Characterisation of non-polar dimers formed during thermo-oxidative degradation of β-sitosterol

Food Chemistry 139 (2013) 464–474

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Food Chemistry journal homepage: www.elsevier.com/locate/foodchem

Characterisation of non-polar dimers formed during thermo-oxidative degradation of b-sitosterol Ewa Sosin´ska a, Roman Przybylski a,⇑, Paul Hazendonk a, Yuan Yuan Zhao b, Jonathan M. Curtis b a b

Department of Chemistry and Biochemistry, University of Lethbridge, Lethbridge, Canada AB T1K 3M4 Department of Agricultural, Food and Nutrition Science, University of Alberta, Edmonton, Canada

a r t i c l e

i n f o

Article history: Received 9 July 2012 Received in revised form 5 November 2012 Accepted 17 January 2013 Available online 29 January 2013 Keywords: b-Sitosterol Dimer Oligomers Thermo-oxidation 3,30 -Sitosteryl ether NMR IR SEC-APCI/MS

a b s t r a c t Thermo-oxidative degradation of sterols at temperature typical for frying leads to the formation of oxidised derivatives, fragmented sterols and oligomers. Recent research on sterol oxidation focuses mainly on the oxysterol derivatives formation to the exclusion of compounds with high molecular mass. The aim of this work was to decipher the chemical structure of non-polar dimers formed during b-sitosterol oxidation at 180 °C in the presence of oxygen. The dimer fraction was separated by size-exclusion chromatography (SEC) after pre-fractionation on silica gel. The chemical structure of the dimers was assessed by 1D and 2D NMR, IR, Raman and MS spectroscopies. NMR data confirmed that the predominant non-polar dimer formed during b-sitosterol oxidative degradation has a configuration of 3b,3b0 -disitosteryl ether. Data from IR and Raman spectroscopies further proved it chemical structure. Applied analytical techniques also confirmed presence of dimers with different configuration than disteryl ethers. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Phytosterols play important roles in plants cells as regulators of membrane fluidity and as precursors of plant hormones (Moreau, Whitaker, & Hicks, 2002). Moreover, they are important constituents of foods, where they are present as free sterols, esterified to fatty or phenolic acids, or as glucosides (Moreau et al., 2002). bsitosterol (Fig. 1) is the most abundant plant sterol found in food products, especially in plant oils, nuts, seeds, cereals, fruits and vegetables (Moreau et al., 2002). Ingestion of plant sterols and stanols can lower amounts of blood low density lipoprotein and with it reduce risk of cardiovascular diseases (Varady, Houweling, & Jones, 2007; Westtrate & Meijer, 1998). Food enriched in plant sta-

Abbreviations: SEC, size-exclusion chromatography; NMR, nuclear magnetic resonance; APCI-MS, atmospheric pressure chemical ionisation mass spectrometry; IR, infrared; ELSD, evaporative light scattering detector; DEPT, distortionless enhancement by polarisation transfer; COSY, two-dimensional proton correlation spectroscopy; NOESY, nuclear overhauser enhancement spectroscopy; HPSEC, high performance size exclusion chromatography; HSQC, heteronuclear single quantum coherence; HMBC, heteronuclear multiple bond correlation; NOE, nuclear overhauser effect; FID, free induction decay; T1, spin-lattice relaxation time; T2, spin–spin relaxation time; TIC, total ion current; XIC, extracted ion chromatogram (reconstructed ion chromatogram). ⇑ Corresponding author. E-mail address: [email protected] (R. Przybylski). 0308-8146/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.foodchem.2013.01.053

nol and phytosterol have been launched to control cholesterol intake and increase amounts of phytosterol in the typical diet to 2 g per day, consumed at this level offers positive health effects (de Jong, Ros, Ocke, & Verhagen, 2008; Westtrate & Meijer, 1998). Phytosterols have also been connected with: inhibition of cancer-cell growth, angiogenesis and apoptosis of cancer cells in lung, stomach, prostate, ovarian and breast cancers, but also with cytotoxicity for aorta endothelial cells (Bradford & Awad, 2007; Rubis et al., 2008; Woyengo, Ramprasath, & Jones, 2009). Phytosterols structurally resembles cholesterol and are susceptible to chemical and enzymatic oxidation. The oxidative stability of sterols is affected by the presence of unsaturation and the steric carbons in the structure, as well as by temperature, time, oxygen availability and the matrix composition (Smith, 1981). The thermo-oxidative degradation of sterols at temperatures typical for frying leads to the formation of oxidised derivatives, fragmented sterols, volatile off-flavor compounds, and oligomers (Rudzin´ska, Przybylski, & Wa˛sowicz, 2009). Oxyphytosterols have been detected in various foods, including coffee, potato chips, French fries, vegetable oils, spreads and infant formulas (Ryan, McCarthy, Maguire, & O’Brien, 2009). Most of the recent research on sterol oxidation focuses on the identification and quantitation of oxysterol derivatives, excluding compounds with lower or higher molecular mass than monomers. A more holistic approach to phytosterols’ thermo-oxidative degradation was recently proposed by

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29 28 21 12 11 19 9

1 2 3

HO

4

5

13

C 8

10

A

B

18

14

20

22

24 23

17

D

26 25 27

16

15

7

6

Fig. 1. Chemical structure of b-sitosterol.

