Lipoprotein Lipase Links Dietary Fat to Solid Tumor Cell Proliferation

Many tumors, including those arising in breast, colon, ovary, and prostate, exhibit a lipogenic phenotype. This features brisk rates of saturated long-chain fatty acid (FA) synthesis driven by enhanced expression of genes coding for the 3 enzymes required to produce palmitic acid from cytosolic citrate [ATP citrate-lyase, acetyl CoA-carboxylase, and fatty acid synthase (FASN)]. Importantly, lipogenic tumor cell growth is slowed in vitro and survival is reduced by FA synthesis inhibitors, whereas nontransformed cells are unaffected (reviewed in refs. 1, 2). Moreover, blocking de novo lipogenesis with FASN inhibitors in vivo exerts potent antitumor effects in rodent models of breast (3) and prostate (4) cancer. These observations, coupled with the low rates of FA synthesis in most normal human tissues (5), have spurred efforts to develop anticancer therapies based on inhibiting lipogenic enzyme activities or silencing the corresponding genes.

Attempts to exploit the metabolic requirements of lipogenic cancers have thus far focused solely on disrupting de novo FA synthesis. Cytotoxicity following inhibition of lipid synthesis, however, may be obviated by the provision of exogenous FA (6–8). This observation, and the improved outcome of breast cancer patients ingesting a low fat diet (9), led us to hypothesize that triglyceride in circulating lipoprotein particles could provide an additional, exogenous source of FA for tumors. This would require triglyceride-rich chylomicrons or very low density lipoproteins (VLDL) as substrate, extracellular lipoprotein lipase (LPL) for hydrolysis, and FA translocase (CD36) for cellular uptake of the free FA (reviewed in ref. 10). As LPL is a secreted enzyme that is bound to the luminal surface of capillary endothelial cells, it could potentially be supplied by tumor cells or by nonmalignant cells in the tumor microenvironment.

Production of the expression dataset has been previously described in detail, as have culture conditions for cell lines (ref. 11; http://cancer.lbl.gov/breastcancer/data.php). RNA from 45 human breast cancer cell lines (ICBP45) grown at subconfluence was harvested, reverse transcribed, and hybridized to Affymetrix U133A gene chips. Resulting Affymetrix image files were normalized (RMA; ref. 12). Unsupervised average linkage cluster analysis of log₂ signal intensities was done using approximately 14,000 probeset IDs of highest variance, using the Cluster Software package, and the resulting dendrogram image produced with Treeview (ref. 13; http://rana.lbl.gov/eisen/?page_id=7). Probeset IDs identifying CD36, LPL, and FASN were identified, median centered, normalized, and a heat map produced indicating the relative hybridization intensity for each sample.

RNA was isolated by the RNeasy Mini Kit (Quiagen). One microgram RNA was reverse transcribed by random hexamer primers with M-MULV reverse transcription (New England Biolabs). PCR was done as described (14). Primers used are described in Supplementary Table S1. Different primers were used for real-time RT-PCR.

RNA was prepared by the PureLink Total RNA purification system (Invitrogen). The purity and concentration of RNA were assessed by a NanoDrop DM-1000 spectrophotometer (NanoDrop Technologies). RNA was converted to cDNA by Superscript II RT and random hexamer primers, according to the manufacturer's protocol (Invitrogen). Primer sequences for LPL were 5′-TATCCGCGTGATTGCAGAGA-3′ (forward) and 5′-GCCTTACTTGGATTTTCTTCATTCA-3′ (reverse). SYBR green was used for detection and 18S rRNA was used as an internal control. Primer sequences for 18S were 5′-CGCCGCTAGAGGTGAAATTC-3′ (forward) and 5′-TTGGCAAATGCTTTCGCTC-3′ (reverse). PCR was in the 7500 Fast Real-Time PCR System (Applied Biosystems). The program used included 2 minutes at 50°C, 1 minute at 95°C, and 40 cycles of 3 seconds at 95°C and 30 seconds at 60°C. The average of the C_t values for each triplicate reaction was expressed relative to the amount of 18S rRNA in the sample.

