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Molecular Cancer Research 5, 801-812, August 1, 2007. doi: 10.1158/1541-7786.MCR-07-0100
© 2007 American Association for Cancer Research
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DNA Damage and Cellular Stress Responses

(Dihydro)ceramide Synthase 1–Regulated Sensitivity to Cisplatin Is Associated with the Activation of p38 Mitogen-Activated Protein Kinase and Is Abrogated by Sphingosine Kinase 1

Junxia Min1, Adi Mesika3, Mayandi Sivaguru1,2, Paul P. Van Veldhoven4, Hannah Alexander1, Anthony H. Futerman3 and Stephen Alexander1

1 Division of Biological Sciences and 2 Molecular Cytology Core, University of Missouri, Columbia, Missouri; 3 Department of Biological Chemistry, Weizmann Institute of Science, Rehovot, Israel; and 4 Katholieke Universiteit Leuven, Departement Moleculaire Celbiologie, Afdeling Farmakologie, Leuven, Belgium

Requests for reprints: Stephen Alexander, Division of Biological Sciences, 303 Tucker Hall, University of Missouri, Columbia, MO 65203. Phone: 573-882-6670; Fax: 573-882-0123. E-mail: alexanderst{at}missouri.edu


   Abstract
Top
 Notes
 Abstract
Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Resistance to chemotherapeutic drugs often limits their clinical efficacy. Previous studies have implicated the bioactive sphingolipid sphingosine-1-phosphate (S-1-P) in regulating sensitivity to cisplatin [cis-diamminedichloroplatinum(II)] and showed that modulating the S-1-P lyase can alter cisplatin sensitivity. Here, we show that the members of the sphingosine kinase (SphK1 and SphK2) and dihydroceramide synthase (LASS1/CerS1, LASS4/CerS4, and LASS5/CerS5) enzyme families each have a unique role in regulating sensitivity to cisplatin and other drugs. Thus, expression of SphK1 decreases sensitivity to cisplatin, carboplatin, doxorubicin, and vincristine, whereas expression of SphK2 increases sensitivity. Expression of LASS1/CerS1 increases the sensitivity to all the drugs tested, whereas LASS5/CerS5 only increases sensitivity to doxorubicin and vincristine. LASS4/CerS4 expression has no effect on the sensitivity to any drug tested. Reflecting this, we show that the activation of the p38 mitogen-activated protein (MAP) kinase is increased only by LASS1/CerS1, and not by LASS4/CerS4 or LASS5/CerS5. Cisplatin was shown to cause a specific translocation of LASS1/CerS1, but not LASS4/CerS4 or LASS5/CerS5, from the endoplasmic reticulum (ER) to the Golgi apparatus. Supporting the hypothesis that this translocation is mechanistically involved in the response to cisplatin, we showed that expression of SphK1, but not SphK2, abrogates both the increased cisplatin sensitivity in cells stably expressing LASS1/CerS and the translocation of the LASS1/CerS1. The data suggest that the enzymes of the sphingolipid metabolic pathway can be manipulated to improve sensitivity to the widely used drug cisplatin. (Mol Cancer Res 2007;5(8):801–12)


   Introduction
Top
 Notes
 Abstract
 Introduction
Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Chemotherapy is frequently used in the treatment of cancer, but drug resistance greatly limits the efficacy of treatment. Some cancers are initially resistant to chemotherapy, whereas others become resistant during the course of treatment. This is true for the drug cisplatin [cis-diamminedichloroplatinum(II)], which is widely used to treat a variety of solid tumors including non-Hodgkin's lymphoma, small cell and non–small cell lung cancers, testicular, ovarian, head and neck, esophageal, and bladder cancers (1). Many different mechanisms of resistance to cisplatin have been investigated; yet it is clear that the resistance to cisplatin is complex and multifaceted, and we are far from having a full understanding of all the relevant signaling pathways involved (2). A better understanding of the mechanisms controlling the sensitivity to cisplatin could be used to make the drug more effective.

In an unbiased genetic examination of genes that are involved with the cellular response and sensitivity to cisplatin in the model eukaryote Dictyostelium discoideum, we showed that disruption of the gene encoding sphingosine-1-phosphate (S-1-P) lyase resulted in decreased sensitivity to the drug (3). This defined the S-1-P lyase as a potential molecular target for controlling sensitivity to cisplatin and suggested a general role for sphingolipids in the cellular response to the drug. S-1-P is synthesized from sphingosine and ATP by two distinct sphingosine kinases, and S-1-P lyase catalyzes the conversion of S-1-P to hexadecenal and phosphoethanolamine at the end of the pathway of sphingomyelin metabolism (Fig. 1 ). S-1-P is a bioactive sphingolipid that is involved in many cellular functions including controlling cell proliferation, cell differentiation, and cell movement (4). The prevailing thought was that S-1-P functions in a rheostat-like mechanism with another bioactive sphingolipid, ceramide, where the relative levels of the two sphingolipids control whether cells proliferate (high S-1-P) or die (high ceramide; refs. 4, 5). This, coupled with the observed decreased cisplatin sensitivity in the S-1-P lyase null mutant, led us to suggest that modulating the levels of the enzymes for the synthesis and degradation of S-1-P would have predictable effects on the sensitivity to cisplatin (6). Indeed, extensive genetic, biochemical, and pharmacologic studies in D. discoideum further established a clear role for the S-1-P lyase as well as the sphingosine kinases in controlling cisplatin sensitivity, where cells overexpressing sphingosine kinase or null for S-1-P lyase have decreased sensitivity to cisplatin, whereas cells null for the sphingosine kinase or overexpressing S-1-P lyase are more sensitive. The change in sensitivity was seen with cisplatin and carboplatin, but not with other drugs (6-8).