Rudzinska, Przybylski, and Wasowicz (2009). In this work, the amount of sterols transformed into volatiles, fragmented sterols, oxysterols and oligomers was established, indicating that at frying temperature majority of sterols is transformed into the latter. HPSEC was successfully used for separation of oligomers formed during thermo-oxidation of sterols (Lampi, Kemmo, Mäkelä, Heikkinen, & Piironen, 2009; Menéndez-Carreño, Ansorena, Astiasarán, Piironen, & Lampi, 2010; Rudzin´ska et al., 2009). Steroid dimer formation was first observed by Windaus and Borgeaud (1928) and reported that under specific conditions, ergosterol undergoes dehydrogenation and condensation forming bisergostatrienol with C7-C70 linkage. Phytosteryl ether dimers were found in bleached oils and table margarine (Kaufman, Vennekel, & Hamza, 1970; Schulte & Weber, 1987). Smith (1981) theorised the formation of 3,30 -dicholesteryl ether along with other oxycholesterols. During thermo-oxidation of sterols it is expected that sterols will form dimers by ether, peroxy or direct carbon–carbon (C–C) linkages (Byrdwell & Neff, 2004; Christopoulou & Perkins, 1989; Muizebelt & Nielen, 1996). Struijs, Lampi, Ollilainen, and Piironen (2010) proposed formation of stigmasterol dimers via direct C–C, ether (C–O–C) and peroxy (C–O–O–C) linkages, whereas Rudzinska, Przybylski, Zhao, and Curtis (2010) suggested formation of sitosterol dimer via ether C3–C70 linkage. In this work we focused on the identification of the dimers’ structures in order to understand the phytosterol thermo-oxidative degradation. Application of mass spectrometry alone is not sufficient to decipher the chemical structures of oligomers formed during the thermo-oxidative degradation of sterols; therefore, we employed NMR, IR and Raman spectroscopies to further explore chemical structure of dimers. 2. Materials and methods 2.1. Chemicals b-Sitosterol (the supplier declared 78.3% purity, containing campesterol and b-sitostanol), deuterated chloroform (CDCl3, 99.8% D), deuterium oxide (D2O, 99.9% D), tetramethylsilane (TMS), copper (II) sulfate were purchased from Sigma–Aldrich (St. Louis, MO, USA). The HPLC grade solvents tetrahydrofuran (THF), n-hexanes, di-isopropyl ether, diethyl ether, chloroform and toluene along with silica gel (60 Å, 70–230 mesh) and ethanol (99%) were purchased from VWR (Mississauga, ON, Canada). Silica gel GF TLC plates were obtained from Analtech Inc. (Newark, DE, USA). 2.2. Heating and chromatographic pre-cleaning Five hundred mg of sitosterol standard was placed in 100 mL glass ampoule, then 50 mL of oxygen was added and closed container was heated at 180 °C for 24 h. Twenty-five mg of heated

465

sample was dissolved in toluene and loaded on silica gel column (1.3 g of silica, pre-heated at 160 °C for 24 h and adjusted water content to 5%, w/w). The nonpolar fraction containing oligomers was eluted with hexanes/diisopropyl ether (88:12, v/v). After solvent evaporation, the nonpolar fraction was dissolved in THF and further separated by SEC. 2.3. Isolation using HPSEC Dimers were separated using high performance SEC (HPSEC) on a Finnigan Surveyor liquid chromatography (Thermo Electron, Waltham, MA, USA). The nonpolar fraction was injected on two Phenogel columns connected in series (500 and 100 Å, 5 lm, 300  7.80 mm with guard column; Phenomenex, Torrance, CA, USA), kept at 30 °C. Tetrahydrofuran was used as mobile phase at a flow rate of 1.0 mL/min. Dimer fractions were collected using a Gilson FC 203B Fraction Collector (Middleton, WI, USA). Components of the nonpolar fraction or heated standard without fractionation were detected by an evaporative light scattering detector (Sedex 75; Sedere, Alfortville, France) operated at 30 °C with purified air at a pressure of 2.5 bar. 2.4. NMR Solution state NMR spectra were recorded on a Bruker Avance II 300 spectrometer, using 5 mm HX BB probe. The spectrometer was operated at 300.13 MHz for 1H and 75.47 MHz for 13C. A 15–50 mg sample was dissolved in 0.7 mL of CDCl3 with TMS (dH 0.00 ppm, dC 0.00 ppm) and placed in the NMR tube. 1H NMR spectra were obtained with 4 transients using a recycle delay of 2 s, covering a spectral width of 4500 Hz. 13C NMR spectra were obtained under inverse-gated proton decoupling conditions using a recycle delay of 10 s, with 3072 transients covering a spectral width of 16,500 Hz. FID’s were apodized using 0.5 Hz line broadening. 13C DEPT 135 and 13C DEPT 90 experiments were acquired with 512 transients with a 2 s recycle delay. 2D spectra: 1H–1H COSY, 1 H–1H NOESY, 1H–13C HSQC and 1H–13C HMBC were recorded with a spectral width covering 4500 Hz for 1H and 16,500 Hz for 13C. The pulsed-field gradient COSY was acquired with 512 increments of 8 transients of 2048 points with a recycle delay of 4 s. The indirect dimension was linear predicted to 1024 further zero-filled to 2048 points. The pulsed-field gradient NOESY was collected with 256 increments of 8 transients of 1024 points using 4 s recycle delay and a mixing time of 1.5 s. The direct dimension was zero-filled to 2048 points, whereas the indirect dimension was linear predicted for 1024 points, and further zero-filled to 2048 points. The pulsed-field gradient HSQC experiment was conducted with 256 increments of 32 transients for 2048 data points, using a recycle delay of 2.8 s and a 1.72 ms polarisation transfer delay. Data in the direct dimension were zero-filled to 8192 points, whereas the indirect dimension was linearly predicted to 512 points and further zero-filled to 1024 points. The a pulsed-field gradient HMBC experiment was acquired with 256 increments in 64 transients of 512 points, using a recycle delay of 1.5 s and a 62.5 ms delay. The results were zero-filled to 1024 points in direct dimension and linear predicted for 1024 points in indirect dimension. 2.5. IR and Raman spectroscopy IR spectra were recorded on a Bruker ALPHA-S spectrophotometer equipped with DTGS detector, and Platinum ATR module with Diamond crystal plate and KBr as a beam splitter. Spectra were acquired in 8 scans at a resolution of 2 cm1 within 400–4000 cm1. Raman spectra were recorded on a Bruker RF3 100/S spectrometer in the room temperature with sample in a melting point capillary using a laser power of 150 mW. Spectra were recorded within 400