LiSa-2 liposarcoma cells were from Martin Wabitsch (University of Ulm, Germany) and we confirmed their identity by the ability to produce lipid droplets that stained with oil red-O on confluence. All other lines were from the American Type Culture Collection except VCaP, which was from ECACC, and these lines were acquired recently and were of low passage number (<10). Cells were grown in DMEM:F12 supplemented with 10% fetal calf serum (FCS; Atlanta Biologicals), 1% penicillin/streptomycin, and 2 mmol/L l-glutamine, in 5% CO₂ at 37°C. Cell growth was monitored in an MTT assay (15). Lipoprotein-deficient FCS was prepared by the method of Goldstein and coworkers (16).

HeLa cells were transfected with 10 nmol/L siRNA targeting LPL (siRNA A: 5′-GGUAGAUAUUGGAGAACUA; siRNA B: 5′-GGAUGGAGGAGGAGUUUAA; Dharmacon) with the use of Lipofectamine RNAiMAX (Invitrogen). A siRNA targeting luciferase (5′-CGUACGCGGAAUACUUCGA) was used as control. RNA was harvested 48 hours later, and cell viability was assessed 96 hours after transfection.

Dialyzed VLDL (0.5 mg; Kalen Biomedical) were incubated at 37°C for 15 hours with 25 μL diI D282 (Invitrogen; 3mg/mL in dimethyl sulfoxide). Free dye was removed by dialysis against PBS. Prior to observation by confocal microscopy, 25 μL labeled VLDLs were added to slide chambers (Tissue Tek #177402) containing 10⁶ cells grown for 3 days in delipidated Dulbecco's Modified Eagle's Medium. Excitation was at 549 nm, and fluorescence was detected at 565 nm.

Mice were immunized with a peptide (Sigma) representing human LPL residues 21–36 (CASRGGVAAAQRRDFID) coupled to keyhole limpet hemocyanin. After fusion of splenocytes to mouse multiple myeloma cells, media from candidate clones were screened for reactivity to bacterially expressed LPL in an enzyme-linked immunosorbent assay. Positive clones were further screened by Western blot of skeletal muscle from transgenic mice expressing a human muscle-specific LPL transgene (MCK-LPL, kindly supplied by Ira Goldberg, Columbia School of Medicine, New York, NY) and against human breast milk. Mouse tissues were homogenized in immunoprecipitation assay buffer containing 10 μg/mL phenylmethylsulfonylfluoride. Samples were centrifuged at 10,000 × g for 10 minutes × 2. Protein content was determined by the BCA assay (Pierce). Samples were boiled in 2X sample buffer and fractionated through 15% acrylamide. Following transfer to polyvinylidene difluoride (PVDF) membranes (Immobilon–FL; Millipore), blocking was with SuperBlock (Pierce). Incubation with 1:200 dilution of the primary antibody in TBS-Tween was overnight at 4°C, followed by 2 TBS washes. Recombinant protein A/G conjugated with horseradish peroxidase (Pierce) was applied for detection at 1:5,000 in TBS-Tween for 1 hour at RT. After 4 TBS washes, membranes were developed with NBT-BCIP (Pierce).

The 2.3 kb EcoRI–HindIII fragment coding LPL was excised from pCMV-SPORT6-LPL (Open Biosystems) and inserted into the pProEx-HTa His-tag vector. Most of the fusion protein could not be solubilized, but those recovered showed reactivity with the anti-His4 antibody (Invitrogen). For immunodot assays 10 ng protein from cleared lysates of Escherichia coli DH5α transformed with empty or LPL plasmid were spotted onto PVDF membranes, blocked, and incubated with antibody as described earlier in the text.

Human milk and conditioned cell culture media were fractionated over heparin sepharose (Sigma) by a procedure modified from Hata and colleagues (17).

