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Carcinogenesis Advance Access originally published online on May 15, 2006
Carcinogenesis 2006 27(10):1950-1960; doi:10.1093/carcin/bgl023
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© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Arachidonic acid-induced gene expression in colon cancer cells

Arta M. Monjazeb, Kevin P. High2, Abbie Connoy3, Lori S. Hart, Constantinos Koumenis1 and Floyd H. Chilton4,*

Department of Cancer Biology, Wake Forest University Baptist Medical Center Medical Center Boulevard, Winston-Salem, NC 27157, USA
1 Department of Radiation Oncology, Department of Internal Medicine, Wake Forest University Baptist Medical Center Medical Center Boulevard, Winston-Salem, NC 27157, USA
2 Section of Infectious Diseases, Wake Forest University Baptist Medical Center Medical Center Boulevard, Winston-Salem, NC 27157, USA
3 Section of Molecular Medicine, Wake Forest University Baptist Medical Center Medical Center Boulevard, Winston-Salem, NC 27157, USA
4 Department of Physiology and Pharmacology, Wake Forest University Baptist Medical Center Medical Center Boulevard, Winston-Salem, NC 27157, USA

*To whom correspondence should be addressed. Tel: +1 336 713 7105; Fax: +1 336 713 7168; Email: schilton{at}wfubmc.edu


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 References
 
It is well documented that arachidonic acid (AA) and its metabolites are intimately linked to cancer biology. However, the downstream mechanism(s) that link AA levels to cancer cell proliferation remain to be elucidated. Initial experiments in the current study showed that exogenous AA and inhibitors of AA metabolism that lead to the accumulation of unesterified AA are cytotoxic to the colon cancer cell line, HCT-116. Additionally, exogenous AA and triacsin C, an inhibitor of AA acylation, induced apoptosis and related caspase-3 activity in a transcriptionally dependent manner. Gene array analysis revealed that both exogenous AA and triacsin C alter the expression of similar genes in HCT-116 cells. For example, both downregulate several genes with well-documented roles in cell survival and apoptotic resistance. Conversely, both upregulate genes encoding activator protein-1 (AP-1) transcription factors, which have known roles in inducing apoptosis, and genes that counteract ras (Erk/MAPK) growth signaling pathways. Real-time polymerase chain reaction and immunoblotting demonstrated that mRNA and protein levels of one of the major AP-1 transcription factors, c-Jun, is markedly elevated by exogenous AA and triacsin C. Additionally, the cyclooxygenase inhibitor, sulindac sulfide, increases c-Jun mRNA levels. Together, these studies reveal that the generation of intracellular AA and its subsequent impact on gene expression probably represents a critical step that regulates colon cancer cell proliferation.

Abbreviations: AA, arachidonic acid; ActD, actinomycin D; AP-1, activator protein-1; CoA-IT, CoA-independent transacylase; COX, cyclooxygenase; cPLA2, cytosolic phospholipase A2; DRB, 5,6-dichloro-1-b-D-ribofuranosylbenzimidazole; FACL-4, fatty acid CoA ligase 4; NSAIDs, non-steroidal anti-inflammatory drugs; PBS, phosphate-buffered saline; PCR, polymerase chain reaction


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 References
 
Arachidonic acid (AA; 20:4, n-6) is a relatively minor polyunsaturated fatty acid found at the sn-2 position of cell membrane glycerolipids (1). In contrast to other more abundant unsaturated fatty acids (i.e. oleic, linoleic or linolenic acid), levels of unesterified AA are stringently controlled within mammalian cells (2). Pathways of AA uptake, incorporation and remodeling in glycerolipids are well documented (2). Two distinct pathways, one that controls exposure of cells to low concentrations of AA and another that controls exposure to high AA concentrations, have been identified. The first is a ‘high-affinity–low-capacity’ pathway that incorporates low concentrations of intracellular AA into glycerolipids; this is thought to be the major pathway responsible for phospholipid–arachidonate remodeling. Then there is a ‘low-affinity–high-capacity’ pathway that incorporates unesterified AA primarily into triglycerides (often seen in lipid bodies) and diarachidonyl phospholipids. This pathway functions primarily when the high-affinity pathway has been saturated when cells are exposed to high AA concentrations (2) (Figure 1). AA is also released from membrane glycerolipids, and once released, AA and its metabolites are important signals that regulate a wide range of cellular functions (3–5).