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to 3500 cm1 with a spectral resolution of 4 cm1. The 1064 nm line of a neodym laser was used for sample excitation. 2.6. HPSEC-APCI/MS The analyses were performed on an Agilent 1200 series HPLC system (Agilent Technologies Inc, Palo Alto, CA, USA) coupled to a QSTAR Elite mass spectrometer (Applied Biosystems/MDS Sciex, Concord, ON, Canada) using APCI in positive ion mode. Sample (4 ll) was injected on two columns connected in series: Styragel Ò 1 THF (5 lm, 300 mm  4.6 mm, Waters) and Phenogel (5 lm, 500 Å, 300  4.6 mm, Phenomenex). THF was used as mobile phase at a flow rate of 0.2 mL/min. The following conditions of APCI/MS were used: positive ion mode, probe temperature 370 °C, needle current 2 lA, DP (declustering potential) 30 V, FP (focus potential) 200 V. Nitrogen was used as an auxillary, nebulizing and curtain gas, with values set at 10, 75 and 25 arbitrary unit, respectively. Mass spectra were recorded within range from m/z 150 to 2000. Analyst QS 2.0 software was employed for data acquisition and analysis. 2.7. Synthesis of 3,30 -disitosteryl ether The ether dimer was synthesised according to Bergenthal, Schulte, and Weber (1990), with the following modification, briefly, 1 g of b-sitosterol standard was heated at 150 °C for 25 min with 1 g of anhydrous copper (II) sulfate. The reaction mixture was extracted three times with hexanes; sitosta-3,5-diene was removed with hot ethanol. 3,30 -Disitosteryl ether was purified by a TLC using mixture of hexanes/diethyl ether (97.5:2.5, v/v) as developing solvent. 3. Results and discussion 3.1. Thermo-oxidation of sitosterol standard and dimer fraction collection Sitosterol standard heated at 180 °C produced compounds spanning a wide range of molecular weights. The SEC chromatogram of the non-polar compounds formed during thermal oxida-

120

tion of sitosterol is shown in Fig. 2. Monomers and fragmented sterol molecules are separated from oligomers, however dimers, trimers and tetramers are not baseline resolved. A non-polar dimer fraction was collected from SEC as marked by shaded bar (Fig. 2). Oligomers were the main products formed during thermo-oxidative degradation of b-sitosterol (Rudzinska et al., 2010; Rudzin´ska et al., 2009) and stigmasterol (Lampi et al., 2009; Struijs et al., 2010). Phytosterols degradation leads to the formation of oligomers with a wide range of polarities (Menéndez-Carreño et al., 2010; Struijs et al., 2010), whereas in this study non-polar dimer fraction was analysed by NMR, IR, Raman and MS spectroscopies.

3.2. NMR spectroscopy Data obtained from 1D NMR including 1H (Table 1), 13C (Table 2), 13C DEPT 135, 13C DEPT 45 and 2D NMR including 1H–13C HSQC (Fig. 3a), 1H–13C HMBC (Fig. 3b), 1H–1H COSY (Table 1) and 1 H–1H NOESY (Fig. 4) confirmed that the most abundant compound of non-polar fraction had 3b,3b0 -sitosterol ether structure (Fig. 5). Chemical shifts (d) for proton and carbon atoms in the dimer fraction were very similar (diff. 60.03 ppm for 1H and 60.3 ppm for 13C) to that in b-sitosterol (Tables 1 and 2), with exception for atoms in the ring A. The carbon at dC 76.34 in the dimer fraction was assigned as C-3/C-30 as it correlated with proton at dH 3.28 (H-3a/H-3a0 ) in the HSQC spectrum (Fig. 3a) and with H4a & b/H-4a0 & b0 , H-3a0 /H-3a, H-2a & b/H-2a0 & b0 and H-1a & b/ H-1a0 & b0 in the HMBC spectrum (Fig. 3b). The cross-peak for H-3a with C-30 (and H-3a0 with C-3) is present in the HMBC spectrum of the dimer fraction and the synthetic sitosteryl ether, but correlation between H-3a with C-3 was not observed in the b-sitosterol HMBC spectrum (Fig. 3c). This proves that H-3a (H-3a0 ) is twoor three-bonds away from the C-30 (C-3) in the dimer molecule. The proton at dH 3.28 was assigned as H-3a/H-3a0 as its part in 1 H spin system was revealed by COSY and NOESY spectra (Table 1, Figs. 4 and 6c). The COSY spectrum confirmed strong through-bond couplings between H-3a/H-3a’ and neighbouring protons: H-4b/ H-4b’ (dH 2.21) and H-2a/H-2a0 (dH 1.81). Whereas, the NOESY spectrum indicated strong through-space-direct dipolar coupling interactions, via NOEs, with H-4a/H-4a0 (dH 2.27), H-2a/H-2a0 and H-1a/H-1a0 (dH 1.03) (Fig. 6c). The change in the chemical shift

Sterols

110

monomers

100

sterol molecules

90

Detector response [mV]

Fragmented

Dimers

80 70 60

Trimers

50 40

Tetramers

30 20 10 0 -10 14.0 14.5 15.0 15.5 16.0 16.5 17.0 17.5 18.0 18.5 19.0 19.5 20.0 20.5 21.0 21.5 22.0

Retention time [min] Fig. 2. HPSEC/ELSD profile of the non-polar fraction of thermo-oxidised b-sitosterol standard. Shaded bar indicates collection window for dimeric fraction.

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Table 1 1 H NMR chemical shifts (dH in ppm) of b-sitosterol, the dimer fraction and the synthesised 3,30 -sitosteryl ether with 1H–1H spin coupling constants. 1H–1H COSY correlations for the dimer fraction are also shown. H

1a1 1b 2a 2b 3a 4a 4b 6 7a 7b 8b 9a 11a 11b 12a 12b 14a 15a 15b 16a 16b 17a 18 19 20 21 22R 22S 23R 23S 24 25 26 27 28 29

b-Sitosterol

3,30 -Sitosteryl ether

Dimer fraction

dH (J in Hz)

dH (J in Hz)

1

1.09 1.85 1.84 1.48 3.51 2.29 2.23 5.35 1.51 1.97 1.45 0.93 1.50 1.44 1.16 2.02 1.02 1.57 1.05 1.84 1.25 1.11 0.68 1.01 1.35 0.92 1.34 1.04 1.20 1.17 0.93 1.66 0.81 0.83 1.26 0.85

1.03 1.84 1.81 1.49 3.28 2.27 2.21 5.33 1.49 1.97 1.45 0.91 1.49 1.41 1.16 2.01 1.00 1.56 1.02 1.84 1.24 1.10 0.67 0.99 1.36 0.92 1.33 1.02 1.18 1.14 0.92 1.66 0.81 0.83 1.25 0.84

