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The chemical composition of Tunisian Nigella sativa L. seeds was investigated. The results showed that linoleic acid (C18:2) was the major fatty acid with 65.1% of total fatty acids (TFAs) followed by oleic acid (C18:1) with 12.7% of TFAs. Neutral lipids (NLs) were mainly composed of triacylglycerols with 98.4% of total NLs. Polar lipids were mainly composed of phosphatidylcholine as the major phospholipid subclass, whereas digalactosyldiacylglycerol was the major galactolipid. Total sterols (TSs) represented 2.2% of the fixed oil and were composed of (-sitosterol as the major sterol representing 60.2% of the TSs. The results indicated that the sterols were mainly present in the esterified and in the free forms with 51.2 and 36.1%, respectively, of TS content. Finally, the aroma composition of the volatile oil from oleoresin exhibited the presence of many bioactive compounds such as p-cymene as the major component followed by ocimene,α-thujene, octen-3-ol with appreciable rates of 1,8-cineole and thymol.


The results obtained showed that Nigella sativa L. seeds were a good source of polyunsaturated fatty acids (PUFAs), phytosterols (PSs) and phospholipids (PhLs) for the human diet. These seeds could be used by the food industry for formulating functional foods enriched with PUFAs and PSs. For pharmaceutical applications, N. sativa L. conjugated sterols could be used as precursors for the hemisynthesis of many hydrosoluble steroids. Also, N. sativa L. seeds are a good source of PhLs and aroma compounds, and therefore, they could be utilized in biscuit manufacturing and in food flavoring. The presence of many bioactive compounds in N. sativa L. essential oil (p-cymene, limonene, α-pinene, linalool and thymol), known for their powerful antimicrobial function, could support the utilization of the essential oil bactericidal agents.


It is actually known that consumers are more and more interested in plants with medicinal interests, which produce natural substances necessary to man's health and have therapeutic effects toward certain illnesses and infections. Indeed, the classic medicine using synthetic medicinal substances with secondary effects is more and more oriented toward an alternative medicine based on the prescription of dietetics and medicines made of medicinal plants.

In this perspective, we judged it useful to carry out our study on the seeds of an oleaginous and medicinal plant that deserves to be developed, and that is black cumin (Nigella sativa L.), an annual herbaceous plant of the Ranunculacea family. This plant is believed to be indigenous to the Mediterranean region but has been cultivated into other parts of the world including Saudi Arabia, northern Africa and parts of Asia. The seeds of this plant have been traditionally used for centuries in the Middle East, northern Africa and India for the treatment of asthma, cough, influenza, eczema, as a diuretic, lactagogue and vermifuge. They are also used in the preparation of a traditional sweet dish composed of black cumin paste, which is sweetened with honey or syrup, and in flavoring of foods, especially bakery products and cheese (Cheikh-Rouhou et al. 2007). Actually, a great deal of attention has been focused on black cumin seeds regarding their nutritional and therapeutic values. The proximate analysis of mature N. sativa seeds showed that the moisture content ranged from 5.52 to 7.43%, crude protein from 20 to 27%, ash from 3.77 to 4.92%, carbohydrates from 23.5 to 33.2% and ether-extractable lipid from 34.49 to 38.72% (Abdel-Aal and Attia 1993; Takruri and Dameh 1998; Salem 2001). N. sativa L. seeds are also a source of potassium, magnesium, calcium, phosphorus, sodium, iron, copper, zinc and manganese (Cheikh-Rouhou et al. 2007). They contain carotene, which is converted by the liver to vitamin A (Saleh Al-Jassir 1992). Recently, many medicinal properties have been attributed to the black cumin seeds and/or its oil, including antineoplastic (antitumor) (Swamy and Tan 2000), antibacterial (Hanafy and Hatem 1991), anti-inflammatory (Al-Ghamdi 2001), analgesic (Al-Naggar et al. 2003; Thabrew et al. 2003), antiulcer (Akhtar et al. 1996; El-Dakhakhny et al. 2002; Rajkapoor et al. 2002) and antispasmodic (Gilani et al. 2001).

