Structural Analysis of Oxidized Cerebrosides from the Extract of Deep-Sea Sponge Aulosaccus sp.: Occurrence of Amide-Linked Allylically Oxygenated Fatty Acids

Abstract

The structural elucidation of primary and secondary peroxidation products, formed from complex lipids, is a challenge in lipid analysis. In the present study, rare minor oxidized cerebrosides, isolated from the extract of a far eastern deep-sea glass sponge, Aulosaccus sp., were analyzed as constituents of a multi-component RP-HPLC (high-performance liquid chromatography on reversed-phase column) fraction using NMR (nuclear magnetic resonance) spectroscopy, mass spectrometry, GC (gas chromatography), and chemical transformations (including hydrogenation or derivatization with dimethyl disulfide before hydrolysis). Eighteen previously unknown β-D-glucopyranosyl-(1→1)-ceramides (1a–a^//, 1b–b^//, 2a–a^//, 2b–b^//, 3c–c^//, 3d–d^//) were shown to contain phytosphingosine-type backbones (2S,3S,4R,11Z)-2-aminoeicos-11-ene-1,3,4-triol (in 1), (2S,3S,4R,13Z)-2-aminoeicos-13-ene-1,3,4-triol (in 2), and (13S*,14R*)-2-amino-13,14-methylene-eicosane-1,3,4-triol (in 3). These backbones were N-acylated with straight-chain monoenoic (2R)-2-hydroxy acids that had allylic hydroperoxy/hydroxy/keto groups on C-17^/ in the 15^/E-23:1 chain (a–a^//), C-16^/ in the 17^/E-23:1 (b–b^//) and 14^/E-22:1 (c–c^//) chains, and C-15^/ in the 16^/E-22:1 chain (d–d^//). Utilizing complementary instrumental and chemical methods allowed for the first detailed structural analysis of a complex mixture of glycosphingolipids, containing allylically oxygenated monoenoic acyl chains.

1. Introduction

Lipid hydroperoxides are labile compounds derived from lipids containing carbon-carbon double bonds. The formation of these primary peroxidation products occurs in enzymatic and non-enzymatic (autooxidation, photo-oxidation) reactions [1]. Lipid hydroperoxides, formed in biological systems, have not only multiple damaging effects on cellular macromolecules, but are also important regulators of many cellular processes [2]. In some pathological situations, lipid hydroperoxides are generated at higher than normal rates. Such overproduction is implicated in several human diseases and exposures including atherosclerosis, cancer, diabetes, acute lung injury, chronic alcohol exposure, and neurodegenerative disorders. The complex nature of lipid peroxidation and its potential biological significance have attracted the attention of scientists across many disciplinary fields, ranging from chemistry and biochemistry to biology and clinical science (see review [3] and references cited herein).

Recent progress in the characterization and quantification of oxygenated fatty acids (FAs) has facilitated a better understanding of lipid oxidation, but the methods currently available still suffer from unresolved sensitivity and selectivity problems [4]. The main reason for these problems is the complexity of the product profile formed by oxidation from a single lipid molecular species. In particular, peroxidation of any unsaturated fatty acyl group may generate a mixture of allylic hydroperoxides with different double bond positions and/or configurations. These hydroperoxides are easily decomposed or go through further reactions to form different secondary peroxidation products including epoxides, allylic alcohols, α,β-unsaturated ketones (enones), and chain-cleavage products [1]. The constituents of these mixtures are difficult to separate, isolate, and identify. Additionally, natural extracts are characterized, as a rule, by extremely low levels of oxidized lipids, causing further difficulties in their isolation and analysis [3,4]. Undoubtedly, structural elucidation of primary and secondary peroxidation products, formed from complex lipids, remains a challenge for lipid analysts.

Monoenes are much less prone to undergo oxidation than polyenes. Respectively, direct addition of peroxyl radicals to monounsaturated lipids is not generally observed with the exceptions of cholesterol [3] and related sterols. In particular, sphingolipids, containing mainly saturated or monounsaturated hydrocarbon chains, are poor substrates for peroxidation. We found only one report on the isolation of peroxidized sphingolipids from a natural source, but these compounds formed due to oxidation of unique sphingoid base moiety with conjugated double bonds. Namely, some stereoisomeric glycosphingolipids, containing endoperoxide and allylic hydroperoxide functions in their dienoic sphingoid base moieties, were isolated from the extract of the sponge Axinella corrugate [5]. The locations of the double bonds and peroxide-containing groups were determined by ¹H,¹H-COSY (proton-proton correlation spectroscopy), НМВС (heteronuclear multiple-bond correlation spectroscopy), and ROESY (rotating-frame nuclear Overhauser effect spectroscopy) experiments due to the proximity of the previously mentioned functionalities to the polar portion of the sphingolipids. Methanolysis (MeOH/HCl) of the peroxidized glycosphingolipids was used to release methyl esters of saturated FAs, but oxidized dienoic sphingoid bases were not obtained, apparently, due to rapid decomposition under acidic conditions.

In mass spectrometric (MS) studies of oxidative stress markers (disease biomarkers), oxidation of some standard glycosphingolipids, containing monoenoic FAs, was induced by a Fenton reaction (H₂O₂/FeCl₂) or UVA (ultraviolet A) irradiation [6,7]. In particular, Couto et al. obtained galactosylceramides with allylic hydroperoxy, hydroxy, or keto groups in FA chains. These cerebrosides were characterized using ESI-MS (electrospray ionization mass spectrometry) and HPLC-MS (high-performance liquid chromatography-mass spectrometry) methods (including tandem mass spectrometry (MS/MS)) [6]. In our continued study on lipids from far eastern marine sponges [8], similar oxidized cerebrosides (Figure 1), presumably derived from glucosylceramides with monounsaturated fatty acyl groups, were found in the extract of a deep-sea glass sponge, Aulosaccus sp. As seen in Figure 1, the depicted compounds contain an allylic hydroperoxy, hydroxy, or keto group in each acyl chain, but their monoenoic sphingoid base moieties are not oxidized. A fraction of these minor cerebrosides, along with fractions of non-oxidized cerebrosides [8], was isolated from the sponge extract using high-performance liquid chromatography on reversed-phase column (RP-HPLC). The present study presents structural elucidation of rare oxidized glycosphingolipids using mass spectrometry (ESI-MS and gas-chromatography-mass spectrometry (GC-MS)), ¹Н-, ¹³С-NMR (nuclear magnetic resonance) spectroscopy, GC, chemical transformations, and optical rotation measurement.

A classical approach to the analysis of any complex lipid includes methanolysis or hydrolysis, which is followed by analyses of liberated derivatized simple lipids and sugar. However, prolonged high temperatures and treatments with acidic or alkaline solutions, required for solvolysis of N-acyl lipids, presents significant problems with respect to the potentially labile allylic oxygen-containing groups of the acyl chains. To solve these problems, we used catalytic hydrogenation to fix the starting positions of the oxygenated groups prior to chemical degradation of the oxidized glycosphingolipids. In addition, attempts were made to fix the double bonds of allylic substructures by reacting with dimethyl disulfide (DMDS) because, in our preliminary research, the DMDS adduct of methyl oleate did not lose S-methyl groups during hydrolysis with HCl in MeCN-H₂O. To minimize possible allylic rearrangements (1,3-isomerizations) and other alterations, we avoided elevated temperatures and strong acid/base conditions in the derivatization reactions before hydrolysis. Thus, our attention was mainly focused on procedures suitable for an initial detailed structural analysis of a complex mixture of glycosphingolipids, containing an allylic hydroperoxy, hydroxy, or keto group in the monoenoic acyl chain.