H-2b, H-1b H-1a H-2b, H-3a H-4b, H-3a, H-2a, H-1a H-4b, H-4a, H-2b, H-2a H-6, H-4b, H-3a H-6, H-4a, H-3a, H-2b H-7b, H-7a, H-4b, H-4a H-6 H-6, H-8b, H-7a, H-14a, H-7b H-11b H-12b, H-12a H-12a, H-9a H-11b, H-12b, H-18, H-12a, H-11a, H-11b H-15a, 8b H-16a, H-15b H-15a H-17a, H-16b, H-15a H-16a, H-15a H-16a H-12a H-1a H-21 H-20 H-23R H-23S H-22R, H-24 H-22S, H-24 H-25, H-23, H-28 H-27, H-26, H-25, H-24 H-25 H-25 H-29, H-24 H-28

m m m m dddd4 (12.02, 11.42, 4.9, 4.22) ddd4 (13.0, 4.9, 2.2) m ddd4 (5.3, 2.22, 1.42) m m m m m m m m m m m m m m s s m d (6.4) m m m m m m d (6.5) d (6.4) m t (7.5)

1

H– H COSY

m m m m dddd4 (12.02, 11.42, 4.6, 4.02) ddd4 (12.8, 4.6, 2.1) m ddd4 (5.4, 2.22, 1.42) m m m m m m m m m m m m m m s s m d (6.2) m m m m m m d (6.6) d (6.6) m t (7.4)

dH (J in Hz) 1.03 1.84 1.81 1.49 3.28 2.27 2.21 5.33 1.49 1.97 1.45 0.91 1.49 1.41 1.16 2.01 1.00 1.56 1.02 1.84 1.24 1.10 0.67 0.99 1.36 0.92 1.33 1.02 1.18 1.14 0.92 1.66 0.81 0.83 1.25 0.84

m m m m dddd4 (12.03, 11.43, 4.62, 4.23) ddd4 (13.0, 4.62, 2.02) m ddd4 (5.4, 2.23, 1.43) m m m m m m m m m m m m m m s s m d (6.3) m m m m m m d (6.6) d (6.6) m t (7.3)

1 For b-sitosterol, dimer fraction and synthesised 3,30 -sitosteryl ether signals were identical for H-n and H-n’ atoms. Accuracy of 1H–1H spin coupling constants is estimated to be ±0.1 Hz except for couplings marked by 2 (±0.2 Hz), or 3 (±0.5 Hz). 4 Long-range couplings (4JHH, 5JHH) with constants 61 Hz were present.

dH from 3.51 ppm (H-3a) and dC from 71.72 ppm (C-3) in b-sitosterol to dH 3.28 (H-3a/H-3a0 ) and dC 76.34 (C-3/C-30 ) in the dimer is characteristic for transition from alcohol to ether (Ning, 2005; Silverstein, Webster, & Kiemle, 2005). The increase in the chemical shift of C-3/C-30 bond in ether molecule by 4.62 ppm results from the substitution of the hydrogen in hydroxyl group by a carbon atom (with higher electronegativity) from other sterol molecule (Fig. 5). Interestingly, the chemical shifts of C-2/C-20 and C-4/C-40 decreased by 2.08 and 2.19 ppm (Table 2), which can be explained by a c-gauche effect, as both carbons are in c-position from the point of substitution. The other, more remote protons and carbons from the substitution site remained less changed in chemical shift (Silverstein et al., 2005). The loss of hydroxyl group at C-3 in formed ether was further validated by comparison of the NOESY spectra (Fig. 6a–d). The correlation between proton in the –OH group with H-3a was only observed in the b-sitosterol NOESY spectrum (dH 2.01 with dH 3.51) (Fig. 6a), but not in the dimer fraction (Fig. 6c) or the synthetic 3,3-sitosteryl ether spectra (Fig. 6d). An experiment with deuterium oxide exchange confirmed that the proton at dH 2.01 was in the hydroxyl group of b-sitosterol (Fig. 6b). Formation of ether with 3a,3a0 configuration has to be excluded, as it would have great impact on the 1H spin system of proton atoms on C-3 and C-4 in comparison with b-sitosterol (Fig. 7a), and it was not observed either in the dimer fraction (Fig. 7b) or the synthetic sitosteryl ether 1H NMR spectrum (Fig. 7c). The double bond at C-5 in the ring B was preserved in the formed ether molecule as con-

firmed by dH at 5.33 (H-6/H-60 ), dC at 121.35 (C-6/C-60 ) and dC at 141.34 (C-5/C-50 ). The assignment of 1H and 13C NMR signals of 3b,3b0 -sitosteryl ether stays in agreement with data published by Bergenthal et al. (1990); they reported full assignment of all 13C NMR signals, but only of eight 1H NMR signals. The HSQC spectrum (Fig. 4a) of the dimer fraction reveals all the 3,30 -sitosterol ether correlations and also some of dimers with campesteryl (i.e. dH-28/ dC-28 0.76/ 15.37) and sitostanol (i.e. dH-18/dC-18 0.64/12.02) units, as well as impurities (e.g. hydrocarbon grease dH 1.26, dC 29.7) (Gottlieb, Kotlyar, & Nudelmanm, 1997; Iida, Tamura, & Matsumoto, 1980). The NMR chemical shifts for synthetic 3,30 -sitosteryl ether are presented in Tables 1 and 2, and all of the signals present in the 1 H and 13C NMR spectra were observed in the spectra of the dimer fraction. Moreover, in spectra of both: synthesised ether and nonpolar fraction, signals arising from dimers containing campesteryl and sitostanyl units were present, as these phytosterols were minor components of b-sitosterol standard. The differences between NMR spectra of the dimer fraction and synthetic 3,30 -sitosteryl ether were some broad low-intensity signals with chemical shifts at 5.39, 4.60 and 4.39 ppm, along with corresponding correlations, i.e., in HSQC spectrum (dH 2.31, m with dC 38.13; dH 4.39, 1H, q, J = 7.1 Hz with dC 77.08; dH 4.6, 1H, m with dC 73.82; dH 5.39, 1H, m with dC 122.7), HMBC spectrum (dH 2.29 with dC 173.3) and COSY spectrum (dH 4.39 with dH 1.39, 3H, s; dH 4.6, 1H, m with dH 2.31, m). In addition, the relative intensity ratio for specific pro-

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Table 2 13 C NMR chemical shifts (dC in ppm) of b-sitosterol, the dimer fraction and the synthesised 3,30 -sitosteryl ether. Dimer fraction