Figure 1
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  		Fig. 1 Schematic of AA metabolic pathways. Inhibitors of AA metabolic enzymes are underlined. In contrast to other abundant unsaturated fatty acids, levels of unesterified AA are tightly controlled by two distinct pathways working in coordination. The first pathway is referred to as ‘high-affinity–low-capacity’ because it incorporates low concentrations of intracellular AA into glycerolipids. This is the predominant pathway utilized. When the burden of unesterified AA overwhelms the low-capacity pathway, the cell increases incorporation of unesterified AA into neutral lipids via de novo glycerolipid biosynthesis utilizing a ‘low-affinity–high-capacity’ pathway. FACL-4 catalyzes an ester bond between AA and coenzyme A, which is the first reaction in both pathways. CoA-IT orchestrates the remodeling of AA between phospholipid pools such as phosphatidylcholine and phosphatidylethanolamine in a CoA-independent manner (2). Thus, PLT 98625, via inhibition of CoA-IT, impedes only the ‘high-affinity–low-capacity’, whereas triacsin C, via inhibition of FACL-4, inhibits both pathways.

 
Over the past decade, it has become apparent that AA metabolism plays a key role in carcinogenesis. Such data are particularly compelling with regard to cyclooxygenase (COX) mediated AA metabolism (6). For example, the role of COX overexpression (7–12) and COX-derived eicosanoids (13–16) in carcinogenesis as well as the antineoplastic effects of non-steroidal anti-inflammatory drugs (NSAIDs) (17–20) are now firmly established. Similar lines of evidence have implicated the overexpression of other AA metabolic enzymes in cancer cell survival (21,22) and have demonstrated the anti-neoplastic properties of inhibitors of these enzymes (23–25).

The capacity of inhibitors of enzymes that metabolize AA to induce cancer cell apoptosis is of therapeutic interest, and the mechanism by which these inhibitors induce apoptosis is still a matter of debate. With regard to NSAIDs, the most well studied of these inhibitors, it is unclear whether their anti-neoplastic effects result from COX inhibition or COX-independent targets. For example, NSAIDs have been shown to effect putative targets such as PDK1, NAG-1, Wnt, Akt and others (reviewed in ref. 26).

One hypothesis that is consistent with much of the data in the literature is that certain inhibitors of AA metabolism induce the intracellular accumulation of unesterified AA, which provides signals for cells to undergo apoptosis (4,24,27) (Figure 1). This notion is supported by studies that show that inhibition of COX, AA–CoA synthetase [also known as fatty acid CoA ligase 4 (FACL-4)] (Figure 1) (2) and CoA-independent transacylase (CoA-IT) (Figure 1) (2) induce the accumulation of unesterified AA in cancer cells at concentrations that lead to apoptosis (24,27,28). Moreover, addition of exogenous AA to cancer cell lines leads to growth inhibition and apoptosis. Furthermore, the overexpression of enzymes such as COX, LOX (21,29) and FACL-4 (22), all of which lower intracellular AA levels, provides an apoptotic escape mechanism in several neoplastic cell types. Interestingly, Dong et al. (30) have recently described a low cytosolic phospholipase A2 (cPLA2)/high cyclooxygenase-2 phenotype in human colon tumors. This phenotype would be predicted to maintain very low levels of intracellular AA and they hypothesize that establishment of this phenotype is a key step in early carcinogenesis. Finally, we have recently demonstrated that NS-398, PLT 98625 and triacsin C (inhibitors of COX, CoA-IT and FACL-4, respectively) induce accumulation of high levels of unesterified AA and apoptosis, and, in combination, act synergistically to produce both AA accumulation and apoptosis (31).

However, a key question remains as to how accumulation of intracellular AA initiates apoptosis. The range of biological processes in which AA and its metabolites participate is vast, and thus an extensive number of possibilities exist. In this study we investigate changes in the gene expression induced by the accumulation of intracellular unesterified AA. AA and its metabolites have been shown to regulate the transcription of several families of genes including heat shock proteins (32,33), genes involved with cell cycle control (34), inflammation (35,36), steroid biosynthesis (37) and proto-oncogenes (38–40). The current study demonstrates that AA-induced apoptosis is transcriptionally dependent and involves the regulation of key families of genes involved in cell survival and apoptosis.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 References
 