2. Results and Discussion

Isomeric allylic hydroperoxides 1a, 1b, 2a, 2b, 3c, 3d, related isomeric allylic alcohols 1a^/, 1b^/, 2a^/, 2b^/, 3c^/, 3d^/, and isomeric enones 1a^//, 1b^//, 2a^//, 2b^//, 3c^//, 3d^// (Figure 1) were constituents of a single RP-HPLC fraction. In the beginning of this study, the fraction contained 46% allylic hydroperoxides, 43% allylic alcohols, and 11% enones, calculated using relative intensities of the characteristic, isolated ¹Н-NMR signals of trans-olefinic protons (see below). The final results of this study yielded 25%, 54%, and 21% of these allylically oxygenated compounds, respectively. Therefore, further transformation of allylic hydroperoxides to allylic alcohols and enones occurred upon analysis (registrations of NMR and MS spectra, preparation of samples, storage, etc.).

In UPLC-MS (ultra-performance liquid chromatography–mass spectrometry) analysis of oxidized cerebrosides, base peak chromatogram, and extracted-ion chromatograms provided limited information because many components eluted simultaneously. In particular, peaks for major isomeric allylic hydroperoxides 1a and 1b were not well-resolved (Supplementary Materials, Figure S1a,b).

2.1. Positive and Negative Ion Mode ESI-MS/MS Analyses of Oxidized Cerebrosides

The molecular formulae, С₄₉H₉₃NO₁₂ for allylic hydroperoxides (1a, 1b, 2a, 2b, 3c, 3d), С₄₉H₉₃NO₁₁ for allylic alcohols (1a^/, 1b^/, 2a^/, 2b^/, 3c^/, 3d^/), and С₄₉H₉₁NO₁₁ for enones (1a^//, 1b^//, 2a^//, 2b^//, 3c^//, 3d^//), were determined by HR-ESI-MS (high resolution ESI-MS) analyses in positive-((+)ESI-MS) and negative-((–)ESI-MS) ion modes. Complementary (+)ESI- and (–)ESI-MS/MS analyses of these glucosylceramides, containing 2-hydroxy acyl chains and phytosphingosine-type backbones, resulted in a series of fragment ions, as shown in Scheme 1.

Many of the fragment ions, shown in Scheme 1, have also been detected in our ESI-MS/MS studies of non-oxidized cerebrosides isolated from Aulosaccus sp. Namely, (+)ESI-MS/MS spectra of sodium adducts from non-oxidized glycosphingolipids have been characterized by prominent peaks, corresponding to [M + Na]⁺ (base peak), Y₀, Z₀, O, and C₁ ions, and by small peaks, representing [M + Na − H₂O]⁺, E, and B₁ ions. However, the (+)ESI-MS/MS spectrum of the [M + Na]⁺ ion of isomeric hydroperoxy cerebrosides (Figure 2) showed other relative abundances for a variety of these ions. In particular, the O ions of m/z 528.35, representing isomeric monoglucosylated monounsaturated С₂₀ sphingoid base backbones 1 and 2, constituted the base peak of this spectrum. The spectrum also exhibited a homologous, less abundant O^/ ion (m/z 542.36), containing a monoglucosylated cyclopropane С₂₁ sphingoid base backbone 3. A relatively low intensity pseudo-molecular ion peak (m/z 910.65, [M + Na]⁺) and very small peaks, corresponding to Y₀ (m/z 748.60) and Z₀ (m/z 730.58) ions (not shown), were observed. The presence of the [M + Na − Н]⁺ peak, comparable with the pseudo-molecular ion peak of the hydroperoxides, was explained by hydrogen atom abstraction, followed by electronic delocalization in the resulting radical, which might yield rearranged products. At the same time, the (+)ESI-MS/MS spectrum revealed more abundant [M + Na − 102]⁺ (m/z 808.55), [M + Na − 88]⁺ (m/z 822.56), [Y₀ − 102] (m/z 646.505), [Y₀ − 88] (m/z 660.52), [Z₀ − 102] (m/z 628.49), and [Z₀ − 88] (m/z 642.50) ions. These fragments and fragment [W − 102] (m/z 275.20), each with a terminal α,β-unsaturated aldehyde, were thought to arise from specific α-cleavages of lipid hydroperoxides (Figure 2). Related fragment ions, formed by C_nH_2n+2O losses from [M + Na]⁺ ions, were also found in the MS/MS studies of some monoenoic [6] and polyenoic [11,12,13] FA moieties or free FAs, in which an allylic hydroperoxy group was between double bond(s) and a terminal methyl group. In general, the [M + Na − C_nH_2n+2O]⁺ ions were also more abundant in MS/MS spectra of those compounds compared to their [M + Na]⁺ ions.

Ions, possibly formed by Hock cleavage [14,15], were insignificant in our (+)ESI-MS/MS analysis of allylic hydroperoxides. These fragments included m/z 796.55 (for 1b, 2b, and 3d) and 782.54 (for the allylic isomers of 1a, 2a, and 3c) ions, as illustrated in Scheme S1 (Supplementary Materials) and Figure 2 (allylic rearrangements for acyl chains a and c are not shown). In contrast, the relatively abundant ions, presumably formed by analogous cleavage from hemiacetal derivatives, were reported for MS/MS fragmentations of [M + Na]⁺ precursors of some free polyenoic FAs in which an allylic hydroperoxy group was between the double bond system and C-1 [12,13].

Favorable cleavage of the isomers, formed by allylic rearrangements of acyl chains b and d (Figure 2), occurred, losing an 88 Da fragment. At the same time, compounds with parent acyl chains b and d underwent other favorable fragmentations, yielding a distinct group of homologous ions (from m/z 668.44 to 738.50), containing the most significant fragment of m/z 724.49. We suggest that the occurrence of these ions may be connected with homolysis of a weak RO-̸-OH bond, formation of an alkoxyl radical, and subsequent formation of a radical centered on a remote and non-activated carbon atom of a saturated hydrocarbon chain. This may lead to fast cyclization that, in turn, leads to MS fragmentations of the resulting cyclic ethers, shown in Scheme 2. The proposed cyclization reaction is reminiscent of the formation of cyclic (mainly, five-membered) ethers from acyclic saturated monohydroxy alcohols, occurring via radical (primarily alkoxyl) intermediates under appropriate chemical and thermal conditions (for reviews, see References [16,17]). In this process, secondary aliphatic alcohols yielded 2,5-dialkyltetrahydrofurans [18]. A possible mechanism for the formation of such products includes 1,5-transposition of the radical center from the oxygen atom of the alkoxyl radical to δ-carbon atom, involving a 1,5-hydrogen transfer through a chair-like six-membered transition state. For a linear hydrocarbon chain with an initial alkoxyl radical, the 1,5-hydrogen atom transfer may be considered the most common reaction, even though intramolecular abstractions of the hydrogen atom from other positions (1,4-migrations, 1,6-migrations, and 1,7-migrations, etc.) may be observed [17]. Similar processes may have occurred in our MS/MS experiment because alkyl hydroperoxides are known to form alkoxyl radicals by thermal or photolytic decomposition [17,19]. Apparently, the ability to undergo favorable cyclic transition states affected the fragmentation process, leading to the formation of major homologous ions, as illustrated in Scheme 2.

Ion intensity profiles, obtained for cerebrosides with allylic hydroxy or keto groups, were, in general, similar to those of non-oxidized cerebrosides found in Aulosaccus sp. In particular, the MS/MS spectra of sodiated molecular ions of allylic alcohols (Figure 3a: m/z 894.66 [M + Na]⁺) and enones (Figure 3b: m/z 892.64 [M + Na]⁺) showed predominant peaks of pseudo-molecular ions, along with peaks of lower intensities, which represented [M + Na − H₂O]⁺, Y₀, Z₀, and O ions. However, unlike [M + Na]⁺ ions of non-oxidized cerebrosides, the sodium adducts of allylic alcohols and enones fragmented to give discernible ions of W-type and minor U and T ions. Additionally, the acyl-containing ions of allylic alcohols had a tendency to lose one, or even two, hydrogen atoms. In this case, a trend was observed toward the increased loss of hydrogen atoms with decreasing ion masses. For example, a significant difference was noted between the relative intensities of [M + Na]⁺ and [M + Na − H]⁺ peaks ([M + Na − 2H]⁺ ions were not even detected), but the intensities of W, [W − H], and [W − 2H] peaks were comparable (Figure 3a).