We used the radiochemical assay of Nilsson-Ehle and Schotz (18) or a colorimetric assay based on determination of glycerol production (BioVision). We used a protocol based on that of Cruz and colleagues (19) for determination of heparin-releasable LPL. Briefly, 5 × 10⁶ cells from 75 cm² flasks were cultured for 72 hours, and scraped pellets were washed 3 times in PBS with or without 100 U/mL heparin. Media and lysed cell pellets were assayed in triplicate for residual LPL activity.

Immunohistochemistry was done as previously described (20). Anti-LPL monoclonal antibody clone 43 was used at a dilution of 1:10, with Citra Plus antigen retrieval (Biogenix). CD36 was assessed by an affinity-purified rabbit polyclonal antibody (Thermo Scientific) according to the supplier's protocol. The Institutional Review Board (IRB)-approved the use of breast cancer tissue and the tissue microarray containing 147 primary breast cancers from postmenopausal women, diagnosed between 2000 and 2007 at Dartmouth-Hitchcock Medical Center, Lebanon, NH. Each case was represented by one tissue core 1.0 mm in diameter. The liposarcoma tissue microarray, also IRB approved, contained 26 liposarcomas diagnosed between 1995 and 2008 at Dartmouth-Hitchcock Medical Center. Each case was represented by two to four 1.0-mm tissue cores. Prostate cancer specimens were acquired at the Katholieke Universiteit Leuven, Belgium, with IRB approval.

We used a cDNA microarray to screen 45 breast cancer–derived cell lines from the dataset of Neve and colleagues (11) for LPL gene expression, and for FASN mRNA as a marker for de novo FA synthesis. We analyzed cell lines because breast tumor samples may contain adipocytes, which express high levels of LPL and FASN. We also sorted the breast cancer lines by their global gene expression signatures (21). These signatures include the luminal type (estrogen receptor–positive; ER⁺), the basal, or triple-negative type that lacks receptors for estrogen, progestin, and trastuzumab (22), and the type with Her2/neu amplification. Only 6 breast cancer cell lines (HCC2157, HCC1008, HCC1599, Du4475, SUM149, and SUM190) expressed high levels of LPL mRNA, and each of these exhibited the aggressive basal gene expression signature (Supplementary Fig. S1). Expression of LPL mRNA by selected cell lines was verified by RT-PCR (Fig. 1A), as was expression of CD36 mRNA (Fig. 1B). LiSa-2 liposarcoma cells, which we previously showed to exhibit the lipogenic phenotype (23), also expressed LPL and CD36, as expected for a tumor cell derived from an adipocytic lineage. All of the cell types expressed substantial FASN mRNA (Fig. 1C), and in the breast cancer cell lines this did not vary among the gene expression signatures (Supplementary Fig. S1). Quantitative real-time RT-PCR of representative lines confirmed that LiSa-2 liposarcoma and triple-negative Du4475 breast cancer cells expressed the highest levels of LPL mRNA (Fig. 1D). In contrast, prostate cancer cells, which are highly lipogenic (24), expressed relatively low levels of LPL mRNA, and ER⁺ T47D and BT474 breast cancer cells expressed essentially none.

We examined conditioned tissue culture media for LPL enzyme activity, and it paralleled the levels of LPL mRNA (Fig. 2A). LPL activity accumulated over time in culture media of LiSa-2 liposarcoma and Du4475 breast cancer cells (Fig. 2B). In contrast, ER⁺ T47D, ER⁺ Her2/neu⁺ BT474 breast cancer cells, and fibroblasts did not secrete detectable lipase activity. Prostate cancer cells produced low levels of the enzyme. LPL activities in breast milk and murine striated muscle were substantially greater than those observed in any of the conditioned (72 hours) media.

We found that available antibodies were not sufficiently specific to analyze LPL protein by immunohistochemistry. We therefore raised a mouse monoclonal antibody using a peptide representing residues 20 to 36 of the human enzyme as antigen. This antibody is highly specific (Supplementary Fig. S2), and permitted detection of heparin sepharose–purified LPL from tissue culture media conditioned by Du4475 breast cancer and LiSa-2 liposarcoma cells (Fig. 2C, top). The band recognized by this antibody in Western analysis of milk was verified to represent LPL by mass spectrometry. We could not detect LPL protein in media from ER⁺ breast or prostate cancer cells. Western analysis of a clinical breast tumor homogenate (50 μg protein) without affinity purification revealed a single band exhibiting the same migration as that observed in milk (Fig. 2C, bottom).