Figure 1
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  		FIGURE 1. Pathway of sphingolipid metabolism. The enzymes of special importance to this study are the sphingosine kinases (SphK1-2) and the dihydroceramide synthases (CerS1-6/LASS1-6). Previous studies had shown that high ceramide promotes cell death, whereas high S-1-P promotes cell proliferation. The current study reveals a more detailed picture. DH, dihydro, PEA, phosphoethanolamine.

 
These studies were extended to human cells. We showed increased drug sensitivity in either HEK293 cells (human embryonic kidney) or A549 cells (human lung cancer) overexpressing the human S-1-P lyase gene, and again, the response was highly biased toward the platinum-based drugs cisplatin and carboplatin. Moreover, we showed that inhibition of the sphingosine kinase with the competitive inhibitor dimethylsphingosine has a synergistic effect on increasing cisplatin sensitivity (9). Mechanistically, we showed that the increase in cisplatin sensitivity resulting from the overexpression of the S-1-P lyase was mediated through the activation of the p38 mitogen-activated protein (MAP) kinase, and that the level of p38 activation following cisplatin treatment was much higher in the S-1-P lyase–expressing cells than in control cells treated with the drug. Inhibition of p38 by small interfering RNA (siRNA) or with a specific pharmacologic inhibitor reduced the sensitivity of the cells to cisplatin. These findings are in agreement with other recent studies that also showed increased activation of p38 in S-1-P lyase–overexpressing cells, a diminished apoptotic response following knockdown of the S-1-P lyase enzyme, and down-regulation of the enzyme in colorectal tumor tissues (10, 11). Thus, in addition to the apparent role of sphingosine kinase as an oncogene (12), its activity regulates sensitivity to chemotherapeutic drugs.

The identification of multiple enzymes involved in the generation of ceramide (dihydroceramide synthases) and S-1-P (sphingosine kinases) has allowed us to further test this model, to ask whether modulation of any, or all, of these enzymes results in an altered cellular response to cisplatin, and to begin to establish the roles of these enzymes relative to one another in controlling this response. Synthesis of S-1-P in human cells is mediated by two sphingosine kinase enzymes, which are reported to have opposing roles in the cell (13). Sphingosine kinase 1 (SphK1) is a prosurvival enzyme, which promotes cell proliferation, whereas sphingosine kinase 2 (SphK2) is believed to be proapoptotic. The SphK2 protein contains a domain with homology to the BH3-only proteins, common to the proapoptotic members of the Bcl-2 family of proteins (14). Both SphK1 and SphK2 use the natural substrate D-erythro-sphingosine with similar Km values (although SphK2 can use a broader range of analogs; refs. 15-17). This suggests the possibility that it is the cellular location and/or specific protein-protein interactions of these enzymes that determines their effect on the cell, rather than the absolute level of S-1-P in the cell (13).

The de novo synthesis of the upstream bioactive molecule ceramide in human cells is executed by a family of six transmembrane dihydroceramide synthase enzymes that are involved in the synthesis of dihydroceramide from sphinganine and fatty acyl CoA. These enzymes were originally termed LASS (longevity assurance) genes to denote that the first gene discovered in yeast (LAG1) extended life span (18), but they have been renamed CerS to more closely reflect their biochemical nature (19). Accumulating evidence suggests that each CerS enzyme has a select substrate specificity whereby it preferentially uses specific (often only one) fatty acyl CoA substrates, which results in the production of specific dihydroceramide species with the cognate fatty acyl chain lengths (19-23). The dihydroceramides are then converted to the corresponding ceramides by dihydroceramide desaturase. An increasing amount of evidence suggests that different ceramide species mediate different responses within cells (24). For example, generation of C16-ceramide has been shown to contribute to tumor necrosis factor-{alpha}–induced apoptosis in hepatocytes (25). Head and neck cancer cells, but not normal cells, have been shown to down-regulate C18 ceramide while elevating the levels of ceramides with other fatty acyl chain lengths (26).

In the current study, we tested the effect of ectopic expression of the SphK or the CerS enzymes on cellular sensitivity to cisplatin. We show that modulating the level of CerS1, CerS4, and CerS5 each has a dramatically different effect on drug sensitivity and on the downstream activation of the p38 MAP kinase. In addition, we show a molecular correlation where the CerS1 enzyme, but not CerS4 or CerS5 enzymes, is translocated from the endoplasmic reticulum (ER) to the Golgi apparatus after exposure to cisplatin. Similarly, the two SphK enzymes had a differential effect on drug sensitivity, where overexpression of SphK1 resulted in decreased sensitivity to cisplatin. In contrast, expression of SphK2 rendered the cells more sensitive to the drug. It was further shown that SphK1 can abrogate the CerS1-mediated increased sensitivity of the cells to cisplatin, as well as the specific translocation of CerS1 to the Golgi apparatus. Taken together, these studies present an increasingly detailed understanding of how the sphingolipid-metabolizing enzymes act together to control drug sensitivity in human cells and show a surprisingly unsuspected level of complexity (compare to the more generalized view in Fig. 1). The results suggest that these enzymes can be considered as potential targets for improving the efficacy of cisplatin therapy.


   Results
Top
 Notes
 Abstract
 Introduction
 Results
Discussion
 Materials and Methods
 Acknowledgements
 References
 
Characterization of Stable Cell Lines
Cells were transfected with the CerS1, CerS4, CerS5, SphK1, or SphK2 cDNAs, and stable cell lines were selected. Figure 2A depicts an analysis of the expressed CerS proteins in the stable cell lines and shows that the stably transfected cells express the CerS proteins of the expected molecular weights.