Materials
All cell culture reagents were purchased from Invitrogen (San Diego, CA). HCT-116 cell lines were maintained in McCoy's medium supplemented with 10% heat-inactivated fetal bovine serum (FBS), L-glutamine and 100 U/ml penicillin–streptomycin. Triacsin C and sulindac sulfide were purchased from Biomol Research Laboratories (Plymouth Meeting, PA). PLT 98625 (formerly SK&F 98625) has been described previously (41) and was provided by Pilot Therapeutics (Winston-Salem, NC). Fatty acids were purchased from Cayman Chemical (Ann Arbor, MI). Etoposide, actinomycin D (ActD), 5,6-dichloro-1-b-D-ribofuranosylbenzimidazole (DRB) and albumin were purchased from Sigma Chemical (St Louis, MO). [3H]AA (200 µCi/nmol) was purchased from American Radiolabeled Chemical (St Louis, MO). Rabbit immunoaffinity-purified antibody to phospho-Jun was purchased from Upstate (Lake Placid, NY). Rabbit anti-Jun IgG was purchased from Cell Signaling Technology (Beverly, MA). Goat anti-rabbit IgG and anti beta-actin antibodies were purchased from Santa Cruz Biotechnologies (Santa Cruz, CA).

MTS cell proliferation assay
Cell proliferation was quantified using the CellTiter 96 Aqueous MTS [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium)] colorimetric assay kit (Promega, Madison, WI). Cells were plated in appropriate medium at 10 000 cells/well in 96-well tissue culture plates and incubated at 37°C for 24 h. For experiments utilizing the transcriptional inhibitors DRB (50 µM) or ActD (1 µg/ml), cells were pre-treated with these inhibitors for 1 h. The appropriate treatments or vehicle were then added to each well and cells were incubated for an additional 48 h. The assay was then developed according to manufacturer's instructions and absorbance at 490 nm was measured.

Labeling of glycerolipids with [3H] AA to isotopic equilibrium and analysis of labeled products
Cellular glycerolipids were labeled to isotopic equilibrium as described previously (28). Briefly, HCT-116 cells at 80% confluency were pulse-labeled with [3H] AA (1 µCi/106 cells) in Hanks' buffered saline containing essentially fatty acid-free human serum albumin (0.25 µg/ml) for 60 min at 37°C. Cells were then washed several times to remove any exogenous [3H]-labeled AA that was not internalized during the pulse. Subsequently, cells were incubated for 48 h to allow [3H] AA to reach constant label to mass ratios within all intracellular glycerolipid AA pools. Cells were then treated with triacsin C, PLT 98625, etoposide or vehicle for 18 h at 37°C. Intra- and extracellular lipids were extracted by the method of Bligh and Dyer (42). Extracted lipids were combined with cold glycerolipid standards, spotted on silica gel G TLC plates (Analtech, Newark, DE) and developed using hexane : ethyl ether : formic acid (90 : 60 : 6 v/v). TLC zones were collected and analyzed by liquid scintillation spectroscopy to determine the amount of radioactive AA in each lipid pool.

RNA isolation
HCT-116 cells were seeded at 2 x 106 cells/dish in a 100 mm culture dish and incubated for 24 h. Cells were then treated with triacsin C (20 µM), AA (200 µM), sulindac sulfide (160 µM) or vehicle alone for 6 or 12 h, and total cellular RNA was isolated using TriZol reagent (Gibco, Grand Island, NY) following the manufacturer's instructions. For gene chip expression analysis each sample consisted of two pooled plates.

Real-time reverse transcription–polymerase chain reaction (RT–PCR)
Reverse transcription of total RNA was performed with the Promega Reverse Transcription kit (Promega, Madison WI) according to manufacturer's instructions. Real-time PCR was performed using the ABI Prism 7700 instrument. PCR consisted of RT cDNA product (20–200 ng), 100 nM primer and 25 µl 2x Sybr Green PCR Master Mix [Sybr Green dye, AmpliTaq Gold, DNA polymerase, dNTP Mix, optimized buffer components; Applied Biosystems, Foster, CA] in a total volume of 50 µl. Amplification was performed for 35 cycles. Following an initial hot start of 10 min, each cycle consisted of 15 s of denaturation and 1 min of annealing at proper melting temperature (Tm). The correct sequence of the amplicon was confirmed by sequence analysis, and the correct size confirmed by gel electrophoresis. mRNA expression levels were determined and standardized to GAPDH (Applied Biosystems, Foster, CA). Primer pairs were forward: 5'-GCATGAGGAACCGCATTGCCGCCTCCAAGT-3' and reverse: 5'-GCGACCAAGTCCTTCCCACTCGTGCACACT-3' for c-Jun; and forward: 5'-TCCTCTGACTTCAACAGCGACACC-3' and reverse: 5'-TCTCTCTTCCTCTTGTGCTCTTGG-3' for GAPDH (43). Both pairs have a Tm of 57°C.

