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. Author manuscript; available in PMC: 2016 Oct 1.
Published in final edited form as: Cancer Prev Res (Phila). 2015 Aug 19;8(10):989–999. doi: 10.1158/1940-6207.CAPR-15-0050

Chemopreventive effects of dietary eicosapentaenoic acid supplementation in experimental myeloid leukemia

Emily R Finch 1,2, Avinash K Kudva 2, Michael D Quickel 2, Laura L Goodfield 1,2, Mary J Kennett 2, Jay Whelan 3, Robert F Paulson 2,†, K Sandeep Prabhu 2,†
PMCID: PMC4596789  NIHMSID: NIHMS717400  PMID: 26290393

Abstract

Current therapies for treatment of myeloid leukemia do not eliminate leukemia stem cells (LSC), leading to disease relapse. In this study, we supplemented mice with eicosapentaenoic acid (EPA, C20:5), a polyunsaturated omega-3 fatty acid, at pharmacological levels, to examine if the endogenous metabolite, cyclopentenone prostaglandin delta-12 PGJ3 (Δ12-PGJ3), was effective in targeting LSCs in experimental leukemia. EPA supplementation for eight weeks resulted in enhanced endogenous production of Δ12-PGJ3 that was blocked by indomethacin, a cyclooxygenase inhibitor. Using a murine model of chronic myelogenous leukemia (CML) induced by bone marrow transplantation of BCR-ABL-expressing hematopoietic stem cells, mice supplemented with EPA showed a decrease in the LSC population, reduced splenomegaly and leukocytosis, when compared to mice on an oleic acid diet. Supplementation of CML mice carrying the T315I mutation (in BCR-ABL) with EPA resulted in a similar effect. Indomethacin blocked the EPA effect and increased the severity of BCR-ABL-induced CML and decreased apoptosis. Δ12-PGJ3 rescued indomethacin-treated BCR-ABL mice and decreased LSCs. Inhibition of hematopoietic-prostaglandin D synthase (H-PGDS) by HQL-79 in EPA-supplemented CML mice also blocked the effect of EPA. In addition, EPA supplementation was effective in a murine model of acute myeloid leukemia. Supplemented mice exhibited a decrease in leukemia burden and a decrease in the LSC colony-forming unit (LSC-CFU). The decrease in LSCs was confirmed through serial transplantation assays in all disease models. The results support a chemopreventive role for EPA in myeloid leukemia, which is dependent on the ability to efficiently convert EPA to endogenous cyclooxygenase-derived prostanoids, including Δ12-PGJ3.

Keywords: CR-ABL, T315I, MLL-AF9, leukemia stem cells, delta 12-prostaglandin J3, cyclooxygenase, prostaglandin D synthase, NSAID, Lovaza®

Introduction

Leukemia remains one of the leading causes of cancer death, despite established well-defined standards of care. Each year, an estimated 43,000 new cases of leukemia will be diagnosed and 22,000 people will die from the disease (1). This observation underscores the need for further research into alternative therapies. Previously, we have demonstrated that certain cyclopentenone prostaglandin (CyPG) metabolites of polyunsaturated fatty acids (PUFA) are effective in treating murine models of leukemia (2). While PUFAs comprised of omega-6 and omega-3 fatty acids are essential for healthy growth and development, the ratio of these lipids is of particular interest in disease (3). This is because prostanoids produced from eicosapentaenoic acid (EPA, C20:5; n-3) show increased anti-inflammatory or ‘pro-resolving’ functions when compared to some of their omega-6 derived counterparts (4–6). Furthermore, alternative pathways produce other bioactive anti-inflammatory lipid mediators, including resolvins and protectins, from EPA and docosahexaenoic acid (DHA, C22:6; n-3), respectively (7–9).

Membrane incorporation of dietary EPA provides the substrate for the production of CyPG, Δ12-PGJ3 that is of particular interest in our laboratory, given its anti-leukemic properties (2). Release of EPA from the membrane by phospholipase A2 commits the PUFA to CyPG synthesis through COX activity; EPA is converted to PGH3 (10). PGH3 is acted upon by PG-synthases, such as PGD synthases hematopoietic (H-PGDS) and lipocalin type (L-PGDS), to produce PGD3 that undergoes dehydration and isomerization to Δ12-PGJ3. However, in cells, arachidonic acid (ARA, C20:4; n-6) is the preferred substrate for COX-2, while EPA is utilized at approximately 30% efficiency of ARA by COX-2 (11). Thus to increase production of 3-series PGs, EPA must be saturated in the membrane by increasing intake of omega-3 fatty acids. It is therefore conceivable that inhibition of COX-2 by non-steroidal anti-inflammatory drugs (NSAIDs) or H-PGDS by HQL-79 can impact leukemia progression. However, the results of epidemiological studies are inconclusive regarding the relationship between non-aspirin NSAID use and incidence of leukemia (12–14), necessitating further research before embarking on new studies.

Myeloid leukemias are myeloproliferative disorders that arise from mutations within the hematopoietic stem cell (HSC) population in the bone marrow (15). These mutations lead to defects in the generation of healthy myeloid-lineage cells. In chronic myelogenous leukemia (CML), the Philadelphia chromosome is the result of a 9:22 chromosomal translocation, which leads to a constitutively active tyrosine kinase fusion protein BCR-ABL that is found in over 90% of cases (15–16). Current therapy for BCR-ABL+ CML includes treatment with tyrosine kinase inhibitors (TKI). However, this maintenance therapy has various negative side effects and targets bulk tumor cells along with healthy cells while leaving leukemia stem cells (LSCs) unharmed. Disease relapse occurs due to LSCs’ ability to self-renew and differentiate into bulk tumor cells (16–17). Additionally, many patients become resistant to the therapy (15,18,19). The T315I mutation in BCR-ABL confers resistance to nearly all TKI therapies, except ponatinib (20). Finding a new mechanism that is independent of ABL kinase inhibition is essential to eliminating the cancer cells. Acute myeloid leukemia (AML) is characterized by immature blood cells that grow quickly and progress to leukemia in a few months. However, AML is a more heterogeneous disease with multiple etiologies including translocations. One translocation involved the MLL locus and multiple partners that lead to pediatric AML. One common MLL translocation generates the MLL-AF9 fusion protein that can confer self-renewal ability on hematopoietic progenitor cells leading to aggressive leukemogenesis (21).

