The protein, fat, moisture, fiber and ash contents of seinat seed flours were determined using the standard AOAC methods 932.06, 925.09 and 923.03, respectively (AOAC, 1990) and carbohydrate content was determined by difference. Iron content was estimated using a UV-Visible spectrophotometer (TECHCOM Co., Shanghai, China) at 480 nm (AOAC, 1995). Magnesium was analyzed by the Ranganna (1986) method. The blue color that developed was read at 650 nm in a UV-Visible spectrophotometer and expressed as magnesium mg·100 g^-1 meal. Other minerals were estimated by atomic absorption spectroscopy (Shimadzu AA 6701F, Atomic Absorption Flame Emission Spectrophotometer equipped with a hollow cathode lamp).

Two hundred g of seeds were milled and mixed with 1 L of n-hexane using a shaker (IKA^® RW 20 digital, UK-plug) at a rate of 440 rpm for 4 h. The mixture was then centrifuged for 20 min at a temperature of (4 °C). The supernatant was then filtered. The extraction process was repeated. The oil was then recovered by evaporating off the solvent using a rotary evaporator (Model N-1, Eyela; Tokyo Rikakikal, Japan), and the solvent was removed under a laboratory fume hood for 30 min at (37 °C). The extracted seinat oil was drained under a nitrogen stream and was then stored in a refrigerator at (4 °C) for analysis.

A Hunter Lab digital colorimeter (TC-PIIG system, Beijing Optical Instrument Co. Ltd., Beijing, China) was used to measure the seinat oil color and color scales as L*a*b* values were recorded. A cylindrical plastic dish (58 mm in diameter and 15 mm in depth) containing the same amount of sample was placed at the light port (50 mm in diameter).

The oxidative stability of seinat seed oil was measured using the Rancimat method (743 Rancimat, Metrohm Co., Basel, Switzerland). Briefly, 3.6 g of oil were weighed into the reaction vessel, which was placed into the heating block kept at 120 °C. Air flow was set at 20 L·h^–1. Volatile compounds released during the degradation process were collected in a receiving flask filled with 60 mL distilled water. The conductivity of this solution was measured and recorded.

American Oil Chemist’s Society (AOCS, 1997) methods were used for the determination of free fatty acids (method Ca 5a-4), peroxide value (method Cd 8-53), saponification value (method Cd 3-25), unsaponifiable matter (method Ca 6a-40), and specific gravity (using a 10 ml pycnometer at 25 °C). The refractive index was determined using an Abbe refractometer at 25 °C. Iodine value was calculated following the procedures of Kyriakidis and Katsiloulis (2000). Specific absorptive values k232 and k270 were determined using a UV spectrophotometer by measuring the absorbance.

Fatty acids were converted to their methyl esters (FAME) following the method of He and Xia (2007) with a slight modification. In brief, 1µL of FAME sample was injected into the gas chromatograph (Series PEG30 M) equipped with a flame ionization detector. GC separation was conducted on a capillary column (PEG30 M; 30m × 0.32mm × 0.50 µm). The carrier gas was nitrogen and the column flow rate was 0.8 mL·min^–1. Initially, the oven temperature was calibrated at 190 °C for 1 min, increased from 190 to 230 °C at a rate of 3 °C·min^–1 and then maintained at 230 °C for 10 min. The temperatures of the injection port and detector were 240 and 250 °C, respectively. The peaks were identified on the chromatogram according to retention data from analyzed standard samples. Finally, fatty acid contents were calculated as percentages (%).

To determine tocopherol composition, the NY/T 1598–2008 method was used. A solution of 250 mg of seinat oil, 25 mL of ethanol, 5 mL of 10% ascorbic acid and 10 mL of a 50% KOH solution were saponified in a capped flask in a water bath at 90 °C for 1 h. After cooling, 100 mL of distilled water were added and mixed. The solution was extracted with 50 mL of diethyl ether. The upper layer was collected and washed with distilled water until a neutral pH was reached. The organic layer was then separated and dried over Na₂SO₄. After filtration of this solution, the solvent was evaporated to dryness under a vacuum at 40 °C. The dry matter was dissolved in 2 mL of ethanol, and then centrifuged at 5.000 rpm for 5 min. The upper layer (10 µL) was injected for the HPLC analysis. The HPLC system (Waters, USA) included a Waters 2996 PAD detector and a Spherisorb Silica column (250 mm × 4.6 mm, 5 µm), operating at 30 °C. The separation of tocopherols was based on a mobile phase of methanol/ water (98:2, v/v) at a flow rate of 1.0 mL·min^–1 with detection at 290 nm. Tocopherols were calculated as mg/g oil.