In (−)ESI-MS/MS experiments with [M − H]⁻ and [M + Cl]⁻ ions (Figure 4a–c), allylic hydroperoxides again produced more fragments than allylic alcohols and enones. The main feature of (−)ESI-MS/MS fragmentation of [M − H]⁻, Y₀, Z₀, Z₀/Q, T, W, and other precursor ions, containing an allylic hydroperoxy group, was the loss of water to produce fragments with enone functionality in acyl chains, as described for [M − H]⁻ ions of hydroperoxy-eicosatetraenoic acids [20]. In particular, the homologous ions of m/z 436.4 ([Z₀/Q − H₂O]) and 422.4 ([Z₀^//Q − H₂O]), containing acyl C₂₃ and С₂₂ chains, respectively (Figure 4a), were also observed in (−)ESI-MS/MS spectrum of enones (Figure 4c). Then, relatively low-intensity pseudo-molecular ion peaks ([M + Cl]⁻ and [M − H]⁻) and very small peaks, corresponding to Y₀ and Z₀ ions (not shown), were seen in the (−)ESI-MS/MS spectrum of allylic hydroperoxides. A discernible [M − 2H] peak was comparable with a pseudo-molecular [M − H] peak. This spectrum also exhibited [M + Cl − 102]⁻, [M + Cl − 88]⁻, [M − H − 102]⁻, [M − H − 88]⁻, and [Z₀/Q − 102] ions, interpreted as α-cleavage ions with a terminal α,β-unsaturated aldehyde. Additionally, the two minor ions of m/z 743.6 and 729.55 could be formed by α-cleavages of compounds in which an allylic hydroperoxy group was between a double bond and C-1^/. In particular, the m/z 743.6 ions could be fragments of isomeric compounds 1b, 2b (C-15^/−C-16^/ bond fission), and 3d (C-14^/−C-15^/ bond fission), while the less abundant m/z 729.55 ions could be fragments of the allylic isomers of compounds 1a, 2a (C-14^/−C-15^/ bond fission), and 3c (C-13^/−C-14^/ bond fission).

Like the (−)ESI-MS/MS spectra of the non-oxidized cerebrosides of Aulosaccus sp., those of allylic alcohols (Figure 4b) and enones (Figure 4c) exhibited significant peaks corresponding to [M − H]⁻ and Z₀/Q ions, with lower intensity peaks representing [M + Cl]⁻, Y₀, Z₀, [Z₀/Q − C₃H₅N], and W ions. The negatively charged acyl-containing ions of allylic alcohols (Figure 4b), like their positively charged acyl-containing counterparts (Figure 3a), tended to lose hydrogen atoms in the MS/MS experiment.

2.2. NMR Characterization of Oxidized Cerebrosides

The ¹Н-NMR and ¹³С-NMR spectra (CD₃OD) of oxidized cerebrosides (

Table 1. ¹Н-(500 MHz) and ¹³С-(125 MHz) NMR data (CD₃OD) ¹ for the RP-HPLC fraction, containing compounds 1a–a^//, 1b–b^//, 2a–a^//, 2b–b^//, 3c–c^//, and 3d–d^//.

Position	δH (mult., J Hz)	δC
Sphingoid base moieties
1	3.80 (dd, 3.9, 10.5), 4.045 (dd, 6.2, 10.5)	70.6
2	4.25 (m)	52.3
3	3.60 (t, 6.0)	76.1
4	3.51 (m)	73.6
5	1.39 (m), 1.64 (m)	33.7
6	1.31 (m), 1.55 (m)	27.6
СН2-pool	1.22−1.42 (m)	30.6–32.0
Allylic СН2 of 1 and 2	2.02 (m)	28.7–28.85
Olefinic СН of 1 and 2	5.34 (m)	131.4–131.5
12, 15 of 3	1.17 (m), 1.37 (m)	30.5
13, 14 of 3	0.67 (m)	17.5
21 of 3	–0.33 (dt, 4.1, 5.3), 0.58 (ddd, 4.1, 8.3, 8.3)	12.2
18	1.28 (m)	33.5
19	1.31 (m)	24.3
20 of 1 and 2 20 of 3	0.895 (t, 6.9) 0.899 (t, 7.0) 2	15.0 15.0
N-Acyl moieties (some general features 3)
1/	−	177.7
2/	4.01 (dd, 4.0, 7.7)	73.6
3/	1.60 (m), 1.74 (m)	36.4
4/	1.42 (m)	26.7
СН2-pool	1.22−1.42 (m)	30.6–32.0
(n−2) CH2	1.28 (m)	33.7
(n−1) CH2	1.32 (m)	24.2
CH3	0.893 and 0.897 (t, 7.0) 2	15.05
β-Glucopyranoside moiety
1//	4.28 (d, 7.8)	105.3
2//	3.17 (dd, 7.8, 9.1)	75.6
3//	3.35 (m)	78.5
4//	3.27 (m)	72.2
5//	3.27 (m)	78.6
6//	3.66 (m), 3.865 (dd, 1.1, 11.9)	63.3

¹ All signals were assigned based on data of ¹Н−¹Н COSY, HSQC, and НМВС experiments. ² The two resolved triplets were observed in 700-MHz spectra. ³ δ_H and δ_C values for acyl fragments with allylic –OOН/–OН/=O are given separately (Figure 5).

, Figures S2 and S3), as well as the corresponding spectra of the non-oxidized cerebrosides of Aulosaccus sp. [8], showed signals of β-glucopyranosyl-(1→1)-ceramides that had monoenoic or cyclopropane-containing phytosphingosine-type backbones, N-acylated with 2-hydroxy FAs. In particular, H-2 (δ_Н 4.25, m) of the sphingoid base moieties displayed a characteristic cross signal with N-acyl C-1^/ (δ_C 177.7), as demonstrated by an НMBC experiment with sphingolipids 1–3. ¹Н,¹Н-COSY diagram indicated that several protons, starting from −O−CH₂− (δ_Н 3.80 and 4.045, dd, CH₂-1) and ending with alkyl −CH₂− (δ_Н 1.31 and 1.55, m, CH₂-6), formed a linear spin system of phytosphingosine-type moieties of 1–3. Another spin system consisted of CH-2^/ (δ_Н 4.01, dd), CH₂-3^/ (δ_Н 1.60 and 1.74, m), and CH₂-4^/ (δ_Н 1.42, m) protons of 2-hydroxy acyl chains. The signals of a β-glucopyranoside moiety were sequentially assigned by ¹Н,¹Н-COSY, HSQC (heteronuclear single-quantum correlation spectroscopy), and НМВС experiments, starting from the signal of anomeric CH-1^// (δ_Н 4.28, d, J = 7.8 Hz; δ_C 105.3). Accordingly, cross signals CН₂-1/С-1^// and CН-1^///С-1 were observed in the НМВС diagram of glycosides 1–3. Then, the ¹Н- and ¹³С-NMR spectra of these sphingolipids showed signals of long hydrocarbon chains (−(CH₂)_n−, δ_Н 1.22–1.42, m, δ_С 30.6–32.0) and terminal methyl groups (δ_Н 0.89–0.90, several overlapping triplets, δ_С 15.0–15.05, broad signal). The δ values of allylic CH₂ (δ_C 28.7–28.85, δ_Н 2.02, m) and olefinic CH (δ_Н 5.34, m) were used to characterize cis-double bonds of backbones 1 and 2. These data were in good agreement with ¹Н-NMR and ¹³С-NMR (CD₃OD) data on some cerebrosides [21] and FA standards [22] with isolated cis-double bonds (cis-isomers: δ_C 28.7–28.9, δ_Н 2.02–2.03 m, allylic CH₂, and δ_H 5.335–5.34 m, olefinic CH, trans-isomers: δ_C 34.15–34.2, δ_Н 1.97–1.975 m, allylic CH₂, and δ_Н 5.37–5.38 m, olefinic CH). A cis-cyclopropane ring in backbone 3 caused three upfield shifted signals at δ_Н −0.33 (dt, J = 4.1, 5.3 Hz, H-21a), 0.58 (ddd, J = 4.1, 8.3, 8.3 Hz, H-21b), and 0.67 (m, H-13, H-14).

