Purification and characterization of a second type of neutral ceramidase from rat brain: A second more hydrophobic form of rat brain ceramidase

Biochimica et Biophysica Acta 1811 (2011) 242–252

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Purification and characterization of a second type of neutral ceramidase from rat brain: A second more hydrophobic form of rat brain ceramidase Faisal Thayyullathil a, Shahanas Chathoth a, Abdulkader Hago a, Mahendra Patel a, Zdzislaw M. Szulc b, Yusuf Hannun b, Sehamuddin Galadari a,⁎ a b

Cell Signaling Laboratory, Department of Biochemistry, Faculty of Medicine and Health Science, UEA University, PO Box 17666, Al- Ain, UAE Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, SC 28425, USA

a r t i c l e

i n f o

Article history: Received 5 September 2010 Received in revised form 29 November 2010 Accepted 26 December 2010 Available online 9 January 2011 Keywords: Ceramidase Ceramide Sphingosine Chromatography Glycosidase F

a b s t r a c t Ceramidases (CDase) are enzymes that catalyze the hydrolysis of N-acyl linkage of ceramide (Cer) to generate sphingosine and free fatty acids. In this study we report the purification and characterization of a novel second type of neutral ceramidase from rat brain (RBCDase II). Triton X-100 protein extract from rat brain membrane was purified sequentially using Q-Sepharose, HiLoad16/60 Superdex 200 pg, heparin-Sepharose, phenylSepharose HP, and Mono Q columns. After Mono Q, the specific activity of the enzyme increased by ~ 15,000fold over that of the rat brain homogenate. This enzyme has pH optima of 7.5, and it has a larger apparent molecular weight (110 kDa) than the previously purified (90 kDa) and characterized neutral rat brain CDase (RBCDase I). De-glycosylation experiments show that the differences in molecular mass of RBCDase I and II on SDS-PAGE are not due to the heterogeneity with N-glycan. RBCDase II is partially stimulated by Ca2+ and is inhibited by pyrimidine mono nucleotides such as TMP and UMP. This finding is significant as it demonstrates for the first time an effect by nucleotides on a CDase activity. The enzyme was also inhibited by both oxidized and reduced GSH. The effects of metal ions were examined, and we found that the enzyme is very sensitive to Hg2+ and Fe3+, while it is not affected by Mn2+. EDTA was somewhat inhibitory at a 20 mM concentration. © 2011 Published by Elsevier B.V.

1. Introduction Ceramidase (CDase, EC3.5.1.23) is an enzyme that catalyzes the hydrolysis of N-acyl linkage of ceramide (Cer) to generate free fatty acids and sphingosine (SPH). Several lines of investigation have clearly established Cer as a mediator of the eukaryotic stress response [1]. Moreover, Cer has been implicated in differentiation, [2], cell cycle arrest [3], apoptosis [4], senescence [5], and in autophagy [6], in several cell types. Sphingosine is not produced by de novo synthesis, and the main source of its cellular generation is by the action of CDase on Cer [7,8]. In turn, SPH has been emerged as a novel sphingolipid second messenger. Several lines of evidence indicated that SPH could modulate cell's fate via inhibiting protein kinase C [9], modulation of

Abbreviations: RBCDase I, rat brain ceramidase 1; RBCDase II, rat brain ceramidase II; Cer, ceramide; SPH, sphingosine; S-1-P, sphingosine-1-phosphate; C12-NBD- Cer, 4nitrobenzo-2- oxa-l,3-diazole ceramide; AMP, adenosine monophosphate; ADP, adenosine diphosphate; ATP, adenosine triphosphate; GPM, guanosine monophosphate; GDP, guanosine diphosphate; GTP, guanosine triphosphate; TMP, thymidine monophosphate; TDP, thymidine diphosphate; TTP, thymidine triphosphate; UMP, uridine monophosphate; UDP, uridine diphosphate; UTP, uridine triphosphate; SDSPAGE, sodium dodicyl suphate polyacrylamide gel electrophoresis; PA, phosphatic acid; PC, phosphatidylcholine; PG, phosphatidylglycerol; PE, phosphatidylethanolamine; PI, phosphatidylinositol; PS, phosphatidylserine ⁎ Corresponding author. Tel.: +97 137137507; fax: +97 137672033. E-mail address: [email protected] (S. Galadari). 1388-1981/$ – see front matter © 2011 Published by Elsevier B.V. doi:10.1016/j.bbalip.2010.12.012

Na+, K+-ATPase [10], activation of p38 MAPK [11], inhibition of p44ERK1/p42-ERK2 [12,13], and caspase 3 activation [14,15]. Furthermore, sphingosine-1-phosphate (S-1-P), a phosphorylated derivative of sphingosine generated by the action of sphingosine kinase [16], is found to be involved in diverse biological processes including cell growth, cell survival, and cell motility [17–19]. Thus, CDase activity plays an important role in the regulation of the level of the bioactive lipids Cer, the intermediate product SPH, and its phosphorylated product S-1-P. Based on the pH optima and primary sequence, CDases are classified in to three groups: acid, neutral, and alkaline CDases. Acid CDase has an acidic pH optimum and is responsible for Faber disease, in which Cer is accumulated in lysosomes due to deficiency of the enzyme [20]. Acid CDase was purified from human urine [21], and cDNAs encoding the enzyme were cloned from human and mouse [22]. Alkaline CDase, which prefers phyto-ceramide over normal Cer containing sphingenine and which shows alkaline pH optimum, is found in yeast and human. Recently, several alkaline CDases were cloned and characterized, including phytoCDase and dihydroCDase from yeast Saccharomyces cerevisiae [23,24] and a homolog of phytoCDase from human [25]. Recently, a human alkaline ceramidase 2 (ACER2), a Golgi enzyme, regulates the maturation of integrin β1 subunit by controlling the generation of sphingosine [26]. Neutral CDases (nCDases) have also been purified and biochemically characterized from various tissues such as mouse liver [27], rat