In the present study, we show that EPA supplementation of mice at pharmacological doses alleviates symptoms and signs of leukemia in models of CML, T315I CML, and MLL-AF9 AML. Our data further support the key role of COX/H-PGDS-pathway-derived endogenous metabolite of EPA to specifically target the LSC population to correct leukocytosis, splenomegaly, and overall disease index in these models of experimental leukemia.

Materials and Methods

Animals

Three-week-old C57BL/6 mice (Taconic Biosciences, Hudson, NY) were randomly assigned to two dietary groups: animals were housed 3–4 mice per cage in a temperature- and humidity-controlled room with a 12-hour light-dark cycle. All mice were 11–16 weeks of age at sacrifice, unless noted otherwise. For most experiments total animal number was 8–12; this was the summation of 3–4 independent experiments run in at least biological triplicate. The Institutional Animal Use and Care Committee (IACUC) at The Pennsylvania State University preapproved all procedures.

Mouse diet

An energetically equivalent mouse diet, in terms of omega-3 fatty acid, equivalent the FDA approved omega-3 supplement Lovaza® was developed (22). Diets remained isocaloric to the base diet (AIN-76A; Research Diets, Inc., New Brunswick, NJ) with 5g of total fat (12% of total calories). A portion of the corn oil in the base diet was replaced with the ethyl esters of oleic acid (OA, C18:1 n-9) (100%) or EPA (>90%) (Nu-Chek Prep, Elysian, MN) at a level of 0.82% (w/w), or 1.8% of total energy (kcal) (Table S1). The OA diet was used as an experimental control diet. The diets were stored in individual portions in airtight Whirl-Pak® bags at −80°C until the day of feeding (to prevent oxidation). The mice were fed ad libitum with fresh diet daily. In all experiments, mice were sacrificed after overnight fast (12–16 hours).

Fatty acid analysis

Fatty acid methyl esters of the tissue phospholipids were prepared and analyzed as described previously (22), with the following modifications. Briefly, tissues were homogenized in ice-cold saline. Three ml of chloroform-methanol (1:2, v/v) were added to each homogenate, and lipids were extracted with chloroform (1ml) plus saline (1ml), followed by chloroform (1ml) (x2). The internal standard 1,2 diheptadecanoyl-sn-glycero-3-phosphocholine (17:0) (Avanti Polar Lipids, Alabaster, AL) was added to each sample prior to lipid extraction. The pooled chloroform extracts were evaporated, resuspended in a small volume of chloroform, and subjected to thin layer chromatography (TLC). The phospholipids were separated by TLC on silica gel 60 HPTLC plates (Merck, Darmstadt, Germany) with chloroform/methanol (8:1 v/v) as the solvent system (phospholipids remained at the origin). Bands corresponding to the phospholipids were scraped, transferred to screw-topped test tubes and saponified in the presence of 14% BF3 in methanol (Thermo Scientific, Bellefonte, PA) at 86°C for 10 min, followed by the addition of 0.9% saline (2ml). Fatty acid methyl esters were isolated with equal volumes of hexane (x2), pooled and evaporated under nitrogen. The fatty acid methyl esters were resuspended in hexane and analyzed using a Hewlett Packard model 5890 series II gas chromatograph equipped with flame ionization detector and a DB23 fused silica capillary column (0.25mm × 30m) (Agilent Technologies, Santa Clara, CA) with hydrogen as the carrier gas, and temperature programming from 160°C to 250°C at 3.5°C/min. The fatty acid methyl esters were identified by comparing the retention times with those of known standards (NuChek Prep, Elysian, MN). The fatty acids are presented as mole percent.

EPA supplementation ex vivo to detect Δ12-PGJ3

Murine macrophage-like cell line RAW 264.7 (ATCC, Manassas, VA) was cultured as previously described (23). In brief, cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) containing 5% fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA), 2mM L-glutamine, 100units/ml penicillin-G (Cellgro, Manassas, VA), and 100µg/ml streptomycin (Invitrogen, Carlsbad, CA). For routine culture, the cells were passaged every three days at a ratio of 1:10 as per the recommendations by ATCC. For experimental assays, 100mm plates were seeded with 2×106 cells. Exogenous fatty acid treatment began when cells were approximately 40% confluent. EPA or OA as ethyl ester (Cayman Chemicals, Ann Arbor, MI) was incubated with cell-culture grade and endotoxin-free bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, MO) in serum free DMEM for one hour at 37°C. Final concentration of EPA or OA in culture was 50µM conjugated to 1% (v/v) BSA. Cells were treated with fatty acid for at least three days, with media changed daily, prior to 50ng/ml lipopolysaccharide (LPS) stimulation for 12 hours. In cultures treated with 10µM indomethacin (Cayman Chemicals, Ann Arbor, MI), the NSAID was added at least two hours prior to addition of fatty acid (as EPA or OA conjugated to BSA). Indomethacin was added to culture daily and was present during LPS stimulation.

Protocol for harvesting and culturing bone-marrow-derived macrophages (BMDMs) was modified from previously described work from our laboratory (23,24). In short, BMDMs were prepared by extracting femoral bone marrow plugs from C57BL/6 mice maintained on either EPA or OA supplemented diet (as described above) for eight weeks. These cells were cultured in the same medium as described earlier with the addition of 20%(v/v) L929 fibroblast-conditioned medium (as a source of macrophage colony-stimulating factor, M-CSF) for at least three days. No additional fatty acid was added when culturing BMDMs. On day three, LPS (50ng/ml) was added for 12 hours. Clarified media was collected for lipid extraction as described below.