The sterol composition of seinat oil was determined by derivatization with N-methyl-N-trimethyl-silyl-heptafluor butyramid as silylation agent. The assignments were made using the retention times of the individual sterols and calculation of the relative retention times in relation to betulin as internal standard (ISO/FIDS, 1999). Briefly, 250 mg of oil were saponified with a solution of ethanolic potassium hydroxide by boiling under reflux. The unsaponifiable matter was extracted by solid-phase extraction on an aluminum oxide column on which fatty acid anions were retained and sterols passed through. The sterol fraction from the unsaponifiable substance was separated by thin-layer chromatography (TLC) on 20×20 cm silica gel, 0.25 mm layer thickness using hexane/diethyl ether as developing solvent was extracted again from the TLC substrate. The sterol fraction was detected by gas liquid chromatography using an internal standard (betulin). Unclear cases were detected by GC mass only. Finally, other parameters were hydrogen which was the carrier gas; split ratio, 1:20; injection and detection temperature calibrated to 320 °C; temperature was programmed between 245–260 °C at 5 °C·min^–1.

TPC was determined with the Folin-Ciocalteu method. In brief, 100 mg oil equivalents·mL^–1 were mixed with 0.5 mL of Folin-Ciocalteu reagent and 2 mL of methanol. The mixture was shaken for 1 min, and then 1.5 mL of 15% Na₂CO₃ was added and the mixture was shaken again. Finally, the solution was brought up to 10 mL by adding distilled water. The mixture was incubated at 50 °C for 20 min and centrifuged at 3,000 rpm for 10 min. The absorbance was recorded at 750 nm. TPC of the oil was calculated using Gallic acid as a standard (Liu and Yao, 2007).

The reducing power of the oil was determined using the method of Zheng et al. (2012) with slight modification. Oil concentrations of 5, 10, 15, 20 and 25 mg oil equivalents·mL^–1 were mixed with a sodium phosphate buffer (200 µL, 0.2 M, pH 6.6) and potassium ferricyanide (200 µL, 10 mg·mL^–1) and incubated at 50 °C for 30 min. Trichloroacetic acid (200 µL, 100 mg·mL^–1) was added and the mixtures were again incubated for 5 min to stop the reaction. Then 680 µL of the reaction mixture were mixed with 680 µL of distilled water and 68 µL of ferric chloride (10 mg·mL^–1). 0.5 mM of Ascorbic acid was used as the reference compound. The absorbance of solution was recorded at 700 nm.

To measure the ABTS radical scavenging activity of seinat oil, an aliquot of oil 5, 10, 15, 20 and 25 mg oil equivalents·mL^–1 concentrations were mixed with 900 µL of 100 mM Tris-HCl buffer (pH 7.4), 40 µL of methanol and 50 µL of 0.5% (w/w) Tween 20 solution (Bruni et al., 2004). An ABTS radical was generated by mixing 7 mM ABTS and 2.45 mM potassium persulphate via incubation at 23 °C in the dark for 12 h. Then, 0.1 mL of the sample solution was mixed with 2.6 mL of a diluted ABTS radical solution. The absorbance of the solution was recorded at 734 nm after incubation at 23 °C for 6 min. ABTS radical scavenging assay was calculated using the formula: scavenging activity% = (A_B–A_S /A_B) ×100, where A_B: absorption of blank sample and A_S: absorption of the sample. BHT was used as the reference compound.