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kidney [28], brain [29], intestine [30], and human intestine [31]. Recently, presence of a novel amidase motif present in the human neutral CDase has been identified [32]. Additionally, nCDases have been cloned from Pseudomonas aeruginosa [33], slime mould [34], Drosophila [35], zebra fish [36], rat [28], mouse [37], and human [38]. Pata et al. have recently reported the molecular cloning and characterization of a plant CDase from rice (Oriza sativa spp.) [39]. Mammalian nCDase appears to play a major role in cell proliferation and apoptosis by regulating the levels of the bioactive lipids ceramide, SPH, and S-1-P. Several reports have shown the involvement of nCDase in the regulation of ceramide, SPH, and/or S-1-P levels in agonistmediated cell responses. Activation of nCDase leading to an increase in SPH and/or S-1-P levels and to responses associated with these lipids has been shown in rat glomerular mesangial cells stimulated with platelet-derived growth factor (PDGF) [40], in rat hepatocytes stimulated with low concentration of interleukin 1 [41], in rat mesangial cells stimulated with nitric oxide donors [42], and in vascular smooth muscle cells treated with oxidized low-density lipoprotein [43]. On the other hand, studies using inhibitors of ceramidase (N-oleolylethanolamine and D-erythro-2-(N-myristoylamino)-1-phenyl-1propanol) have also shown that inhibition of these enzymes causes an elevation in the endogenous level of ceramide, which is either sufficient to inhibit growth or augments the effect of other inducers of growth arrest [44]. Recently, it has been reported that targeted expression of nCDase rescued the retinal degradation in arrestin and phospholipase C mutants of Drosophila through modulation of endocytosis of rhodopsin in photoreceptors [45,46]. Moreover, it was also reported that the gene knockdown of zebra fish nCDase during embryogenesis results in a defect of blood cell circulation [36]. Taken together, these observations underscore the potential importance of CDases and their roles in cellular signaling of biological processes such as apoptosis and cell proliferation. In this study, we describe the purification and characterization of a second type of neutral CDase from rat brain having a molecular weight of 110 kDa. The RBCDase II, which is described in this study, is novel and different from the previously purified RBCDase I [29] in several aspects: molecular mass, glycosylation state, pH optima, metal dependence, and phospholipid dependence.

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25 μl of 100 μM of C12-NBD-Cer, 20 ng the enzyme (in 10 μl) and the reaction volume was adjusted to 50 μl with 50 mM Tris buffer (pH 7.5) containing 0.3% Triton X-100. The reaction was stopped by adding 100 μl chloroform/methanol (1:1). After drying in a speed vacuum concentrator (Savant Instruments, Inc.), the sample was dissolved in 25 μl of chloroform/methanol (2:1) and applied to a TLC plate, which was developed with chloroform, methanol, and ammonia (75:15:0.9). The spot corresponding to NBD-dodecanoic acid and C12-NBD-Cer were scraped, incubated with ethanol at 37 °C for 5 min to extract the compounds from Silica and their fluorescence was measured at (485/ 535 nm) excitation/emission wave length in a Perkin-Elmer Spectrofluorophotometer. The compounds were quantified using a standard curve of known amounts of C12-NBD-Cer and NBD-dodecanoic acid. One enzyme unit is defined as the amount capable of catalyzing the release of 1 μmol of NBD-dodecanoic acid/min from C12-NBD-Cer. For the pH optimum determination, the substrate was dissolved in the following buffers: pH 3–5, 100 mM acetate buffer; pH 6–7, 100 mM phosphate buffer; pH 7–8, 100 mM Tris or Hepes buffer and pH 8–10, 100 mM glycine buffer. In some experiments, HPLC assay for CDase was used to quantitate the release of sphigoid base as described previously with little modification [48]. Ceramide species were dissolved in 50 mM Tris (pH–7.4) containing 0.4% IGEPAL CA 630. The final concentration of IGEPAL CA 630 in the assay was 0.2%. The reaction was started by adding 20 ng enzyme (10 μl), and the incubation time was for 1 h at 37 °C. The reaction was stopped by adding 55 μl of stopping buffer (1:9, 0.07 M potassium hydrogen phosphate buffer: methanol). The released SPH was derivatized with o-phthaladehyde (OPA) reagent. After stopping the reaction add 25 μl of freshly prepared OPA reagent (12.5 mg OPA dissolved in 250 μl ethanol and 12.5 μl β-mercaptoethanol and made up to 12.5 ml with 3% (w/v) boric acid). The mixture was allowed to stand for 30 min. An aliquot of 25 μl injected in the HPLC. HPLC analysis was done using Waters 1525 binary pump system. Waters XTerra C18 column (5 μm, 3.9 mm× 150 mm) was equilibrated with a mobile phase (20% methanol, 80% 1:9, 0.07 M potassium hydrogen phosphate buffer: methanol) at a flow rate of 0.5 ml/min. The fluorescence detector (Waters 2475) was set at an excitation wavelength of 340 nm and an emission wavelength of 455 nm.

2. Experimental procedures 2.3. Fractionations and Triton X-100 extraction 2.1. Materials Hitrap Q-Sepharose high-performance HiLoad16/60 Superdex 200 pg, Hitrap heparin-Sepharose, phenyl-Sepharose HP, Mono Q, and PD-10 column were purchased from GE Healthcare (Uppsala, Sweden). Centriprep and Centricon sample concentrators were from Amicon Inc. (Beverly, MA 01915 USA). D-erythro-C12-NBD-ceramide was kindly provided by the Lipidomics Core Facility at the Medical University of South Carolina. All other lipids were from Avanti polar lipids. Pre-coated Silica Gel 60 TLC plates were obtained from Whatmann (Germany). Rabbit polyclonal anti-nCDase antibodies were generated at the MUSC antibody facility (Charleston, SC, USA), against the peptide sequence: KNRGYLPGQGPFVANFA. This peptide sequence was deduced as described previously [47]. Goat anti-rabbit horseradish peroxidase-conjugated secondary antibody was from sigma (St. Louis, MO, USA). All SDS-PAGE reagents were purchased from Bio-Rad (USA). Silver staining kits, Gel Code Blue staining kit, Triton X-100, Extracti-Gel D detergent removing gel, BCA proteins assay kits, and ECL were from Pierce (Rockford. IL, USA). Glycosidase F was purchased from Calbiochem (USA). All other chemicals used were purchased from Sigma (St. Louis, MI, USA). 2.2. Neutral CDase enzyme assay and biochemical characterization CDase activity was measured by using C12-NBD-Cer as a substrate as described previously [32]. Briefly, the reaction mixture contained

Tissue fractionation and Triton X-100 extraction were carried out as previously described [29]. Briefly, 30–40 frozen Wistar rat brains (provided by animal facility at Faculty of Medicine and Health sciences, United Arab Emirate University) were thawed and homogenized in homogenization buffer (150 ml of 20 mM cold phosphate buffer (pH 7.4) containing 0.25 M sucrose, 1 mM EDTA, and 0.2 mM phenylmethylsulphonyl fluoride). The homogenate was centrifuged at 1000×g for 10 min, and the pellet of this centrifugation was homogenized further using 100 ml of homogenization buffer. After centrifugation at 1000×g for 10 min the pellet was washed twice with 75 ml homogenization buffer and all supernatants were combined (the post nuclear supernatant fraction). The post nuclear supernatant fraction was then centrifuged at 10,000×g for 30 min and pellet of this centrifugation was resuspended in solubilization buffer (150 ml of 20 mM Tris buffer (pH 7.4), 1 mM EDTA, 0.2 mM phenylmethylsulfony fluoride, 0.5% Triton X-100). After mixing for 1 h, the Triton X-100 solubilized fraction was obtained by centrifugation of the mixture at 10,000×g for 30 min. The supernatant (Triton X-100 extract) was used as a source for ceramidase purification. All the steps were carried out at 4 °C. 2.3.1. Q-Sepharose The Triton X-100 extract (148 ml) was applied to Hitrap Q-Sepharose high-performance column (25 ml) equilibrated with buffer A (20 mM Tri, pH 7.4, 1 mM EDTA, 0.2 mM phenyl methyl sulfonyfluoride, 0.005%