These therapeutic properties and nutritional values of N. sativa seeds were because of the presence in the seeds of many bioactive compounds such as essential fatty acids (linoleic and linolenic acids), polar lipids ([PLs] mainly phospholipids [PhLs] and galactolipids [GLs]), sterols, aroma compounds and many other constituents. According to Rathjen and Steinhart (1997), PhLs are widely distributed in food, and pro- as well as antioxidant effects have been attributed to them. Their technologic functions and nutritional benefits enable them to be used as multifunctional additives in food as well as for pharmaceutical and industrial applications (Endre and Szuhaj 1996). The development of new or better sources of PhLs is potentially a very important research area for the future (Cherry and Kramer 1986). As for plant GLs, they are thought to be nutrients in the human diet, but little is known about their intestinal digestion and absorption in mammals (Anderson et al. 1995). On the other hand, phytosterols (PSs) in vegetable oils are hypocholesterolemic and their antioxidant activity has been attributed to the formation of an allylic free radical and its isomerization to other relatively stable free radicals (Wang et al. 2002). Plant sterols also may inhibit colon cancer development (Awad et al. 1998). The analysis of sterols provides a powerful tool for the quality control of foods and vegetable oils and for the detection of oil and mixtures otherwise not recognized by the fatty acids profile. Along with free sterols, the seeds also contain sterol derivatives with the 3-hydroxyl group either esterified with a long-chain fatty acid (steryl esters) or alternatively (-linked to the 1-position of a free monosaccharide (steryl glycosides [SG]) or of an acylated monosaccharide (acylated steryl glycosides [ASG]) in which preferably, the 6-position of the sugar is esterified by a fatty acid (Breinhölder et al. 2002). Conjugated sterols are of great importance to the pharmaceutical industry because they constitute the precursors for the hemisynthesis of many hydrosoluble steroids such as saponins, alkaloids, provitamins D2 and D3, and cardiotonic glycosides. In addition to the components listed above, N. sativa seeds contain appreciable amounts of essential oil composed of many bioactive compounds with antioxidant, antibacterial and antifungal properties. As mentioned in the literature, the essential oil was generally extracted either by hydrodistillation (D'Antuono et al. 2002; Nickavar et al. 2003) or by steam distillation (Burits and Bucar 2000). These hot methods of extraction could affect the oil properties and may induce partial or total alteration of many compounds. Finally, the study of oilseeds for their minor constituents is useful in order to use both the oil and minor constituents efficiently.

Literature data on the chemical composition of N. sativa L. seeds are very limited, and the investigation on lipid classes, total sterols (TSs) and their repartition in four sterol classes (free sterols and conjugated ones) in Tunisian N. sativa L. seeds has not yet been done. In addition, to our knowledge, there is no data reporting the biochemical composition of the aroma fraction extracted from N. sativa seeds by the technique of dynamic headspace. The advantage of this technique is that it preserves the real composition of the aroma fraction without any artifacts or secondary products that may be produced by heating like in the other techniques such as hydrodistillation or steam distillation.

In order to strengthen the valorization of N. sativa seeds as a new source of edible oil with nutritional, industrial and pharmaceutical importance, we investigated in the present work, neutral lipids (NLs), PLs and total PSs of the fixed oil and biochemical composition of the aroma fraction extracted from the seeds by the technique of dynamic headspace. Information on triacylglycerols (TAGs), PLs, PSs and aroma compounds is important in processing and utilizing both the fixed and volatile oils and by-products.


Plant Material, Standards and Reagents

Fully ripened seeds used in the present work were harvested in the year 2003 from the agricultural province of El Gobba located in the region of Menzel-Temime in the northeast of Tunisia. The latitude of the region is 36.74° and the altitude is 45 m. The precipitation average was 500 mm/year and the monthly temperature average was 19.9C. After harvest, the seeds were stored at 4C until extraction. Silica gel plates used for thin-layer chromatography (TLC) were from Merck (Darmstadt, Germany). BSTFA (N,O-Bis(trimethylsilyl)trifluoroacetamide) + TMCS (Trimethylchlorosilane) (99:1) used for the sylilation of sterols and berberine used for sterol identification were purchased from Fluka (Buchs, Switzerland). Betulin (Lup-20[29]-en-3β, 28diol) used as the internal standard for sterol quantification was obtained from Aldrich (St. Louis, MO). The highly purified standards used for the aroma characterization of N. sativa L. volatiles were the following: α-pinene, β-pinene and 1,8-cineole from Supelco (Bellefonte, PA); sabinene, α-phellandrene, myrcene, p-cymene, terpinolene, octen-3-ol, terpinene-4-ol, carvacrol and ocimene from Fluka; thymol from Merck; and limonene, γ-terpinene and α-terpineol from Aldrich. All reagents and chemicals used in the study were analytical grade.