3,30 -Sitosteryl ether

dC

dC

dC

37.26 31.50 71.72 42.24 140.75 121.65 31.80 31.80 50.10 36.45 21.08 39.77 42.20 56.70 24.25 28.26 56.00 11.86 19.33 36.10 18.71 33.86 26.04 45.77 29.05 18.96 19.75 22.98 11.99

37.42 29.42 76.34 40.05 141.34 121.35 31.97 31.90 50.24 36.86 21.06 39.78 42.32 56.79 24.30 28.26 56.04 11.86 19.40 36.14 18.78 33.93 26.02 45.81 29.12 19.03 19.83 23.05 11.98

37.43 29.44 76.35 40.06 141.35 121.36 31.97 31.91 50.25 36.87 21.07 39.79 42.33 56.80 24.31 28.26 56.05 11.86 19.40 36.15 18.79 33.94 26.03 45.83 29.13 19.03 19.83 23.06 11.99

C

b-Sitosterol

11 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

1 For b-sitosterol, dimer fraction and synthesised 3,30 -sitosteryl ether signals were identical for C-n and C-n0 atoms.

ton atoms H-18: H-6: H-3a (Fig. 5) was at 1.00: 0.90: 0.99 in the synthesised 3,30 -sitosteryl ether, whereas for ether in the dimer fraction was at 1.00:0.75:0.65. 1H NMR spectra were obtained with longer delay time (d1 = 5 s) to fulfill condition d1 P 5T1; and values represent summation of integrals for the H-18 and H-3a protons in dimers with sitosterol, sitostanol or campesterol units. The lower intensity of H-6 signal in the synthesised ether is due to the presence of saturated sitostanyl ether next to unsaturated ethers. Whereas in the dimer fraction the lower intensities for both H-6 and H-3a, as well as listed above additional signals in 1D and 2D NMR spectra indicate the presence of other components than disteryl ethers. Presence of dimers with linkages different than specific C3–O–C30 results in lower contribution of the H-3a signal, whereas presence of saturated units in dimers configuration decreased the contribution of the H-6 signal. Moreover other molecules, containing one monomer unit connected with some fragmented sterol molecule could increase the H-18 signal. Furthermore presence of other components in the dimer fraction could influence a molecular mobility, and the T1 and T2 relaxation times of ether atoms leading to changes in line shapes and intensities of signals in spectra. 3.3. IR and Raman spectroscopy The NMR results were further substantiated by IR measurements. In the IR spectrum of b-sitosterol, a very broad band at 3350 cm1 and an intense band at 1050 cm1, typical for O–Hand C–O-stretching vibrations, are clearly visible (Fig. 8a). These are absent in the IR spectrum of the dimer fraction (Fig. 8b) and the synthesised 3,30 -sitosteryl ether (Fig. 8c), indicating the loss of the hydroxyl group as effect of dimerisation. The medium intense absorption band at 1093 cm1 in the dimer fraction and syn-

thesised ether (Fig. 8b and c) is due to a C–O–C-stretching vibration, demonstrating the presence of the ether linkage. The presence of the double bond in both the dimer fraction and the synthesised ether can be confirmed by an absorption band at 1669 cm1 (C@C-stretching vibration) in their Raman spectra (Fig. 9b and c), the same as in b-sitosterol spectrum (Fig. 9a). The IR and Raman spectra of the dimer fraction (Figs. 8b and 9b) were similar to the spectra of the synthesised ether (Figs. 8c and 9c), the only differences were: an additional broad band at 1606 cm1 in Raman and a moderately intense band at 1731 cm1 in IR spectrum of the dimer fraction (Figs. 9b and 8b). The absorption band at 1606 cm1 in the Raman spectrum could be attributed to C@Ostretching vibration of a saturated ketone or C@C-stretching vibration in a five-member ring, whereas the band at 1731 cm1 in IR spectrum to the C@O-stretching vibrations in a carbonyl group (Ning, 2005). Presence of both t(C@O)- and t(C–O)-bands can be due to presence of ester in the dimer fraction. The presence of another carbonyl group configuration is less likely because the typical additional bands at 2820 and 2720 cm1 for aldehydes or at 3000– 2500 cm1 for acids, are absent from the spectrum. For a ketone, the IR absorption would be around 1730 cm1 and the 13C NMR spectrum would provide a signal with chemical shift near to or greater than 200 ppm (Ning, 2005; Silverstein et al., 2005). An aldehyde group would have a proton with a chemical shift in the range of 9.5–10 ppm, and a carbon with d above 180 ppm. Similarly, the proton and carbon atoms of the carboxylic group would result in chemical shifts in the range of 10–13 ppm and above 165 ppm, respectively. An ester group would give signals for 13C NMR with shifts at around 170 ppm, and for proton chemical shifts at 3.3–4.5 ppm (Ning, 2005; Silverstein et al., 2005). Almost none of the aforementioned signals and correlations were found in the 1D and 2D NMR spectra, except for the signals and correlations leading to the hypothesis that some steryl ester was present in the dimer fraction. On the basis of data presented by Wilson et al. (1996) and Lou, Liu, Qi, Wu, and Zhang (2006), the additional signals, discussed in subsection 3.2, at dH 4.6 and dC 73.82 can be assigned as H-3 and C-3 (numeration in steryl ester molecule, where ester group is on C-3 atom); dC 173.3 as C-10 (-CH-COO-steryl unit); dH 2.29 as H-20 (–CH–COO-steryl unit); including dH 2.31 and dC 38.13 as H-4 and C-4; and dH 5.39 and dC 122.7 as H-6 and C5. The thermo-oxidative degradation of sterol leads to the formation of various hydrocarbons and fragmented sterols, therefore the presence of ester with steryl unit and fragment of an oxidised sterol molecule is possible (Rudzin´ska et al., 2009). 3.4. Mass spectrometry Sterols ionised with APCI produce a mass spectrum with a base peak configuration [MH2O+H]+ and a low intensity protonated molecular peak [M+H]+ (Huang, Zhong, Chen, Ye, & Chen, 2007; Rozenberg et al., 2003). Rozenberg et al. (2003) and Huang et al. (2007) reported the presence of these ions with two additional ones: [M2H+H]+ and [M4H+H]+ formed by a dehydrogenation and cyclisation occurring during the APCI ionisation of sterols. In the experiment described here, HPSEC was combined with APCIMS resulting in the chromatograms presented in Fig. 10a, where TIC (total ion current) is superimposed with extracted ion chromatogram (XIC) within m/z range at 780–900. The APCI mass spectrum of the major peak of the dimer fraction eluting at 27.3 min, is shown in Fig. 11a. The most abundant ion in the dimer fraction spectrum had m/z at 397.4 (C29H49,), which represents a dehydrated b-sitosterol backbone. Other observed ions of dimer fragments had: m/z at 395.4 (C29H47) for dehydrated stigmasterol and/or dehydrogenated, dehydrated sitosterol; m/z at 411.4 (C29H47O) and 413.4 (C29H49O) for dehydrogenated sitosterol; and m/z at 383.4 (C28H47) for dehydrated campesterol. Next to