Apart from the previously mentioned NMR resonances, the signals of trans-monoenoic acyl moieties with an allylic hydroperoxy, hydroxy, or keto group (Figure 5a–c) were observed in the NMR spectra of the RP-HPLC fraction, containing oxidized cerebrosides. The δ_H values (CD₃OD) of the signals for –HC(OOH)– (4.16, dt), –HC(OH)– (3.94, q), and −CH=CH–CO– (6.105, d) were close to δ_H values (CDCl₃) of signals of corresponding protons in the trans-monoenoic allylic hydroperoxides (4.2, q), allylic alcohols (4.0, q), and enones (6.1, d), respectively, prepared from oleic acid [23]. Deviations of δ_H values (this study) from those in other experiments [23] arose from solvent effects. According to ¹Н,¹Н-COSY correlations, a spin system, assigned to acyl chains a–d (Figure 5a), included protons of allylic hydroperoxide, especially two olefinic CH (δ_Н 5.35 and 5.665, m) and allylic CH (δ_Н 4.16, dt, J = 6.6, 7.0 Hz), bearing −OOH. Allylic CH (δ_Н 3.94, q, J = 6.8 Hz), bearing –OH, and two olefinic CH (δ_Н 5.395 and 5.59, m) groups, belonging to allylic alcohol, were part of a spin system within acyl chains a^/–d^/ (Figure 5b). In experiments, involving selective irradiation of allylic protons, the coupling constants J = 15.6 Hz and J = 15.4 Hz for the trans-olefinic protons of the allylic hydroperoxides and allylic alcohols, respectively, were detected. The characteristic NMR resonances of two trans-alkenyl CH (δ_Н 6.105, d, J = 15.9 Hz, and 6.91, dt, J = 7.0, 15.9 Hz), conjugated C=O (δ_С 204.3), and α-CH₂ (δ_Н 2.565, t, J = 7.4 Hz) groups were used for determining substructures of enones in acyl chains a^//–d^// (Figure 5c). The structures of the allylic hydroperoxides, allylic alcohols, and enones were confirmed by HMBC correlations, as depicted in Figure 5.

Some very weak signals in the ¹Н-NMR, ¹Н,¹Н-COSY, and HSQC spectra of oxidized cerebrosides found in the present study were attributed to cis-double bonds of allylic hydroperoxides and allylic alcohols (Appendix A, Figure A1). The complete structures of these compounds could not be elucidated due to their trace amounts.

2.3. Analyses of FAs, Sphingoid Bases, and Sugar Obtained from Oxidized Cerebrosides

The RP-HPLC oxidized cerebroside fraction was divided into two parts (parts 1 and 2), which were treated using different chemical procedures before hydrolysis. Then, we applied MeCN/HCl hydrolysis [24] for chemical degradation of cerebrosides. In our experience [8], this procedure causes less disruption of spingoid bases than methanolysis (MeOН/HCl), which is most widely used in studies of complex lipids.

Analysis of FAs from Part 1. Part 1 of the oxidized cerebrosides was subjected to hydrogenation (with Adams’ catalyst) to fix the positions of allylic oxygen-containing groups before hydrolysis. Upon hydrolysis, liberated FAs were acetylated and methylated. The ¹Н-NMR spectrum (CDCl₃) of FA derivatives showed proton signals of mid-chain substructures, including −H₂C−CO−CH₂− at δ_Н 2.38 (t, J = 7.4 Hz) and −СН(OAс)– at δ_H 4.855 (m). GC-MS analysis (electron impact ionization) of these derivatives revealed methyl esters of 2-acetyloxy C₂₃ and C₂₂ acids, containing an isolated keto or acetyloxy group or no additional oxygenated group. Similar products, namely keto, hydroxy, and non-oxygenated acid derivatives, have been previously reported for the hydrogenation (in EtOH over Adams’ catalyst) of allylic 9-hydroperoxides and 10-hydroperoxides, obtained from methyl oleate [25].

In the present report, mass spectra exhibited base peaks at m/z 339 and 325 ([M − MeOCO − CH₂CO]⁺) for methyl esters of 2-acetyloxy keto C₂₃ (440 [M]⁺) and C₂₂ (426 [M]⁺) acids, respectively (Figures S4–S7). Ions produced by cleavage β to a keto group and ions formed from the methyl end of the molecules by cleavage α to the keto group were prominent in the mass spectra, as described for MS fragmentations of several methyl esters of oxo (keto) FAs [26]. In particular, mass spectra of methyl esters of C₂₃ and C₂₂ acids displayed homologous pairs of m/z 127/142 and 113/128 fragments, containing methyl ends of the molecules with keto groups on the (n–8) or (n–7) carbon, respectively. These keto group positions were confirmed by a number of ions containing the polar end of the FA esters that also produced daughter ions due to loss of AcOH, CH₂CO, or MeOH (Figures S4–S7).

Hydrogenation of part 1, followed by hydrolysis, acetylation, and methylation, also yielded the methyl esters of 2-acetyloxy C₂₃ and C₂₂ acids, containing an additional isolated acetyloxy group. Expectedly, [M]⁺ peaks were absent in the mass spectra of the methyl esters of these diacetylated acids, but [M − CH₃CO]⁺ and [M − AcOH]⁺ ions were observed in high mass regions (Figures S8–S11). Additionally, these compounds, as derivatives of 2-acetyloxy FAs, fragmented to give abundant [M − MeOCO − AcOH]⁺ ions of m/z 365 and 351 for methyl esters of C₂₃ and C₂₂ acids, respectively. Isomers, containing an isolated acetyloxy group in different positions, were discerned based on the presence of diagnostic α-cleavage ions and more abundant product ions, formed by elimination of CH₂CO or AcOH from α-fragments, as described for acetates of secondary alcohols [27]. In particular, the α-fragments included the ions of m/z 385 and 399 (Figures S8 and S9) for the derivatives of 2,16-, and 2,17-diacetyloxy C₂₃ acids, respectively, and the ions of m/z 371 and 385 (Figures S10 and S11) for the 2,15- and 2,16-diacetyloxy C₂₂ acid derivatives, respectively. Moreover, some homologous ions, which arose from C-1–⁄–C-2 bond fission and cleavage α to an isolated acetyloxy group, were specific for the different positions of this group in acyl chains. For example, the abundant ions of m/z 283 and 297 (Figures S8 and S9), which were detected in the mass spectra of the methyl esters of isomeric 2-acetyloxy C₂₃ acids, confirmed the presence of second acetyloxy groups on C-16 and C-17, respectively. Similarly, m/z 269 and 283 ions in the mass spectra of the methyl esters of 2-acetyloxy C₂₂ acids (Figures S10 and S11) indicated a second acetoxy group on C-15 and C-16, respectively. Additionally, α-fragmentation of isomers with an acetyloxy group in the (n–8) or (n–7) positions and subsequent loss of AcOH gave rise to m/z 111 and 97 ions, respectively, containing the methyl end of acyl chains.

As a result of the FA analyses, hydrogenated minor derivatives of allylically oxygenated FAs were also detected. While the major derivatives formed from fatty acyl groups containing hydroperoxy/hydroxy/keto groups in the (n–8) or (n–7) positions, the minor derivatives were rearranged products of these FA moieties (Appendix B, Figures S12–S20).

Analysis of FAs from Part 2. In this investigation, we tried to use S-methyl groups as markers for oxidized cerebroside double bonds. However, enones with polarized double bonds that did not react with DMDS under mild conditions, and labile allylic hydroperoxides were not suitable for this purpose. Therefore, enones and allylic hydroperoxides were converted into allylic alcohols that were expected to add to DMDS.