Gene chip expression analysis
RNA isolation, preparation and hybridization to the Human Genome U133A Array (Affymetrix, Santa Clara, CA) were performed according to the manufacturer's instructions. Isolated total cellular RNA was purified using an RNeasy Mini Kit (QIAGEN, Valencia, CA). Isolated and purified RNA (10 µg) was reverse-transcribed to cDNA using a Superscript II RT cDNA Synthesis Kit (Invitrogen, Carlsbad, CA) and T7 Oligo(dT) gene chip primers (Affymetrix). Biotin-labeled anti-sense cRNA was then synthesized using a BioArray HighYield RNA Transcript Labeling Kit (ENZO Biochem, New York, NY). Biotin-labeled cRNA was then purified and fragmented using the Gene Chip Sample Cleanup Module (Affymetrix) according to manufacturer's instructions. The fragmented labeled cRNA was hybridized to the Human Genome U133A Array (Affymetrix), which contains 22 215 human gene cDNA probes. After washing and staining, the arrays were scanned using an Agilent GeneArray Scanner (Germantown, MD). The gene expression levels of samples were normalized to internal standards and analyzed using Microarray Suite v5, MicroDB v3 and Data Mining Tool software v3 (Affymetrix). The difference of 22 215 gene expressions between samples from cells treated with AA or triacsin C were compared with samples from cells treated with vehicle alone.

Western blot analysis
HCT-116 cells were seeded in 100 mm culture dishes at 2 x 106 cells/dish. After 24 h, cells were treated with AA, triacsin C or vehicle alone and incubated for an additional 12 h. Adherent and floating cells were harvested and total protein was isolated in NP40 lysis buffer, and 50 µg of total protein was separated by gel electrophoresis in 10–20% SDS–polyacrylamide precast gels (Bio-Rad) and transferred to polyvinylidene diflouride membranes (Bio-Rad). After blotting, bands were visualized using ECL-Plus (Amersham Pharmacia Biotech, UK).

Caspase activity assay
Caspase activity was quantified using the Apo-ONE Homogenous Caspase Assay Kit (Promega). Cells were plated in a 96-well culture dish at 5000 cells/well in 100 µl of medium and incubated for 24 h at 37°C. Cells were then pre-incubated for 1 h with vehicle alone, DRB (50 µM) or ActD (1 µg/ml) and then treated with AA, triacsin C or vehicle alone and incubated for an additional 36 h. Floating and attached cells were then harvested and caspase activity was assayed according to the manufacturer's instructions.

Immunocytochemistry
HCT-116 cells (2 x 105) were seeded in 35-mm-diameter cell culture dishes. Cells were incubated for 24 h and then pre-treated with vehicle alone, DRB (50 µM) or ActD (1 µg/ml). After 1 h, cells were then treated with vehicle alone, AA or triacsin C for an additional 36 h. Floating cells from the treated dishes were then harvested using cytospin, fixed in 3.7% formaldehyde for 15 min and permeabilized using 0.1% Triton X-100 in phosphate-buffered saline (PBS) for 5 min. After blocking in 1% bovine serum albumin (BSA) in PBS for 10 min, the cytospins were incubated with rabbit anti-cleaved PARP [poly(ADP-ribose)polymerase] antibody in 1% BSA for 30 min. The cytospins were then washed with PBS and incubated with 25 µg/ml goat anti-rabbit IgG conjugated with rhodamine for 30 min. After washing with PBS and formaldehyde post-fixation, the cytospins were stained with DAPI in methanol and mounted under a coverslip in glycerol. Images were prepared using a Zeiss Axioplan 2 microscope equipped with phase-contrast optics and UV excitation filters.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 References
 
Inhibitors of AA metabolism decrease cell viability and increase unesterified AA
It has been previously reported that inhibition of AA uptake and remodeling enzymes, FACL-4 and CoA-IT, respectively, reduces the proliferation of certain cancer cell lines (24,44). We recently demonstrated that, in the SK-MES-1 non-small cell lung cancer cell line, AA remodeling inhibitors, COX inhibitors and exogenous AA induce apoptosis, which is dependent upon the intracellular accumulation of unesterified AA (31). Others have demonstrated similar results in a variety of cell lines (4,24,27). In the colon cancer cell line, HCT-116, inhibitors of AA remodeling enzymes also induced AA accumulation and decreased cell survival (Figure 2). Specifically, inhibition of the ‘high-affinity–low-capacity’ pathway of AA remodeling by the CoA-IT inhibitor PLT 98625, or inhibition of both the ‘high-affinity–low-capacity’ and the ‘low-affinity–high-capacity’ by the FACL-4 inhibitor, triacsin C, significantly reduces the viability of HCT-116 cells (Figure 2A). Further studies were pursued with triacsin C, as it inhibits both pathways of AA remodeling. This reduction of cell viability in HCT-116 cells was associated with activation of caspase-3 activity (Figure 3).