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Triton X-100) at 1 ml/min. The unbound proteins were eluted by washing the column with 75 ml of buffer A. The bound proteins were then eluted with a 200 ml linear gradient of NaCl from 0 to 0.3 M in buffer A. The salt concentration was then increased to 1.5 M for 100 ml (buffer B1: buffer A + 1.5 M NaCl). After washing the column with B1 buffer the column was again washed with A1 buffer for 50 ml, and then the tightly bound proteins were eluted by a linear gradient of buffer B2 (B1 + 0.5% Triton X-100) to wash out tightly bound proteins. 5 ml fractions were collected, ceramidase activity was measured in these fractions (Triton X-100 fractions), and fractions with a peak activity were pooled. 2.3.2. HiLoad16/60 Superdex 200 pg The pooled Q-Sepharose fractions were concentrated with centriprep and then load on to Hi Load 16/60 superdex 200 pg column equilibrated with buffer A at flow rate of 0.2 ml/min and 1 ml were collected. Fractions containing peak activity were pooled. 2.3.3. Heparin Sepharose The active CDase fractions from Hi Load superdex 200 pg were collected together and then applied to Hitrap heparin column equilibrated with A1 buffer at a flow rate of 1 ml/min. After sample loading, the column was washed with 50 ml of buffer A followed by a 60 ml linear gradient of 0–0.75 M NaCl in buffer A1. Then the salt concentration was increased to 1.5 M for 30 ml. Fractions of 1 ml were collected and those containing CDase activity were pooled. 2.3.4. Phenyl Sepharose HP The fractions of Hitrap heparin were diluted twice with A1 buffer and adjusted to 1.5 M NaCl and then applied to phenyl Sepharose HP (5 ml) column equilibrated with buffer B (buffer A + 1.5 M NaCl) at a flow rate 0.2 ml/min. After the sample was applied, the flow rate was increased to 0.5 ml/min, and the column was washed with decreasing concentration of buffer B (1.5–0.3 M). A stepwise elution was applied using 35 ml of buffer C (buffer A plus 0.3 M NaCl) then 35 ml of 10 mM Tris (pH 7.5) and 1 mM Tris (pH 7.5). Finally 50 ml gradient from 0 to 1% Triton X-100 in 10 mM Tris (pH 7.5) was applied. Fractions of 1 ml were collected, and ceramidase activity was measured. Fractions containing CDase activity (recovered in the Triton X-100 gradient) were combined. 2.3.5. Mono Q The pooled fractions from the phenyl Sepharose high performance were concentrated by using centriprep centricon, and the Triton X-100 was removed by Extracti-Gel D detergent removing gel and then diluted three times with buffer A and applied to a Mono Q column (1 ml) equilibrated with buffer A at a flow rate of 1 ml/min. After washing the column with 5 ml of buffer A to remove unbound protein, ceramidase activity was eluted with a 20 ml linear gradient of NaCl (from 0 to 0.6 M), and then the column was finally washed with 15 ml of 1.5 M NaCl in buffer B. One milliliter fractions were collected, and ceramidase activity was measured in this fraction. 2.4. Protein assay and SDS-PAGE Protein concentration was determined using the Bradford assay or the BCA assay for samples containing Triton X-100. SDS-PAGE of reducing and non-reducing condition was carried out according to the method of Laemmli [49]. The proteins were visualized by staining with silver staining or Gel Code Blue staining. 2.5. Western blotting analysis The purified enzyme preparation was subjected to SDS-PAGE (8% or 10%). The separated protein transferred electrophoretically on to a nitrocellulose membrane. After blocking with 5% non-fat milk in Tris-

buffered saline containing 0.1% Tween 20, the membrane was incubated with anti-nCDase antibody followed by secondary antibody conjugated horseradish peroxidase. Proteins were visualized by using enhanced chemiluminescence system. 2.6. Glycosidase F treatment The glycosylation treatment was performed according to manufacturer's protocol (Calbiochem). Briefly, CDase was denatured in a SDS-PAGE 6× sample buffer for 3 min. The denatured enzyme was then incubated at 37 °C for 18 h with 0.5 milliunit of glycosidase F in the presence of 0.5% Triton X-100. After incubation, the samples were subjected to SDS-PAGE and Western blot analysis using anti-nCDase antibody. 3. Results 3.1. Purification of the neutral rat brain ceramidase II Previously, a second ceramidase activity was detected during the purification of a rat brain membrane bound non-lysosomal ceramidase (RBCDase I). It was reported that this second ceramidase activity was only eluted when Triton X-100 (0.5%) was applied in combination with salt [29]. This activity represented approximately 50–60% of the total activity applied to the Q-Sepharose column. However, it was not determined as to whether this second peak of ceramidase activity was representing a more hydrophobic conformation of the salt-only eluted neutral ceramidase, or that it represented another isoenzyme. Therefore, we named this activity as rat brain neutral ceramidase peak II (RBCDase II), and we decided to purify and characterize this rat brain neutral ceramidase II activity. The purification procedure of RBCDase II is summarized in Table 1. The enzyme was solubilized with 0.5% Triton X-100, and in order to separate ceramidase I and ceramidase II, this solubilized extract was applied to a Q-Sepharose column. Both ceramidase activities bound to this column. The purification procedure using this column was split in two phases; first the well-characterized and identified neutral ceramidase I and other protein contaminants were removed by using a linear shallow NaCl gradient as shown in Fig. 1A. Next, the column was washed with 1.5 M NaCl in order to remove as much unwanted bound proteins as possible. After this wash, a linear gradient of detergent and salt buffer was used to finally elute neutral RBCDase II, at a salt and detergent concentration of 1.5 M NaCl and 0.5% Triton X-100 respectively (Fig. 1A). The peak activity fractions of the Q-Sepharose column were pooled, concentrated and loaded on to a Superdex-200 pg column, and the peak ceramidase activity was eluted at ~110 kDa equivalent molecular weight (Fig. 1B).

Table 1 Purification of rat brain CDase II.a Step

Total protein

Total activity

Mg

milliunits milliunits/mg %

Post nuclear 4176 supernatant Triton X-100 1846 extract Q-Sepharose 520 Superdex 200 pg 320 Heparin Sepharose 98 Phenyl Sepharose 0.48 Mono Q 0.036

Specific activity

4614

1.1

4038

2.1

3864 3456 2081 1239 602

7.4 10.8 21.2 2581 16722

Recovery Purification

100 87.5 83.7 74.9 45.10 26.85 13.05

-fold 1 1.9 6.7 11.6 19.2 2336 15134

a Purification of neutral CDase II from rat brain: RBCDase II was purified from rat brains (70–75 g) as described under Experimental procedures.