The DPPH radical scavenging assay of seinat oil was conducted using a method described by Brand et al. (1995) with a slight modification. Oil concentrations of 5, 10, 15, 20, 25 and 30 mg oil equivalents/ mL were mixed with 3.5 mL of DPPH solution with absorbance at 517 nm. The mixtures were incubated for 30 min at 25 °C. Then, the absorbance was recorded at 517 nm. The DPPH radical scavenging activity was calculated by the formula: scavenging activity% = (A_C–A_S /A_C) ×100, where A_S: absorbance of oil sample and A_C: absorbance of control. Ascorbic acid was used as the reference.

An amount of one mL of β-carotene solution in chloroform (3.34 mg·mL^–1) was pipetted into a flask containing 50 mg linoleic acid and 500 mg Tween 20. The chloroform was removed by rotary evaporation at 40 °C for 5 min and 100 mL of distilled water were slowly added to the solution with vigorous agitation to form an emulsion. A 5 ml aliquot of the emulsion was added to a tube containing 0.2 mL of the antioxidant solution at 200 mg·L^–1 and the absorbance was measured at 470 nm, immediately, against a blank consisting of the emulsion without β-carotene. The tubes were placed in a water bath at 40 °C and the absorbance measurements were made again at 15 min intervals (Lu and Foo, 2000). BHT was used for comparison.

The thermal parameter of seinat oil was measured by using a DSC (DSC-Q2000, V24.9, Instruments, Newcastle, USA). The seinat oil sample (4.8 mg) was weighed directly into a DSC-pan. The oil was rapidly cooled to -60 °C with a speed of 15 °C·min^–1, maintained for 15 min and heated to 120 °C with a heating speed of 15 °C·min^–1. The heating process was repeated and the DSC thermographs were recorded during the second melting. An empty DSC pan was used as reference.

As shown in Table 1, fat, protein, fiber, total sugars, and ash were 31.13%, 196 28.58%, 24.75%, 6.94% and 4.33% on dry weight, respectively, and the moisture content was 4.27%. The high level of oil makes seinat seeds convenient for oil industry application. Despite high percentages of oil in the recorded sample seeds, there are variations in these levels that have been reported. The variations in the oil content can be related to differences in plant species, cultivation climate, ripening level and the isolation method used (Nyam et al., 2009). Since seinat (Cucumis melo var. tibish) seeds also contain a good level of crude fiber, they could be a source of dietary fiber for animals. Seinat seeds were also found to contain high levels of minerals (Table 1). Potassium was the most abundant element (9,548.33 mg·100 g^-1), while calcium was found at the lowest level.

As shown in Table 2, the chemical composition of seinat seed oil was studied. The Relative density value (0.91) was similar to that found by El-Adawy and Taha (2001). The iodine value (110.99) was similar to that reported by Mariod et al., (2009), for Cucumis. melo var. flexuosus. The high iodine value is due to its high content of unsaturated fatty acids (80.18 mg·100 g^–1 oil); this indicates that the seed oil has good edible and drying oil qualities (Eromosele et al., 1997). The peroxide value and free fatty acid composition was 2.50 meq O₂·kg^–1 oil and 1.51%, respectively. The peroxide values indicate that the oil started to degrade since it was identified as highly unsaturated oil. The saponification value of seinat seed oil was recorded as 186.20 mg of KOH·g^–1 of oil. This value was lower than that reported by El-Adawy and Taha, (2001) in their research on watermelon (Citrullus sp) seed kernel oil and higher than that of Rezig et al. (2012) on pumpkin (Cucumis maxima) seed oil. Seinat oil also showed unsaponifiable matter of 1.20%. K232 and k270 were calculated from the absorbance recorded at 232 and 270 nm as 3.06 and 1.54, respectively.

The CIE Lab coordinate values (L*, a*, b*) of seinat seed oil were 64.60, 1.21 and 29.31. Seinat (C. melo var. tibish) seed oil showed a slightly lower L* value and lower a* and b* values than (C. melo var. agrestis) seed oil studied by Mariod et al., (2009), this means that seinat seed oil was not as light in color. The CIE-Lab (L*, a*, b*) values of other vegetable oils, such as palm, soybean, sunflower, olive, and corn ranged from 63.4 to 69.5, 3.8 to 4.4 and 9.2 to 10.4, respectively (Hsu and Yu, 2002). This indicates the presence of yellow pigments like carotenoid compounds.