It seemed possible that expression of heparanase could inactivate LPL, and thus could vitiate the metabolic relevance of LPL expression by tumors. We assessed expression of the heparanase gene (HPSE) using cDNA microarray data from 45 human breast cancer cell lines. This showed that the cells generally express very low levels of heparanase mRNA, as was the general case for LPL mRNA. We were intrigued to note that the subgroup of triple-negative cell lines exhibiting substantial LPL expression also expressed the lowest levels of heparanase mRNA. Indeed, linear regression of the relationship between LPL and heparanase mRNAs in lines with the basal A signature revealed a statistically significant inverse correlation (P = 1.27 × 10⁻⁵, R² = 0.38). Thus, the coupling of high LPL with low heparanase expression seems to provide an advantage to the subset of cells that produce substantial LPL. Our examination of total and heparin-releasable LPL activity in a freshly prepared breast tumor homogenate also reflects on this question, as heparin-releasable activity was readily detectable, arguing against depletion of a cell surface–bound LPL pool in breast tumors (see in the following text).

We carried out 2 experiments to determine whether cancer-associated LPL is bound to tumor cells by noncovalent interactions with cell surface heparan sulfate proteoglycans, using a protocol based on that of Cruz and colleagues (19). First, we homogenized freshly resected invasive breast cancer tissue shown to contain LPL immunoreactivity (Fig. 2C, bottom), and extracted equal aliquots with buffer containing or not containing heparin. LPL activity in the control sample was 1,032 ± 8 without heparin, 768 ± 4 with heparin treatment (mean ± SE, nmol/L glycerol produced/g tumor/h, measured in triplicate; P < 0.0001). This represented a heparin-releasable fraction of 26% of the total tumor-associated LPL activity (represented by the portion of the bar labeled HR, Fig. 2D, left).

Second, we determined the heparin-releasable fraction of LPL in HeLa cells, and calculated turnover rates for cellular LPL pools (Fig. 2D, right). Residual LPL activity in cell pellets was 13,260 ± 1,080 without, and 9,360 ± 820 with heparin exposure (units are nmol/L glycerol produced/flask/h, mean ± SEM, n = triplicate measurements/group; P < 0.04). We thus estimate that 29% of the HeLa cell–associated pool of LPL is heparin-releasable (indicated by HR on the graph), a fraction similar to that observed in the breast tumor sample. Measurement of LPL activity in culture media indicated that 36,000 ± 4,000 units of LPL activity were secreted per 24 hours. We therefore estimate that the total cellular LPL pool turns over more than 2.7 times/d, whereas the heparin-labile pool (3,900 units/well) turns, presumably by secretion, more than 9.2 times/d.

Our FCS contained 660 μg triglyceride/mL. LPL secreted by cells is removed when culture media are replaced, so the enzyme content in tissue culture never approaches that observed in tissues. We therefore assessed the functional significance of LPL by adding the enzyme to media containing 10% FCS and measuring cell accumulation. LPL activity under these culture conditions approximated that observed in mouse muscle. LPL enhanced the growth of T47D breast cancer cells, which do not express LPL, and of LiSa-2 liposarcoma cells, which express LPL (Fig. 3A and B). This effect of LPL was greatly reduced in media containing FCS that was nearly depleted of trigyceride (20 μg/mL).

LNCaP prostate cancer cell growth was not accelerated by LPL addition. The ability of these cells to use exogenous triglyceride-derived FA to maintain growth was revealed, however, in the presence of soraphen A, a potent inhibitor of the lipogenic enzyme acetyl CoA-carboxylase (7). The cells were rescued from Soraphen A–induced cytotoxicity by provision of LPL in the presence, but not in the absence, of lipoproteins (Fig. 3C). Experiments using PC3 prostate cancer cells yielded similar results (Fig. 3D).