Figure 2
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  		FIGURE 2. Characterization of cell lines stably transfected with the CerS1, CerS4, and CerS5 genes and the SphK1 and SphK2 genes. A. Immunoprecipitation using anti-hemagglutinin antibody coupled to agarose beads followed by immunoblotting shows the expression of the CerS proteins of correct molecular weight in each stable cell line. B. In vitro activity of the CerS enzymes using the different acyl-CoA substrates as described in Materials and Methods. n = 3; bars, SD. C. Western blot analysis of the SphK1 enzyme using anti-FLAG antibody and SphK2 enzyme using anti-His antibody shows the expression of the SphK enzymes of correct molecular weight. D. In vitro activity of the SphK enzymes at two KCl concentrations with their respective vector controls as described in Materials and Methods. n = 2; bars, SD.

 
Extracts of each of the CerS-expressing cell lines were then tested for dihydroceramide synthase activity in vitro using a panel of fatty acyl CoA substrates (Fig. 2B). Each CerS enzyme showed a high level of specificity toward a specific substrate. CerS1-overexpressing cells show a 3.2-fold increase in activity using stearoyl CoA (C18) as substrate, the CerS5-overexpressing cells show a 6.7-fold increase using palmitoyl CoA (C16), and the CerS4 cells show a 6.3-fold increase in activity when tested in the presence of arachidonyl CoA (C20). The basal (endogenous) levels of activity for each substrate are seen in the vector-transformed cells and the cells transformed with the other CerS genes. The profiles of substrate preference are consistent with previous studies analyzing HEK293 cells transiently transfected with the CerS genes (21, 22, 27, 28) and with CerS5 purified to homogeneity (20).

Figure 2C shows the analysis of the SphK1 and SphK2 proteins in the stably expressing cell lines, and Fig. 2D shows the in vitro sphingosine kinase activity in the cell lines. There is a 6- and 5-fold increase in expression over the control cells for the SphK1 and SphK2 cells, respectively, when assayed in their preferred salt conditions. Consistent with previous reports, SphK1 activity is higher in low-salt conditions, and SphK2 is more active in high-salt conditions (29, 30).

The CerS1, CerS4, and CerS5 Enzymes Differentially Regulate Sensitivity to Drugs
Earlier work both in the model organism D. discoideum and in human cells showed that the modulation of the enzymes that metabolize S-1-P alters the cellular sensitivity to chemotherapeutic drugs, with a bias toward platinum-based drugs (7-9). To test whether modulating de novo ceramide synthesis also affects drug sensitivity, each CerS-expressing cell line was tested for its sensitivity to cisplatin, carboplatin, doxorubicin, and vincristine (Fig. 3 ). CerS1 expression rendered the cells more sensitive to cisplatin and carboplatin, doxorubicin, and vincristine (P < 0.001). In contrast, expression of CerS4 did not have any effect on the cellular sensitivity to any of the agents tested, whereas CerS5 expression increased the sensitivity only to doxorubicin and vincristine, but not to cisplatin and carboplatin. The results reveal a previously unsuspected complexity in the signaling mechanisms that respond to exposure to drugs and strongly support the idea the CerS genes are not equivalent in function.


Figure 3
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  		FIGURE 3. CerS gene expression alters cellular sensitivity to drugs. CerS1-, CerS4-, and CerS5-overexpressing HEK293 cells and vector control cells were treated with the indicated concentrations of drugs (A, cisplatin; B, carboplatin; C, doxorubicin; D, vincristine), and viability was assayed at 72 h as described in Materials and Methods. The results for each cell line are presented as percent over its own untreated control culture at the same time. SDs were calculated using the statistics tools in Microsoft Excel. Each experiment was repeated thrice. P < 0.0015 for CerS1 and CerS5 versus control.

 
CerS1, but not CerS4 or CerS5, Expression Results in the Increased Activation of the p38 MAP Kinase in Response to Cisplatin
The results presented above suggested that each CerS gene functions through different signaling pathways that are involved in modulating the cellular response and sensitivity to drugs. We and others have shown that the cellular response to cisplatin is mediated through the activation of p38, and that expression of the S-1-P lyase causes increased activation of p38 (9, 11, 31). The lack of response to cisplatin and carboplatin in the CerS4- and CerS5-overexpressing cell lines was very intriguing and suggested the possibility of a differential effect of the CerS enzymes on the activation of p38. Thus, to determine if p38 was mechanistically involved in the specific CerS1-mediated increase in sensitivity to cisplatin, we tested the activation of p38 in the CerS1-, CerS4-, and CerS5-expressing cell lines after cisplatin treatment by directly measuring the p38 kinase enzyme activity. The results in Fig. 4A show that following cisplatin treatment, there is a significant increase of p38 enzyme activity in the CerS1-expressing cells. Significantly, there is no activation of p38 in CerS4- or CerS5-expressing cells in response to cisplatin. The activation of p38 occurs early (30 min), peaks at 6 h, and is diminished at 24 h (Fig. 4B). Interestingly, activation of p38 is also specifically seen in the CerS1 cells after treatment with doxorubicin (Fig. 4C). This CerS1-specific activation of p38 by both drugs, and its absence in the CerS4- and CerS5-overexpressing lines, strongly supports the hypothesis that the different CerS genes interact differentially with unique downstream regulatory pathways.


Figure 4
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  		FIGURE 4. CerS1 specifically mediates the activation of p38 MAP kinase in response to cisplatin. A. Parallel cultures of CerS1-, CerS4-, and CerS5-overexpressing HEK293 cells and vector control cells were treated either with 50 µmol/L cisplatin or buffer in 6-cm Petri dishes for 3 h. These conditions are based on the sensitivities of the assay, as have been documented previously for in vitro p38 MAP kinase assays to generate a robust and synchronous response (31, 51). Cells were harvested and directly assayed for activated p38 MAP kinase enzyme activity by immunoprecipitation of phosphorylated p38 and using this isolated enzyme to phosphorylate ATF-2-GST fusion protein in vitro. The phosphorylated ATF-2 fusion protein is electrophoresed, blotted, and immunodetected. Phosphorylated ATF-2-GST has a molecular weight of 34,000. B. Time course for the activation of p38 following cisplatin treatment in the CerS1-expressing cells. C. CerS1-, CerS4-, and CerS5-overexpressing HEK293 cells and vector control cells treated with either 50 µmol/L doxorubicin or buffer in 6-cm Petri dishes for 3 h. CerS1 specifically enhances the activation of p38 in the presence of cisplatin (A, B) and doxorubicin (C). D. Stable CerS1-expressing cells transfected with SphK1, SphK2, control vector, or untransfected cells (nt) were assayed for the activation of p38 with and without cisplatin.