Determination of Fresh Matter and Dry Matter Weights

After sampling, 30 seeds were weighed in order to determine their fresh matter weight. Thereafter, these same seeds were dried at 103C until constant weight was reached in order to evaluate their moisture content and their dry matter weight.

Total Lipid Extraction

Samples of 200 seeds were first fixed during 3 min with 10 mL of boiling water in order to inactivate the phospholipases (Douce 1964). Then, they were finely ground in a china mortar into the chloroform–methanol–hexane (3:2:1, v/v/v) mixture (Marzouk and Cherif 1981). After the addition of water for fixation followed by a centrifugation during 10 mn at 2,000 rpm (3,000 × g), the organic phase containing total lipids, was recovered, dried under a nitrogen stream and conserved at −20C for further analyses.

Glycerolipid Fractionation by TLC

Lipid classes were separated by TLC using glass plates of 20 × 20 cm, covered with silica gel (G60, Merck) at a thickness of 0.25 mm. For this, the plates were activated at 120C for 2 h immediately before use, and approximately 30 mg of total lipids per gram of adsorbent was fractioned. NLs were separated according to the method of Mangold (1964) using a mobile phase of petroleum ether–ethanol–acetic acid (70:30:0.4, v/v/v). PLs were separated using a mixture of chloroform–acetone–methanol–acetic acid–water (50:20:10:10:5, v/v/v/v/v) as described by Lepage (1967). Lipid spots were detected after a brief exposure of the plates to iodine vapors saturating a tightly closed vat. The identification of lipid classes was made by comparing their Rf values with those of authentic standards chromatographed under the same conditions. After the detection of the lipid classes, the plates were submitted to a nitrogen stream in order to eliminate iodine, and individual bands were scraped from the plates and corresponding glycerolipids were recovered from the silica gel by elution with 5 mL of hexane.

Fatty Acid Methylation

Total fatty acids (TFAs) and those of main glycerolipids were methylated using the sodium methylate at 3% in methanol according to the method described by Cecchi et al. (1985). Methyl heptadecanoate (C17:0) was used as the internal standard.

TS Extraction

TSs of N. sativa L. seeds were extracted by Soxhlet apparatus according to the method described by Brenac and Sauvaire (1996). For this, 10 g of finely ground seed samples was placed in cellulosic extraction cartridges of 30 × 80 mm. Two successive extractions were realized using the Soxhlet extractor: the first one for 12 h using hexane, permitting to recover majority of the free and esterified sterols, and the second extraction using a mixture of isopropanol–water (70:30, v/v) for 18 h, allowing to recover majority of the SG and ASG (Brenac and Sauvaire 1996).

Saponification and Unsaponifiable Extraction

Two milliliters from each extract (hexanic and isopropanolic) was transferred into a hydrolyze ball, and 1 mL of betulinol (Lup-20[29]-en-3β, 28diol), used as the internal standard in the solution at 0.5 mg/mL in absolute ethanol, was added. After evaporation under vacuum, 8 mL of H2SO4 (0.18 N in ethanol 95%) was added, and the ball content was heated under ebb during 12 h in order to hydrolyze the sterol–sugar bond of SG and ASG. To saponify the sterol–fatty acid bond of esterified sterols, the extract was heated under ebb for 90 min after the addition of 16 ml of KOH (10% in ethanol 95%). After these two steps (hydrolysis and saponification), the obtained unsaponifiable contained only free sterols coming from the conjugated forms.

After decreasing the alcoholic title of the obtained extract by the addition of an equal volume of distilled water, the unsaponifiable was extracted three times with 20 mL of ethylic ether. The obtained extracts were mixed and washed with distilled water until neutrality, detected by the addition of some drops of phenolphtalein to the solution, recuperated after washing.