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Fig. 1. Purification of RBCDase II. (A) Triton X-100 solubilized fractions were applied to a Q-Sepharose column equilibrated with buffer A. After washing the column, RBCDase I was eluted with a linear gradient from 0 to 0.3 M NaCl in buffer A, and RBCDase II activity was eluted with buffer A containing 1.5 M NaCl and 0.5% Triton X-100. Fractions of 5 ml were collected. (B) The active fractions obtained from the Q-Sepharose (in Triton X-100 gradient) were loaded in to Hi Load 16/60 superdex 200 pg column equilibrated with buffer A at a flow rate of 0.2 ml/min. Fractions of 1 ml were collected. (C) The active fractions obtained from the Hi Load 16/60 superdex 200 pg were applied directly to Hi-Trap heparin equilibrated with buffer A. After washing the column, CDase activity was eluted with a linear gradient of NaCl from 0 to 0.75 M. Fractions of 1 ml were collected. (D) The active fractions obtained from the heparin column were adjusted to 1.5 M NaCl and applied to phenyl Sepharose column equilibrated with buffer B. After washing the column with buffer B and eluting more protein by decreasing the NaCl concentration (see Experimental procedures), CDase activity was eluted with a Triton X-100 gradient (0–1%) in 10 mM Tris buffer, pH 7.5. Fractions of 1 ml were collected. (E) The active phenyl fractions were applied to a Mono Q column equilibrated with buffer A1. CDase activity was eluted with a linear gradient of NaCl from 0 to 0.6 M. Fractions of 1 ml were collected. CDase activity and protein concentration were measured as explained in the Experimental procedures.

The active fractions of the gel filtration step were pooled and then loaded onto a Hitrap heparin-Sepharose column, and the enzyme was eluted with a linear NaCl gradient of 0–0.75 M. The peak CDase activity was eluted at about 0.375 M NaCl (Fig. 1C). The active fractions of the heparin column were combined and adjusted to a salt concentration of 1.5 M. They were then subjected to phenyl Sepharose hydrophobic interaction chromatography on a phenyl Sepharose HP column. The column was washed with decreasing salt concentration, and proteins were eluted, but the CDase activity did not

elute from the column under these conditions (Fig. 1D); however, the enzyme was then eluted with a Triton X-100 gradient (0–1%) in 10 mM Tris buffer as shown in Fig. 1D. In this step a 50-fold increase in the specific activity of the enzyme was obtained. Unlike RBCDase I, the isoelectric point of the RBCDase II was found to be 7.2 by using gel chromatofocusing on a Pharmacia fast gel system (data not shown). Based on this finding, the active phenyl sepharose fractions were subjected to anion exchange chromatography using Mono Q column, and the ceramidase activity was eluted with a linear

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NaCl gradient 0–0.6 M (Fig. 1E). This purification procedure resulted in ~15,000-fold purification (Table 1) of the neutral ceramidase II enzyme to almost homogeneity (Fig. 2A). 3.2. Physical properties of neutral CDase II Gel filtration chromatography of the partially purified neutral CDase II showed an apparent molecular weight of ~ 110 kDa (Fig. 1B), this is confirmed when the pure enzyme was subjected to silver staining as can be seen in Fig. 2A. However, it should be noted that two very faint and minute bands were observed in these fractions that were not possible to remove, and it is likely that they may be degradation products of this final step purification. In order to determine the relationship of RBCDase II to RBCDase I, Western blot analysis was conducted, and it was found that the enzyme reacted robustly with an antibody raised against purified RBCDase I (Fig. 2B). To investigate whether the enzyme is glycosylated, we performed a glycosidase assay on RBCDase II using a Western blot analysis for the visualization of the deglycosylated sample using a specific anti-CDase antibody. Interestingly, the RBCDase II band had shifted, and it now ran as a clear and sharp band at about 95 kDa (Fig. 2C). This result shows that the 110 kDa RBCDase II seems to be a glycoprotein with N-glycan, since the treatment of the enzyme with glycosidase F resulted in the generation of ~95 kDa protein (Fig. 2C, lanes 1 and 2). Indeed, when the same experiment was repeated on the purified RBCDase I, whose apparent molecular weight was found to be ~95 kDa, the molecular weight was reduced to ~70 kDa upon deglycosylation (Fig. 2C, lanes 3 and 4). The molecular mass of deglycosylated RBCDase II is clearly different from that of RBCDase I (Fig. 2C), suggesting that RBCDase II is not a differently glycosylated form of the previously purified RBCDase I. 3.3. Characterization of RBCDase II pH optimum The purified enzyme had a pI value of 7.2 (data not shown) and was active over a relatively sharp pH range with the pH optimum

being at 7 as seen in Fig. 3. An acid activity was observed when acetate buffer was used, although, no such activity was observed with phosphate buffer at acidic pH 5. Moreover, no such secondary activity peak was present on the alkaline pH scale when using glycine as well as Tris buffers (Fig. 3). 3.4. Effects of metal cations The effects of a number of metal ions on the RBCDase II activity were determined, and it was observed that Ca2+ stimulated the enzyme dose dependently, although, in a bell-shaped manner. The maximum stimulation was observed at 5 mM Ca2+concentration, whereby the activity almost doubled; thereafter, the Ca2+-induced activation was attenuated dose dependently up to 10 mM concentration as seen in Fig. 4A. All other cations either exhibited no effect as with Mg2+ and Mn2+, or were dose dependently inhibitory as with Zn2+, Cu2+, Fe3+, and Hg2+. The most potent inhibitor of RBCDase II activity was Hg2+ since it totally inhibited the enzyme activity even at 1 mM concentration. Fe3+ inhibited the enzyme by 50% at 1 mM concentration, and both Zn2+ and Cu2+ were inhibitory in a similar manner but with less potency (Fig. 4A). The monovalent cation Li+ (Fig. 4A) and Na+ (Fig. 4B) did not show any effect on the RBCDase II activity. However, EDTA inhibited the activity to 50% of control at 20 mM concentration (Fig. 4C). 3.5. Effect of reducing agents on enzyme activity Next, the effects of reducing agents on RBCDase II activity were investigated, and both dithiothreitol (DTT) and β-mercaptoethanol (β-ME) dose dependently inhibited neutral CDase activity (Fig. 5A). Although β-ME was not as effective as DTT in inhibiting the enzyme activity (by almost 80% at 100 mM concentration, it inhibited the RBCDase II activity by more than 50% (Fig. 5A). Similarly, glutathione in both the reduced and oxidized form inhibited RBCDase II activity in a dose-dependent manner. Similar effects were also observed with the purified RBCDase I (data not shown). Surprisingly, the inhibition

Fig. 2. SDS-PAGE of Mono Q fractions. (A) SDS-PAGE of the Mono Q fractions was stained with silver staining. (B) Western blot analysis of RBCDase I and II: lane 1 Q-Sepharose fractions of CDase1 (NaCl gradient fractions); lane 2, Q-Sepharose fractions of CDase II (Triton X-100 fractions). (C) Western blot analysis of RBCDase II (lanes 1 and 2) and RBCDase I (lanes 3 and 4). The samples were treated with glycosidase F as described in the Experimental procedures (lanes 2 and 4).

