CELL CULTURE

The colonic adenocarcinoma cell line HT29 was obtained from the European Collection of Cell Cultures (Salisbury, UK). HT29 cells were grown in Dulbecco's modified Eagle's medium (DMEM; Sigma) supplemented with 5% heat inactivated fetal calf serum (Sigma), 2 mM glutamine, 50 units/ml penicillin, and 50 μg/ml streptomycin (all from Imperial Laboratories, Andover, UK). Cells were grown at 37°C, in 5% vol/vol CO₂ in a humidified atmosphere, and harvested using phosphate buffered saline (PBS) containing 0.05% wt/vol trypsin and 0.02% wt/vol EDTA (Imperial Laboratories). EPA (cis, 5, 8, 11, 14, 17) was purchased from Sigma in sealed ampoules containing 25 mg. The contents were dissolved in ethanol to give a 25 mg/ml stock solution, and crimp sealed in an amber vial under nitrogen. The EPA-ethanol mix was further diluted in DMEM to give final concentrations of 15, 30, and 45 μM. EPA has previously been shown by us to cause apoptosis in HT29 cells at these concentrations14 which are comparable with levels of approximately 65 μM observed in the plasma of unsupplemented human subjects, and substantially lower than concentrations of about 400 μM achieved in subjects supplemented with 3 g of fish oil per day for 18 weeks.23 The final concentration of ethanol was approximately 0.06% vol, a level which was found to have no effect on cell phenotype in control experiments.

CELL VIABILITY ASSAY

The effects of EPA and other treatments on the survival and proliferation of HT29 cells were quantified by measuring the relative numbers of viable adherent cells after culture using a neutral red incorporation assay applied to 96 well plates.24 Cells were seeded into Nunclon 96 well plates (Fisher, Loughborough, UK) at a density of 23×10³ cells per well, allowed 24 hours to adhere, and treated with media containing 15, 30, and 45 μM EPA for 24 or 48 hours, with or without antioxidants or caspase inhibitors, as described below. At least 16 replicate wells were used for each treatment. The relative numbers of viable adherent cells were determined at the end of each experiment by incubation for three hours with 50 μM neutral red (3-amino-7-dimethylamine-2-methylphenozine hydrochloride; Sigma) diluted in DMEM. Plates were then drained, fixed in 40% (vol/vol) formaldehyde/10% (wt/vol) calcium chloride hexahydrate (BDH, Poole, UK) for 30 seconds, washed three times in PBS, and air dried. The neutral red was solubilised from the fixed cells by addition of 200 μl of ethanol (50%)/acetic acid (1%) per well, and absorbance was measured at 550 nm using a Dynatech MR5000 automated plate reader (Dynatech Lab Inc, Alexandria, Virginia, USA). In separate experiments absorbance due to neutral red was found to be linearly proportional to the numbers of adherent HT29 cells over the range 1×10³–50×10³ cells per well (r ²=0.98; data not shown). Results were expressed as a percentage of the absorbance values obtained for control cultures.

CELLULAR LIPID COMPOSITION

Total lipids were extracted from 10⁷ HT29 cells or from isolated mucosal crypts (see below) according to the method of Bligh and Dyer.25 Chloroform extracts were dried under N₂ (g) at 40°C and stored at −20°C in glass vials. The polar phospholipid fraction was extracted using a Sep-Pak column (Waters Corporation, Milford, Massachusetts, USA) according to the manufacturer's instructions. The methanol washes containing the polar phospholipids were collected, and dried under N₂ (g) at 50°C before storage. Conversion of extracted total lipid and phospholipid to fatty acid methyl esters (FAMES) was performed by a cold methylation procedure.26 Changes in fatty acid composition of the whole cell and phospholipid fractions associated with treatment with EPA were assessed by gas chromatographic analysis using a Hewlett Packard 6890 gas chromatograph, equipped with a flame ionisation detector and split injector. Individual FAMES were identified by comparison of retention times with those of standards comprising all of the major fatty acids from C12 to C24 (Nu-Check Prep Inc., Elysian, Minnesota, USA).