LC-MS/MS-MRM analysis

The estimation of Δ12-PGJ3 was carried out as reported previously (25). Briefly, Δ12-PGJ3 was quantified using a HPLC system consisting of LC-20AD UFLC pumps with a SIL-20AC autosampler (Shimadzu Corporation, Columbia, MD), a Luna™ phenyl-hexyl analytical column (2×150mm, 3 µm) (Phenomenex, Torrance, CA) developed with a 30 min isocratic elution with methanol/ water (70:30 v/v) containing 0.1 % acetic acid at a flow rate of 150µL/min and injection volume of 50µL. The negative ion electrospray tandem mass spectrometric analysis was carried out using API 2000 triple quadruple mass spectrometer (AB Sciex, Foster City, CA) at unit resolution with multiple reaction-monitoring mode (MRM). The source temperature was maintained at 450°C, electrospray voltage was −4500V and the declustering potential was set at −16V. Nitrogen was used as collision gas at −20eV and the dwell time was 150ms/ion. During MRM, Δ12-PGJ3 was measured by recording the signal for the transition of the deprotonated molecule of m/z 331 to the most abundant fragment ion with m/z 269. Data were acquired and analyzed using Analyst software program version 1.5 (AB Sciex, Foster City, CA).

Generation of experimental leukemias

To induce experimental CML and T315I CML, retroviral stocks were generated by transfecting HEK293T cells with either pMIG-BCR-ABL (a gift from Dr. Warren Pear, University of Pennsylvania, Philadelphia) or MIGR-BCR-ABL-315T/I (a gift from Dr. Ravi Bhatia, University of Alabama, Birmingham), respectively, with pEco for packaging using TransIT 293 reagent (Mirus Bio, Madison, WI). For experimental AML, pMIG-MLL-AF9 (a gift from Dr. Hong-Gang Wang, Penn State College of Medicine, Hershey) was used. Isolation and transduction of HSCs was performed as described earlier (2, 26). HSCs transduced with BCR-ABL virus are hereafter referred to as BCR-ABL LSCs (BCR-ABL+GFP+Kit+Sca1+Lin-). T315I CML LSCs are similarly defined. AML LSCs have not been defined with regard to cell surface markers. For all AML experiments, total bone marrow was transduced and transplanted (1×106 total transduced cells) to establish the model. Studies using GFP (C57BL/6-Tg(UBC-GFP)30Scha/J) (27) bone marrow transplanted directly into OA and EPA supplemented mice clearly indicated that EPA does not negatively impact engraftment (Fig. S1).

BCR-ABL LSCs were isolated from spleen and bone marrow by FACS using the Influx Cell Sorter (BD Biosciences, San Jose, CA) as described earlier (2, 28). LSCs (5×105 cells) in 100µl of sterile phosphate buffered saline (PBS) were transplanted by retro-orbital injection into lethally irradiated (950 rads) recipient mice maintained on EPA or OA diet groups for at least eight weeks. Disease progression was monitored by complete blood count (CBC) analysis, evaluating peripheral blood for leukocytosis on a Hemavet 950FS equipped with a veterinary software program. All mice were sacrificed by day-14 post-transplant. Spleen and bone marrow were isolated and used for characterization as described below. Any deviation from this transplantation system will be described in figure legends. In all experimental models, transduced bone marrow cells were passaged through mice at least two times prior to transplantation into OA or EPA supplemented mice.

Inhibition of Δ12-PGJ3 synthesis in vivo

Inhibition of COX-1/2 was accomplished by use of indomethacin (Cayman Chemicals, Ann Arbor, MI). Indomethacin (0.00325%, w/v) was dissolved in ethanol (0.5%, v/v) and administered by means of oral drinking water for one week prior to bone marrow transplantation with BCR-ABL LSCs, as described previously (23, 28, 29). In a subset of Δ12-PGJ3 rescue experiments, administration of exogenous Δ12-PGJ3 was performed as described previously (2). In brief, mice were given daily intraperitoneal (i.p.) injection of Δ12-PGJ3 (0.025mg/kg) in 500 µl of sterile PBS for seven days, starting at day-7 post-transplant until sacrifice at day-14. HQL-79 (Cayman Chemicals, Ann Arbor, MI) was used to inhibit the activity of H-PGDS. HQL-79 (30 mg/kg body weight) was administered by i.p. injection every other day (3–4 times/week) from day zero (day of transplant) until day 14 (total of seven injections). HQL-79 was resuspended in a solution of 0.05 mM citric acid and 11.1 mM hydroxypropyl beta-cyclodextrin (HPBCD; Sigma, St. Louis) and incubated for >10 minutes at 37 °C, until dissolved. Mice were injected with this formation within 10 minutes of preparation. All mice were euthanized on day 14 post-transplant.

Flow cytometry of CML LSCs

Whole spleen and bone marrow were collected and red blood cells (RBCs) were lysed with ACK lysis buffer (1.5mM NH4Cl, 100µM KHCO3, 10µM EDTA-2Na). Lineage negative cells were isolated following manufacturer’s protocol (Stem Cell Technologies, Vancouver, British Columbia). Cells were stained for Ly-6A/E (sca-1) and CD117 (c-kit) (BD Biosciences, San Jose, CA). Lineage negative white blood cells (WBCs) and the presence of LSCs were analyzed on Accuri C6 or Fortessa LSR flow cytometers (BD Biosciences, San Jose, CA). KSL (c-kit+sca-1+Lin-) LSCs were defined as Lin- selected cells that stained positive for c-kit, sca-1, and GFP (present in BCR-ABL plasmid).

LSC-Colony Forming Units (LSC-CFU)

Total splenocytes or bone marrow from leukemic mice were plated in M3231 Methocult (Stem Cell Technologies, Vancouver, British Columbia). Cell density is described in figure legends. Media was supplemented with IL-3 (2.5ng/ml; R&D Systems, Minneapolis, MN), stem cell factor (SCF; 50ng/ml; Gold Biotechnology, St. Louis, MO), growth differentiation factor 15 (GDF-15; 30ng/ml; Biomatik, Wilmington, DE), and sonic hedgehog (Shh; 25ng/ml; Gold Biotechnology, St. Louis, MO). On days 7–10, colonies were counted. LSC-colony forming units (LSC-CFU) were defined based on their size (large: >100 cells at high magnification), shape (perfectly circular), density (dark brown/black color), and edge (clearly defined edge with limited differentiation). Any variation from LSC-CFU formation was also identified: in some cases granulocyte-macrophage progenitor colony forming units (CFU-GM) were observed (30). Each biological sample was plated in triplicate. The average number of LSC colonies from each biological sample was tabulated for analysis.