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(Fig. 6A). Moreover, the optimum concentration of Triton X-100 and CHAPS for RBCDase II was found to be 0.2–0.3 and 0.9–1.2, respectively, which increases the enzyme activity about 2.5-fold in comparison with that in the absence of the detergent (Fig. 6B). As shown in Fig. 6A and B, both RBCDase I and II are inhibited by the detergent β-octyl-glucoside. 3.7. Effects of phospholipids and sphingolipids

Fig. 3. pH dependence of RBCDase II activity. CDase activity was measured as described in the Experimental procedures by incubating an aliquot of purified enzyme with C12NBD-Cer. After 1-h incubation, the spot of NBD-dodecanoic acid and C12-NBD-Cer identified on TLC were scraped, incubated with ethanol at 37 °C for 5 min and their fluorescence was measured at (485/535 nm) excitation/emission wave length in a Perkin-Elmer Spectrofluorophotometer. The pH value was adjusted by the addition of the indicted buffer at a final concentration of 100 mM. The data are the averages of duplicates of at least three independent experiments.

was most pronounced with the oxidized glutathione where the activity was inhibited almost totally at 20 mM concentration (Fig. 5B). 3.6. Effect of detergents The enzyme activity of both RBCDase I and II was greatly enhanced by the by the addition of detergent such as Triton X-100 and CHAPS (Fig. 6A and B). The optimum concentration of detergents for the activation both enzymes is differed markedly depending on the detergent used. For RBCDase I optimum concentration of Triton-X100 and CHAPS were found to be 0.3–0.6 and 0.6–0.9, respectively

The effects of various phospholipids were tested on RBCDase I and II by using C12-NBD-Cer as the substrate. These lipids were added at the indicated concentrations with the substrate as can be seen in (Fig. 7A, B, C, and B). It is noteworthy that of the acidic phospholipids that were tested, PA dose dependently inhibited RBCDase I and II activity (Fig. 7A and C), while PC and PE had only a very small inhibitory effect at low concentration. Other phospholipids such as PS and PI were activating RBCDase I at a concentration of 0.25 mM and 1 mM, respectively, whereas PS and PI were inhibiting RBCDase II activity even at low concentration (Fig. 7B and D). Moreover both CDase were inhibited by PG (Fig. 7B and D). When sphingolipids were tested, it was observed that sphingomyelin and SPH dose dependently inhibited RBCDase II activity (Fig. 7E). However, S-1-P had no inhibitory effect. In fact S-1-P appeared to be somewhat stimulatory with respect to CDase activity. In addition, DHSph at 0.25 mM concentration inhibited CDase activity by up to 40%; however, this effect was not dose-dependent and this inhibition was much less than that of SPH (Fig. 7E). 3.8. Effects of nucleotides The effects of various nucleotides (and nucleosides) on the RBCDase II enzyme activity were tested. It is noteworthy that in this highly purified enzyme preparation, ATP, initially stimulated CDase

Fig. 4. Effects of cations and metal chelators on RBCDase II. (A) CDase activity was measured as described in the Experimental procedures by incubating an aliquot of purified enzyme with C12-NBD-Cer. Metal ions were added as the respective chloride salt at different concentrations. The data are the averages of duplicates of at least three independent experiments. (B) Effects of indicated concentration NaCl. (C) Effects of indicated concentration of EDTA.

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Fig. 5. Effects of reducing agents on RBCDase II activity. (A) CDase from Mono Q fraction was preincubated with the indicated concentrations (DTT, dithiothreitol, and βmercaptoethanol) for 5 min at 37 °C before assayed in the presence of C12-NBD-Cer as described under Experimental procedures. (B) Effects of increasing concentration of GSH and GSSG. The data are the averages of duplicates of at least three independent experiments.

activity by more than 50% at 2.5 mM concentration. Thereafter, it inhibited CDase activity, with maximum inhibition being more than 60% at 24 mM concentration (Fig. 8A). ADP and AMP had no significant effects on the enzyme activity. Interestingly, guanine nucleotides did not have any significant effect (Fig. 8B). Importantly, the pyrimidine mono phosphates TMP and UMP both exerted significant and dose-dependent inhibitory effects on the RBCDase II activity (Fig. 8C and D). At 12 mM both TMP and UMP inhibited the enzyme activity by more than 95%. This effect of the purines was specific to the monophosphates, as TDP, TTP, UDP, and UTP had no significant effect on the enzyme activity (Fig. 8C and D). Similar result were obtained when RBCDase I is treated in the presence of these nucleotides (data not shown). 3.9. Substrate specificity of RBCDase II For obtaining substrate specificity of RBCDase I and II, a HPLC assay of CDase was optimized by measuring the release of SPH as described by El Bawab et al. [48]. Among the various Cer tested C12:0-Cer (Nlauroyl-D-erythro-sphingosine) was most efficiently hydrolyzed by both RBCDase I and II followed by C14:0-Cer (N-myristoyl-D-erythrosphingosine), C16:0-Cer (N-palmitoyl-D-erythro-sphingosine) and C18:0-Cer (N-stearoyl-D-erythro-sphingosine) (Table 3). Furthermore,

Cers containing sphinganine such as Dh-C14-Cer was efficiently hydrolyzed by RBCDase II (55%) rather than RBCDase I (31%) as shown in the Table 3. Long chain dihydro Cer (Dh-C16-Cer) was somewhat resistant to the both enzymes. 4. Discussion Previously, a second ceramidase activity was detected during the purification of a rat brain membrane bound non-lysosomal ceramidase (RBCDase I) [29]. However, it was not determined as to whether this second peak of ceramidase activity was representing a more hydrophobic conformation of the salt-only eluted neutral ceramidase, or that it represented another isoenzyme. In this study, we have purified almost to homogeneity the membrane-associated second type of neutral CDase from rat brain, which we have named as rat brain neutral ceramidase peak II (RBCDase II).The RBCDase II described in this study is clearly different from RBCDase I in several aspects: molecular mass, glycosylation, pH optima, metal dependence, and dependence on phospholipids. In order to separate RBCDase I and II, the Triton X-100 extract was applied to a Q-sepharose column as previously reported [29]. From this first purification step, we were able to separate the two activities as shown in Fig. 1A. The difference in the solubility between the

Fig. 6. Effect of detergents on RBCDase I and II. RBCDase I (A) and RBCDase II (B) activity was measured as described in the Experimental procedures by incubating an aliquot of purified enzyme with C12-NBD-Cer, which had been dissolved in 50 mM Tris–HCl pH 7.5 containing indicated concentration of detergent. The data are the averages of duplicates of at least three independent experiments.

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Fig. 7. Effects of phospholipids and sphingolipids on RBCDase I and II. The aliquots of Mono Q fraction were assayed in the presence or absence of phospholipids (A and B for RBCDase I and C and D for RBCDase II) and sphingolipids (E for RBCDase II). Individual lipids were dried with the substrate, and the combined mixture was resuspended in reaction buffer containing Triton X-100 at a final concentration of 0.3% and the CDase assay were performed as described under Experimental procedures. The data are the averages of duplicates of at least three independent experiments.