Unsaponifiable Fractionation by TLC

The obtained unsaponifiable was concentrated under vacuum, diluted with 0.5 mL of chloroform and separated on silica gel plates (G60, Merck) using a developmental mixture of chloroform–ethylic ether (9:1, v/v). After migration, TSs and betulinol spots were visualized under UV light at 365 nm after spraying with berberine (0.1% in ethanol 95%). Sterol spots were then scrapped and eluted from the silica gel with 2 mL of chloroform.

Derivatization of Sterols

Before analysis with gas chromatography (GC), all the samples were dried under a nitrogen stream and derivatized to their trimethylsilylethers. For this, samples were generated for 1 h at 45C in 50 µL of silyl 911 (99% BSTFA + 1% TMCS) according to Wubert et al. (1997).

Separation of Sterols and Sterol Conjugates by TLC

Free sterols, esterified sterols, SG and ASG were separated by TLC according to the method described by Hartmann and Benveniste (1987). For this, 100 µL from hexaneic and isopropanolic extracts were mixed and evaporated to dryness. The obtained residue was then diluted into 100 µL of a mixture composed of chloroform–methanol (2:1, v/v) and fractionated using silica gel plates with alumina support (Silicagel G60, Merck) activated during 1 h at 110C. The plates were submitted to a double development using two solvent combinations: the first one using a mixture of hexane–ethyl acetate (92:8, v/v) and the migration was stopped at 18 cm, followed by the second one using a mixture composed of dichloromethane–methanol–water (90:10:0.5, v/v/v) and stopped at 11.7 cm. After spraying with berberine (0.1% in 95% ethanol), sterolic forms were visualized under UV light at 365 nm. The identification of sterolic forms was achieved by co-chromatography of authentic standards under the same conditions.

Quantitative Estimation of Sterols in the Sterolic Forms by Spectrophotometry

The method utilized for the quantification of sterols in each sterolic form was inspired from the work of Sabir et al. (2003) based on the estimation of sterols by the Liberman–Burchard method: after identification, sterolic forms were scrapped and eluted from the silica gel with 2 mL of chloroform. The obtained eluates were then reduced to dryness and an additional 8 mL of chloroform was added and stirred well until complete dissolution. After the addition of 2 mL of reagents, their absorbance was determined by a spectrophotometer. The Liberman–Burchard reagent reacts with sterols to produce a characteristic green color whose absorbance is determined by a spectrophotometer 259 (Sherwood Scientific Ltd, The Paddocks, Cambridge, U.K.) at 640 nm.

In order to quantify the sterols in each sample, a standard range was prepared from a standard solution of stigmasterol at 1 mg/mL in the chloroform. For this, we put successively 0.5, 1, 1.5, 2 and 2.5 mL of the standard solution into six tubes marked from 1 to 6, while tube 6 was kept blank. Then, 2 mL of the Liberman–Burchard reagent was added to all six tubes and the final volume was made equal in all tubes by adding chloroform. The tubes were covered with black carbon paper and kept in the dark for 15 min. Then, the absorbance of all the standards at 640 nm was determined by a spectrophotometer, and a standard graph was plotted. Finally, the absorbance of the sample solutions was determined after the addition of the Liberman–Burchard reagent under the same conditions.

Aroma Extraction by Dynamic Headspace

Ten grams of finely ground seed samples was transferred into cartridges made of cellulose and their oleoresin was extracted by Soxhlet for 6 h using isopentane. After this, the extract obtained was concentrated under ebb at 37C. The concentrated oleoresin was then introduced in headspace tube and aroma compounds were extracted by the method of dynamic headspace using nitrogen as carrier gas. The adsorbent used was activated charcoal (50 mg). After 3 h, the adsorbent was recuperated and aroma compounds were eluted from active charcoal with 2 mL of ethyl ether.

Gas Chromatographic Analyses of Fatty Acids, Sterols and Aroma Compounds

Fatty acid methyl esters (FAMEs) and aroma analyses were achieved by gas chromatography using an HP 6890 chromatograph (Agilent, Palo Alto, CA) equipped with a Flame Ionization Detector (FID) and a capillary column (HP Innowax): 30 m × 0.25 mm × 0.2 µm with a stationary phase made of polyethylene glycol.