Histopathology and TUNEL staining of tissue sections

Whole spleen tissue was fixed in 4% paraformaldehyde at time of sacrifice. Sections were prepared (5 micron thickness) and stained with hematoxylin and eosin (H&E) at the Animal Diagnostic Laboratory, Penn State University, University Park, PA. Experimental groups were blinded prior to being examined and scored by a board certified laboratory animal veterinarian with training in pathology. Six characteristics were evaluated: disruption of splenic architecture (DSA), tumor infiltrates (TI), mitotic figures (MF; scored as 0= rare to none, 1= 1–3 per high power field (hpf), 2= 3–6 per hpf, 3= 6–9 per hpf, 4= >7 per hpf), megakaryocytes (MK), extramedulary hematopoiesis (EMH), and necrosis. With the exception of MF, tissue characteristics were scored on a 0–4 scale as follows: 0= within normal limits, 1= minimally increased/elevated/affected, 2= mildly increased/elevated/affected, 3= moderately increased/elevated/affected, 4= extensively or severely increased or affected.

TUNEL assay was subsequently performed on representative tissues following histopathological examination using the In Situ Cell Death Detection Kit (Roche Diagnostics Corp., Indianapolis, IN). Tissue sections were deparaffinized in Histo-Clear™ (National Diagnostics, Atlanta, GA), washed in 100% ethanol, 95% ethanol, 70% ethanol, and distilled water. Slides were then boiled in 10mM sodium citrate pH6.0 with 0.05% Tween-20 and allowed to cool. TUNEL staining was performed according to the manufacturer’s instructions. Images were collected using an Olympus BX51 fluorescence microscope (Olympus America, Center Valley, PA). Total fluorescent area (TUNEL+ cells) of stained slides was determined using Image J software program (NIH, Bethesda, MD).

Statistical analysis

Statistical analysis was performed using GraphPad Prism version 6 (GraphPad Software, San Diego, CA). Unless noted, nonparametric tests, including one-way ANOVA and un-paired one-tailed student t-test, were used where appropriate. The OA group was used as the control, unless otherwise noted, for t-tests (comparing EPA only to OA only). For ANOVA, all groups were compared. Variation from these analyses is described in figure legends. Bars represent mean ± SEM: *p<0.05, **p<0.01, ***p<0.001, #p<0.0001. The Biostatistics, Epidemiology, and Research Design (BERD) Core of Penn State Clinical and Translational Sciences Institute (CTSI) provided statistical consulting.

Results

Effect of EPA diet and omega-3 index

Two experimental diets were designed such that the only difference between dietary groups was the replacement of a subset of the base diet (AIN76-A) corn oil with either EPA or OA as ethyl esters at 0.82% (w/w), or 1.8% of total kcal (Table S1). Mice on either diet consumed a similar amount of diet and gained weight at the same rate (Fig. 1A). A widely accepted method for assessing omega-3 status is the omega-3 index, which is the relative abundance of EPA + DHA in red blood cells (RBCs) (Palozza, 1996; Luxwolda, 2011). To examine if EPA supplementation led to changes in the membrane lipid composition in the spleen, liver, and RBCs, the omega-3 index was determined by GC-MS analysis (22, 31). The amount and type of omega-3 fatty acid esters increased with EPA supplementation, as shown by the membrane lipid composition of the spleen, liver and RBCs (Fig. 1B). Compared to OA control diet, EPA supplementation led to a significant increase in the omega-3 index in the membrane of all examined tissues (Fig. 1C and Table S2). The content of omega-3 in the liver amounted to 19.4% of the membrane with EPA supplementation versus 7.8% in OA supplemented mice, indicating a 2.5-fold increase. In the spleen, there was a 3.5-fold increase in the omega-3 in EPA mice; the omega-3 index was 11.1% in EPA mice of compared with 3.2% in OA mice. The largest increase was in the RBCs where EPA supplementation increased the omega-3 index to 5.5% versus 1.0%, a 5.5 fold increase. These data demonstrate that an EPA supplemented diet increases the omega-3 content of cells, which will allow us to determine whether these changes lead to changes in Δ12-PGJ3 production, which in turn will affect CML and AML progression.

Figure 1. EPA-supplemented diet increases the omega-3 index.

Figure 1

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A, Food intake and relative body weight (compared to day 0) were monitored daily for eight weeks. n= 12 per diet group. B, Fatty acid composition of spleen, liver, and RBCs from OA- and EPA-supplemented mice by GC-MS. The fatty acids are presented as mole %. Data are represented as mean. C, Percent of membrane made of omega-3 fatty acids i.e. omega-3 index calculated from mean ± SEM. n= 7–12 per diet group. #p< 0.0001.

Endogenous Δ12-PGJ3 production

Based on our previously reported studies demonstrating the ability of Δ12-PGJ3 to specifically target LSCs (2), endogenous production of Δ12-PGJ3 was measured. EPA supplemented cells and mice were treated with LPS to mobilize membrane esterified EPA and to upregulate COX-2 activity (Fig. 2). As expected, EPA supplementation without LPS stimulation did not lead to detectable levels of Δ12-PGJ3 in either RAW 264.7 cells or BMDMs (data not shown). Primary BMDMs from mice on either OA or EPA supplemented diets were cultured ex vivo and stimulated with LPS. No additional fatty acid was added to the cultures. BMDMs derived from EPA supplemented mice produced significantly higher levels of Δ12-PGJ3 than OA derived BMDMs (Fig. 2A). We next tested whether Δ12-PGJ3 production was COX dependent. RAW 264.7 cells treated with EPA generated Δ12-PGJ3, in response to LPS. In contrast, cells treated with BSA (used as a carrier) alone or OA-conjugated BSA failed to produce any Δ12-PGJ3 (data not shown). As shown in Fig. 2B, indomethacin treatment blocked the production of Δ12-PGJ3 by EPA treated RAW 264.7 cells (Fig. 2B). Although EPA treated macrophages produced Δ12-PGJ3 in vitro in response to LPS treatment, we needed to determine whether mice fed on EPA supplemented diet produced significantly higher Δ12-PGJ3, in vivo, following LPS injection. Lipid extracts were isolated from plasma and subjected to LC-MS/MS analysis (Fig. 2C). While there was a non-significant increase in Δ12-PGJ3 in EPA mice versus OA mice without LPS treatment, stimulation with LPS for 12 hours led to a 3.5 fold increase in Δ12-PGJ3 in the plasma of EPA supplemented mice (Fig. 2C). Taken together, these results correlate an increase in the omega-3 index in EPA-fed mice with increased endogenous Δ12-PGJ3 production.