ACTIVATION AND INHIBITION OF CASPASES

Caspase 3-like protease activity (DEVD specific) was assessed in adherent HT29 cells grown in 80 cm² flasks with 0, 15, 30, or 45 μM EPA for 24 or 48 hours. DEVDase activity was assayed in cell pellets (2×10⁶ cells) using a colorimetric kit (Clontech, Basingstoke, UK) which detects cleavage of the chromophore p-nitroanilide (pNA) from a consensus DEVD tetrapeptide sequence. The DEVDase inhibitor Z-DEVD-FMK (Calbiochem, Nottingham, UK) was added to one treatment sample as a negative control to confirm that liberation of pNA was DEVDase specific. Positive control cells were treated with 25 μM C₂ ceramide for three hours (Calbiochem) or UV light for 30 minutes (12000 J/cm²); negative control cells were incubated in the absence of EPA. EPA and DEVD-pNA were also incubated together in a reaction (for 1, 12, 24, and 48 hours) to confirm that EPA alone had no capacity to cleave pNA from the DEVD tetrapeptide.

For caspase inhibition studies, cells were incubated with 30 μM EPA in 96 well plates in the presence of cell permeable caspase inhibitors for 24 or 48 hours. Caspase inhibitors (Calbiochem) included Z-DEVD-FMK (caspase 3), IETD-CHO (caspase 8), Z-VAD-FMK (broad spectrum), and Boc-D(OMe)-FMK (BAF; broad spectrum). Cell viability was assessed as described above.

ANTIOXIDANT STUDIES

To establish if oxidative processes were involved in EPA induced reductions in cell viability, HT29 cells were cultured for 48 hours with EPA (0, 15, 30, or 45 μM) in 96 well plates in the presence of the antioxidants N-acetylcysteine (Nac 1 mM; Calbiochem), ebselen (40 μM; Calbiochem), or U74389G (5 μM; Calbiochem). Media were refreshed with EPA and antioxidants after 24 hours and cell viability was determined as described above.

DEPLETION OF INTRACELLULAR GLUTATHIONE

The effects of reduced glutathione (GSH) depletion on the response of HT29 cells to treatment with EPA were investigated usingdl-buthionine sulphoximine (BSO), a specific γ-glutamyl-cysteine synthetase inhibitor that suppresses GSH neosynthesis. To determine the effect of BSO on cellular GSH levels, HT29 cells were grown in 80 cm² flasks (10⁷cells per flask) with 30 μM EPA+100 μM BSO for 48 hours, washed twice with PBS, and collected into microfuge tubes as pellets of 10⁷ cells. Total GSH (reduced GSH and oxidised GSH (GSSG)) was assayed as intracellular glutathione (GSX) units according to the enzymatic recycling method of Baker and colleagues27 which was adapted for an automated biochemistry analyser inhouse (Cobas Mira; Roche, Welwyn, UK). For viability studies, cells were cultured in 96 well plates for 24 or 48 hours in media containing EPA (0–45 μM), EPA+100 μM BSO, or EPA+BSO+ebselen (40 μM) for 24 or 48 hours, and cell viability assayed as described previously.

IN VIVO STUDY

To explore the role of GSH metabolism in induction of apoptosis by n-3 fatty acids in vivo, 72 male Wistar rats (125–150 g) were housed individually in polypropylene containers with wire bottoms and tops in a temperature (21°C) and humidity controlled animal unit with a daily 12 hour light-dark cycle. Prior to the start of the experiment, rats were allowed to consume a semisynthetic powdered diet ad libitum (g/kg: starch 280; sucrose 380; casein 200; corn oil 80; balanced mineral and vitamin mix 60). After four weeks rats were assigned randomly to one of four treatment groups of 18. Two groups continued to receive the basal diet containing corn oil (CO) and two received a similar diet in which corn oil was entirely replaced with fish oil (FO: 96.7% triacylglycerols of which 18.7% C20:5 and 8.0% C22:6; Callanish, Breasclete, UK). One CO and one FO group received distilled water ad libitum for the duration of the experiment and the remaining groups (CO-BSO; FO-BSO) received distilled water supplemented with BSO (Sigma) at a final concentration of 5 mM. Weight gain and food and water intake were measured every 24 or 48 hours. After 24 hours, 48 hours, and one week, six rats from each group were killed by intraperitoneal injection of pentabarbitone sodium (1.0 ml) followed by cervical dislocation under deep anaesthesia. The entire colon was removed via a midline abdominal incision and tissues were processed as described below.

To determine the effects of the experimental treatments on mucosal lipids, a parallel experiment was performed using a further 20 male Wistar rats assigned to four groups of five each, and fed CO or FO, with or without 5 mM BSO in their drinking water, for seven days. They were then killed, and intact colonic crypts were isolated for lipid analysis as described for HT29 cells.