The oxidized cerebrosides from part 2 were acetylated to increase solubility of these relatively polar compounds in DMDS and other low-polarity or non-polar organic solvents. In this process, allylic hydroperoxides were transformed into enones (Scheme 3), as reported by Porter and Wujek [23]. The mixture obtained after acetylation was treated with NaBH₄/CeCl₃ [28] to convert enones into allylic alcohols. The resulting derivatives, containing an allylic hydroxy or acetyloxy group, were treated with DMDS, and the products of this reaction were hydrolyzed in MeCN/HCl. Liberated 2-hydroxy FAs were acetylated, methylated, and analyzed by a GC-MS method that revealed major mono(methylthio) and minor tris(methylthio) derivatives (Scheme 3 and Scheme 4). The mono(methylthio) compounds, containing an allylic methylthio group, were characterized by ¹H-NMR resonances (CDCl₃) of two trans-olefinic CH (δ_Н 5.175 dd, J = 9.0, 15.2 Hz, and 5.40 dt, J = 6.9, 15.2 Hz) and one CH (δ_Н 2.99 m), bearing a –SMe group (δ_H 1.97 s). The presence of the mid-chain allylic substructure was confirmed by a TOCSY (total correlation spectroscopy) experiment.

Cleavage patterns for methyl esters of 2-acetyloxy С₂₃ (470 [M]⁺) and С₂₂ (456 [M]⁺) acids, containing an allylic methylthio group, are depicted in Scheme 4. The positional isomers of these allylic thioethers were only partially GC-separated. The mass spectra of isomeric allylic thioethers (Figures S21–S28) were characterized by diagnostic peaks, corresponding to ions produced by α-cleavage. The fragments, formed by cleavage α to the carbon carrying an allylic methylthio group on the side remote from the carboxyl group included m/z 385 and 399 ions for the methyl esters of 2-acetyloxy C₂₃ acids with –SMe group in the (n–7) and (n–6) positions, respectively (Figures S22 and S24). The peaks at m/z 371 and 385, observed in the mass spectra of methyl esters of homologous 2-acetyloxy C₂₂ acids, represented fragments of the same origin (Figures S26 and S28). α-fragmentation of other mono(methylthio) derivatives gave rise to the ions at m/z 171 and 157, containing the methyl end of isomers with an allylic –SMe group in the (n–9) and (n–8) positions, respectively (Figures S21, S23, S25, and S27). Relative abundances of α-fragments in average mass spectrum were used to quantify isomer distribution, and approximately equal amounts of the four isomeric allylic thioethers were found. This finding may reflect the fact that these S-methyl derivatives were possibly products of allylic rearrangements, occurring prior to GC-MS analysis.

The minor tris(methylthio) derivatives of the methyl esters of 2-acetyloxy C₂₃ (564 [M]⁺) or C₂₂ (550 [M]⁺) acids produced more diagnostic fragments than the previously mentioned major mono(methylthio) derivatives. Expectedly, the cleavages of minor S-methylated compounds occurred between methylthio-carrying carbons to yield substantial fragment ions, as illustrated in Scheme 4 and Figures S29–S32. A cluster of four GC peaks for isomeric tris(methylthio) derivatives was observed. According to fragmentation patterns, there were two peaks representing positional isomers and two peaks that represented stereoisomers of these regioisomers on the chromatogram.

The results of the transformations of allylic alcohols and their acetates into methylthio derivatives (Scheme 3) were confirmed by experiments with model compounds, methyl esters of 11-hydroxy and 8-acetyloxy elaidic acids (prepared from methyl oleate, Appendix C, Scheme A1, Figures S33–S37). Under the conditions used here, the allylic alcohol acetates reacted with DMDS to give major allylic thioethers and minor tris(methylthio) derivatives. Allylic alcohols reacted with DMDS to give major DMDS adducts and minor allylic thioethers and tris(methylthio) derivatives. However, in contrast to the DMDS adducts of monoenes with an isolated double bond, the bis(methilthio) derivatives of allylic alcohols were destroyed during MeCN/HCl hydrolysis.

Previously, the synthesis of allylic thioethers from allylic alcohols and thiols was reported by Zhang et al. ([29]: iodine-catalyzed process) and Tabarelli et al. ([30]: catalyst-free approach). Regio-isomeric mixtures of allylic thioethers were produced when the allylic alcohol contained two different substituents. To explain the presence of regio-isomer products of 1,3-isomerization, the allylic cation, formed by water loss from the allylic alcohol, was proposed to be an intermediate in the reaction pathway [30]. The existence of a similar mechanism explains the formation of regio-isomeric allylic thioethers and their tris(methilthio) derivatives in the iodine-catalyzed reaction of allylic alcohols and their acetates with DMDS reported here. Additionally, allylic thioethers, which could undergo 1,3-isomerization under acidic conditions, were possibly formed from the bis-DMDS adducts of allylic alcohols during MeCN/HCl hydrolysis. As a result of these rearrangements, the FA methylthio derivatives, obtained from oxidized cerebrosides, gave characteristic mass spectra, permitting locations of three-carbon allylically oxygenated substructures, rather than double bonds in the starting acyl chains.

We clarified the acyl structures (Figure 1) using a logical approach. The GC-MS analyses of the methyl esters of 2-acetyloxy methylthio FAs (Scheme 4) revealed that the three-carbon allylically functionalized substructures of oxidized cerebrosides included С-15^/–С-16^/–С-17^/ and С-16^/–С-17^/–С-18^/ fragments for 2-hydroxy С₂₃ acyl chains and С-14^/–С-15^/–С-16^/ and С-15^/–С-16^/–С-17^/ fragments for 2-hydroxy С₂₂ acyl chains. According to the GC-MS analyses of the hydrogenated FA derivatives, the amide-linked FAs of oxidized cerebrosides contained hydroperoxy, hydroxy, or keto groups in the (n–8) or (n–7) positions, more specifically in the 16^/ and 17^/ positions of 2-hydroxy С₂₃ acyl chains and the 15^/ and 16^/ positions of 2-hydroxy С₂₂ acyl chains. A priori, an allylic hydroperoxy, hydroxy, or keto group should be located in the terminal points of the previously mentioned three-carbon substructures. Consequently, the С-15^/–С-16^/–С-17^/ fragments of the С₂₃ acyl chains (a–a^//) contained such groups in position С-17^/ and the double bond between С-15^/ and C-16^/, while the С-16^/–С-17^/–С-18^/ fragments of the other C₂₃ acyl chains (b–b^//) had 16^/-hydroperoxy/hydroxy/oxo groups and 17^/,18^/-double bonds. Similarly, the С-14^/–С-15^/–С-16^/ fragments of the С₂₂ acyl chains (c–c^//) contained 16^/-hydroperoxy/hydroxy/oxo groups and 14^/,15^/-double bonds, while the С-15^/–С-16^/–С-17^/ fragments of the other C₂₂ acyl chains (d–d^//) had 15^/-hydroperoxy/hydroxy/oxo groups and double bonds between C-16^/ and C-17^/.

The mono(methylthio) and tris(methylthio) derivatives of allylically oxygenated FAs were also used to explain other structural peculiarities of the starting material. Upon hydrogenation, all the methylthio derivatives lost S-methyl groups, giving saturated hydrocarbon chains. In particular, the transformations, shown in Scheme 3, with subsequent hydrodesulfurization allowed us to convert –CH=CH–CH(OOH/OH)– and –CH=CH–CO– substructures to –СН₂–СН₂–СН₂– chains without reducing other oxygen-containing groups in the molecules. As a result, the unbranched structures of the parent amide-linked FAs were clarified using retention times of the 2-acetyloxy tricosanoic and docosanoic acid methyl esters, obtained from the methylthio derivatives. Then, these methyl esters were converted into (2S)-oct-2-yl esters of 2-hydroxy acids. The resulting (2S)-oct-2-yl esters of 2-hydroxy tricosanoic and docosanoic acids coeluted in GC analyses with the reference (2S)-oct-2-yl ester of (2R)-2-hydroxy tricosanoic or docosanoic acids, respectively, indicating (2R)-configurations of 2-hydroxy acids liberated from oxidized cerebrosides.