 
Expression of the Two SphK Enzymes Has Differential Effects on Drug Sensitivity
As stated above, there are two SphK enzymes, and we wished to determine their individual roles in regulating sensitivity to drugs. Our results show that the expression of SphK1 enzyme decreased sensitivity to cisplatin and carboplatin and, to a lesser extent, to vincristine and doxorubicin, as compared with control cells (Fig. 5A ). These results with SphK1 are in agreement with our previous studies on drug sensitivity that showed that the overexpression of the S-1-P lyase resulted in increased sensitivity (9).


Figure 5
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  		FIGURE 5. The SphK1 and SphK2 enzymes have opposite effects on drug sensitivity. Cells stably expressing SphK1, SphK2, or the respective vector control cells were treated with the indicated concentrations of drugs, and viability was assayed at 72 h as described in Materials and Methods. The results for each cell line are presented as percent over its own untreated control culture at the same time. SDs were calculated using the statistics tools in Microsoft Excel. Each experiment was repeated thrice. P < 0.0015 versus control. A. SphK1 decreases sensitivity (increases resistance); B. SphK2 increases sensitivity.

 
In contrast, expression of SphK2 rendered the cells more sensitive to the tested drugs (Fig. 5B). Again, there was a bias toward the platinum-based cisplatin and carboplatin and, to a lesser extent, to doxorubicin. Overexpression of SphK2 did not alter the response to vincristine. The findings presented here are in agreement with the reported opposing roles of SphK1 and SphK2 in the cells and the suggested proapoptotic activity of sphingosine kinase 2 and extends this idea to the control of drug sensitivity by the SphK enzymes.

SphK1 and CerS1 Have Antagonistic Influences on Cellular Sensitivity to Cisplatin
The opposing effects of overexpression of SphK1 (decreased sensitivity) and CerS1 (increased sensitivity) on the cellular response to cisplatin raised the central question of the relationship between the functions of these two enzymes. To this end, we tested the influence of SphK1 and CerS1 on each other, with regard to cisplatin sensitivity. The results show that these enzymes can balance the effects of each other with respect to drug sensitivity. Thus, transient transfection of SphK1 into cells stably expressing CerS1 abrogated the increased sensitivity of the CerS1-expressing cells to cisplatin (Fig. 6A ). In similar fashion, the reciprocal experiment of transient transfection of CerS1 into cells stably expressing the SphK1 abrogated the decreased sensitivity to cisplatin in the SphK1 cells (Fig. 6B). Assays of the SphK1 and CerS1 enzymes were done in these experiments, and the results confirm that there is an increase in the activity of the enzyme that was transiently expressed without a corresponding change in the enzyme that was stably expressed in the host cells. Transient transfection of CerS1 into cells stably expressing SphK1 increases the CerS1 enzyme activity 2.2-fold over the untreated or vector-transfected cells, whereas the level of SphK1 enzyme activity is unaffected. In contrast, transient transfection of SphK1 into cells stably expressing CerS1 increases the SphK1 enzyme activity 9-fold over untreated or vector-transfected cells, whereas the level of CerS1 enzyme activity is unaffected. This is a direct demonstration of the opposing roles of SphK1 and CerS1 in regulating sensitivity to cisplatin.


Figure 6
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  		FIGURE 6. SphK1 and CerS1 have antagonistic effects on the cellular response to cisplatin. A. Transient transfection of SphK1 or vector into cell lines stably expressing CerS1. Following transfection, the cells were treated with cisplatin at the indicated concentrations and assayed for survival as described in Materials and Methods. Transfection with SphK1 decreases the sensitivity of the cells to cisplatin, but transfection with vector alone has no effect. B. Transient transfection of CerS1 or vector into cell lines stably expressing SphK1. Following transfection, the cells were treated with cisplatin at the indicated concentrations and assayed for survival as described in Materials and Methods. Transfection with Cers1 increases the sensitivity of the cells to cisplatin, but transfection with vector alone has no effect.

 
We further tested the effect of expressing the SphK genes in the CerS1-expressing cells. SphK1 and SphK2 were shown to have different effects on the activation of p38 (Fig. 4D). Expression of SphK2 in the CerS1-expressing cell resulted in the activation of p38 even without the addition of cisplatin and enhanced activation after cisplatin treatment. Interestingly, SphK1 did not abrogate p38 activation by cisplatin in the CerS1-expressing cells. This is in agreement with reports that show that SphK1 activates the survival MAP kinase extracellular signal-regulated kinase, which, in turn, is antagonistic to p38 activation (32-35).