In complementary studies we assessed the impact of siRNA-mediated knockdown of LPL mRNA on the growth of HeLa cells, which we previously reported to express the LPL gene (25), and its interaction with inhibition of lipogenesis by soraphen A. Two different siRNAs each caused greater than 90% disappearance of LPL mRNA, whereas a nonspecific siRNA was without effect (Fig. 3E). Soraphen A caused a major inhibition of HeLa cell accumulation, and this effect was prevented by provision of LPL to the cultures (Fig. 3F). Transfection of LPL siRNA A or B, but not of the nonspecific siRNA, significantly inhibited HeLa cell growth, and the anticancer effects of the 2 LPL siRNAs were further enhanced by exposure to soraphen A.

We used immunohistochemistry to assess the relevance of our findings in cultured cells to human tumors. We assessed the expression of markers of de novo lipogenesis [FASN, THRSP (Spot 14, S14)], lipolysis (LPL), and exogenous FA uptake (CD36) in a panel of 147 breast, 24 liposarcoma, and 10 prostate tumor tissues (examples in Fig. 4). FASN was cytosolic, in agreement with previous studies. S14, which promotes expression of the FASN gene (26, 27), was primarily nuclear, as reported (20).

In contrast to our findings in breast cancer cell lines, LPL immunoreactivity was observed in all of the breast tumors examined, and, also in contrast to the cell lines, expression was not limited to triple-negative tumors. Similarly, all liposarcoma and prostate tumors examined expressed readily detectable LPL by immunohistochemistry. Intracellular LPL showed an asymmetric, perinuclear distribution suggestive of localization to the Golgi apparatus, as predicted for a glycosylated and secreted protein (Fig. 4C, insets). As expected, extracellular LPL was found on the luminal surfaces of capillaries (Supplementary Fig. S2C, left). We stained tonsil tissue as a negative control, based on previous work showing undetectable LPL mRNA in lymphoid cells (25). The lymphoid stroma indeed showed no staining except for scattered isolated monocytes, whereas the highly proliferative basal (stem cell) layer of the mucosal epithelium overlying the tonsil unexpectedly showed a strong signal (Supplementary Fig. S2C, right).

The majority of tumors also stained for CD36 (Fig. 4D). Interestingly, 2 distinct staining patterns were observed in breast cancer tissue. Of the 144 evaluable cores, 42 exhibited diffuse cytoplasmic staining without accentuation at the plasma membrane (Fig. 4D, left, top), whereas 100 also showed a strong cell surface signal (Fig. 4D, left, bottom). Only two breast cancer cases were devoid of staining. A statistically significant difference in the prevalence of the membranous staining pattern between the triple-negative and ER⁺ breast cancers was shown by χ² analysis (42% vs. 76%, P < 0.02).

Of the 25 liposarcoma cases, 21 stained for CD36, almost all in a mixed cytoplasmic plus plasma membrane pattern (Fig. 4D, middle), including all 9 cases of well-differentiated liposarcoma. Of the 9 evaluable prostate cancers, 4 showed focally positive staining in a mixed cytoplasmic and plasma membrane pattern (Fig. 4D, right), whereas 5 cases scored negative for CD36.

Expression of LPL by breast cancer cells suggested the possibility that the cells could use the enzyme not only to hydrolyze extracellular triglyceride, but also for receptor-mediated endocytosis of triglyceride-rich lipoproteins. This process uses LPL as a bridge between the cell surface receptor syndecan-1 and the lipoprotein (28). RT-PCR revealed readily detectable syndecan-1 mRNA from DU4475 breast cancer cells, whereas LiSa-2 and T47D exhibited a faint signal (Supplementary Fig. S3A). We incubated fibroblasts and DU4475 cells with fluorescently labeled VLDL particles, and assessed for cellular uptake by confocal microscopy. Abundant uptake was observed in fibroblasts (Supplementary Fig. S3B) but not in DU4475 cells (Supplementary Fig. S3C). Occasional fluorescence was detected on the cell surface (Supplementary Fig. S3D) but never within the breast cancer cells.