Figure 2
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  		Fig. 2 Inhibitors of AA-uptake and AA-remodeling enzymes reduce cell viability and generate intracellular unesterified AA in HCT-116 cells. (A) HCT-116 cells were incubated with vehicle, exogenous AA, triacsin C, PLT 98625 or etoposide at the indicated concentrations for 48 h and then subjected to cell viability assay. All treatments result in significant loss of HCT-116 cell viability. Results shown are the mean of three independent experiments expressed as a percentage of the viability of vehicle treated cells as determined by metabolic conversion of MTS to formazan. Error bars indicate the standard deviation from the mean (*P < 0.05 cells treated cell versus vehicle treated controls). (B) Unesterified AA accumulates in HCT-116 cells treated with inhibitors of AA metabolic enzymes. HCT-116 cells labeled to isotopic equilibrium with [3H] AA were incubated with triacsin C, PLT 98625, etoposide or vehicle for 18 h. Cellular and extracellular lipids were then extracted and separated by TLC. Increases in extracellular unesterified AA are expressed as fold increase over vehicle treated cells and are the mean of three independent experiments. Error bars represent the standard deviation from the mean (*P < 0.05 versus vehicle treated cells).

 

Figure 3
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  		Fig. 3 Transcription is required for caspase activity induced by AA or triacsin C. HCT-116 cells were pre-incubated for 1 h with vehicle (control), DRB (50 µM) or ActD (1 µg/ml). Cells were then treated with AA (200 µM), triacsin C (20 µM) or vehicle (control) and incubated for an additional 36 h. Apoptosis was quantified by fluorometric assessment of caspase-3 activity. Values shown are expressed as a fold increase in capase-3 activity over cells pre-incubated and treated with vehicle alone, and are the mean of three independent experiments. Error bars represent the standard deviation from the mean, and calculations of significance were made within each treatment group by comparing cells pre-incubated with vehicle alone with those pre-incubated with transcriptional inhibitors (*P < 0.05).

 
Both the CoA-IT and FACL-4 inhibitors also induced the accumulation of unesterified AA. Within 18 h, 32 and 24% of the cell's total AA were released as free fatty acid into the medium in response to PLT 98625 (40 µM) and triacsin C (20 µM), respectively (Figure 2B). This effect was not due to cytotoxicity alone, as etoposide, a reagent that does not affect AA metabolism, reduced cell viability but failed to induce AA accumulation (Figure 2). Addition of 200 µM exogenous AA also reduced HCT-116 cell viability (Figure 2A). This effect is not observed with other control fatty acids (24,27).

AA-induced apoptosis is transcriptionally dependent
Although there is a growing body of evidence supporting the concept that apoptosis induced by inhibitors of AA metabolism is mediated by the accumulation of intracellular AA, little is known about the downstream pathways by which unesterified AA signals apoptosis. To test the hypothesis that AA induces apoptosis in a transcriptionally dependent manner, transcriptional inhibitors DRB and ActD were added to HCT-116 along with inhibitors of AA metabolism, and caspase-3 activity was measured (Figure 3). Both DRB and ActD were able to reverse caspase-3 activation induced by exogenous AA or triacsin C in HCT-116 cells (Figure 3), suggesting that transcription is required for apoptosis induced by AA or FACL-4 inhibition. This result was confirmed using immunocytochemistry, which demonstrated that PARP cleavage, nuclear condensation and morphological changes of apoptosis in cells treated with AA and triacsin C are diminished by pre-treatment with ActD (Figure 4). Transcriptional inhibition was, however, unable to improve the viability of HCT-116 cells, as these transcriptional inhibitors are cytotoxic after 48 h (Figure 5). On the basis of these experiments it was postulated that transcriptional regulation of genes involved in cell survival and apoptosis were downstream mechanisms by which AA accumulation signals apoptosis.


Figure 4
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  		Fig. 4 Pre-treatment with transcriptional inhibitors reduces immunocytochemical and morphological markers of apoptosis in HCT-116 cells treated with AA (200 µM) or triacsin C (20 µM). Changes associated with apoptosis such as cell blistering, blebbing and a shrunken morphology were examined by phase-contrast microscopy (top panel); nuclear condensation and fragmentation was measured by DAPI staining (middle panel); and the presence of cleaved PARP in the nuclear and peri-nuclear regions was measured by dark field nuclear immunostaining (bottom panel). Cells were pre-treated with ActD (1 µg/ml) for 1 h before the addition of AA and triacsin C.