The results from the Rancimat test are presented in Table 2. The stability of seinat seed oil expressed as the oxidation induction time was about 4.28 h. This value may be justified by the fact that the oil was crude. A linear regression based on the linoleic ratio and the contents of phenols and tocopherols in virgin olive oil showed a good correlation with the oxidative stability measured by Rancimat (Aparicio et al., 1999). Similarly, the Cucumis. melo var. agrestis seed oil obtained by Mariod and Matthäus, (2006) reported an oxidation stability index of 5.7 to 5.9 h under the same conditions.

The fatty acid composition of seinat seed oil is shown in Table 3. Of the nine fatty acids determined, five were unsaturated. Linoleic acid was found in the highest content (61.10%), followed by oleic acid (18.75%). These percentages are near to those reported by El-Adawy and Taha, (2001), for watermelon seed oil. It has been suggested that its high level of linoleic acid makes seed oil specifically prone to oxidation. This fatty acid may have favorable nutritional implications and beneficial physiological effects in the prevention of diseases such as cancer and coronary heart disease (Oomah et al., 2000).

The tocopherol contents (α, δ and γ) of the crude oil are shown in Table 3. δ-tocopherol was the highest content and was about 63.43% of the total tocopherols, followed by γ-tocopherol and α-tocopherol (30.32% and 6.25%, respectively). Oils and derived compounds are a main source of tocopherols. Alpha-tocopherol is benefitial to human nutrition because it has a higher biological activity than other tocopherols (Fatnassi et al., 2009). The total tocopherol content in this crude oil was 43.20 mg·100g^–1 oil, higher than that of Cucumis. melo var. flexuosus seed oil as reported by Mariod et al., (2009).

The Sterols content of this crude oil is illustrated in Table 4. The main sterol in the seed oil was β-sitosterol (289 mg·100g^–1 oil), followed by Δ5-Avenasterol (3.85 mg·100g^–1 oil) from the total sterols (302.70 mg·100g^–1 oil). The Δ5-avenasterol can act as an antioxidant and anti-polymerization agent in frying oils. Sterols with an ethylidene group in the side chain are most effective as antioxidants, and a synergistic effect of the sterols with other antioxidants could be occurring. The total sterols of seinat (Cucumis melo var. tibish) seed oil is higher than that reported by Rezig et al., 2012, for pumpkin (Cucurbita maxima) seed oil and close to the results obtained by Mariod and Matthäus, 2006, for (Cucumis melo var. agrestis) oil. Seinat seed oil compared with other oils could be a good source for human nutrition because it contains a high content of total sterols.

As shown in Figure 1, the DSC of seinat seed oil was measured. The oil sample exhibited four transitions when heated from -60 °C to 120 °C. The first and the last transition respectively occurred at -53 °C and 0 °C, and showed approximately the same temperature range. Commonly, a crude oil sample with a high amount of saturated fatty acids illustrates a DSC melting profile at higher temperature zones when compared to an oil sample with a high amount of unsaturated fatty acids (Nyam et al., 2009). No endothermic phenomenon was shown beyond 0 °C. Such a feature may confirm the liquid state of the seed oil at room temperature (25 °C). It is worth noting that the asymmetry of the recorded peak at -27.17 °C can be regulated by two components of different weights. Actually, the thermogram seemed to correspond to a number of components higher than the visible ones, which suggested the presence of triglyceride fractions with melting points too near to be differentiated under the conditions used. (Herrera and Aňón, 1991).

As illustrated in Figure 2, the antioxidant activity of seinat seed oil was measured by four different assay systems: reducing power, ABTS radical scavenging activity, DPPH radical scavenging activity, and β-carotene bleaching inhibition activity. Edible vegetable oils contain many natural antioxidants such as phenolic compounds, phytosterols, tocopherols (Vitamin E) and carotenoid compounds that might prevent biological systems from reactive oxygen species, preventing chronic diseases such as cancer and cardiovascular diseases (Castelo and Torres, 2009).