´ ska et al. / Food Chemistry 139 (2013) 464–474 E. Sosin

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a 20 40

C3/3’

80 100

δC [ppm]

60

H3α/3α’

H6/6’ 120

C6/6’

140 5.5

5.0

4.5

4.0

3.5

3.0

δH [ppm]

2.5

2.0

1.5

1.0

0.5

b 20 40

C3/3’

80

H3α’/3α

H4α/4α’ H2β/2β’ H1α/1α’ H2α/2α’ H1β/1β’

100

δC [ppm]

60

H4β/4β’

120

5.5

5.0

4.5

4.0

3.5

3.0

2.5

2.0

1.5

1.0

δH [ppm]

H4α H4β

H2β

H1α

C-3

71 72

H2α H1β 5.0

4.5

4.0

3.5 3.0 2.5 F Chδ [ppm] i l Shif ( )

2.0

1.5

δC [ppm]

c

1.0

H

Fig. 3. 1H–13C HSQC (a) and 1H–13C HMBC (b) spectrum of the dimer fraction and a fragment of 1H–13C HMBC spectrum of the b-sitosterol standard (c). Key correlations are also shown.

listed above monomer ions, produced as effect of dimers fragmentation in the APCI source, other ions with masses typical for dimers were detected. The most abundant ions, corresponding to the potential dimers, had m/z at: 807.7, 823.7, 843.7 and 829.7, 827.7, 841.7, 825.7. Ions with m/z below 813 would be expected for disitosteryl dimers with one oxygen in the molecule, whereas dimers with two, three or four oxygen atoms would produce ions with m/z at: 827–829, 843–845 or 859–861, respectively. In fact in the spectrum of the major peak from the dimer fraction were found ions most likely representing mono-, di- and tri-oxygenated dimers (Fig. 11a). The retention time of the synthesised 3,30 -sitoste-

ryl ether was at 27.3 min (Fig. 10b), the same as dimer fraction and produced mass spectrum (Fig. 11b) with the base peak at m/z 397.4. This clearly indicates that under APCI conditions disitosteryl ether was unstable producing monomer fragment ion as a base peak, along with low intensity ions typical for dimers, as shown in Fig. 10a, where TIC is superimposed with extracted chromatogram for m/z range at 780–900 (XIC). The most abundant ion within m/z range at 780–900 was observed with at m/z 883.8, which is proposed to be a THF solvent adduct of disitosteryl ether [M+THF+H]+ (Fig. 11b). We have observed similar phenomena when other solvents such as acetonitrile/dichloromethane and ace-

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with low intensity molecular ions [M+H]+ (0.5%) and [M]+ (0.3%). Struijs et al. (2010) reported that during APCI ionisation the distigmasteryl ether was unstable. Generally, it is observed that ether linkages present in thermo-oxidised lipids are thermolabile and usually breaking during APCI ionisation (Muizebelt & Nielen, 1996).

H H CH3

H H

H

11

19

H

H 10

1

8

2

H

O

9

6

4

3

3'

5

H

H

H

H

H

4. Discussion

H

7

Using NMR, IR and Raman spectroscopies, we have established that 3b,3b0 -sitosteryl ether was formed during thermo-oxidative degradation of sterol standard as the main component. We confirmed that a complex mixture of sterol dimers is formed during thermo-oxidative degradation of sterol, similarly to the reports by Rudzinska et al. (2010) and Struijs et al. (2010). Listed above spectroscopies, together with APCI/MS, further corroborated the formation of dimers with other configuration than ethers. High intensity of ions within m/z 780–900 in the dimer fraction compared to the synthesised ether further substantiated the presence of dimers bonded with more stable bond than ether bridges, particularly compounds with C–C or ester linkages (Moreau, Scott, & Haas, 2008; Muizebelt & Nielen, 1996; Rozenberg et al., 2003). The complex pattern of ions in the dimer region of APCI/MS spectrum indicates the presence of: (1) dimers without and with one or more oxygen atoms in the molecule; (2) dimers with different sterol units than b-sitosterol, since the standard used for the thermooxidation contained other phytosterols and (3) dimers adducts formed in the APCI source (Ma & Kim, 1997).

H

Fig. 4. Key 1H–1H NOESY correlations of the 3,30 -sitosteryl ether in the dimer fraction.

tonitrile/2-propanol were used, which resulted in formation of solvent derived adducts of disitosteryl ether under APCI conditions (data not shown). Formation of mobile phase adducts of lipids, in particular sterol oxides, has been reported before (Ma & Kim, 1997; Manini, Andreoli, Careri, Elviri, & Musci, 1998). Moreover, when sample of synthesised disteryl ether was analysed by FIA (flow injection analysis) we have observed molecular ion of disitosteryl ether at m/z 811.7 [M+H]+ (<0.1%) and solvent adduct at m/z 883.8 [M+THF+H]+ (0.1%) (data not shown). Similarly, Bergenthal et al. (1990) reported EI-MS spectra of disitosteryl ether with ions representing fragmentation of ether molecule at m/z 397 and 411

29'

29 28'

26'

28

24' 25'

22'

21' 20'

23'

27'

17'

18'

18 12'

13'

12 11'

16'

11 19'

15'

22

21

14'

10'

7'

5' 6'

1 2' 3'

4'

9

4

14 8

26

27

15

7

5

3

O

25

16

10

2

24 23

17

19 1'

9' 8'

13

20

6

Fig. 5. Structure of the 3b,3b0 -sitosteryl ether.

b

H1α

c H1α/1α’

H1α 1.5

H2α -OH

1.5

3.60 3.55 3.50 3.45

2.5

3.60 3.55 3.50 3.45

2.5

H3α/α’

3.35 3.30 3.25 3.20

1.0

1.5

H2α/2α’ 2.0

H4α/4α’ H3α

H1α/1α’

1.5

2.0

H4α H3α

1.0

H2α/2α’

H2α 2.0

H4α

d

2.0

δH [ppm]

a

H4α/4α’ 2.5

H3α/α’

2.5

3.35 3.30 3.25 3.20 3.15

δH [ppm] Fig. 6. Fragments of 1H–1H NOESY spectra of the b-sitosterol standard (a), the b-sitosterol standard with deuterium oxide (b), the dimer fraction (c) and the synthesised 3,30 sitosteryl ether (d).