Figure 2. Δ12-PGJ3 is produced in vivo and in vitro from exogenous EPA via the COX pathway.

Figure 2

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A, Primary BMDMs were harvested from the femurs of EPA- or OA- supplemented mice (on diet for >8 weeks). Cells were cultured for four days and stimulated with 50ng/ml LPS for 12 hours prior to harvesting cells and media. Data represent one of five independent experiments. B, RAW 264.7 macrophages were treated for three days with 50µM fatty acid ethyl ester as OA or EPA- conjugated to BSA. Cells were treated with or without addition of 10µM indomethacin (Indo) for at least two hours prior to addition of fatty acid. LPS (50ng/ml) was added for 12 hours prior to collection of cells and media. Data represent two of five independent experiments. C, Plasma from healthy mice on supplemented diet was harvested after eight weeks on diet. LPS (1mg/kg body weight) was administered by i.p. injection 12 hours prior to harvesting blood. n= 8–12. Lipids were extracted from clarified media or whole plasma for LC-MS/MS analysis (MRM method) of Δ12-PGJ3. Individual data points are represented with error bars as the mean ± SEM. *p< 0.05, **p< 0.01, ***p< 0.001.

Supplementation with EPA affects CML

To relate increases in the omega-3 index and the ability of in vivo EPA supplementation to form endogenous Δ12-PGJ3 during CML, mice on the specific diets were given a bone marrow transplant of BCR-ABL+ LSCs. As reported previously, transplantation with BCR-ABL LSCs leads to leukocytosis and splenomegaly (2, 28, 31). Interestingly, mice on an EPA supplemented diet showed less severe symptoms of BCR-ABL induced CML. Splenomegaly was decreased in EPA mice when compared to OA mice (Fig. 3A, Fig S2A), whereas healthy mice on diet showed no difference in spleen weight (Fig. S1A). Furthermore, H&E stained sections of whole spleen were scored based on six parameters of tissue health as mentioned in the “Materials and Methods” section (Fig. 3B). OA supplemented LSC-transplanted mice had deterioration of normal spleen architecture with no clear demarcation between the red and white pulp areas (Fig. 3B, Fig. S2B). In contrast, H&E staining of splenic sections prepared from EPA supplemented LSC-transplanted mice showed normal structure in the spleen with well demarcated red and white pulp areas (Fig. 3B, Fig. S1B, Fig. S2B). To further demonstrate the effect of EPA in BCR-ABL induced CML, we examined the BCR-ABL LSC population in the spleen and bone marrow by flow cytometric analysis on day-14 post-transplantation. In both the spleen and bone marrow, OA-supplemented mice showed an approximate two-fold increase in the percentage of LSCs compared with EPA-supplemented mice (Fig. 3C). This increase in BCR-ABL LSCs was further corroborated with ex vivo investigation of BCR-ABL LSC-colony forming units (CFU) from the spleen and bone marrow. EPA mice failed to produce LSC-CFU colonies, whereas OA mice formed numerous LSC-CFUs (Fig. 3D). EPA mice exhibit dense colonies consistent with CFU-GM progenitor colonies (30). These assays showed that there were not only a limited number of LSCs in the EPA-supplemented group (as shown from flow cytometric results), but also these cells were not functional nor able to expand ex vivo.

Figure 3. EPA-supplemented mice show less severe signs and symptoms of experimental CML.

Figure 3

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A-D: BCR-ABL+ LSC transplanted mice; E-H: Secondary transplant model of total BCR-ABL bone marrow (See Supplementary Fig. 2C for details); I-J: T315I transplant of 5×105 total T315I bone marrow. A, Spleen size in OA- and EPA-supplemented BCR-ABL+ LSC transplanted mice. n= 8–12. Healthy spleen size: small dashed line (0.1g). B, Total histology scores of H&E stained spleen sections from OA- and EPA-supplemented BCR-ABL+ LSC transplanted mice. Six parameters (DSA, TI, MF, MK, EMH, and necrosis) were scored from 0–4 and total scores are represented. n= 5–8 per diet group. C, Flow cytometric analysis of percentage of Lin GFP+c-kit+sca-1+ BCR-ABL LSCs in spleen and bone marrow. n= 6. D, Representative image of CML LSC-CFU assay from both spleen and bone marrow of OA- and EPA-supplemented BCR-ABL+ LSC transplanted mice. Cells were plated at a concentration of 2×104 cells per mL. n= 3. E, Weight loss, presented as % body weight relative to week 0 post-secondary transplant of total BCR-ABL bone marrow. n= 4. F, Total WBC counts in peripheral blood (at 28 days post-secondary transplant of total BCR-ABL bone marrow). Shaded area (1.8–10.7 k/µl) indicates healthy range of WBC counts. n= 4. G, Spleen weight in secondary transplanted BCR-ABL mice. n= 4. H, Total BCR-ABL LSC-CFU counts (per 1×106 cells) in secondary transplanted BCR-ABL mice. Too numerous to count (TNC). n= 4. I, Spleen size in OA- and EPA-supplemented T315I total bone marrow transplanted mice. n= 5. J, Total T315I LSC-CFU counts in spleen and bone marrow from OA- and EPA-supplemented T315I transplanted mice. Cells were plated at the concentration of 2.5×103 cells per mL. n=4. Data points represent individual mice and bars represent mean ± SEM *p< 0.05, **p< 0.01, ***p< 0.001.