RBCDase I and RBCDase II is not unique to the RBCDase protein. In fact, a similar situation has been observed in the case of membrane bound epidermal alkaline ceramidase from guinea pig skin [50]. Before application to the heparin column, the sample was passed through gel filtration chromatography to remove the high concentration of salt and the detergent Triton X-100. This step also removed a large part of other protein impurities. In the next step, the combined fractions from the heparin column were applied to phenyl Sepharose high-performance column. The enzyme was tightly bound to the

column and was eluted with 1% Triton X-100 (Fig. 1C). This also highlights the strong hydrophobic character of RBCDase II. After Mono Q ion exchange chromatography, a single band with molecular mass of ~ 110 kDa was identified by both silver staining and Western blot analysis. The molecular mass of the purified RBCDase II is about ~ 110 kDa, which is close to neutral CDases purified from rat kidney [28], rat intestine [30], and human intestine [31]. In contrast, the size of RBCDase I described by Bawab et al. [29] has the molecular weight of

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Fig. 8. Effect of nucleotides on RBCDase II. (A) CDase activity was determined in the presence of the indicated concentrations AMP, ADP, and ATP. (B) GMP, GDP, and GTP. (C) TMP, TDP, and TTP. (D) UMP, UDP, and UTP.

95 kDa (see Table 2). Interestingly, upon deglycosylation both RBCDase I and II bands shifted from their original molecular weights of ~ 95 kDa and ~110 kDa to ~70 kDa and ~ 95 kDa, respectively (Fig. 2C). This result implies that both CDases are glycosylated. Importantly, this result shows that the difference in molecular mass on SDS-PAGE between RBCDase II and I is not due to the difference in glycosylation (Table 2). The pH optimum of the purified RBCDase II is similar to those of neutral ceramidases purified from rat kidney [28], intestine [30], and human intestine [31], all of which show broad pH with maximum activity at pH 7–8. Hence, we suggest a classification of this enzyme as neutral CDase. In contrast, the pH optima for the RBCDase I was in a broad range of pH 7–10. This different in pH profile demonstrates another difference between the two enzymes and suggests that these two CDase enzymes may be regulated independently by pH in the rat brain.

The metal dependence of the 95 kDa RBCDase I published by Bawab et al. [29] does not match the metal dependence profile of the 110 kDa RBCDase II (Table 2). The RBCDase I activity was inhibited by Mn2+ and had no effect with Zn2+ and Cu2+. In contrast, Mn2+ has no effect on the RBCDase II activity, whereas both Zn2+ and Cu2+ exert inhibitory effects (Fig. 4A). Unlike RBCDase I, 20 mM EDTA inhibits the RBCDase II activity by about 50% (Fig. 4D). Similar to Bawab et al.'s RBCDase I [29], the purified RBCDase II is also inhibited by reducing agents such as β-mercaptoethanol and DTT (Fig. 5A). Interestingly, RBCDase II is inhibited by both oxidized (GSSG) and reduced (GSH) forms of glutathione. This is the first time that the inhibitory action of glutathione on CDase enzyme is reported (Fig. 5B). A similar effect was also seen in the case of RBCDase I (data not shown). These data lead to the insight that GSH may function as a specific allosteric regulator of RBCDase II. It has been proposed that during oxidative stress, depletion of glutathione causes the activation

Table 2 Properties of neutral CDases. Neutral CDase

Molecular mass, SDS-PAGE (kDa)

Deglycosylated molecular mass (kDa)

Reverse reaction

Hydrolysis pH

Reference

RB CDase 1 RB CDase II Rat kidney CDase Rat intestinal CDase Mouse Liver CDase Human intestinal CDase

95 110* 112 116 94 116

70* 95* 89 97 75 97

+ + + ND + +

7.0–10.0 6.5–7.0* 6.5–7.0 6.0–8.0 6.0–8.0 6.0–8.0

El Bawab et al. [29] *Present study Mitsutake et al. [28] Olsson et al. [30] Tani et al. [27] Ohlsson et al. [31]

ND, not determined.

F. Thayyullathil et al. / Biochimica et Biophysica Acta 1811 (2011) 242–252 Table 3 Substrate specificity of RBCDase I and II. Various substrates were dissolved in 50 mM Tris–HCl buffer, pH 7.5, containing 0.2% IGEPAL CA 630 and then incubated with 20 ng CDase (in 10 μl) at 37 °C for 1 h. The extent of hydrolysis of substrate was determined as described under Experimental procedures. The percent value represents relative activity for each substrate when the hydrolysis value for C16:0-Cer is taken as 100%. Results are expressed as means of two separate experiments. CDase I

CDase II

Substrate

Hydrolysis (%)

Hydrolysis (%)

C6:0-Cer C12:0-Cer C14:0-Cer C16:0-Cer C18:0-Cer C24:0 -Cer Dh-C6-Cer Dh-C14-Cer Dh-C16-Cer

86.7 285.3 244.7 100.0 39.0 12.0 0.0 31.3 5.9

100.3 293.2 212.2 100.0 98.5 25.1 0.0 55.9 4.5

[3]

[4]

[5] [6]

[7] [8]

[9]

[10]

of nSMase for the execution of ceramide mediated cellular signaling [51,52]. At this stage RBCDase II activity may be modulated in order to protect from ceramide mediated cellular stress. It is tempting to speculate that under these cellular stresses, RBCDase II activity is modulated to clear out the Cer formed from the nSMase activity. The effect of various phospholipids on RBCDase II is also revealing. PA, PC, PE, and PG moderately inhibit the CDase activity. Indeed, among the various lipids tested, we found PA is the most effective inhibitor of RBCDase I and II activity. Interestingly, PS and PI activated the RBCDase I activity, while these phospholipids are showing inhibitory effects towards the RBCDase II. PS is abundant in the inner leaflet of the plasma membrane, and PA is formed by the action of either phospholipase D or diacylglycerol kinase during signal transduction [53]. Therefore, it is possible to propose that both PS and PA may selectively regulate the activity and/or localization of these two RBCDases. Both RBCDase I and II are inhibited by purine monophosphate nucleotides. This is also the first report on the inhibitory action of nucleotides towards nCDase activity. The finding that pyrimidine monophosphates such as TMP and UMP inhibit nCDase sheds light on one regulatory mechanism for this enzyme. Further study is required to prove the exact role and biological significance of nucleotides on nCDase. In conclusion, we have purified a second type of nCDase from rat brain and describe many of its biochemical properties. We have also found that the purified RBCDase II is clearly different from those reported data of RBCDase I in several aspects such as molecular mass, glycosylation, pH optima, metal dependence, dependence on phospholipids, and substrate specificity. The enzyme was inhibited by reduced and oxidized forms of glutathione and the pyrimidine nucleotides such as TMP and UMP. Acknowledgments We wish to thank the lipidomics core facility at the Medical University of South Carolina, USA, for their kind provision of the substrate and other rare sphingolipids. This work was financially supported by grant from The Emirates Foundation (13 - 2008/075), and in parts, from The Terry Fox Foundation for Cancer Research, and The Terry Fox Foundation for medical Research. References [1] Y.A. Hannun, Functions of ceramide in coordinating cellular responses to stress, Science 274 (1996) 1855–1859. [2] S. Furuya, J. Mitoma, A. Makino, Y. Hirabayashi, Ceramide and its interconvertible metabolite sphingosine function as indispensable lipid factors involved in survival