´ ska et al. / Food Chemistry 139 (2013) 464–474 E. Sosin

a

3.5

c

b

Η4α Η4β

Η4α/4α’ Η4β/4β’

Η4α/4α’ Η4β/4β’

Η3α/3α’

Η3α

3.6

//

2.3

471

2.2

3.4

3.3

3.2

Η3α/3α’

//

2.3

2.2

3.4

3.3

3.2

//

2.3

2.2

δH [ppm] Fig. 7. Fragments of 1H NMR spectra of the b-sitosterol standard (a), the dimer fraction (b) and the synthesised 3,30 -sitosteryl ether (c).

Fig. 8. IR spectra of the b-sitosterol standard (a), the dimer fraction (b) and the synthesised 3,30 -sitosteryl ether (c).

Fig. 9. Raman spectra of the b-sitosterol standard (a), the dimer fraction (b) and the synthesised 3,30 -sitosteryl ether (c).

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27.3

Intensity [cps]

a

3.8e4

2.4e4

1.4e4

0.4e4 2

Intensity [cps]

b

6

10

14 18 22 Time [min]

26

30

34

TIC XIC 38

27.3 2.4e4

1.6e4

0.8e4

0.2e4 2

6

10

14

18 22 Time [min]

26

30

34

38

TIC XIC (x200)

Fig. 10. HPSEC-APCI/MS chromatogram of the dimer fraction (a) and the synthesised 3,30 -sitosteryl ether (b) presented as superimposed total ion current (TIC) and extracted ion chromatogram within m/z range at 780–900 (XIC).

Smith (1981) hypothesised that after conjugated steradiene (cholesta-3,5-diene) and various oxycholesterols, 3,30 -cholesteryl ether will be the most abundant products formed during oxidative degradation of cholesterol when heated at elevated temperatures. Recently, Struijs et al. (2010) suggested that distigmasteryl ether along with many other dimers was present in apolar dimer fraction, formed during stigmasterol thermo-oxidative degradation. Rudzinska et al. (2010) proposed the formation of sterol dimers through ether 3,70 -linkage, whereas Struijs et al. (2010) hypothesised formation of stigmasterol dimers mainly through direct C7– C70 bond. Both hypotheses were supported by the preferential oxidation at the C7, where 7-hydroxy- and 7-ketosterols are the most abundant oxyphytosterols formed during oxidation of phytosterols (Kemmo, Ollilainen, Lampi, & Piironen, 2008; Rudzin´ska et al., 2009). The hypothesis about direct C7–C70 linkage was based on photochemical dimerisation of ergosterol (Windaus & Borgeaud, 1928). Photodimerisation of steroids via carbons in ring B and B0 requires two unsaturation sites in ring B, as is in the ergosterol (D5,7) and 20-hydroxyecdysone (D7, 6-ketone) (Harmatha, Budsíensky, & Vokác, 2002). If this configuration is not present, it was proposed that dimers are formed by connexions between ring A and A0 , and A and D0 (DellaGreca, Iesce, Previtera, Temussi, & Zarrell, 2002). During thermo-oxidation the dimerisation via C7–C70 bond (ring B and B’) is less feasible than linkage between C3–C30 in ring A and A0 , because formation of an intermediate 7-ketosterol is required to form C7–C7’ linkage while formation of ether bond is a simple condensation process. The presence of sitosteryl dimers with carbon–carbon bonds at C3–C70 , C7–C30 or C7–C70 in analysed

non-polar dimer fraction is possible only if all hydroxyl groups in sterol molecules are exhausted. However, if one or both sitosteryl units were dehydrated (D3,6), signals should appear at 4.3, 5.6 and 5.9 ppm for 1H and at 125.9, 127.9, 131.7 and 145 ppm for 13 C in NMR spectra (Wilson et al., 1996). Formation of sterol oligomers during thermo-oxidative degradation can lower sterols’ bioavailability and thus diminish the positive effects of sterol consumption. Yet, studies under possible harmful effects of sterol oligomers have not been conducted, besides disteryl ethers. Weber, Benning, and Schulte (1988) observed no effects on feed intake, weight gain, organ weights, and abnormalities in feces or urine when mice were gavage fed at 400 mg/ kg body weight per day for 4 weeks with dicholesteryl and disitosteryl ether. Steryl ethers were found to be neither absorbed nor metabolised in the gastrointestinal tract (Weber et al., 1988). Dicholesteryl ether implanted subcutaneously into mice, remained intact with no tissues reaction and exhibited no carcinogenic properties on rats (Kaufman et al., 1970; Larsen & Barrett, 1944).

5. Conclusions Presented work proves the formation of disteryl ethers along with other dimers when phytosterols are heated up to frying temperature in the presence of oxygen for a prolonged time. In this work, for the first time the spectroscopic methods such as NMR, IR, Raman and SEC/APCI/MS spectroscopies were applied to decipher chemical dimers structure. Further investigation is required

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Fig. 11. HPSEC-APCI/MS mass spectra of the dimer fraction (a) and the synthesised 3,30 -sitosteryl ether (b) eluting at 27.3 min.