 

Figure 5
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  		Fig. 5 Transcriptional inhibitors reduce HCT-116 cell viability and do not block the cytotoxicity of AA or triacsin C. HCT-116 cells were pre-treated with vehicle (control), or transcriptional inhibitors DRB (50 µM), or ActD (1 µg/ml) for 1 h and then incubated with vehicle (control), exogenous AA, triacsin C, PLT 98625 or etoposide at the indicated concentrations for 48 h. Results shown are the mean of three independent experiments expressed as per cent survival of vehicle treated cells as determined by metabolic conversion of MTS to formazan. Error bars indicate the standard deviation from the mean (*P < 0.05 transcriptional inhibitor pre-treated cells versus vehicle pre-treated controls).

 
AA accumulation influences gene expression
To further investigate whether specific genes may play a role in AA-induced apoptosis, gene chip technology was employed. HCT-116 cells were incubated with triacsin C or exogenous AA for 6 or 12 h and then total cellular RNA was extracted, transformed to biotinylated cRNA and hybridized to Affymetrix Human Genome U133A array, which contains 22 215 human gene cDNA probes. A 6 h time point was chosen to examine early changes in gene expression, as unesterified AA accumulation in the extracellular media is seen as early as 6 h and presumably the intracellular accumulation occurs even earlier. A 12 h time point was also chosen to examine genes whose expression may change later, but still before the morphological changes associated with apoptosis. At 6 h, 769 and 437 targets were upregulated ≥2-fold by AA and triacsin C, respectively (Figure 6A). Two hundred and nineteen of these were commonly upregulated by both AA and triacsin C. At 12 h, 832 and 189 targets were upregulated ≥2-fold by AA and triacsin C, respectively (Figure 6A). Of these, 107 were commonly upregulated by both AA and triacsin C (Figure 6A). Of the 22 215 targets, 29 (0.13%) were upregulated by both triacsin C and AA at both the 6 and 12 h time points (Figure 6A, Table I). The large overlap of upregulated genes by triacsin C- and AA-treated cells suggested a common mechanism of transcriptional regulation in the two treatments.


Figure 6
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  		Fig. 6 Selection strategy for genes with a ≥2-fold change in expression in response to AA or triacsin C. HCT-116 cells were seeded in 100 mm dishes at 2 x 106 cells/plate, and two plates were pooled per sample. Cells were incubated with AA (200 µM) or triacsin C (20 µM) for 6 or 12 h and then total RNA was extracted and analyzed on Affymetrix Human Genome U133A chip. Changes in mRNA expression were determined for samples receiving treatment versus vehicle alone. (A) Shows genes that are upregulated ≥2-fold by AA and triacsin C at 6 and 12 h. (B) Shows genes that were downregulated ≥2-fold by AA and triacsin C at 6 and 12 h.

 

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  		Table I Genes upregulated ≥2-fold by both triacsin C and AA at both 6 and 12 h

 
Similarly, genes whose expression was downregulated by >2-fold were examined at 6 and 12 h (Figure 6B). Downregulation of expression was minimal at 6 h. At 12 h, 40 of 22 125 targets (0.18%) were downregulated ≥2-fold by both AA and triacsin C (Table II).


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  		Table II Genes downregulated by both triacsin C and AA after 12 h

 
Activator protein-1(AP-1) transcription factors are primary target genes affected by AA accumulation
Interestingly, the majority of upregulated targets function in signal transduction or as transcription factors and are implicated in apoptosis and cell cycle progression (Table I). Of the 29 common upregulated targets, at both 6 and 12 h, 6 code for AP-1 proteins implicated in Jun dimerization and apoptotic signaling and 2 more code for the early growth response protein 1 (EGR-1) zinc finger transcription factor, which has been shown to physically interact with c-Jun in neuronal apoptosis (45,46) (Table I). Together, these data suggested that perturbations of cellular AA levels may signal apoptosis through the AP-1 transcription factors. To confirm these results, mRNA and protein levels of one of these AP-1 transcription factors, c-Jun, which was upregulated on the gene array, was measured by real-time PCR and protein immunoblotting (Figure 7). As predicted by gene array experiments, c-Jun was upregulated in response to both triacsin C and AA at both the transcript and protein levels. Furthermore, triacsin C and AA also induce c-Jun phosphorylation (Figure 7), which is an important regulatory step in its transcriptional activation. The COX-2 inhibitor, sulindac sulfide, is also known to reduce cell viability and induce intracellular unesterified AA accumulation in HCT-116 (27). Real-time PCR demonstrated that incubation of HCT-116 with sulindac sulfide upregulates c-Jun expression levels 6.82-fold (P < 0.001).