CerS1 Translocates to the Golgi Apparatus in Response to Drugs
The activation of several of the enzymes of the sphingolipid metabolic pathway has been shown to depend on cellular translocation of the enzymes in response to stimuli. For example, various agonists cause SphK1 to translocate from the cytoplasm to the plasma membrane (33), and stress has been shown to cause SphK2 to translocate from the cytoplasm to the ER (13). In this regard, we asked whether an intracellular relocalization of the CerS enzymes was associated with cisplatin treatment. The CerS1, CerS4, and CerS5 enzymes in the stable cell lines are all localized in the ER (Supplementary Fig. S1; CerS proteins, green; protein disulfide isomerase, red) in agreement with previous reports (21, 22, 27). However, upon treatment with cisplatin, there is a dramatic translocation of CerS1, but not CerS4 or CerS5, from the ER to the Golgi apparatus. The translocation is rapid, is evident as early as 30 min, and occurs even at low cisplatin concentrations (Fig. 7 ; CerS proteins, green; Golgi marker GM-130, red). We further show that this result is not restricted to cisplatin by demonstrating that doxorubicin also specifically causes the specific translocation of CerS1, but not CerS4 or CerS5, to the Golgi apparatus (Fig. 8 and Supplementary Fig. S2). Overall, these data suggest that the specific translocation of the CerS1 enzyme from the ER to the Golgi apparatus is mechanistically related to the increase in cisplatin sensitivity observed in the CerS1-expressing cells.


Figure 7
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  		FIGURE 7. CerS1, but not CerS4 or CerS5, translocates from the ER to the Golgi apparatus after treatment with cisplatin. CerS1-, CerS4-, and CerS5-expressing cells were treated with 5 or 50 µmol/L cisplatin and examined at 3 h after the addition of cisplatin. All three CerS proteins localize in the ER in untreated cells (see 0 time samples and Supplementary Fig. S1). Following cisplatin treatment, the CerS1 protein (stained green with anti-hemagglutinin antibody) specifically translocates and colocalizes with the Golgi apparatus (stained red with anti-GM130 antibody) where the combination appears yellow. Blue, nuclei. Arrows, translocation of the CerS1 protein. No translocation of CerS4 or CerS5 was seen. Bar, 10 µm.

 

Figure 8
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  		FIGURE 8. CerS1 translocates from the ER to the Golgi apparatus after treatment with doxorubicin. CerS1-expressing cells were treated with 5 or 50 µmol/L doxorubicin and examined at 3 h after the addition of the drug. As with cisplatin (Fig. 7), following doxorubicin treatment, the CerS1 protein (stained green with anti-hemagglutinin antibody) specifically translocates and colocalizes with the Golgi apparatus (stained red with anti-GM130 antibody) where the combination appears yellow. Blue, nuclei. Arrows, translocation of the CerS1 protein. Supplementary Fig. S2 shows that in parallel experiments done the same day, there was no translocation of CerS4 or CerS5. Bar, 10 µm.

 
Support for this idea comes from the experiment depicted in Fig. 9 , which shows that expression of SphK1, but not SphK2, completely abrogated the translocation of CerS1 in response to stress. Thus, in the cells transfected with SphK1 (middle), the Golgi appear red because the CerS1 protein is not colocalized there as it is in untransformed cells (CerS1) or the SphK2-transfected cells. (Supplementary Fig. S3 shows that the SphK1 and SphK2 transfections were very efficient.) This, coupled with the fact that overexpression of SphK1 abrogates the increased sensitivity of CerS1-expressing cells to drugs (Fig. 6A), supports the hypothesis that the CerS1 translocation is controlled by the balance of the functions of the CerS1 and SphK1 enzymes and further supports the suggestion of a mechanistic connection between the specific ER to Golgi apparatus translocation of CerS1 and the regulation of cisplatin sensitivity.


Figure 9
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  		FIGURE 9. Expression of SphK1 specifically blocks the ER to Golgi apparatus translocation of CerS1 following treatment with cisplatin. Cells stably expressing CerS1 were transiently transfected with either the SphK1 or SphK2 genes and treated with cisplatin. The CerS1 protein is stained green with anti-hemagglutinin antibody, and the Golgi apparatus is stained red with anti-GM130 antibody. Blue, nuclei. CerS1 clearly translocates from the ER to the Golgi apparatus in both the CerS1-expressing cells (A) and the SphK2-transfected cells (C), where the Golgi is yellow, but not the SphK1-transfected cells (B), where the Golgi apparatus remains red.

 

   Discussion
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
Materials and Methods
 Acknowledgements
 References
 
Cellular processes such as proliferation, differentiation, or apoptosis are influenced by the balance between the bioactive sphingolipids ceramide and S-1-P (4, 36, 37). Several studies of the S-1-P lyase have shown that modulation of this enzyme is a key way to regulate the sensitivity of cells to chemotherapeutic drugs, and in particular the platinum-based drug cisplatin (7, 9, 11). The work presented here greatly expands on these earlier studies by determining the roles of the members of the families of sphingosine kinase and dihydroceramide synthase enzymes in regulating drug sensitivity in human cells. The results reveal an important and unexpected level of complexity and directly show an antagonistic interaction between the sphingosine kinase 1 enzyme and the CerS1 dihydroceramide synthase. The data will be important in determining the potential of these enzymes as targets for improving cisplatin therapy.

Dihydroceramide Synthase (CerS) Enzymes
Six genes encoding different isoforms of mammalian dihydroceramide synthase have been identified based on homology to the yeast LAG1 gene (19, 21, 38). Interestingly, yeast (18) and Dictyostelium (crsA DDB0230136) seem to have only one dihydroceramide synthase gene. The mammalian enzymes have unique but greatly overlapping tissue distributions (21, 22, 28), but the precise roles of these enzymes and their unique products are not known. The preponderance of evidence indicates that each enzyme has a preference toward a specific fatty acyl CoA and therefore produces dihydroceramide with the corresponding fatty acyl chain (38). In this study, we focused on the CerS1, CerS4, and CerS5 genes, and stably expressing cell lines were generated for each of the genes. Consistent with previous reports, we showed that CerS1 enzyme synthesizes predominantly C18 ceramide, CerS5 preferentially synthesizes C16 ceramide, and the stable CerS4-expressing cells had a clear preference for synthesizing C20 ceramide (20-23, 27).