Thus, we used two complementary approaches for determining oxidized acyl chain structures in glycosphingolipids. Analysis of the FA derivatives from part 1 indicated the allylic oxygen-containing group positions, but hydrogenation resulted in the loss of information regarding the positions of double bonds in the starting material (approach 1). In the analysis of part 2 (approach 2), the data on the locations of three-carbon allylically oxygenated substructures in the FA derivatives and the information on their straight-chain structures and (2R)-configurations were obtained. Although there was no direct information about the position of the double bond in the FA esters, the combined data of approaches 1 and 2 allowed for determination of the double bond and allylic hydroperoxy, hydroxy, or keto group locations in each acyl chain.

Analyses of Sphingoid Bases and Sugar from Parts 1 and 2. The sphingoid bases, liberated by hydrolysis of oxidized cerebrosides, were obtained as acetylated derivatives. The ¹Н-NМR data (δ_Н values of CH₂-1–CH-4; CDCl₃) and optical rotation value ([α]²⁵_D = + 27.9, CHCl₃) of the hydrogenated sphingoid base acetates, isolated from the hydrolysate of part 1, indicated their (2S,3S,4R)-configuration [31]. The ¹Н-NМR spectrum of these compounds also showed signals of the terminal methyl groups of dominant normal-chain (δ_Н 0.88, t, J = 6.9 Hz) and minor cyclopropane-containing (δ_Н 0.89, t, J = 6.9 Hz) constituents. Under our conditions of hydrogenation, ring opening was not the dominant process for the minor constituent, so the signals of cis-cyclopropane protons at δ_Н −0.33 (dt, J = 4.2, 5.5 Hz), 0.56 (ddd, J = 4.2, 8.3, 8.3 Hz), and 0.645 (m) [32] were observed. The hydrolysis of derivatized oxidized cerebrosides from part 2 with subsequent acetylation of products gave three sphingoid base derivatives. Two spingoid base derivatives were the DMDS adducts of acetylated isomeric monoenoic С₂₀ compounds. The mass spectrum of the DMDS adduct of major acetylated C₂₀ monoene exhibited significant peaks at m/z 432 [M − H₃CSC₉H₁₈]⁺, 372 [M − H₃CSC₉H₁₈ − AcOH]⁺ (base peak), and 173 [H₃CSC₉H₁₈]⁺, indicating 11,12-double bond in backbone 1. Key peaks in the mass spectrum of the DMDS adduct of isomeric minor acetylated C₂₀ monoene, which have a longer retention time in GC-MS, were observed at m/z 460 [M − H₃CSC₇H₁₄]⁺, 400 [M − H₃CSC₇H₁₄ − AcOH]⁺ (base peak), and 145 [H₃CSC₇H₁₄]⁺, indicating Δ13 unsaturation in backbone 2. A third acetylated sphingoid base, derived from cyclopropane-containing backbone 3, did not give a DMDS adduct. In the mass spectrum of this compound, an [M − AcOH]⁺ ion fragmented to give discernible peaks at m/z 380 [M − AcOH − (CH₂)₆СH₃]⁺ (α cleavage to a cyclopropane ring, at the C-14–C-15 position) and 394 [M − AcOH − (CH₂)₅СH₃]⁺ (β-cleavage, at the C-15–C-16 position) that was characteristic of the peracetate of the С₂₁ sphingoid base containing a ring between C-13 and C-14.

Methylthiolation of cis-monoenes and trans-monoenes with DMDS, as an anti-addition, leads to the threo-adducts and erythro-adducts, respectively [33,34]. The threo-diastereomers and erythro-diastereomers can be easily distinguished by NMR shifts of protons and carbons in and close to the 1,2-bis(alkylthio) moiety [35]. In addition to the data presented in the NMR study of Knothe and Steidley [35], we used δ_Н values of the DMDS adducts of two standards, methyl palmitoleate (Figure 6a) and its trans-isomer (Figure 6b), to confirm the configurations of double bonds in the monoenoic sphingoid base moieties of the starting oxidized cerebrosides. The δ_H values for CH₂, α to the –CH(SMe)–CH(SMe)– moiety, were obtained through correlations in ¹Н,¹Н-COSY diagrams. The ¹Н-NМR and ¹Н,¹Н-COSY spectra (CDCl₃) of the DMDS derivatives of acetylated sphingoid bases, derived from backbones 1 and 2, showed superimposed resonances of two vicinal CH groups (δ_Н 2.685, m), linked to –SMe (δ_H 2.10, s), and two α-CH₂ groups (δ_Н 1.84, m; 1.32, m), characteristic of threo-diastereomers. Consequently, cis-monoenes (1 and 2) were precursors of these compounds.

GC analyses of peracetylated (2R)- and (2S)-oct-2-yl glucosides showed a D-configuration of glucose, released from parts 1 and 2 [36].

2.4. Oxidized Cerebrosides from the Extract of Aulosaccus sp.: Structures and Possible Origins

As a result of our study, structures of 18 previously unknown compounds, found in the complex mixture of the oxidized cerebrosides from the extract of Aulosaccus sp., were elucidated. These β-D-glucopyranosyl-(1→1)-ceramides (1a–a^//, 1b–b^//, 2a–a^//, 2b–b^//, 3c–c^//, 3d–d^//) were shown to contain phytosphingosine-type backbones, (2S,3S,4R,11Z)-2-aminoeicos-11-ene-1,3,4-triol (in 1), (2S,3S,4R,13Z)-2-aminoeicos-13-ene-1,3,4-triol (in 2), and (13S*,14R*)-2-amino-13,14-methylene-eicosane-1,3,4-triol (in 3). These backbones were N-acylated with straight-chain monoenoic (2R)-2-hydroxy acids that had allylic hydroperoxy/hydroxy/keto groups on C-17^/ in the 15^/E-23:1 chain (a–a^//), C-16^/ in the 17^/E-23:1 (b–b^//) and 14^/E-22:1 (c–c^//) chains, and C-15^/ in the 16^/E-22:1 chain (d–d^//). Cerebrosides, having backbones 1, 2, and 3, comprised, respectively, 60%, 20%, and 20% of the mixture. The percentages were calculated from the integration of signals of a cyclopropane ring in the ¹H-NMR spectra of this mixture (backbone 3) and from relative intensities of GC peaks, represented by DMDS derivatives of two acetylated isomeric monoenoic sphingoid bases (backbones 1 and 2, Δ¹¹:Δ¹³ ≈ 3:1). The GC-MS analysis of the hydrogenation products of amide-linked FAs indicated that the a:b, c:d, a^/:b^/, c^/:d^/, a^//:b^//, and c^//:d^// isomer ratios were approximately 1:1.Therefore, the employed complementary instrumental and chemical methods clarified structures of oxidized cerebrosides in a complex mixture, without requiring isolation or complete separation.

Previously, the possible precursors of the oxidized cerebrosides were found in the major RP-HPLC fraction of glycosphingolipids, isolated from the extract of Aulosaccus sp. These potential precursors were β-d-glucopyranosyl-(1→1)-ceramides that contained backbones 1 (60% in the fraction) and 2 (20%), N-acylated with (2R,16Z)-2-hydroxytricos-16-enoic acid, and backbone 3 (20%), N-acylated with (2R,15Z)-2-hydroxydocos-15-enoic acid [8]. Perhaps, peroxidation of the amide-linked FAs occurred symmetrically about the cis-(n–7) double bond, so a hydroperoxy, hydroxy, or keto group, found in the major trans-monoenoic peroxidation products, was located at each of the carbon atoms, which originally formed the double bond (namely, in the (n–8) and (n–7) positions, ≈1:1). In particular, structures a–a^// and b–b^// could be formed from С₂₃Δ^16Z acyl chain while structures c–c^// and d–d^// could be formed from the С₂₂Δ^15Z acyl chain.