Figure 7
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  		Fig. 7 c-Jun transcript, protein and phosphorylation levels are upregulated by exogenous AA and triacsin C. (A) RNA was extracted from HCT-116 cells after 12 h incubation with AA (200 µM) or triacsin C (20 µM) and analyzed for c-Jun expression by real-time PCR analysis using the primers noted in Materials and methods. Results are the mean of three independent experiments and error bars are the standard deviation from the mean (*P < 0.05 treated versus vehicle treated controls). (B) and (C) Cellular proteins were harvested from HCT-116 cells incubated for 12 h with AA (200 µM) or triacsin C (200 µM). Proteins were examined by western blot analysis. Results are representative of three independent experiments.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
References
 
Tumor cell carcinogenesis is often accompanied by dramatic changes in the levels of key enzymes that take up, remodel, release and metabolize AA. This observation taken together with the fact that inhibitors of AA metabolism can markedly affect the viability of a number of cancer cells implicate that AA metabolism may play a pivotal role in the survival of certain cancers. Our laboratory has previously shown that inhibitors of AA uptake, remodeling and metabolism induce apoptosis in SK-MES-1 cells and that this apoptosis is probably mediated by the accumulation of unesterified AA (31). Other studies have also drawn similar conclusions with regard to COX inhibition (27), FACL-4 inhibition (22) and CoA-IT inhibition (28).

Nowhere have these observations been more dramatic than in models of colon cancer. In fact, the original observations of the chemopreventive effects of COX inhibition were made in colon cancer incidence (47). Colon cancer cells appear to have developed elegant strategies to control levels of AA and AA metabolites produced within the cell. For example, it has been recognized for sometime that elevation of COX-2 expression and prostaglandin-E2 (PGE2) production is often associated with cancer cell survival. Moreover, a high COX-2/low cPLA2 phenotype has recently been described and suggested to play a key role in early colon carcinogenesis (30). Such a phenotype would maintain very low levels of intracellular AA, as little would be generated by cPLA2 and that which was generated would be rapidly metabolized by the high levels of Cox-2. This is a characteristic that would be predicted to protect cell survival by preventing apoptosis secondary to intracellular AA accumulation.

As expected, the current study revealed that inhibition of AA metabolism reduces cell viability and induces accumulation of unesterified AA in the colon cancer cell line, HCT-116. Accumulation of unesterified AA was not simply an artifact of cell death, as etoposide, which does induce cell death, did not increase AA levels. Furthermore, AA accumulation was seen as early as 6 h post-treatment, whereas the morphological changes of apoptosis were not observed until 24 h post-treatment (data not shown).

If the accumulation of unesterified AA mediates this cytotoxicity, then treatment of cells with unesterified AA, at concentrations high enough to overcome its incorporation into cellular sinks, should also be cytotoxic. As predicted, exogenous unesterified AA (200 µM) was also cytotoxic to HCT-116 cells. Human serum AA concentrations are normally in the 300–400 µM range (48) and can be significantly elevated in specific human diseases. However, the vast majority of AA in serum exists esterified to the glycerol backbone of phospholipids and triglycerides, and normal physiological concentrations of unesterified AA are probably well <200 µM. The necessity of supra physiological concentrations of unesterified AA to induce cytotoxicity is to be expected, as these concentrations would be required to overwhelm the coenzyme A-dependent and coenzyme A-independent enzymatic steps that incorporate AA into cellular glycerolipids. The response to exogenous AA does not appear to be a detergent effect, as other unsaturated fatty acids do not induce this same response. Furthermore, it is likely that detergent effects would induce necrosis rather than apoptosis. These results, coupled with similar findings in numerous other studies (3,24,27,28,49), indicate that the response of HCT-116 cells to AA metabolic inhibitors is probably mediated by an increase in unesterified AA.

Deciphering a common downstream pathway of apoptotic signaling by perturbations in AA metabolism is a critical next step in understanding the role of AA metabolism in carcinogenesis. Initial experiments in this study have showed that cytotoxicity induced by AA accumulation is caspase-3- and gene transcription-dependent. A transcriptionally dependent apoptotic process coupled with the knowledge that AA and its metabolites have been shown to alter the transcription of certain genes in other cell types (50) led us to further examine the role of transcriptional regulation in apoptosis induced by AA and triacsin C.