Analysis of the CerS-expressing cell lines showed a fascinating pattern of drug sensitivity. CerS4 overexpression did not alter sensitivity to any of the drugs tested, although the immunofluorescent staining and the in vitro assays showed a substantial level of CerS4 enzyme activity. Cells transfected with CerS5 showed an increase in sensitivity to doxorubicin and vincristine (naturally occurring drugs or derivatives), but not to cisplatin or carboplatin (DNA-damaging/alkylating agents). In contrast, CerS1-overexpressing cells showed an increase in sensitivity to all the drugs tested. Take together, these data clearly indicate that the function of the three CerS enzymes studied with respect to modulating drug sensitivity is not equivalent and suggests that there might be a relationship between drug sensitivity and the individual substrate preferences (and, therefore, the specific ceramide products) of each of the CerS enzyme, although establishing this in a definitive way will require substantial additional experimentation. These results tie in well with other work showing that specific ceramide species are altered in various tumors (25, 26).

Sphingosine Kinase Enzymes
There are two sphingosine kinase enzymes in mammalian cells, and although they both catalyze the synthesis of S-1-P, there is an accumulating body of evidence supporting the idea that they have unique functions. Specifically, SphK1 is believed to promote survival, whereas surprisingly, SphK2 has been shown to be proapoptotic, possibly due to the BH3-only–like domain. Expression of SphK1 results in a decrease in ceramide levels, whereas SphK2 results in an increase in ceramide levels (39). These data suggested that the two enzymes might play unique roles in regulating sensitivity to drugs, and we tested that hypothesis in cell lines stably expressing the two genes.

The expression of SphK1 results in a decrease in the sensitivity to drugs. In contrast, SphK2 has the opposite effect and increases sensitivity to the drug. Thus, similar to what was observed with the CerS enzymes, the SphK enzymes each have unique effects on drug sensitivity. There is a great deal of interest in the SphK enzymes in cancer and chemotherapy (17, 40-42), and it is becoming clear that their intracellular localization may be leading to the unique functions of the two enzymes, although the precise details of this are still unclear at this time (13, 43). It is possible that by altering the intracellular localization of the two SphK enzymes, it would be possible to show that this is a mechanism by which the differential effects of the enzymes on drug sensitivity is effected.

The opposing functions of the CerS1 and SphK1 enzymes were shown in experiments where changes in drug sensitivity caused by expression of one enzyme were abrogated by the expression of the other. These experiments support the rheostat model in the context of regulation of drug sensitivity.

The Role of p38 MAP Kinase
Significantly, the sensitivity to cisplatin was only altered in the cell line expressing CerS1, but not in cells expressing CerS4 or CerS5. Previously, we had definitively shown by a combination of measuring the enzyme phosphorylation, enzyme activation, and pharmacologic or siRNA inhibition that p38 was required for the increased drug sensitivity observed in human cells expressing the S-1-P lyase. These findings are supported by other studies which also show increased activation of p38 in S-1-P lyase–overexpressing cells, a diminished apoptotic response following knockdown of the S-1-P lyase enzyme, and down-regulation of the enzyme in colorectal tumor tissues (10, 11). The general model arising from these studies (6) was that the increase in S-1-P lyase in these cells results in an increase in p38, which, when activated in the wake of cisplatin exposure, results in increased cell death (i.e., increased drug sensitivity).

In this study, we showed that CerS1-expressing cells were more sensitive to cisplatin, whereas CerS4- or CerS5-expressing cells were not more sensitive to cisplatin than control cells. This suggested that the underlying reason for this differential change in cisplatin sensitivity was that p38 was differentially activated in the CerS-expressing cell lines. The data clearly reveal that only the expression of CerS1 increases the activation of p38 in the presence of cisplatin. Interestingly, doxorubicin also activates p38 specifically in the CerS1-expressing cells. Given that CerS5 also sensitizes cells to doxorubicin, this result indicates that CerS5 functions through another signaling pathway. The identification of the molecular details of how CerS1 activates p38 is an important goal for the future.

Specific Translocation of the CerS1 Protein from the ER to the Golgi Apparatus
In the course of these studies, we asked if the intracellular location of the CerS proteins was altered in response to drugs. There was precedent for this in the case of the SphK enzymes, which have been observed to translocate between cellular compartments in response to stress (13, 33). The general idea is that this allows the enzymes to be matched up with their substrates when needed and thereby activate the reaction (43). Nevertheless, it was surprising to find that there was a specific translocation of CerS1, but not CerS4 or CerS5, from the ER to the Golgi apparatus after treatment of the cells with cisplatin or doxorubicin. These data suggest the possibility that the translocation is mechanistically related to the change in drug sensitivity. This hypothesis is testable by creating a series of mutant CerS1 proteins that are unable to translocate and testing their ability to induce changes in drug sensitivity. Alternatively, making a catalytically inactive CerS1 will allow the testing whether the change in drug sensitivity requires ceramide synthase catalytic activity or whether another mechanism is involved to induce changes in drug sensitivity. Overall, this finding opens an exciting new field and again clearly shows the unique nature of each of the CerS enzymes.

The present study has greatly expanded on previous work on the roles of S-1-P lyase and p38 MAP kinase in regulating sensitivity to cisplatin. The data show unique roles for the members of two other families of sphingolipid-metabolizing enzymes in regulating drug sensitivity and further clarify the role of the p38 MAP kinase with this pathway. Direct evidence is presented supporting the sphingolipid rheostat model in the context of the regulation of drug sensitivity. However, the data suggest that the specific enzymes and their unique products (in the case of the CerS enzymes), as well as their intracellular localizations, all play pivotal roles in balancing the rheostat. The data are important in assessing the potential of modulating these enzymes and their products as targets for improving the efficacy of individual chemotherapeutic drugs.