[11]

[12]

[13]

[14]

[15] [16]

[17] [18] [19] [20] [21]

[22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

251

and dendritic differentiation of cerebellar Purkinje cells, J. Neurochem. 71 (1998) 366–377. X.F. Zhu, Z.C. Liu, B.F. Xie, G.K. Feng, Y.X. Zeng, Ceramide induces cell cycle arrest and upregulates p27kip in nasopharyngeal carcinoma cells, Cancer Lett. 193 (2003) 149–154. Y. Uchida, A.D. Nardo, V. Collins, P.M. Elias, W.M. Holleran, De novo ceramide synthesis participates in the ultraviolet B irradiation-induced apoptosis in undifferentiated cultured human keratinocytes, J. Invest. Dermatol. 120 (2003) 662–669. M.E. Venable, L.Y. Lee, M.J. Smyth, A. Bielawska, L.M. Obeid, Role of ceramide in cellular senescence, J. Biol. Chem. 270 (1995) 30701–30708. S. Daido, T. Kanzawa, A. Yamamoto, H. Takeuchi, Y. Kondo, S. Kondo, Pivotal role of the cell death factor BNIP3 in ceramide-induced autophagic cell death in malignant glioma cells, Cancer Res. 64 (2004) 4286–4293. S. Gatt, Enzymic hydrolysis and synthesis of ceramides, J. Biol. Chem. 238 (1963) 3131–3133. C. Michel, G. van Echten-Deckert, J. Rother, K. Sandhoff, E. Wang, A.H. Merrill Jr., Characterization of ceramide synthesis. A dihydroceramide desaturase introduces the 4,5-trans-double bond of sphingosine at the level of dihydroceramide, J. Biol. Chem. 272 (1997) 22432–22437. Y.A. Hannun, C.R. Loomis, A.H. Merrill Jr., R.M. Bell, Sphingosine inhibition of protein kinase C activity and of phorbol dibutyrate binding in vitro and in human platelets, J. Biol. Chem. 261 (1986) 12604–12609. K. Oishi, B. Zheng, J.F. Kuo, Inhibition of Na, K-ATPase and sodium pump by protein kinase C regulators sphingosine, lysophosphatidylcholine, and oleic acid, J. Biol. Chem. 265 (1990) 70–75. S.C. Frasch, J.A. Nick, V.A. Fadok, D.L. Bratton, G.S. Worthen, P.M. Henson, p38 mitogen-activated protein kinase-dependent and -independent intracellular signal transduction pathways leading to apoptosis in human neutrophils, J. Biol. Chem. 273 (1998) 8389–8397. W.D. Jarvis, F.A. Fornari Jr., K.L. Auer, A.J. Freemerman, E. Szabo, M.J. Birrer, C.R. Johnson, S.E. Barbour, P. Dent, S. Grant, Coordinate regulation of stress- and mitogen-activated protein kinases in the apoptotic actions of ceramide and sphingosine, Mol. Pharmacol. 52 (1997) 935–947. W.D. Jarvis, F.A. Fornari Jr., R.M. Tombes, R.K. Erukulla, R. Bittman, G.K. Schwartz, P. Dent, S. Grant, Evidence for involvement of mitogen-activated protein kinase, rather than stress-activated protein kinase, in potentiation of 1-beta-D-arabinofuranosylcytosine-induced apoptosis by interruption of protein kinase C signaling, Mol. Pharmacol. 54 (1998) 844–856. W.C. Hung, H.C. Chang, L.Y. Chuang, Activation of caspase-3-like proteases in apoptosis induced by sphingosine and other long-chain bases in Hep3B hepatoma cells, Biochem. J. 338 (1999) 161–166. H.C. Chang, L.H. Tsai, L.Y. Chuang, W.C. Hung, Role of AKT kinase in sphingosineinduced apoptosis in human hepatoma cells, J. Cell. Physiol. 188 (2001) 188–193. W. Stoffel, E. Bauer, J. Stahl, The metabolism of sphingosine bases in Tetrahymena pyriformis. Sphingosine kinase and sphingosine-1-phosphate lyase, Hoppe Seylers Z Physiol. Chem. 355 (1974) 61–74. S. Spiegel, A.H. Merrill Jr., Sphingolipid metabolism and cell growth regulation, FASEB J. 10 (1996) 1388–1397. S. Spiegel, D. Foster, R. Kolesnick, Signal transduction through lipid second messengers, Curr. Opin. Cell Biol. 8 (1996) 159–167. D.M. zu Heringdorf, C.J. van Koppen, K.H. Jakobs, Molecular diversity of sphingolipid signaling Meyer Test, FEBS Lett. 410 (1997) 34–38. W. Chen, A.B. Moser, H.W. Moser, Role of lysosomal acid ceramidase in the metabolism of ceramide in human skin fibroblasts, Arch. Biochem. Biophys. 208 (1981) 444–455. K. Bernardo, R. Hurwitz, T. Zenk, R.J. Desnick, K. Ferlinz, E.H. Schuchman, K. Sandhoff, Purification, characterization, and biosynthesis of human acid ceramidase, J. Biol. Chem. 270 (1995) 11098–110102. J. Koch, S. Gartner, C.M. Li, L.E. Quintern, K. Bernardo, O. Levran, D. Schnabel, R.J. Desnick, E.H. Schuchman, K. Sandhoff, Molecular cloning and characterization of a full-length complementary DNA encoding human acid ceramidase. Identification Of the first molecular lesion causing Farber disease, J. Biol. Chem. 271 (1996) 33110–33115. C. Mao, R. Xu, A. Bielawska, L.M. Obeid, Cloning of an alkaline ceramidase from Saccharomyces cerevisiae. An enzyme with reverse (CoA-independent) ceramide synthase activity, J. Biol. Chem. 275 (2000) 6876–6884. C. Mao, R. Xu, A. Bielawska, Z.M. Szulc, L.M. Obeid, Cloning and characterization of a Saccharomyces cerevisiae alkaline ceramidase with specificity for dihydroceramide, J. Biol. Chem. 275 (2000) 31369–31378. C. Mao, R. Xu, Z.M. Szulc, A. Bielawska, S.H. Galadari, L.M. Obeid, Cloning and characterization of a novel human alkaline ceramidase. A mammalian enzyme that hydrolyzes phytoceramide, J. Biol. Chem. 276 (2001) 26577–26588. W. Sun, W. Hu, R. Xu, J. Jin, Z.M. Szulc, G. Zhang, S.H. Galadari, L.M. Obeid, C. Mao, Alkaline ceramidase 2 regulates {beta}1 integrin maturation and cell adhesion, FASEB J. 23 (2009) 656–666. M. Tani, N. Okino, K. Mori, T. Tanigawa, H. Izu, M. Ito, Molecular cloning of the fulllength cDNA encoding mouse neutral ceramidase. A novel but highly conserved gene family of neutral/alkaline ceramidases, J. Biol. Chem. 275 (2000) 11229–11234. S. Mitsutake, M. Tani, N. Okino, K. Mori, S. Ichinose, A. Omori, H. Iida, T. Nakamura, M. Ito, Purification, characterization, molecular cloning, and subcellular distribution of neutral ceramidase of rat kidney, J. Biol. Chem. 276 (2001) 26249–26259. S. El Bawab, A. Bielawska, Y.A. Hannun, Purification and characterization of a membrane-bound nonlysosomal ceramidase from rat brain, J. Biol. Chem. 274 (1999) 27948–27955. M. Olsson, R.D. Duan, L. Ohlsson, A. Nilsson, Rat intestinal ceramidase: purification, properties, and physiological relevance, Am. J. Physiol. Gastrointest. Liver Physiol. 287 (2004) G929–G937.