ANALYSIS OF CRYPT CELL PROLIFERATION AND APOPTOSIS

The isolated colon was everted on a 3 mm diameter metal rod and cut in half. Full thickness tissue samples (approximately 5 mm) from the distal end were fixed in ethanol:acetic acid (75:25) and stored at 4°C in 2 dram glass vials. For analysis of numbers and spatial distribution of apoptotic and mitotic crypt cells, tissue samples were rehydrated in distilled water, hydrolysed in 1 M HCl for seven minutes at 60°C, and stained with Fuelgen's reagent. Tissues were microdissected under low power microscopy to yield thin strips of crypts which were gently squashed beneath a coverslip and viewed under a light microscope (×400). The length of each crypt was determined by comparison with a calibrated linear eyepiece graticule (Nikon UK Ltd), apoptotic and mitotic cells were identified by morphological features as previously described,8 and their positions were recorded and allocated among five equally spaced longitudinal compartments. A total of 20 crypts were analysed for each rat and data were expressed as total number of mitotic or apoptotic nuclei per crypt or crypt compartment. The origin of the samples was unknown to the histologist during the analysis.

MEASUREMENT OF EPITHELIAL GLUTATHIONE LEVELS

Intact colonic crypts were isolated from segments (approximately 4.0 cm) of freshly excised colon. The segments were stirred gently for 30 minutes in PBS (pH 7.4; 0°C) containing EDTA (3 mM; BDH) and dithiothrietol (1.0 mM; Sigma), transferred to fresh buffer in polythene tubes (25.0 ml), and shaken by hand to disaggregate the crypts. The denuded muscle tissues were discarded, the crypt suspension was pelleted by centrifugation (1000 rpm), and the pellets (500 μl) were frozen in liquid N₂. Frozen pellets were later thawed and probe sonicated for 15 seconds prior to analysis. One subsample (50 μl) was assayed for total protein (Micro-BCA Protein Assay Kit; Sigma) and a second (200 μl) for total GSH (reduced and oxidised: GSH+GSSG=GSX), as described above for HT29 cells.

STATISTICAL ANALYSIS

The significance of differences between the various treatment groups in this study was assessed by one way analysis of variance coupled with either Dunnet's test to compare means of treatments with those of untreated cultures or Tukey's test for comparison of individual means. In some experiments, two way analysis of variance using the general linear model was also used. Probability values of less than 5% (p<0.05) were regarded as significant. All analyses were carried out using the Minitab statistical package (State College, Pennsylvania, USA).

EFFECTS OF EPA ON VIABILITY AND LIPID COMPOSITION OF HT29 CELLS

Exposure of cells to EPA (15, 30, and 45 μM) caused statistically significant reductions in cell viability compared with untreated cells at 24 and 48 hours after initial exposure (fig 1). Cell viability was inversely associated with increasing concentrations of EPA from 15 to 45 μM at both time points (p<0.05). Two way analysis of variance revealed a significant interaction between treatment with EPA and duration of exposure (24 or 48 hours). The percentage of EPA (C20:5) in the phospholipid fraction of HT29 cells treated with 15 μM EPA for 24 hour rose from an initial value of 0.91% (n=2; range 0.02%) to 18.2% (n=2; range 1.76%). There was no further rise after treatment for 48 hours.

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Figure 1

Adherent cell viability of HT29 cells following treatment with increasing concentrations of eicosapetaenoic acid (EPA 15–45 μM) for 24 or 48 hours. Cell viability was assessed using the neutral red assay, and data are expressed as per cent of the optical density (OD)^550nm of untreated control cultures (mean (SEM)). A minimum of 16 OD^550nm readings were taken for each treatment. EPA significantly reduced cell viability after 24 and 48 hours at all concentrations tested (p<0.05) compared with untreated cells. Reduction in viable cell number was significantly associated with EPA treatment and time (p<0.001).

INDUCTION OF CASPASE ACTIVITY

Exposure of cells to EPA at 15, 30, and 45 μM induced significant increases in DEVDase activity expressed as nmol pNA/h per 10⁶ cells compared with untreated cultures after 24 and 48 hours of exposure (fig 2). DEVDase activity also increased significantly (p<0.05) with increasing concentrations of EPA over the range 0–45 μM and there was a significant interaction between treatment with EPA and time. DEVDase activity was found to be specific for caspase 3-like proteases as liberation of pNA after exposure to EPA at 45 μM (10.8 (1.2) DEVDase units) was blocked by addition of a DEVD consensus caspase inhibitor to the reaction (3.0 (0.1) DEVDase units; p<0.05).