The analyses of FAMEs were performed in split mode under an oven temperature program: isotherm at 150C for 1 min, from 150 to 225C at the rate of 15C/min, from 200 to 225C at the rate of 2C/min and isotherm at 225C for 2 min. Injector and detector temperatures were held at 250 and 275C, respectively. Carrier gas was N2 and flow rate was at 1.6 mL/min. As for the aroma compounds, they were analyzed under the following temperature programs: oven isotherm at 35C for 10 min, from 35 to 205C at the rate of 3C/min and isotherm at 205C for 10 min. The identification of aroma compounds was made by the comparison of their retention times to those of authentic standards analyzed in the same conditions.

The sterol samples were analyzed by GC/MS using an FID HP 5890 chromatograph equipped with an HP5 capillary column (25 m; 0.32 nm; 0.52 µm) made of 5% diphenyl and 95% dimethylpolysiloxane. Helium was used as the carrier gas at a flow of 1 mL/min. The analyses were performed in split mode under the following temperature programs: isotherm at 150C for 1 min, from 150 to 300C at the rate of 6C/min, and isotherm at 300C for 12 min. Injector and detector temperatures were 250 and 280C, respectively.

The GC was coupled to an HP 5972 mass spectrometer operating in the electron-impact mode. The electron energy was set to 70 eV; the source temperature was set to 150C. Spectra were obtained by electron-impact ionization within a mass range of 50–550 m/z (threshold = 150). The scan speed was 1 s/decade. Peaks were identified by comparing their mass spectra with those of authentic compounds. The analyses were performed in split mode with split rate: 60:1, carrier gas: nitrogen and total flow: 1.6 mL/min. Injector and detector temperatures were 250 and 300C, respectively.


Oil Content and Glycerolipids Composition

The oil content of N. sativa L. seeds calculated from the hexane extract on the basis of dry matter weight was of 31.7%. This result was slightly higher than that obtained by Cheikh-Rouhou et al. (2007) who reported an oil content of 28.48% in N. sativa L. seeds from the same location, using the same solvent extraction (hexane). However, they proceeded to the extraction of oil by Soxhlet apparatus during only 8 h, whereas in our case, the extraction time reached 12 h. On the other hand, Atta (2003) reported that the extraction of oil with petroleum ether from the seeds of N. sativa L. originated from Egypt yields 34.78% of crude oil. Nevertheless, this author reported that oil extracted by cold press was lower than that gained by solvent extraction.

In fully ripened seeds, TFAs account for 483.35 mg/g DMW. As shown in Table 1, the main fatty acids detected in N. sativa L. seeds were myristic (C14:0 = 3.2 g/100 g of TFA), palmitic (C16:0 = 12.2 g/100 g of TFA), stearic (C18:0 = 6.3 g/100 g of TFA), oleic (C18:1 = 12.7 g/100 g of TFA) and linoleic (C18:2 = 61.3 g/100 g of TFA), which represents the major fatty acid. It is well known that dietary fats, rich in linoleic acid, prevent cardiovascular disorders such as coronary heart diseases, atherosclerosis and high blood pressure. Also, it was reported that the nutritional value of linoleic acid is because of its metabolism at the tissue levels, which produces the long-chain polyunsaturated fatty acids and prostaglandins (Sayanova et al. 2003).

Table 1. 
Fatty acidPercent of total fatty acids
  1. Values are given as means of three replicates ± SD.

Myristic (C14:0)3.2 ± 0.1
Palmitic (C16:0)12.2 ± 0.1
Stearic (C18:0)6.3 ± 0.3
Oleic (C18:1)12.7 ± 0.2
Linoleic (C18:2)61.3 ± 0.4
Linolenic (C18:3)1.5 ± 0.2
Arachidic (C20:0)0.2 ± 0.05
Eicosenoic (C20:1)0.4 ± 0.1
Behenic (C22:0)2.2 ± 0.1
Saturated fatty acids24.1 ± 0.8
Unsaturated fatty acids75.9 ± 0.8