A defining characteristic of LSCs is their ability to cause relapse of disease upon serial transplantation (32, 33). To further test the role of EPA in limiting experimental BCR-ABL induced CML, whole bone marrow from BCR-ABL LSC-primary-transplanted OA or EPA supplemented mice was used for a secondary transplant in mice fed on normal chow (Fig. S2C). The disease was allowed to progress for one month prior to analysis. This time point was chosen based on humane endpoints. At the time of sacrifice, recipients of leukemic OA bone marrow had lost 20% of body weight (Fig. 3E). Bone marrow from EPA supplemented BCR-ABL leukemic mice caused less severe disease as compared to OA supplemented mice, as shown by inhibition of leukocytosis (Fig. 3F, Fig S1C) and reduction in splenomegaly (Fig. 3G). Mice receiving OA leukemic bone marrow showed increased proliferation of splenocytes into LSC-CFUs (Fig. 3H). The number of colonies was too numerous to count (TNC) in all but one OA group, whereas the EPA group consistently showed limited cell growth (Fig. 3H). Taken together, these results supported our findings that EPA supplementation limited the LSC population in the spleen.

The T315I mutation in BCR-ABL is resistant to most current TKI therapeutics (20). These therapies are not only ineffective, but are also extremely damaging to healthy cells. EPA-supplemented mice transplanted with T315I bone marrow showed a decrease in disease, as compared to OA controls (Fig. 3I–J, Fig. S2D–F). Mice were monitored for overall survival and were euthanized based on humane endpoints, including drastic changes in behavior (data not shown) and in body weight (Fig. S2D). The experiment was ended at day 18 post-transplant. EPA mice showed a significant decrease in splenomegaly at time of sacrifice (Fig. 3I). Of particular interest, the EPA diet significantly limited the LSC population, as seen from LSC-CFU assay (Fig. 3J) and from flow cytometric analysis (Fig. S2F). No T315I LSC-CFUs formed ex vivo from EPA-supplemented bone marrow or spleen (Fig. 3J). These results suggest a potential breakthrough in limiting TKI-resistant CML with dietary change, where a COX-derived metabolite of EPA, such as Δ12-PGJ3, was likely mediating this process.

Inhibition of cyclooxygenase and the downstream CyPG cascade blocks the antileukemic effect of EPA

We hypothesized that COX activity was critical in mediating the antileukemic effects of EPA supplementation. To examine such a relationship, we used the NSAID indomethacin, a non-specific inhibitor of COX-1 and COX-2, to limit the endogenous production of prostanoids, including Δ12-PGJ3. Mice were given indomethacin in drinking water ad libitum one week prior to transplantation of BCR-ABL LSCs. Interestingly, the protective property of EPA supplementation in BCR-ABL LSC-transplanted mice was blocked by indomethacin (Fig. 4A–E). Indomethacin-treated mice on both OA and EPA diet showed the same severity of disease as measured by the development of splenomegaly, leukocytosis, and disruption of normal splenic architecture, represented by total histology scoring (Fig. 4A–C, Fig. S3A, B). We previously reported Δ12-PGJ3 as mediating apoptosis in LSCs (2). To determine the level of apoptosis in EPA-supplemented LSC-transplanted mice, TUNEL staining was preformed on splenic sections. EPA-supplemented BCR-ABL mice displayed increased apoptosis in the spleen compared to OA-supplemented mice (Fig. 4D). In contract, indomethacin treatment eliminated the antileukemic property of EPA as measured by the decreased TUNEL+ cells following NSAID treatment. There was no difference in apoptosis between dietary groups in healthy (non-leukemic) splenic tissue (Fig. S1D). These data suggest that a metabolite of the COX pathway was responsible for the increased apoptosis, in EPA-supplemented CML mice. Furthermore, flow cytometric analysis of LSCs showed that indomethacin blocked the protective effects of EPA. EPA-supplemented CML mice treated with indomethacin showed no difference in the percentage of LSCs in the spleen and bone marrow when compared to OA alone and OA + indomethacin groups (Fig. 4E). However, treatment of EPA-fed mice on indomethacin with exogenous Δ12-PGJ3 starting one-week post-transplant of LSCs rescued the anti-leukemia effect in these mice as seen by significantly lowered BCR-ABL LSCs in the spleen and bone marrow, which were similar to that observed in EPA-treated mice (Fig. 4E). Surprisingly, the general phenotype of these mice was only partly restored to that of EPA-supplemented mice, as seen by decreased splenomegaly (Fig. S3C, D).

Figure 4. Generation of CyPGs downstream of the COX enzyme are essential for antileukemic effect of EPA supplementation.

Figure 4

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A-E: EPA- or OA-supplemented mice were on diet for seven weeks prior to indomethacin (0.00325% w/v). Mice were transplanted with BCR-ABL+ LSCs at week 8; F-H: Mice were irradiated with 450 rads and transplanted with 1×106 total BCR-ABL cells. Mice were treated with HQL-79 (30mg/kg) by i.p. injection throughout bone marrow transplant. A, Spleen size of EPA- or OA-supplemented mice with or without indomethacin (indo) treatment. Healthy spleen size: small dashed line (0.1g). Data compiled from six independent experiments. n= 8–16 per group. Data were compared to OA only group for analysis. B, WBC counts in peripheral blood of BCR-ABL+ LSC transplanted mice (12–14 days post-transplant). Shaded area (1.8–10.7 k/µl) indicates healthy range of WBC counts. n= 3. C, Total histology scores of OA- and EPA-supplemented BCR-ABL+ LSC transplanted mice treated with or without indomethacin (0.00325% w/v). Six parameters (DSA, TI, MF, MK, EMH, and necrosis) were scored from 0–4 and total scores are represented. n= 6–8. D, TUNEL assay of the splenic sections from BCR–ABL+ LSC transplanted mice on EPA and OA supplemented diets ± indomethacin. Representative images with TUNEL-positive staining are shown. Total area of TUNEL+ staining is shown below each panel. E, Flow cytometric analysis of percentage of Lin GFP+c-kit+sca-1+ LSCs in spleen and bone marrow of EPA- or OA-supplemented BCR-ABL+ LSC transplanted mice. Mice were treated with or without indomethacin (indo; 0.00325% w/v). i.p. injection of Δ12-PGJ3 (0.025mg/kg body weight) began seven days post-transplant and continued until sacrifice at day 14. Data are representative of four independent experiments (OA/ EPA); two independent experiments (OA/EPA + indo); one independent experiment (Δ12-PGJ3). Overall sample sizes: bone marrow: OA: n=12, EPA n= 9, OA + Indo: n=6, EPA + Indo: n= 5, EPA + Indo + Δ12-PGJ3: n= 3; spleen: OA: n= 12, EPA n= 9, OA + Indo: n= 7, EPA + Indo: n= 5, EPA + Indo + Δ12-PGJ3: n= 3. F, WBC counts in peripheral blood of BCR-ABL transplanted mice on day 16 post-transplant. n= 4. G, Total CML LSC-CFU counts in spleen and bone marrow from BCR-ABL transplanted mice. n= 4. H, Representative images of CML LSC-CFU from BCR-ABL transplanted mice in spleen and bone marrow. Data represent individual mice with error bars of mean ± SEM. *p< 0.05, **p< 0.01.