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Figure 2

Activity of DEVDase caspases (chiefly caspase 3) in HT29 cells following treatment with increasing concentrations of eicosapetaenoic acid (EPA 15–45 μM) for 24 or 48 hours. Positive control cells were also treated with 25 μM C₂ ceramide (three hours) or 12 000 J/cm² (UV, 30 minutes). Data are expressed as mean (SEM) pNA liberated from a DEVDase consensus cleavage site per hour per 10⁶ cells of four samples per treatment group. Caspase activity was significantly higher in all EPA treated cells than in untreated cells after 24 and 48 hours (p<0.05).

CASPASE INHIBITION

The adverse effects of EPA on HT29 cell viability were blocked or greatly diminished by cell permeable caspase inhibitors. Addition of 30 μM EPA to HT29 cultures significantly reduced cell viability compared with untreated cultures at 48 hours (p<0.05) but when cells were treated with EPA and the broad range caspase inhibitors Z-VAD-FMK or BAF, cell viability was significantly higher than after treatment with EPA alone (table 1). The caspase 3 inhibitor Z-DEVD-FMK and the caspase 8 inhibitor IETD-CHO also significantly increased cell viability after 48 hours in the presence of 30 μM EPA. There were no significant effects of any of the caspase inhibitors in the absence of EPA on cell viability of control cultures.

EFFECT OF ANTIOXIDANTS ON CELL VIABILITY IN THE PRESENCE OF EPA

Ebselen alone had no significant effect on the viability of HT29 cells but it proved highly effective at blocking the reductions in cell viability caused by treatment with EPA. Indeed there were no significant differences in the viability of cells cultured for 48 hours without EPA or with EPA (15–45 μM) in the presence of ebselen (table2). Similarly, U74389G had no effect on cell viability in the absence of EPA but at 48 hours cell numbers in cultures co-treated with U74389G and EPA at 15 and 30 μM were not significantly different from control values (table 2). Nac alone caused an approximate 10% reduction in viable cell number compared with untreated cells (p<0.05) and was less effective than ebselen at blocking EPA associated reductions in cell viability (table 2). Nevertheless, viable cell numbers were significantly higher in all cultures co-treated with Nac and EPA after 48 hours than in corresponding EPA treated cultures (p<0.05).

EFFECTS OF GLUTATHIONE DEPLETION ON CELL VIABILITY IN THE PRESENCE OF EPA

Exposure to EPA alone caused a 20% reduction in the level of GSX in HT29 cells at 48 hours (p<0.05) whereas treatment with BSO reduced GSX content of cells by approximately 90% (p<0.05). In the first 24 hours of EPA+BSO exposure, cell viability was similar to that of cultures treated with EPA alone (data not shown) but there was an approximate 95% reduction in viable cell number after 48 hours (fig3). The major reductions in cell viability occurred within a window of approximately five hours (45–50 hours after initial exposure) where widespread cell death occurred rapidly in the whole population. Addition of the antioxidant ebselen to EPA+BSO treated cells restored cell viability at all concentrations of EPA tested (p<0.05). However, the broad range caspase inhibitor Z-VAD-FMK, which inhibits caspases 1, 3, 4, and 7, was unable to block EPA induced cell death in the presence of BSO, indicating that this gross cytotoxicity was not dependent on these caspases and therefore was probably necrotic rather than apoptotic.

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Figure 3

Adherent HT29 cell viability following treatment with eicosapetaenoic acid (EPA) (15–45 μM), EPA+100 μM buthionine sulphoximine (BSO), or EPA+BSO+40 μM ebselen for 48 hours. Cell viability was assessed using the neutral red assay, and data are expressed as mean (SEM) per cent of the optical density (OD)^550nm of untreated control cultures. A minimum of eight OD^550nm readings were taken for each treatment. EPA significantly reduced cell viability after 48 hours at all concentrations tested (p<0.05) compared with untreated cells. BSO alone did not significantly affect cell viability. Exposure to EPA (15–30 μM)+BSO significantly reduced cell viability (p<0.05) but addition of ebselen to EPA+BSO cultures significantly increased cell viability (p<0.05).