Other fatty acids present in small amounts in the seeds of N. sativa L. were linolenic (C18:3 = 1.5 g/100 g of TFA), arachidic (C20:0 = 0.2 g/100 g of TFA), eicosenoic (C20:1 = 0.4 g/100 g of TFA) and behenic (C22:0 = 2.2 g/100 g of TFA) acids. In fully ripened seeds, saturated fatty acids represent 24.1% of TFAs, while unsaturated ones form 75.9%. In our study, a few amount of eicosenoic acid was detected (0.4%) in the chloroform–methanol extract and not reported by Babayan et al. (1978). These results are in agreement with those of Cheikh-Rouhou et al. (2007), whereas myristic (C14:0), linolenic (C18:3) and arachidic (C20:0) acids were not detected by Ramadan and Mörsel (2002) in the seeds of N. sativa L. from Turkey. On the other hand, few amounts of myristoleic (C14:1) = 0.18% and lignoceric (C24:0) = 1.08% acids were detected by Saleh Al-Jassir (1992) in N. sativa L. seeds from Saudi Arabia. These two fatty acids were also detected in very few amounts (in trace for myristoleic acid and 0.3% of TFA for lignoceric acid) by Atta (2003) in Egyptian N. sativa L. seeds. This author reported also small rates of palmitoleic acid (C16:1) = 0.7%. Only one study conducted by Cheikh-Rouhou et al. (2007) reported the presence of margaric (C17:0) and margaroleic (C17:1) acids in traces in the seeds of both Tunisian and Iranian N. sativa L. In our study, both of these fatty acids were not detected even in traces, which makes our study in agreement with the majority of previous works. The source of this variability in fatty acid composition may be genetic (plant cultivar, variety grown), environmental, seed quality (maturity, harvesting-caused damage and handling/storage conditions), oil-processing variables or accuracy of detection and quantitative techniques (Ramadan and Mörsel 2002).

Table 2 shows the proportions of NL classes in N. sativa L. seed oil. The data showed that NLs, which represent 93.5% of the total lipids, were mainly formed of TAGs (96%), free fatty acids (FFAs) with 2% and finally, diacylglycerols representing 1.9% of NLs, whereas monoacylglycerols were the less represented with 0.1% of NLs. Nevertheless, these results conflict with those reported by Ramadan and Mörsel (2002) who found that NL profile of N. sativa L. seeds was characterized by exceptionally high levels of FFAs (14.3–16.2% of the total NL). In these studies, the high amounts of FFAs could be the consequence of a high TAG lipolytic activity.

Table 2. 
 g/100 g of NLsg/100 g of PhLsg/100 g of GLs
  1. Values are given as means of three replicates ± SD.

  2. NLs, neutral lipids; PhLs, phospholipids; GLs, galactolipids; MAGs, monoacylglycerols; DAGs, diacylglycerols; FFAs, free fatty acids; TAGs, triacylglycerols; PI, phosphatidylinositol; PC, phospatidylcholine; PG, phosphatidylglycerol; PS, phytosterol; PE, phosphatidyl ethanolamine; PA, phosphatidic acid; MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; SQDG, sulfoquinovosyldiacylglycerol.

MAGs0.1 ± 0.02
DAGs1.9 ± 0.04
FFAs2.0 ± 0.04
TAGs96.0 ± 0.09
PI9.3 ± 0.03
PC37.2 ± 0.05
PG2.1 ± 0.03
PS12.6 ± 0.04
PE21.5 ± 0.06
PA17.3 ± 0.04
MGDG43.7 ± 0.08
DGDG55.9 ± 0.05
SQDG0.4 ± 0.02

Widely distributed in food, the PhLs have both pro- and antioxidant effects (Rathjen and Steinhart 1997; Boyd 2001). Even though they are used as food emulsifiers worldwide and although at the same time, they have a very positive image, their use in functional foods is still limited. Considering the amount of clinical data, there is no doubt that PhL will become a standard ingredient for this rapidly expanding category of food (Schneider 2001). As shown in Table 2, PLs from N. sativa L. seeds account for 6.5% of total lipids (TL) and were constituted of a major percentage of PhLs and a few proportions of GLs representing 0.4 and 0.09%, respectively, of total lipids. PhLs from N. sativa L. seed oil were separated into six classes by TLC and were identified as phosphatidylinositol (PI), phosphatidylcholine (PC), phosphatidylglycerol (PG), phosphatidylserine (PS), phosphatidyl ethanolamine (PE), and phosphatidic acid (PA) by comparing their retention times with those of authentic standards analyzed in the same conditions. PC was the most abundant class with 37.2% of total PhL content followed by PE, PA and PS, respectively. PC and PE together make 58.5% of the total PhL content. As for PG and PI, they were found to be present in insignificant amounts. According to Ramadan and Mörsel (2002), PC and PE together make up to 75% of the total PhLs from the seeds of N. sativa L., which originated from Turkey. On the other hand, these authors have shown that solvent and/or mixtures used in lipid extraction process play an important role in the amount and composition of recovered lipids.