to establish the chemical structure of other oligomers formed during oxidative degradation of sterols applying high resolution mass spectrometry and more efficient ionisation to prevent degradation of dimers during ionisation. Acknowledgments This research was funded by the Alberta Value Added Corporation and the Agriculture and Food Council through the Alberta Agriculture Funding Consortium. We thank Spectral Service AG, Germany, especially Dr. Bernd W. K. Diehl, for supplementary NMR analysis and comments. We thank Dr. Felix Aladedunye for helpful discussions. References Bergenthal, D., Schulte, E., & Weber, N. (1990). Formation and characterization of 0 0 D5,6/5 ,6 -unsaturated disteryl ethers. Chemistry and Physics of Lipids, 53, 77–84. Bradford, P. G., & Awad, A. B. (2007). Phytosterols as anticancer compounds. Review. Molecular Nutrition & Food Research, 51, 161–170. Byrdwell, C. W., & Neff, W. E. (2004). Electrospray ionization MS of High M.W. TAG oligomers. Journal of the American Oil Chemists’ Society, 81, 14–26. Christopoulou, C. N., & Perkins, E. G. (1989). High performance size exclusion chromatography of monomer, dimer and trimer mixtures. Journal of the American Oil Chemists’ Society, 66, 1338–1343. de Jong, N., Ros, M. M., Ocke, M. C., & Verhagen, H. (2008). A general post launch monitoring framework for functional foods tested with the phytosterol/-stanol case. Trends in Food Science & Technology, 19, 535–545. DellaGreca, M., Iesce, M. R., Previtera, L., Temussi, F., Zarrell Mattia, C. A., et al. (2002). Solid-state photodimerization of steroid enones. Journal of Organic Chemistry, 67, 9011–9015. Gottlieb, H. E., Kotlyar, V., & Nudelmanm, A. (1997). NMR chemical shifts of common laboratory solvents as trace impurities. Journal of Organic Chemistry, 62, 7512–7515. Harmatha, J., Budsíensky, M., & Vokác, K. (2002). Photochemical transformation of 20-hydroxyecdysone: Production of monomeric and dimericecdysteroid analogues. Steroids, 67, 127–135.

Huang, L., Zhong, T., Chen, T., Ye, Z., & Chen, G. (2007). Identification of b-sitosterol, stigmasterol and ergosterin in A. roxburghii using supercritical fluid extraction followed by liquid chromatography/atmospheric pressure chemical ionization ion trap mass spectrometry. Rapid Communications in Mass Spectrometry, 21, 3024–3032. Iida, T., Tamura, T., & Matsumoto, T. (1980). Proton nuclear magnetic resonance identification and discrimination of side chain isomers of phytosterols using a lanthanide shift reagent. Journal of Lipid Research, 21, 326–338. Kaufman, H. P., Vennekel, E., & Hamza, Y. (1970). Über die veränderung der sterine in fetten and ölenbei der industriellen bearbeitungderselben I. Fette Seifen Anstrichmittel, 72, 242–246. Kemmo, S., Ollilainen, V., Lampi, A. M., & Piironen, V. (2008). Liquid chromatography mass spectrometry for plant sterol oxide determination in complex mixtures. European Food Research and Technology, 226, 1325–1334. Lampi, A. M., Kemmo, S., Mäkelä, A., Heikkinen, S., & Piironen, V. (2009). Distribution of monomeric, dimeric and polymeric products of stigmasterol during thermooxidation. European Journal of Lipid Science and Technology, 111, 1027–1034. Larsen, C. D., & Barrett, M. K. (1944). Administration of 3,5-cholestadiene and dicholesteryl ether to mice and rats. Journal of the National Cancer Institute, 4(587), 584. Lou, Y., Liu, Y., Qi, H., Wu, Z., & Zhang, G. (2006). Steryl esters and phenylethanol esters from Syringakomarowii. Steroids, 71, 700–705. Ma, Y.-C., & Kim, H.-Y. (1997). Determination of steroids by liquid chromatography/ mass spectrometry. Journal of The American Society for Mass Spectrometry, 8, 1010–1020. Manini, P., Andreoli, R., Careri, M., Elviri, L., & Musci, M. (1998). Atmospheric pressure chemical ionization liquid chromatography/mass spectrometry in cholesterol oxide determination and characterization. Rapid Communications in Mass Spectrometry, 12, 883–889. Menéndez-Carreño, M., Ansorena, D., Astiasarán, I., Piironen, V., & Lampi, A. M. (2010). Determination of non-polar and mid-polar monomeric oxidation products of stigmasterol during thermo-oxidation. Food Chemistry, 122, 277–284. Moreau, R. A., Scott, K. M., & Haas, M. J. (2008). The identification and quantification of steryl glucosides in precipitates from commercial biodiesel. Journal of the American Oil Chemists’ Society, 85, 761–770. Moreau, R. A., Whitaker, B. D., & Hicks, K. B. (2002). Phytosterols, phytostanols, and their conjugates in foods: Structural diversity, quantitative analysis, and healthpromoting uses. Progress in Lipid Research, 41, 457–500. Muizebelt, W. J., & Nielen, M. W. F. (1996). Oxidative crosslinking of unsaturated fatty acids studied with mass spectrometry. Journal of Mass Spectrometry, 31, 545–554.

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Smith, L. L. (1981). Cholesterol autoxidation. New York: Plenum Press. Struijs, K., Lampi, A. M., Ollilainen, V., & Piironen, V. (2010). Dimer formation during the thermo-oxidation of stigmasterol. European Food Research and Technology, 231, 853–863. Varady, K. A., Houweling, A. H., & Jones, P. J. (2007). Effect of plant sterols and exercise training on cholesterol absorption and synthesis in previously sedentary hypercholesterolemic subjects. Translational Research, 149, 22–30. Weber, N., Benning, H., & Schulte, E. (1988). Intestinal absorption, metabolism and nutritional effects of dietary disteryl ethers in mice. Journal of Agricultural and Food Chemistry, 36, 788–791. Westtrate, J. A., & Meijer, G. W. (1998). Plant sterol-enriched margarines and reduction of plasma total- and LDL-cholesterol concentrations in normocholesterolaemic and mildly hypercholesterolaemic subjects. European Journal of Clinical Nutrition, 52, 334–343. Wilson, W. K., Sumpter, R. M., Warren, J. J., Rogers, P. S., Ruan, B., & Schroepfer, G. J. (1996). Analysis of unsaturated C27 sterols by nuclear magnetic resonance spectroscopy. Journal of Lipid Research, 37, 1529–1555. Windaus, A., & Borgeaud, P. (1928). Über die photochemische dehydrierungdes ergosterins. Liebigs Annalen, 460, 235. Woyengo, T. A., Ramprasath, V. R., & Jones, P. J. H. (2009). Anticancer effects of phytosterols. European Journal of Clinical Nutrition, 63, 813–820.