To further examine if PGD3-derived CyPGs mediated the EPA-effect, BCR-ABL mice were given i.p. injections of HQL-79, a specific H-PGDS enzyme inhibitor, from time of transplantation to experiment end (Fig. S3E). At day 14 post-transplant, HQL-79 treatment of EPA-fed mice had significantly higher total WBCs, as compared to EPA-supplemented mice (Fig. 4F, Fig. S3F) and displayed similar disease index to OA-supplemented and OA + HQL-79 BCR-ABL mice. The BCR-ABL LSC population was analyzed by flow cytometry and LSC-CFU analysis. HQL-79 treatment of EPA mice doubled the LSC population in the spleen (Fig. S3G). BCR-ABL LSC-CFU formation was significantly increased in the EPA + HQL-79 group compared to EPA only. In the bone marrow there was a 6.5 fold increase in LSC-CFU formation, and splenocytes formed 5.5 fold more LSC-CFUs (Fig. 4G, H). There was no difference in LSC-CFU formation between OA diet only and OA diet + HQL-79 in BCR-ABL transplanted mice (Fig. 4G, H). HQL-79 treatment eliminated the protective effect of EPA supplementation and returned BCR-ABL LSCs to levels as in OA controls. These studies reveal that COX and H-PGDS play an important role in the antileukemic function of EPA in BCR-ABL transplanted mice.

EPA supplemented mice display decreased formation of LSC-CFUs in AML

The MLL-AF9 bone marrow transplant model was used to explore whether EPA was protective in AML. At day 13 post transplant, analysis of WBC showed severe leukocytosis in all OA-supplemented mice, while two EPA-supplemented mice had WBC counts within the normal healthy range (Fig. 5A). Further analysis of mice showed a slight increase in spleen weight in OA- as compared to EPA-supplemented AML mice (Fig. 5B). However, of greatest interest was the effect of EPA on suppressing AML LSC-CFUs. There was a significant decrease in the formation of AML LSC-CFUs from EPA-supplemented spleen and bone marrow. OA splenocytes and bone marrow showed a nearly two-fold increase and a four-fold increase, respectively, compared to EPA (Fig. 5C, D). Though EPA colonies were formed, the morphology of these colonies appeared to be CFU-GM like in nature (Fig. 5D).

Figure 5. EPA-supplemented mice display decreased signs and symptoms of AML progression.

Figure 5

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A-D: Primary AML transplant of 1×106 total AML bone marrow. Mice were euthanized on day 13 post-transplant; E-H: Secondary AML transplant of 1×106 pooled bone marrow from primary OA or EPA supplemented AML mice (See Supplementary Fig. 2C for details). Mice were euthanized at day 14 post-transplant‥ A, WBC counts in peripheral blood of primary OA and EPA supplemented mice. B, Spleen weight of primary OA- and EPA-supplemented AML mice. C, Total AML LSC-CFU counts in the bone marrow and spleen of primary transplanted OA- and EPA-supplemented mice. D, Representative image of AML LSC-CFU in the bone marrow and spleen of primary transplanted OA and EPA supplemented mice. E, WBC counts in peripheral blood of secondary transplanted AML mice. F, Spleen weight of mice transplanted with AML cells from OA- or EPA-supplemented mice. G, Total AML LSC-CFU counts in the bone marrow and spleen of secondary transplanted AML mice. H, Representative image of AML LSC-CFU from the bone marrow and spleen of secondary transplanted AML mice. Data represent individual mice with error bars of mean ± SEM. n= 4. *p< 0.05, **p< 0.01, ***p<0.001.

Secondary transplantation with either OA or EPA total bone marrow was used to determine the functionality of AML LSCs in EPA-supplemented mice (see Fig. S2C for experimental set-up). Though the peripheral WBC count was not different between diets at 14 days post-transplant, there was a general decrease in WBCs in mice transplanted with EPA-supplemented bone marrow (Fig. 5E). There was a significant decrease in the spleen size in secondary transplanted EPA mice (Fig. 5F). AML LSC-CFUs were lower in the bone marrow of mice transplanted with EPA-supplemented AML cells, but the differences did not reach significance (Fig. 5G, H). In contrast, there was a significant decrease in AML LSC-CFUs from the spleen of secondary EPA-transplanted mice (Fig. 5G). Representative examples of AML LSC-CFUs from secondary transplanted mice showed clear differences in the morphology of the colonies (Fig. 5H). OA colonies were similar to those from primary transplant, whereas EPA colonies are clearly differentiated from the typical AML LSC-CFU. These results also suggest that the antileukemic effect of EPA supplementation is not limited to a specific oncogene.

Discussion

This study demonstrates a novel role for EPA at pharmacological levels in three experimental models of myelogenous leukemia, with a focus on BCR-ABL induced CML. To our knowledge, no previous study has investigated the in vivo role of EPA supplementation on CML or AML progression. Moreover, previous studies cite the beneficial role of omega-3 in limiting COX activity (34), whereas our data indicate that the activity of COX in EPA-supplemented conditions contributes toward the endogenous production of CyPGs, such as Δ12-PGJ3. Mice on an EPA-supplemented diet displayed decreased signs and symptoms of leukemia, including a decrease in the LSC population when compared to mice on an OA-supplemented diet. Furthermore, indomethacin and HQL-79 blocked the antileukemic effect of EPA. This supports our hypothesis that EPA supplementation limits CML due to generation of endogenous CyPGs, most importantly those derived downstream of COX-2 and H-PGDS, namely Δ12-PGJ3. The dose of EPA used in this study was selected specifically to translate to the human pharmaceutical dosage. The FDA approved drug Lovaza®, comprised of 1.86g EPA (55%) and 1.5g DHA (45%), provides an equivalent dosage of EPA as in our study. We limited the supplementation only to EPA to understand the role of EPA-derived CyPGs via the COX-H-PGDS pathway.