According to the product composition, photo-oxidation and autooxidation [1] are possible mechanisms involved in the formation of the oxidized cerebrosides from the extract of Aulosaccus sp. However, we would like to point to another possible origin of the oxidized cerebrosides in Aulosaccus sp., taking into account the relationship between these oxidation products and other compounds isolated from the same sponge sample. In particular, some bacterial branched-chain, cyclopropane-containing FAs, and their monoenoic precursors were present in significant amounts in Aulosaccus sp. [32], and an overwhelming number of the sterols (stanols, Δ⁵-, Δ⁷-, and Δ⁸⁽¹⁴⁾-sterols) of this sponge were oxidized to the corresponding 3-ketosteroids [37]. The occurrence of these FAs and steroids in Aulosaccus sp. suggested this sponge was associated with actinobacteria, known as sponge-specific microorganisms [38] and sterol degraders [39]. Cholesterol oxidase, produced by a variety of actinobacteria [40], could catalyze the transformations of the previously mentioned sterols into 3-ketosteroids [41] with the generation of H₂O₂. We suggest that H₂O₂ production in the enzymatic oxidation of Aulosaccus sp. sterols led to oxidative transformations of a certain part of cerebrosides, located in the membranes of eukaryotic cells together with sterols.

3. Materials and Methods

3.1. General Procedures

¹H-, ¹³C-NMR, ¹Н,¹Н-COSY, HSQC, HMBC, NOESY (Nuclear Overhauser Effect Spectroscopy), and TOCSY spectra (in CD₃OD or CDCl₃) were recorded on Bruker Avance III HD 500 and Bruker Avance III 700 spectrometers (Bruker BioSpin, Bremen, Germany) at 125 MHz (¹³C), 500 (¹H), and 700 (¹H) MHz. Chemical shifts (ppm) were internally referenced to the corresponding residual solvent signals δ_H 3.306/ δ_C 49.6 for CD₃OD. For the spectra of compounds in CDCl₃, TMS was used as an internal standard. ESI mass spectra were recorded on a Q-TOF 6510 mass spectrometer (Agilent Technologies, Santa Clara, CA, USA). GC-MS analyses were carried out on a Hewlett Packard HP6890 GC System (Hewlett-Packard Company, Palo Alto, CA, USA) with an HP-5MS (J&W Scientific, Folsom, CA, USA) capillary column (30.0 m × 0.25 mm), helium as the carrier gas, and 70 eV ionizing potential. GC analyses were done on an Agilent 6850 Series GC System chromatograph (Agilent Technologies, Santa Clara, CA, USA) equipped with an HP-1 (Agilent technology, Santa Clara, CA, USA) capillary column (30 m × 0.32 mm), the carrier gas was helium (flow rate 1.7 mL/min), and the detector temperature was 300 °C (or 280 °C for per-acetylated (2R)-oct-2-yl glucosides). The GC-MS analyses of FA esters and per-acetylated sphingoid bases were done using the injector temperature of 270 °C and the temperature program 100 °C (1 min)–10 °C/min–280 °C (30 min). The GC analyses of per-acetylated (2R)-oct-2-yl derivatives of D-Glc and L-Glc were carried out using the injector temperature of 150 °C and the temperature program 100 °C (0.5 min)–5 °C/min–250 °C (10 min). Optical rotation was measured on a Perkin–Elmer polarimeter, model 343 (Perkin-Elmer GmbH, Überlingen, Germany). Column chromatography was performed using silica gel (50/100 μm, Sorbpolimer, Krasnodar, Russia). RP-HPLC separations were performed using a Du Pont Series 8800 Instrument (DuPont, Wilmington, DE, USA) with an RIDK-102 refractometer (Laboratorni pristroje, Praha, Czechoslovakia). Two columns (Agilent Technologies, Santa Clara, CA, USA), Agilent ZORBAX Eclipse XDB-C8 (4 × 150 mm), and Agilent ZORBAX SB-C18 (4.6 × 250 mm), were used for the HPLC.

3.2. Animal Material

The sample of the genus Aulosaccus (phylum Porifera, class Hexactinellida, order Lyssacinosida, family Rossellidae), 930 g (wet weight), was collected in July 2011 by dredging (with small Sigsbi trail) from a 505-m depth near Iturup Island (Kuril Islands, 45°01.05^/N, 147°00.03^/E) during a cruise onboard the r/v “Academik Oparin”. The species was identified by Dr A.L. Drozdov (A.V. Zhirmunskii Institute of Marine Biology, Far Eastern Branch of RAS, Vladivostok, Russia). The collected sponge was stored at −15 °C.

3.3. Extraction and Isolation

The isolation of total cerebroside sum (238 mg) from the EtOH extract of Aulosaccus sp. was described in our previous article [8]. This sum was separated by RP-HPLC (Agilent ZORBAX Eclipse XDB-C8 column, EtOH-H₂O, 85:15, v/v). A mixture of the most polar minor cerebrosides (22.4 mg) was re-chromatographed (EtOH-H₂O-CHCl₃, 75:25:5, v/v) to give a major fraction (4.4 mg) containing oxidized cerebrosides: (+)HR-ESI-MS: 910.6606 ([M + Na]⁺ of isomeric allylic hydroperoxides, C₄₉H₉₃NO₁₂Na⁺; calc. 910.6590), 894.6649 ([M + Na]⁺ of isomeric allylic alcohols, C₄₉H₉₃NO₁₁Na⁺; calc. 894.6641), and 892.6534 ([M + Na]⁺ of isomeric enones, C₄₉H₉₁NO₁₁Na⁺; calc. 892.6484); (−)HR-ESI-MS: 922.6387 ([M + Cl]⁻ of isomeric allylic hydroperoxides, C₄₉H₉₃NO₁₂Cl⁻; calc. 922.6386), 906.6425 ([M + Cl]⁻ of isomeric allylic alcohols, C₄₉H₉₃NO₁₁Cl⁻; calc. 906.6437), and 904.6353 ([M + Cl]⁻ of isomeric enones, C₄₉H₉₁NO₁₁Cl⁻; calc. 904.6281); ¹H- and ¹³C-NMR (CD₃OD): see Table 1 and Figures S2 and S3. This fraction was divided into two parts. The constituents of parts 1 and 2 were analyzed using different chemical procedures.