Interestingly, a large proportion of genes affected by AA and triacsin C are involved in apoptosis and growth inhibition and have known implications in carcinogenesis (Table I). A number of the upregulated genes alter Erk/MAP kinase signaling by ras family members, which is one of the most well-defined and widely implicated pathways in carcinogenesis. 14-3-3 zeta is a member of 14-3-3 family signal transduction proteins known to interact with ras and raf in regulation of cell cycle progression and apoptotic induction (51). RAPGEF2 is a rap guanine nucleotide exchange factor known to activate rap (52), which functions to counteract ras signaling pathways (53). ARHGDIA is a Rho-GDP disassociation inhibitor known to inactivate Rho signaling pathways (54) that function in cell proliferation (55). Additionally, SQSTM1 (P62) is a negative regulator of Ras signaling pathways (56).

Of the 29 upregulated targets identified by the selection strategy outlined in Figure 6A, 6 code for transcription factor AP-1 proteins implicated in Jun dimerization and apoptotic signaling and 2 more code for the EGR-1 zinc finger transcription factor, which has been shown to physically interact with c-Jun in apoptotic signaling (45,46) (Table I). AP-1 was among the first mammalian transcription factors identified (57) and regulates a broad spectrum of processes including cell proliferation, death, survival and differentiation (58). In fact, many members of the AP-1 family are proto-oncogenes. Structurally, the AP-1 transcription factor consists of a dimer of two basic region-leucine zipper proteins of the AP-1 family. Members of this family include the Jun subfamily (c-Jun, JunB, JunD), the Fos subfamily (c-Fos, FosB, Fra-1, Fra-2), the Maf subfamily (c-Maf, MafB, MafA, MafG/F/K) and the ATF subfamily (ATF-2, ATF-3, B-ATF, JDP1, JDP2) (58). Other studies also demonstrated upregulation of AP-1 transcription factors by AA (40,59). Together, these data suggest that intracellular burdens of unesterified AA may signal apoptosis through the AP-1 transcription factors.

The role of c-Jun expression was further examined and it was demonstrated that perturbations in AA lead to both elevated transcript and protein levels. AA and triacsin C also caused phosphorylation of c-Jun, a known marker of activation. Future studies are under way in our laboratory to further investigate these results and their mechanistic significance. Given the prominent role COX inhibitors have played in colon cancer prevention, the effects of a COX inhibitor, sulindac sulfide, was also examined. Sulindac sulfide induced the upregulation of c-Jun expression at concentrations that induce marked AA accumulation and apoptosis. It is possible that these effects of sulindac are COX-independent, as it is unclear whether HCT-116 cells express COX-2. HCT-116 cells do, however, produce low levels of PGE2 and can be induced to produce higher levels (60). Additionally, at concentrations used (160 µM) in the current study, sulindac sulfide inhibits both COX isoforms (61).

When focusing on genes whose expression was downregulated, fewer genes were altered. However, the majority of these genes are also associated with cancer biology (Table II). For example, a number of these genes act as growth factors (i.e. BDNF) (62–64), growth factor receptors (i.e. fibroblast growth factor receptor 3) (65–68), signal transduction molecules (i.e. P45) (69) and proteins involved in DNA and RNA metabolism (i.e. topoisomerase 2A) (70), whose downregulation could disrupt the growth potential of malignant cells. Given the growing collection of data suggesting a chemoprotective effect of NSAIDs and the link between AA metabolism and carcinogenesis, it is essential to understand the mechanism(s) by which perturbing AA metabolism affects cancer cell growth. While large amounts of data have been generated by this array study, more detailed experiments focusing on specific gene families will be necessary to better determine the role of specific signaling pathways. However, given the broad array of genes and pathways that are altered, it is unlikely that a single pathway or gene will be responsible for induction of apoptosis by AA. Perhaps, more importantly, this study points out the potential for perturbations in cellular AA levels to have broad and coordinated effects on the growth of cancer cells. For almost four decades now, AA metabolism has been shown to play a crucial role in the biology of processes that range from inflammation to reproduction. Studies such as this one build on an ever-growing literature that suggests that AA metabolism is a promising target for cancer therapy.


   Acknowledgments
 
This work is supported by NIH grant RO1AI42022 and P50AT0027820. A.M.M. received support through US Army Research Acquisition Activity award number DAMD 17-00-1-0489.

Conclict of Interest Statement: None declared.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received October 25, 2005; revised February 28, 2006; accepted March 18, 2006.


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