro, D., and Flowers, H. M.(1961) J. Am. Chem. Soc. 83, 3327-3331
  29. Dole, V. P. J.(1956) Clin. Invest. 35, 350-353
  30. Fujino, Y., and Ito, S.(1968) Biochim. Biophys. Acta 152, 627-629 [Medline] [Order article via Infotrieve]
  31. Laemmli, U. K.(1970) Nature 227, 680-685 [Medline] [Order article via Infotrieve]
  32. Okazaki, T., Bielawska, A., Bell, R. M., and Hannun, Y. A.(1990) J. Biol. Chem. 265, 15823-15831 [Abstract/Free Full Text]
  33. Merrill, A. H., Jr.(1992) Nutr. Rev. 50, 78-80 [Medline] [Order article via Infotrieve]
  34. Hannun, Y. A.(1994) J. Biol. Chem. 269, 3125-3128 [Free Full Text]
  35. Obeid, L. M., Linardic, C. M., Karolak, L. A., and Hannun, Y. A.(1993) Science 259, 1769-1771 [Medline] [Order article via Infotrieve]
  36. Rosenwald, A. G., and Pagano, R. E.(1993) J. Biol. Chem. 268, 4577-4579 [Abstract/Free Full Text]
  37. Ballou, L. R., Chao, C. P., Kolness, M. A., Barker, S. C., and Raghow, R.(1992) J. Biol. Chem. 267, 20044-20050 [Abstract/Free Full Text]
  38. Ballou, L. R.(1992) Immunol. Today 13, 339-341 [Medline] [Order article via Infotrieve]
  39. Maret, A., Potier, M., Salvayre, R., and Douste-Blazy, L.(1983) FEBS Lett. 160, 93-97 [CrossRef][Medline] [Order article via Infotrieve]
  40. Maret, A., Potier, M., Salvayre, R., and Douste-Blazy, L.(1984) Biochim. Biophys. Acta 799, 91-94 [Medline] [Order article via Infotrieve]
  41. Merrill, A. H., Jr., and Stevens, V. L.(1989) Biochim. Biophys. Acta 1010, 131-139 [Medline] [Order article via Infotrieve]
  42. Wertz, P. W., Swartzendruber, D. C., Madison, K. C., and Downing, D. T. (1987) J. Invest. Dermatol. 89, 419-425 [Abstract]
  43. Wertz, P. W., and Downing, D. T.(1989) Biochim. Biophys. Acta 1002, 213-217 [Medline] [Order article via Infotrieve]
  44. Wertz, P. W., and Downing, D. T.(1990) J. Invest. Dermatol. 94, 159-161 [Abstract]
  45. Blank, I. H.(1939) J. Invest. Dermatol. 2, 231-235
  46. Anderson, D. S.(1951) J. Dermatol. 63, 283-287

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