We observed a wide range of detectable Δ12-PGJ3 extracted from the cell culture media and plasma following EPA supplementation. While the trend of EPA leading to increased Δ12-PGJ3 remained consistent between independent experiments, the total amount of the compound was variable. This could be due to the extractability of Δ12-PGJ3 and/or the ability of this Michael acceptor electrophile to bind to proteins, thus limiting efficient extraction. In addition, metabolic transformation of Δ12-PGJ3 could also contribute to the observed variability since such metabolic products were not part of LC-MS/MS-based quantitation.

Although targeting inflammation with NSAID treatment has been an attractive proposition in certain types of cancer, our studies demonstrate a potential detrimental role for NSAID use in CML, which may also be true in AML. Most epidemiological studies, to date, are inconclusive regarding the association between non-aspirin NSAID use and leukemia (12–14). Given the controversial nature of the role of NSAIDs in CML, future studies to associate NSAIDs as a potential confounder in omega-3 PUFA trials may shed more light on the role of endogenous CyPGs.

Current chemotherapies are able to control CML for a period of time, but are unable to cure the disease, as a result of the sustained LSC population (16–28, 35). We have previously demonstrated selective targeting of CML LSCs by exogenous treatment with Δ12-PGJ3 (2). By making a change in dietary composition, we demonstrate that Δ12-PGJ3 can be synthesized at levels capable of limiting the disease. It is intriguing to note that the effect of EPA of reducing CML was not due to general differences in the ability of these mice to engraft a foreign bone marrow following irradiation. Engraftment and general parameters of health were not different between diets (Fig. S1E–I). Thus, suppression of the LSC population was primarily due to dietary manipulation, as confirmed by secondary transplantation of bone marrow from EPA-fed leukemic mice into mice fed on normal chow. These data suggest that dietary supplementation is an attractive way to attenuate the relapse of CML, particularly in TKI resistant CML that only responds to ponatinib, a TKI dispensed with a black box warning. Surprisingly, even in the experimental model of MLL-AF9-induced AML that represents aggressive leukemogenesis, high dose EPA proved to be effective. Thus, based on our studies, it appears that the use of pharmacological doses of EPA in combination with existing standard of care, could open new avenues for the successful management of myeloid leukemias.

Our previous work showed that treatment of BCR-ABL LSCs with EPA failed to cause any appreciable increase in apoptosis, supporting the role of the microenvironment in CyPG production (2). The microenvironment, comprised of immune and non-immune cells, has been shown to be important in understanding cancer outcomes (36, 37). Macrophages have been implicated as being major producers of COX-derived metabolites, including Δ12-PGJ3 (2). As shown in Fig. 6, Δ12-PGJ3 is thought to act through paracrine signaling mechanisms on LSCs to activate pathways of apoptosis; this effect does not appear to be limited to CML alone.

Figure 6. Summary of antileukemic properties of dietary EPA supplementation in myeloid leukemia transplanted mice.

Figure 6

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Dietary EPA is incorporated into the phospholipid bilayer of cells. Upon stress, EPA is mobilized by PLA2 that is acted upon by COX-2 to form PGH3. NSAIDs, including indomethacin, inhibit the activity of COX, thus down-regulating the production of all downstream mediators. H-PGDS acts on PGH3 to produce PGD3. Non-enzymatic dehydration of PGD3 leads to formation of Δ12-PGJ3. Inhibition of H-PGDS by HQL-79 blocks the anti-leukemic property observed in EPA-supplemented mice. Δ12-PGJ3 is transported from the cell (depicted as macrophage) and targets the LSC for apoptosis.

Our data support the pivotal role of COX and H-PGDS in the antileukemic effect of EPA, suggesting that any variation in gene expression and/or activity of either or both of these enzymes could potentially impact the outcome. This observation is not limited to leukemia. In fact, several single nucleotide polymorphisms (SNPs) have been identified in the COX-2 gene, PTGS2, and meta-analysis has suggested a possible association to breast cancer (38). Therefore, the translatability of these studies into human therapy would depend on the effect the SNPs have on the expression and/or activity of COX and H-PGDS. Few genome-wide association studies have explored SNPs in the COX gene and resulting leukemia outcomes (39), highlighting an area that should be more closely studied in the future.

In summary, our studies use well-established rodent models of CML, T315I CML, and MLL-AF9 AML to demonstrate that dietary EPA decreases the severity of disease. In the case of CML, the antileukemic effect of EPA was blocked by both indomethacin and HQL-79, therefore confirming the essential role of the COX pathway in the metabolism of EPA into a novel class of endogenous CyPGs, including Δ12-PGJ3. Although deemed safe, the success of this therapy rests on the efficient metabolism of EPA to endogenous levels of CyPGs that target LSCs. Future clinical trials with EPA are warranted to conclusively provide a compelling case for its use as an adjuvant therapy in myeloid leukemia patients.

Supplementary Material

1

Acknowledgments

We thank Dr. Shailaja Hegde for irradiation of mice, Dr. Naveen Kaushal for assisting with bone marrow transplantation, Dr. Ramesh Ramachandran for help with microscopy, Flow Cytometry and Microscopy Facility (Drs. Ruth Nissly and Ningchen Xu), and all current and former members of the Prabhu laboratory for valuable suggestions and help.

Financial Support: This work was supported, in part, by grants from the National Cancer Institute (NIH RO1 CA 175576 to K. S. Prabhu and R. F. Paulson) and the National Center for Advancing Translational Sciences (NCATS) in the form of NIH UL1 TR000127 and TL1 TR000125 to E. R. Finch.

Footnotes

Conflict of Interest: K. S. Prabhu and R. F. Paulson are inventors on a patent [Δ12-Prostaglandin J3] assigned to Penn State Research Foundation and are scientific founders of Nemean Pharma Corporation. K. S. Prabhu and R. F. Paulson’s interests were reviewed and are managed by the Penn State Office for Research Protections in accordance with their conflict of interest policies.

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