GLs were separated by TLC according to Lepage (1967) into three subclasses: monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG) and sulfoquinovosyldiacylglycerol (SQDG). The most abundant subclass is DGDG, representing 55.9% of the total GL content, followed by MGDG with 43.7%, while SQDG amount is only 0.4%. The average daily intake of GLs in human has been reported to be 90 mg of MGDG and 220 mg of DGDG (Sugawara and Miyazawa 1999). Therefore, it is noteworthy that black cumin seed oil could be an excellent source of GLs in the human diet.

Sterol Composition

To our knowledge, there is only one reported study about the TS composition of N. sativa L. seeds (Zeitoun and Neff 1995). In fact, most previous works have described only the free sterol composition of black cumin seeds. As for the extraction of TSs, the most formerly method used is that of Bligh and Dyer (1959), utilizing chloroform and methanol. But more recently, Moreau et al. (2002) demonstrated the deficiency of this method and reported that the mixture chloroform–methanol is unable to extract all sterols contained in the sample. For this reason, we have used, in the present work, the method described by Brenac and Sauvaire (1996). In this method, the authors proceeded to extract lipids by hexane followed by a second extraction with isopropanol–water (70:30, v/v). Applying this method to our samples gave a majority of free sterols (FS) and esterified sterols (ES) and lower amounts of SG and ASG in the hexane extract, while the isopropanol-water extract contained a majority of SG and ASG, and smaller amounts of FS and ES. Comprising these two extracts allowed recovering the totality of sterols contained in the studied sample. On the other hand, before saponification, we have proceeded, in the present work, to a hydrolysis step using H2SO4 (0.18 N in ethanol 95%) according to Grunwald (1970) and permitted the glycosilic forms of sterols (SG and ASG) to hydrolyze, which releases their sterols. Finally, after saponification with KOH (10% in 95% ethanol), the extract contained only free sterols as the sum of sterols coming from the four subclasses (FS + ES + SG + ASG).

According to our results, TSs of N. sativa L. seeds account for 6.931 mg/g of dry matter weight, which is the equivalent of 2.2% of the fixed oil. As shown in Table 3, the GC/MS analyses showed that N. sativa L. sterols were composed of cholesterol representing a minority with 2.2% of TS, campesterol with 10.4%, Δ5-avenasterol forming 2.4%, two nonidentified sterols representing together 5.1%, β-sitosterol and stigmasterol as major sterols representing 60.2 and 19.6%, respectively, of TSs. As for cholesterol, it has been also detected by Atta (2003) in black cumin seeds from Egypt where it represented 7.2% of TSs, whereas this component is not detected by Ramadan and Mörsel (2002) in N. sativa L. seeds from Turkey. However, these authors reported the presence of lanosterol, representing 3.4% of TSs. On the other hand, it is known that the ratio of β-sitosterol/campesterol could be used as an index to identify the purity and the authenticity of oil. In the present investigation, the β-sitosterol/campesterol ratio is of 5.8, which is the same value found by Nergiz and Otles (1993) in black cumin seeds from Turkey.

Table 3. 
  1. Values are given as means of three replicates ± SD.

Sterols (mg/g in oil)
 β-Sitosterol13.24 ± 0.05
 Stigmasterol4.31 ± 0.03
 Campesterol2.28 ± 0.04
 Cholesterol0.48 ± 0.02
 Δ5-Avenasterol0.52 ± 0.02
 Unknown a0.68 ± 0.03
 Unknown b0.44 ± 0.01
Sterolic forms (g/100 g of total sterol)
 FS36.1 ± 0.07
 ES51.2 ± 0.09
 Steryl glycosides7.8 ± 0.07
 Acylated steryl glycosides4.9 ± 0.05

As shown in Table 3, N. sativa L. sterols were mainly present in the esterified form with 51.2% of the TSs (3.122 mg/g of dry matter weight). In addition, free sterols accounted for 36.1% (2.201 mg/g of dry matter weight) and a few amounts of sterols were distributed between the SG and the ASG, representing together 12.7% of the TSs (0.775 mg/g of dry matter weight).