Our data show that cancer cells may use two different mechanisms to acquire FA to fuel proliferation. Breast and liposarcoma tumors are equipped for both lipid synthesis and for LPL-mediated extracellular lipolysis followed by FA uptake via CD36. Prostate cancer cells, which have a very high capacity for de novo lipogenesis (24), express very little LPL. The low LPL expression could be explained in part by the reported loss of heterozygosity at the LPL locus in 47% of prostate tumors, owing to the presence of a nearby tumor suppressor gene (29). These cells, however, can acquire sufficient exogenous FA to maintain growth in the face of FA synthesis inhibition when they are supplied with LPL and triglyceride-rich lipoprotein particles.

LPL expression has been shown to be a marker of poor prognosis in chronic B-cell lymphocytic leukemia (B-CLL; refs. 30, 31). The single reported examination of the functional significance of LPL in B-CLL was difficult to interpret because Orlistat, a compound that inhibits both LPL and FASN (4), was used to inhibit LPL in those studies (32). To our knowledge, these are the first experiments to show widespread expression of LPL by solid tumors. We find that, in contrast to cultured breast cancer cell lines, where substantial LPL is found only in a subset with a triple-negative gene expression signature, the enzyme is a universal component of breast tumors, irrespective of biomarker status. Moreover, we also find that all liposarcoma and prostate tumors examined also express LPL.

Several plausible explanations exist for the discrepancy between cell lines and tumors with respect to LPL expression. First, the cell lines have been passaged over time in culture systems lacking vascular endothelium, which is the physiologic site for LPL action, or reliably fixed concentrations of triglyceride-rich lipoprotein substrate, whereas cell culture media generally contain high concentrations of glucose. Thus, de novo synthesis, rather than lipolysis or receptor-mediated endocytosis, may have been selected as the preferred mechanism for FA acquisition in cell culture. Second, it is possible that interactions with stroma elicit LPL expression. Third, each of the breast cancer cell lines that we find to express substantial levels of LPL are not only triple-negative, but are also nonadherent to tissue culture plasticware. In view of reports that cellular detachment provokes major metabolic adaptations in Her2/neu-expressing breast cancer cells (33), we examined the hypothesis that cellular detachment (72 hours) would provoke enhanced LPL mRNA expression. This proved, however, not to be the case (data not shown). Irrespective of the cause of the discrepancy, it is important to recognize that tissue culture experiments may not faithfully recreate in vivo physiology.

Efficient utilization by cancer cells of FA released by extracellular lipolysis would require the expression of both LPL and CD36. It was therefore not surprising to find CD36 expression in the majority of tumor tissues examined. CD36 is known to traffic from cytoplasm to the plasma membrane in response to insulin stimulation of adipocytes (34). We observed cell surface localization in ∼70% of breast cancers, whereas ∼30% exhibited only a cytoplasmic signal. On the basis of our observation that cell surface staining was significantly less frequent in triple-negative tumors, we speculate that CD36 trafficking may be driven by cell surface acting growth factors and/or sex steroids in breast cancers.

Although further experiments are required to delineate the precise roles of lipogenesis and lipolysis in transformation, proliferation, and metastasis, recent studies have advanced this area. Previous work established a tight linkage of enhanced FA synthesis to transformation (35), and recent studies have defined the role of an intracellular lipase, monoacyl glycerol lipase in promoting tumorigenesis. Monoacyl glycerol lipase provides, by de-esterification, a stream of intracellular free FA to fuel proliferation, growth, and migration (36). This study shows a complementary role for LPL, an extracellular lipase, in providing a stream of FA to fuel cancer cell proliferation.