EFFECTS OF FISH OIL AND BSO ON CRYPT CELL LIPID AND GLUTATHIONE CONTENT IN VIVO

Consumption of the FO diet was associated with significant increases in levels of C20:5 in crypt cell phospholipids, and reductions in C18:2 and C20:4 in FO and FO-BSO animals (p<0.05) compared with those fed CO (table 3). The C20:3 content of phospholipids was also reduced in FO fed animals (p<0.05). There was no significant effect of BSO on crypt cell lipids within the dietary groups. After 24 hours total levels of GSH in isolated colonic crypts from animals fed CO (0.20 (0.04) GSH/mg protein) and FO (0.18 (0.02) mg GSH/mg protein) were not significantly different but both differed significantly (p<0.05) from animals treated with BSO (CO-BSO 0.08 (0.01); FO-BSO 0.07 (0.02) mg GSH/mg protein). At 48 hours and one week there were no further effects of treatment on GSH levels in crypts.

EFFECTS OF LIPID AND BSO ON CRYPT CELL MITOSIS AND APOPTOSIS

Total numbers of mitotic cells per crypt were approximately 30% lower in the distal colon of animals fed the FO diet compared with those fed CO after 24 hours, 48 hours, and one week (p<0.001) (fig 4). To put this into perspective it should be noted that a difference in mitotic frequency of this magnitude would equate to a difference in mitotic rate of approximately three cell divisions per crypt per hour.28 Assuming a crypt density of approximately 300 crypts/mm², this would lead to a deficit in crypt cell production in rats fed FO of about 900 cell divisions/mm²of mucosal surface per hour.

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Figure 4

Mitotic cells per crypt expressed as mean (SEM) in crypts isolated from the distal colon of rats given corn oil (CO) or fish oil (FO) in the diet with or without 5 mM buthionine sulphoximine (BSO) in drinking water. There was a significant reduction in mitotic cells/crypt associated with FO consumption at all time points (p<0.001) but no significant effect of BSO in the drinking water.

There were no significant additional effects of BSO treatment on levels of mitosis in either CO or FO fed rats. In all treatment groups, crypt cell mitosis was maximal within the basal zone, declined in zones 2 and 3, and was entirely absent in the superficial zones 4 and 5 (data not shown). There was no significant difference in crypt length between CO and FO fed groups at 24 hours. However, after seven days mean crypt length in FO fed rats (243 μM) was significantly lower than that of CO fed rats (262 μM; p<0.05) and both BSO treated groups had significantly shorter crypts than their respective controls at this time (about 3%; p<0.05). When data for mitotic cells per crypt were normalised using crypt length as an index of cell number, and re-expressed as mitotic cells per unit crypt length, the distribution of differences between groups remained essentially unchanged and one way analysis of variance confirmed that levels of mitosis were significantly lower in FO and FO-BSO rats than in CO or CO-BSO rats, respectively (p<0.05).

Apoptotic nuclei were rarely seen (<0.05 apoptotic cells per crypt) in the distal colonic tissue of animals fed the CO diet (fig 5). Treatment of CO fed rats with BSO significantly increased levels of crypt cell apoptosis compared with those fed only CO (p<0.05) but there was a greater increase associated with consumption of the FO diet (p<0.001). At each time point, significantly higher levels of apoptosis were found in the colonic crypts of FO fed animals than in CO or CO-BSO animals (p<0.05). Treatment with 5 mM BSO in the drinking water of FO fed animals (FO-BSO) further increased levels of crypt cell apoptosis compared with the FO group (p<0.05), and there was a trend towards an interaction between BSO and the FO diet, and levels of apoptosis (p<0.08). Apoptotic nuclei were found throughout the crypt. When the data were normalised using crypt length as an index of cell number, and re-expressed as apoptotic nuclei per unit crypt length, the distribution of differences between groups remained essentially unchanged.

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Figure 5

Apoptotic cells per crypt expressed as mean (SEM) in crypts isolated from the distal colon of rats given corn oil (CO) or fish oil (FO) in the diet with or without 5 mM buthionine sulphoximine (BSO) in the drinking water. The total number of apoptotic cells is represented by total column height; the number in the top 40% of the crypt is represented in black and in the bottom 60% as the shaded region. Both FO and BSO significantly increased the total number of apoptotic cells/crypt at all time points (p<0.001).