   Materials and Methods
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
Acknowledgements
 References
 
Vectors and Generation of Stable Cell Lines
The COOH-terminal hemagglutinin-tagged murine CerS1, CerS4, and CerS5 cDNAs were cloned into the pcDNA3 vector (22, 27). A construct coding for NH2-terminal FLAG-tagged human SphK1 (pPVV200) was generated by subcloning a 1,215-bp BamHI-PstI fragment of IMAGE clone 1946069 into BamHI-PstI–restricted pCMVTag2A (Stratagene; ref. 44). A plasmid encoding COOH-terminal His-tagged murine SphK2 (pVB201) was obtained by subcloning a KpnI/EcoRI-restricted PCR amplicon using IMAGE clone 2650442 (UK-HGMP Resource Center) as template and selective primers into pcDNA4/HisMaxB (Invitrogen). Human embryonic kidney (HEK) 293 cells [grown in high glucose/DMEM/10% fetal bovine serum (FBS)] were transfected with 25 µg of purified plasmid (Endo-free kit, Qiagen) using LipofectAMINE 2000 (Invitrogen) according to the manufacturer's instructions. Control cells were transfected with the cognate vector DNA. G418 (500 µg/mL) or zeocin (200 µg/mL) in DMEM/10% FBS was added 2 days after transfection, and the cells were incubated in the selection medium for 2 weeks. Replicate vials of each line were stored at 1 x 107 cells/mL, and new cultures for experiments were started monthly. No experiment was done on cells that had been growing for more than 4 weeks. Transient transfections were done as described above, but without G418 selection, and the cells were harvested 36 h after transfection.

Chemicals, Drugs, and Antibodies
All drugs were from Sigma-Aldrich. The ER marker, anti-PDI (protein disulfide isomerase), was from BD Biosciences. Alexa Fluor 488–conjugated goat anti-rabbit antibody, Prolong Gold anti-fade reagent, and monomeric cyanine nucleic acid stain TO-PRO-3 for nuclei visualization were from Invitrogen. Anti-hemagglutinin, anti-His, anti-FLAG, anti-PDI (ER marker), and anti-GM130 (Golgi marker) antibody, poly-L-lysine, defatted bovine serum albumin, phenylmethylsulfonyl fluoride, leupeptin, antipain, aprotonin, sodium fluoride, and sodium orthovanadate were from Sigma-Aldrich. Micro-O-protect was from Roche. D-Erythro-4,5-[3H]sphinganine (specific activity of 11 Ci/mmol) was synthesized as described (45). D-Erythro-stearoyl-sphingosine, D-erythro-palmitoyl-sphingosine, and D-erythro-arachidonyl-sphingosine were from Matreya. Palmitoyl CoA (C16), stearoyl CoA (C18), and arachidonyl CoA (C20) were from Avanti Polar Lipids. [{gamma}-32P]ATP (3,000 Ci/mmol) was from Perkin-Elmer.

Cell Viability
All drug treatments were done as described (9). Cell-Titer Glo reagent (Promega) was used to measure viable cell number in cell cultures (46-49), and the luminescence was measured in a Turner Biosystems luminometer. All results are presented as percent of the untreated culture. Each experiment represents six replica samples (n = 6), and each experiment was repeated at least thrice. Data analyses were done with the statistical tool package in Microsoft Excel.

Immunoprecipitation and Western Blotting
Cells were collected, washed twice in PBS, and resuspended in 750 µL homogenizing buffer [20 mmol/L HEPES (pH, 7.4), 25 mmol/L KCl, 250 mmol/L sucrose, 2 mmol/L MgCl2, protease inhibitor cocktail (1 mmol/L phenylmethylsulfonyl fluoride, 1 µg/mL antipain, 1 µg/mL leupeptin, and 100 kIU/mL aprotonin)]. The cells were homogenized 40 times using a Potter Elvehjem homogenizer at 250 rpm. Digitonin was added to a final concentration of 1% (w/v) and incubated for 1 h at 4°C. The homogenates were centrifuged at 100,000 x gav for 30 min in an Optima ultracentrifuge at 4°C, and the resulting supernatants were collected and incubated with prewashed anti-hemagglutinin agarose-conjugated beads (Sigma-Aldrich) overnight at 4°C. The beads were then washed thrice with homogenizing buffer, and 40-µL beads were mixed with 10 µL of 5x SDS-PAGE sample buffer and loaded on gels for Western blotting with anti-hemagglutinin antibody (20).

Enzyme Assays
Cells at 70% confluence were treated with drugs for the indicated times and concentrations, harvested into cold PBS, washed twice in PBS, and frozen at –80°C.

Dihydroceramide synthase enzyme assays were done as previously described (20, 22, 27, 45). Briefly, pellets were resuspended in 1 mL homogenizing buffer and homogenized as described above, and protein concentration was determined using the Bradford reagent. Homogenates (50–250 µg protein in 0.25 mL final volume) were incubated with 0.25 µCi of 4,5-[3H] sphinganine/7.5 µmol/L sphinganine/20 µmol/L defatted BSA, and 50 µmol/L fatty acyl-CoA (C16, C18, or C20) for 20 min at 37°C. Lipids were extracted, and the level of dihydroceramide synthesis was analyzed by TLC using chloroform/methanol/2 mol/L ammonium hydroxide (40:10:1; v/v/v) as the developing solvent and using N-palmitoyl-sphingosine, N-stearoyl-sphingosine, and N-arachidonyl-sphingosine as authentic standards. Lipids were visualized using a phosphor imager (Fuji), recovered from the TLC plates by scraping the silica directly into scintillation vials, and quantified by liquid scintillation counting. Specific enzyme activity is presented as picomoles of dihydroceramide per minute per milligram of protein.