252

F. Thayyullathil et al. / Biochimica et Biophysica Acta 1811 (2011) 242–252

[31] L. Ohlsson, C. Palmberg, R.D. Duan, M. Olsson, T. Bergman, A. Nilsson, Purification and characterization of human intestinal neutral ceramidase, Biochimie 89 (2007) 950–960. [32] S. Galadari, B.X. Wu, C. Mao, P. Roddy, S. El Bawab, Y.A. Hannun, Identification of a novel amidase motif in neutral ceramidase, Biochem. J. 393 (2006) 687–695. [33] N. Okino, S. Ichinose, A. Omori, S. Imayama, T. Nakamura, M. Ito, Molecular cloning, sequencing, and expression of the gene encoding alkaline ceramidase from Pseudomonas aeruginosa. Cloning of a ceramidase homologue from Mycobacterium tuberculosis, J. Biol. Chem. 274 (1999) 36616–36622. [34] H. Monjusho, N. Okino, M. Tani, M. Maeda, M. Yoshida, M. Ito, A neutral ceramidase homologue from Dictyostelium discoideum exhibits an acidic pH optimum, Biochem. J. 376 (2003) 473–479. [35] Y. Yoshimura, N. Okino, M. Tani, M. Ito, Molecular cloning and characterization of a secretory neutral ceramidase of Drosophila melanogaster, J. Biochem. 132 (2002) 229–236. [36] Y. Yoshimura, M. Tani, N. Okino, H. Iida, M. Ito, Molecular cloning and functional analysis of zebrafish neutral ceramidase, J. Biol. Chem. 279 (2004) 44012–44022. [37] M. Tani, N. Okino, K. Mori, T. Tanigawa, H. Izu, M. Ito, Molecular cloning of the fulllength cDNA encoding mouse neutral ceramidase. A novel but highly conserved gene family of neutral/alkaline ceramidases, J. Biol. Chem. 275 (2000) 11229–11234. [38] Y.H. Hwang, M. Tani, T. Nakagawa, N. Okino, M. Ito, Subcellular localization of human neutral ceramidase expressed in HEK293 cells, Biochem. Biophys. Res. Commun. 331 (2005) 37–42. [39] M.O. Pata, B.X. Wu, J. Bielawski, T.C. Xiong, Y.A. Hannun, C.K. Ng, Molecular cloning and characterization of OsCDase, a ceramidase enzyme from rice, Plant J. 55 (2008) 1000–1009. [40] E. Coroneos, M. Martinez, S. McKenna, M. Kester, Differential regulation of sphingomyelinase and ceramidase activities by growth factors and cytokines. Implications for cellular proliferation and differentiation, J. Biol. Chem. 270 (1995) 23305–23309. [41] M. Nikolova-Karakashian, E.T. Morgan, C. Alexander, D.C. Liotta, A.H. Merrill Jr., Bimodal regulation of ceramidase by interleukin-1beta. Implications for the regulation of cytochrome p450 2C11, J. Biol. Chem. 272 (1997) 18718–18724.

[42] A. Huwiler, J. Pfeilschifter, H. van den Bosch, Nitric oxide donors induce stress signaling via ceramide formation in rat renal mesangial cells, J. Biol. Chem. 274 (1999) 7190–7195. [43] N. Auge, M. Nikolova-Karakashian, S. Carpentier, S. Parthasarathy, A. Nègre-Salvayre, R. Salvayre, A.H. Merrill Jr., T. Levade, Role of sphingosine 1-phosphate in the mitogenesis induced by oxidized low density lipoprotein in smooth muscle cells via activation of sphingomyelinase, ceramidase, and sphingosine kinase, J. Biol. Chem. 274 (1999) 21533–21538. [44] A. Bielawska, M.S. Greenberg, D. Perry, S. Jayadev, J.A. Shayman, C. McKay, Y.A. Hannun, (1S,2R)-D-erythro-2-(N-myristoylamino)-1-phenyl-1-propanol as an inhibitor of ceramidase, J. Biol. Chem. 271 (1996) 12646–12654. [45] U. Acharya, S. Patelm, E. Koundakjianm, K. Nagashima, X. Han, J.K. Acharya, Modulating sphingolipid biosynthetic pathway rescues photoreceptor degeneration, Science 299 (2003) 1740–1743. [46] U. Acharya, M.B. Mowen, K. Nagashima, J.K. Acharya, Ceramidase expression facilitates membrane turnover and endocytosis of rhodopsin in photoreceptors, Proc. Natl Acad. Sci. USA 101 (2004) 1922–1926. [47] S. El Bawab, P. Roddy, T. Qian, A. Bielawska, J.J. Lemasters, Y.A. Hannun, Molecular cloning and characterization of a human mitochondrial ceramidase, J. Biol. Chem. 275 (2000) 21508–21513. [48] S. El Bawab, J. Usta, P. Roddy, Z.M. Szulc, A. Bielawska, Y.A. Hannun, Substrate specificity of rat brain ceramidase, J. Lipid Res. 43 (2002) 141–148. [49] U.K. Laemmli, Cleavage of structural proteins during the assembly of the head of bacteriophage T4, Nature 227 (1970) 680–685. [50] Y.K. Yada, K. Higuchi, G. Imokawa, Purification and biochemical characterization of membrane-bound epidermal ceramidases from guinea pig skin, J. Biol. Chem. 270 (1995) 12677–12684. [51] B. Liu, D.F. Hassler, G.K. Smith, K. Weaver, Y.A. Hannun, Purification and characterization of a membrane bound neutral pH optimum magnesiumdependent and phosphatidylserine-stimulated sphingomyelinase from rat brain, J. Biol. Chem. 273 (1998) 34472–34479. [52] B. Liu, Y.A. Hannun, Inhibition of the neutral magnesium-dependent sphingomyelinase by glutathione, J. Biol. Chem. 272 (1997) 16281–16287. [53] N. Okino, X. He, S. Gatt, K. Sandhoff, M. Ito, E.H. Schuchman, The reverse activity of human acid ceramidase, J. Biol. Chem. 278 (2003) 29948–29953.