Various hypotheses have been proposed to explain the dependence of tumors on lipogenesis, but it is clear that the primary metabolic fate of FA in proliferating tumor cells is incorporation into phospholipids destined for membrane biosynthesis (37, 38). As mitochondrial production and export of citrate are the key steps required to maintain de novo lipogenesis in the cytosol, this begs the question of how such mitochondrial metabolism may be maintained under the hypoxic (but not anoxic) conditions that prevail in tumors. Indeed, hypoxia-induced factor-1, a key mediator of the cellular response to hypoxia, reduces the fractional entry of glucose-derived carbon into mitochondria by downregulating pyruvate dehydrogenase, thus driving the increased lactate production that is the most well-recognized aspect of intermediary metabolism in tumors (39). However, net flux of carbon through the glycolytic pathway is substantially elevated in glucose-avid tumor cells, because of increased uptake and trapping. The reduced amount of carbon directed to mitochondria is thus sufficient to provide an estimated 60–85% of the ATP generated (40). Brisk citrate export from mitochondria seems to be favored by incomplete combustion, as a consequence of the truncated Krebs cycle in tumor mitochondria (41), which also may serve to reduce oxygen use by reducing carbon flux through steps downstream from citrate in the cycle. Thus, the competing oxygen-sparing and anabolic demands on tumors are met by a balanced set of metabolic alterations, the former favored by hypoxia-induced factor-1, and the latter driven by oncogenes (reviewed in ref. 42). Overall, it seems that the uptake of exogenous FA, for which this study shows most tumors to be equipped, would be an advantageous response to the metabolic dilemma of hypoxic, proliferating cancer cells.

Our findings have several implications. First, therapeutic efforts aimed solely at inhibition of long-chain FA synthesis may not be effective for tumors that are provided with LPL and express CD36. Such tumors may be sensitive to agents that inhibit the enzymes for both lipogenesis and lipolysis, such as Orlistat (4) or the dietary supplement conjugated linoleic acid, which can suppress the genes required for both pathways (8, 43). Efforts to target LPL will need to take into account the possibility that prolonged systemic suppression of LPL activity could result in hypertriglyceridemia and consequent pancreatitis, particularly if dietary fat intake is not curtailed. Second, the ability of nearby nonmalignant cells to provide LPL to the tumor microenvironment may favor the ability of tumor cells, particularly those with low lipogenic potential, to establish metastases in LPL-rich tissues such as lung or fatty bone marrow. To benefit from LPL provided by tumor stroma, the expression of CD36 by the tumor would be required. Third, the presence of LPL in the tumor vasculature may mediate the reported effects of dietary fat intake on outcome (9). In addition to the well-characterized lipogenic tumor phenotype, our studies indicate the expression of a previously unappreciated lipolytic pathway active in cancer cells as well.

No potential conflicts of interest were disclosed.

MCK-LPL transgenic mice were kindly supplied by Ira Goldberg, Columbia School of Medicine, New York, NY. We thank Martin Wabitsch (University of Ulm, Germany) for the LiSa2 cells. Soraphen A was kindly provided by Klaus Gerth and Rolf Jansen, Helmholtz-Zentrum für Infektionsforschung, Braunschweig, Germany. Triglyceride measurements were kindly provided by Hong K. Lee, Department of Pathology, Dartmouth-Hitchcock Medical Center, Lebanon, NH. We specially thank Rebecca O'Meara MT (ASCP), Pathology Translational Research Laboratory at Dartmouth Medical School, for doing the immunohistochemistry.

This work was supported by NIH Grant RO1CA126618 (W.B. Kinlaw), NIH Training Grant DK07508 (N.B. Kuemmerle), a Howard Hughes Medical Foundation Fellowship 52005870 (A.J. Flanagan), Norris Cotton Cancer Center Prouty grants (B.L. Eisenberg, W.B. Kinlaw), Grant G.0590.08 (J.V. Swinnen), a fellowship (E. Rysman) from the Research Foundation-Flanders (FWO), the N.C.I. Bay Area Breast Cancer SPORE P50 CA58207 (L.A. Timmerman), and the Program in Experimental and Molecular Medicine at Dartmouth Medical School (C.J. Fricano).

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