Sphingosine kinase enzyme activity was assayed as described (34). Frozen pellets of 5 x 106 cells were lysed in 500 µL SK buffer with 0.2 mol/L KCl, followed by six cycles of freezing and thawing, and centrifugation for 30 min at 150,000 x g at 4°C (Beckman TL-100). Supernatants were transferred to clean tubes, and the protein concentration was determined by BCA (bicinchoninic acid; Pierce Chemical Co.). About 20 to 40 µg of protein extract were brought to 180 µL with SK buffer containing either 0.2 or 1.0 mol/L KCl and incubated with 10 µL of 1 mmol/L sphingosine (delivered in 5% Triton X-100) and 10 µL ATP mix [9 µL of 20 mmol/L ATP in 200 mmol/L MgCl2 + 1 µL [{gamma}-32P]ATP (3,000 Ci/mmol); New England Nuclear] for 90 min at 28°C. The reaction was stopped with 20 µL of 1 N HCl and 800 µL of chloroform/methanol/HCl (50:100:1) for 10 min at room temperature. Then 250 µL chloroform and 250 µL 2 mol/L KCl were added, and the samples were mixed and centrifuged to separate the phases. The top aqueous layer was aspirated, and 100-µL aliquots of the organic phase were spotted onto TLC plates (Silica Gel 60, Merck). The plates were developed in chloroform/acetone/methanol/acetic acid/H2O (10:4:3:2:1) and visualized on a Fuji PhosphorImager. Spots were then scraped from the plates and quantitated in a scintillation counter. One unit activity was defined as the number of picomoles of S-1-P generated per minute per milligram of protein extract.

p38 MAP kinase enzyme activity was assayed using the Nonradioactive p38 MAP Kinase Assay Kit (Cell Signaling) according to the manufacturer's instructions. Cells were harvested in nondenaturing buffer, lysed by sonication, and centrifuged to remove cell debris. Protein concentration was determined by BCA. About 250 µg of extract were used for each assay of p38 activity by immunoprecipitating the phosphorylated p38 protein and subsequently using the isolated protein to phosphorylate the p38 substrate ATF-2-GST fusion protein in vitro. Phosphorylated ATF-2 fusion protein has a molecular weight of 34,000.

Immunofluorescence Staining
Sterilized poly-L-lysine–coated coverslips were place in 24-well plates. Cells were seeded at the appropriate density and allowed to attach overnight. Following drug treatment, the cells were washed thrice with ice-cold PBS [10 mmol/L Na3PO4, 150 mmol/L NaCl (pH, 7.4)]. Cells were fixed by incubating in PBS/4% paraformaldehyde/0.1% glutaraldehyde (Electron Microscopy Sciences) at room temperature (RT) for 30 min, followed by washing thrice with PBS. Cells were permeabilized by incubating in PBS/0.1% Triton X-100 for 25 min at RT, followed by washing twice with PBS. The slides were preincubated for 30 min in PBS/0.1% BSA/0.1% Micro-O-protect, blocked in blocking buffer (PBS/5% BSA/5% normal goat serum/0.1% Micro-O-protect) for 2 h at RT, rinsed thrice with PBS, and incubated with primary antibody diluted in 10% blocking buffer overnight at RT. The slides were then washed thrice with PBS, incubated with fluorescence-conjugated secondary antibody diluted in 10% blocking buffer for 2 h at room temperature, washed thrice with PBS, and mounted inverted onto a drop of Prolong Gold anti-fade reagent. The slides were sealed with nail polish and stored at 4°C.

Confocal Microscopy
Fluorescence was visualized under a sequential scan setting, using 488 nm (green, for hemagglutinin-tagged CerS1) and 568 nm (red, Golgi apparatus or ER) excitation lasers from a krypton/argon-mixed gas laser on a Radiance 2000 confocal system (Bio-Rad Cell Sciences) and 637 nm red diode solid-state laser excitation (blue, nuclei). The confocal laser system is coupled to an Olympus IX70 inverted microscope, and all images were acquired using a 60x (1.2 numerical aperture) water immersion objective with 1-2 electronic/digital zoom at the median plane of focus (single optical sections or series at 300 nm interval) for either individual or groups of cells. All acquisition parameters were kept constant within a given experiment across different treatments. The raw data sets were deconvolved under 20 constrained iterations using a three-dimensional blind algorithm (50) using Autodeblurr software (Autoquant Inc.). The product of the deconvolution planes was uploaded in Metamorph (Universal Imaging Corp.), and individual channels were pseudocolored and merged. All confocal microscopy experiments were repeated at least twice with two independent replicas for each sample.


   Acknowledgements
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
References
 
We thank the Alexander and Futerman lab members for help and comments on this work.


   Notes
Top
 Notes
Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Grant support: Work in the Alexander lab is supported by NIH grant GM53929. Work in the Futerman lab is supported by the Israel Science Foundation, grant 1047/03, and by the Minerva Foundation, Munich, Germany. A.H. Futerman is the Joseph Meyerhoff Professor of Biochemistry at the Weizmann Institute of Science. S. Alexander and A.H. Futerman are the corecipients of United States–Israel National Science Foundation grant. P.P. Van Veldhoven was supported by grants from the Flemish Fonds voor Werenschappelijk Onderzoek (G.405.02) and from the Belgian Ministry of Federaal Wetenschapsbeleid Interuniversitarie Attractiepolen (IAP-P5/05).

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

Note: Supplementary data for this article is available at Molecular Cancer Research Online (http://mcr.aacrjournals.org/).

Current address for J. Min: Brigham and Woman's Hospital, Department of Medicine/Division of Genetics, Harvard Medical School New Research Building, 77 Avenue Louis Pasteur, Boston, MA 02115.

Current address for M. Sivaguru: Institute for Genomic Biology, 1206 W. Gregory Dr., University of Illinois, Urbana, IL 61801.

Received 2/27/07; revised 5/31/07; accepted 6/12/07.


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 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 

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