The predominance of the esterified and the free forms in seeds has been also reported by Katayama and Katoh (1973) during the ripening of soybean seeds. Davis and Poneleit (1974) revealed in corn seeds high proportions of free sterols at the beginning of seed development, which decreased during ripening in favor of esterified sterols probably by the activation of the acyl glycerol: sterol acyltransferase.

Aroma Composition

The optimization of the extraction yield of the volatiles contained in the oleoresin by the technique of dynamic headspace showed that 3 h is the time that permits the extraction of the maximum number of compounds, which is equivalent to 18 volatiles illustrated in Table 4. These compounds belong to four chemical classes: monoterpenic hydrocarbons forming 86.3% of the total volatiles with p-cymene as the major compound (53.1%) followed by ocimene (18.5%); monoterpenic alcohols representing 7.1% of the total volatiles and with octen-3-ol as the major compound (6.5%); terpenic ethers represented only by 1,8-cineole with a percentage of 1.9%; and monoterpenic phenols represented only by thymol with a rate of 1.8% of total compounds.

Table 4. 
Volatile compoundsRetention indexes in HP Innowax capillary column% of total aroma compounds
  1. Values are given as means of three replicates ± SD.

Terpenic hydrocarbons
 α-Pinene11,009.101.4 ± 0.03
 α-Thujene21,015.637.2 ± 0.04
 β-Pinene31,088.351.8 ± 0.03
 Sabinene41,107.890.7 ± 0.03
 α-Phellandrene51,150.600.1 ± 0.02
 Myrcene61,165.372.1 ± 0.04
 Limonene71,187.620.1 ± 0.01
 Ocimene81,236.1618.5 ± 0.05
 γ-Terpinene91,241.121.2 ± 0.02
 p-Cymene101,263.7753.1 ± 0.07
 Terpinolene111,269.070.1 ± 0.01
Terpenic alcohols
 Octen-3-ol121,281.696.5 ± 0.03
 Linalool131,562.520.1 ± 0.02
 Terpinene-4-ol141,590.070.4 ± 0.01
 α-Terpineol151,698.130.1 ± 0.01
Terpenic ethers
 1,8-Cineole161,183.081.9 ± 0.03
Terpenic phenols
 Thymol17>2,0001.8 ± 0.02
Nonidentified  2.9 ± 0.02

Most of these compounds have been identified by D'Antuono et al. (2002) in the essential oil of black cumin originated from Morrocco and extracted by hydrodistillation. Also, these authors have mentioned that p-cymene is the major compound. On the other hand, a less important percentage of p-cymene (14%) was detected by Nickavar et al. (2003) in the essential oil of Iranian N. sativa L. extracted by steam distillation of the oleoresin. These authors reported that trans-anethole was the major component representing 38.3% of the total aroma.

These results confirm that N. sativa essential oil is a good source of bioactive compounds such as p-cymene, limonene, α-pinene, linalool and thymol, which were considered as powerful bactericides (Knobloch et al. 1989). These results may justify and support the utilization of this plant in traditional medicine for the treatment of certain infections.


The present results show that N. sativa L. is a good source of vegetable oil rich in bioactive compounds. This oil of linoleic–oleic type is a good source of essential fatty acids for human nutrition and also could be of an interest in the cosmetic industry because of the effect of linoleic acid on the skin's moisture balance. PL fraction has a high PC and PE proportions, making this oil a good source of these components. As for GLs, their presence in appreciable amounts makes black cumin seeds an excellent source of these compounds in the human diet. On the other hand, the TS fraction in the seed oil, which to our knowledge have not been reported before, could be successfully extracted using the conditions mentioned, which allowed us to consider N. sativa L. as a good source of PSs as well as for the human diet and the pharmaceutical industry. Finally, the aroma composition of the volatile fraction extracted from N. sativa L. oleoresin by the technique of dynamic headspace showed the presence of many bioactive compounds, which has been proven to exhibit antibacterial and antioxidant activities. These particularities could find a large field of utilization.


The authors are grateful to Prof. Mohamed Hammami and for Imed Cherif from the Unity of Mass Spectrometry of the Faculty of Medicine in Monastir for their help and their cooperation.