3.4. Analyses of FAs, Sphingoid Bases, and Sugar from Oxidized Cerebrosides (Parts 1 and 2)

The constituents of part 1 (3.0 mg) were hydrogenated in EtOH over Adams’ catalyst at room temperature (10 h). The resulting hydrogenation product was hydrolyzed in MeCN (0.45 mL) and 5 N HCl (0.05 mL) for 4 h at 75 °C in a capped vial. Hydrolysate was concentrated in vacuo, partitioned between CHCl₃ (1.0 mL) and H₂O (3 × 1.0 mL). The H₂O extract was evaporated to dryness to give sugar-containing subfraction. The CHCl₃ layer was concentrated in vacuo, and a dry residue was acetylated with Ac₂O (0.2 mL) in pyridine (0.2 mL), overnight. The acetylated material was subjected to chromatography over silica gel column (3.0 cm × 1.2 cm), using hexane-ethylacetate (5:1→2:1→1:1, v/v) system, and then CHCl₃-EtOH (1:1, v/v) system. Elution with 60 mL of hexane-ethylacetate (2:1, v/v) gave a subfraction of per-acetylated sphingoid bases (0.8 mg). 2-Acetyloxy FA (0.7 mg) was eluted with 20 mL of CHCl₃-EtOH (1:1, v/v) system. The subfraction of the acetates of sphingoid bases was re-purified by column chromatography (SiO₂, 3.5 cm × 1.2 cm) eluting with 60 mL of hexane-ethyl acetate (2:1, v/v). The eluate was evaporated in vacuo yielding a mixture (0.5 mg) of (2S,3S,4R)-2-aminoeicosane-1,3,4-triol (n-t20:0, using shorthand nomenclature [42]) and (2S,3S,4R,13S*,14R*)-2-amino-13,14-methylene-eicosane-1,3,4-triol (cis-13,14-methylene-t21:0): [α]²⁵_D = +27.9 (c = 0.03, CHCl₃), ¹H-NMR (500 MHz, CDCl₃): 5.94 (d, J = 9.4 Hz, NH), 5.10 (dd, J = 3.0, 8.1 Hz, H-3), 4.94 (dt, J = 3.0, 9.9 Hz, H-4), 4.475 (m, H-2), 4.29 (dd, J = 4.8, 11.6 Hz, H-1b), 4.01 (dd, J = 3.0, 11.6 Hz, H-1a), 2.08 (s, −OCOCH₃ at C-3), 2.05 (two s, −OCOCH₃ at C-4 and at C-1), 2.025 (s, −NHCOCH₃), 1.64 (m, H₂C-5), 1.45−1.05 (m, –(CH₂)_n–), 0.88 (t, J = 6.9 Hz, H₃C-20 of n-t20:0), 0.89 (t, J = 6.9 Hz, H₃C-20 of cis-13,14-methylene-t21:0), 0.65 (m, H-13, H-14 of cis-13,14-methylene-t21:0), 0.565 (ddd, J = 4.1, 8.3, 8.3 Hz, H-21b of cis-13,14-methylene-t21:0), −0.33 (dt, J = 4.1, 5.5 Hz, H-21a of cis-13,14-methylene-t21:0); (2S,3S,4R)-2-aminoeicosane-1,3,4-triol MS, m/z (relative intensity, %): 452/453/454 [M − AcOH − H/M − AcOH/M − AcNH₂]⁺ (1.0/0.3/0.9), 380 [M − AcOCH₂ − AcOH]⁺ (15), 338 [M − AcOCH₂ − AcOH − CH₂CO]⁺ (14), 333 [M − 3AcOH]⁺ (19), 320 [M − AcOCH₂ − 2AcOH]⁺ (23),144 [AcOCH₂CH(NHAc)]⁺ (90), 102 [AcOCH₂CH(NHAc) − СН₂СO]⁺ (45), 84 [AcOCH₂CH(NHAc) − AсOН]⁺ (100); (13S*,14R*)-2-amino-13,14-methylene-eicosane-1,3,4-triol MS, m/z (relative intensity, %): 525 [M]⁺ (2), 465 [M − AcOH]⁺ (21), 406 [M − AcOH − AcNH₂]⁺ (9), 394 (6), 380 (5), 332 [M − AcOCH₂ − 2AcOH]⁺ (12), 144 [AcO-CH₂CH(NHAc)]⁺ (52), 102 [AcOCH₂CH(NHAc) − СН₂СO]⁺ (52), 84 [AcOCH₂CH(NHAc) − AсOН]⁺ (100). The components of the subfraction of 2-acetyloxy FAs were methylated by a standard method with N-nitroso-N-methylurea. The resulting compounds were purified by column chromatography (SiO₂, 3.5 cm × 1.2 cm) eluting with 44 mL of hexane-ethyl acetate (10:1, v/v). The elution gave a mixture of the methyl esters of 2-acetyloxy FAs (0.5 mg): ¹H-NMR (CDCl₃, 700 MHz): 4.99 (dd, J = 6.6, 6.9 Hz, H-2), 4.855 (m, mid-chain –СН(OAс)– of diacetyloxy acids), 3.74 (s, H₃CO– at C-1), 2.38 (t, J = 7.4 Hz, mid-chain –H₂C–CO–CH₂– of keto acids), 2.135 (s, −OCOCH₃ at C-2), 1.82 (m, H₂C-3), 1.45–1.15 (m, –(CH₂)_n–), 0.88 (a series of triplets about this point, J ≈ 7.0 Hz), the methyl esters of keto C₂₃ and C₂₂ acids MS: Figures S4–S7, S12–S16; the methyl esters of diacetyloxy C₂₃ and C₂₂ acids MS: Figures S8–S11, S17–S20, the methyl ester of saturated 2-acetyloxy C₂₃ acid MS: 426 [M]⁺ (0.04%), 384 [M − CH₂CO]⁺ (100%), 352 [M − CH₂CO − MeOH]⁺ (34%), and the methyl ester of saturated 2-acetyloxy C₂₂ acid MS: 412 [M]⁺ (0.04%), 370 [M − CH₂CO]⁺ (100%), 338 [M − CH₂CO − MeOH]⁺ (35%).

Components of part 2 (1.4 mg) were acetylated with Ac₂O (0.2 mL) in pyridine (0.2 mL), overnight. After vacuum drying, CHCl₃ (two drops) and CeCl₃·7H₂O (18.6 mg) were added to the acetylated material, and the mixture was dissolved in MeOH (0.125 mL). NaBH₄ (5 mg, excess) was slowly added with stirring. After 5 min, the reaction was quenched by the addition of NH₄Cl water solution (5.4 mg in 0.2 mL H₂O). A dry residue was obtained after complete evaporation of the reaction mixture. It was extracted with CHCl₃ to remove most inorganic admixtures. After vacuum drying of the CHCl₃ extract, the resulting dry material was dissolved in DMDS (0.1 mL), and a solution (0.025 mL) of iodine in diethyl ether (60 mg/mL) was added. The mixture was kept for 4 days at room temperature. Then the reaction was stopped by adding aqueous solution of Na₂S₂O₃ (5%, 0.2 mL), and the mixture was extracted with hexane (5 × 0.5 mL). The hexane extract was evaporated to dryness, and the resulting compounds were hydrolyzed in MeCN/HCl. Then, subfractions, containing sugar, peracetylated sphingoid bases (0.4 mg), and 2-acetyloxy FAs (0.3 mg), were obtained, as described above. The subfraction of peracetylated sphingoid bases consisted of peracetylated (13S*,14R*)-2-amino-13,14-methylene-eicosane-1,3,4-triol and DMDS derivatives of peracetylated (11Z)-2-aminoeicos-11-ene-1,3,4-triol and (13Z)-2-aminoeicos-13-ene-1,3,4-triol: ¹H-NMR (500 MHz, CDCl₃): 5.93 (d, J = 9.5 Hz, NH), 5.095 (dd, J = 3.1, 8.3 Hz, H-3), 4.94 (dt, J = 3.1, 9.9 Hz, H-4), 4.47 (m, H-2), 4.29 (dd, J = 4.8, 11.8 Hz, H-1b), 4.00 (dd, J = 3.1, 11.8 Hz, H-1a), 2.685 (m, –CH(SCH₃)–CH(SCH₃)–), 2.10 (s, –CH(SCH₃)–CH(SCH₃)–), 2.08 (s, −OCOCH₃ at C-3), 2.05 (two s, −OCOCH₃ at C-4 and at C-1), 2.025 (s, −NHCOCH₃), 1.84 (m; H-b, –H₂CCH(SCH₃)–CH(SCH₃)CH₂–), 1.64 (m, H₂C-5), 1.42−1.18 (m, –(CH₂)_n–), 1.32 (m, H-a, –H₂CCH(SCH₃)–CH(SCH₃)CH₂–), 0.88 (t, J = 6.9 Hz, H₃C-20 of n-t20:0), 0.89 (t, J = 6.9 Hz, H₃C-20 of cis-13,14-methylene-t21:0), 0.645 (m, H-13, H-14 of cis-13,14-methylene-t21:0), 0.56 (ddd, J = 4.2, 8.2, 8.2 Hz, H-21b of cis-13,14-methylene-t21:0), −0.33 (dt, J = 4.2, 5.4 Hz, H-21a of cis-13,14-methylene-t21:0). The DMDS derivative of per-acetylated (11Z)-2-aminoeicos-11-ene-1,3,4-triol MS, m/z (relative intensity, %): 557 [M − HSMe]⁺ (1.4), 432 [M − H₃CSC₉H₁₈]⁺ (24), 372 [M − H₃CSC₉H₁₈ − CH₃COOH]⁺ (100), 330 (11), 312 (6), 173 [H₃CSC₉H₁₈]⁺ (30), 102 [AcOCH₂CH(NHAc) − СН₂СO]⁺ (25), 84 [AcOCH₂CH(NHAc) − AсOН]⁺ (45). The DMDS derivative of per-acetylated (13Z)-2-aminoeicos-13-ene-1,3,4-triol MS, m/z (relative intensity, %): 557 [M − HSMe]⁺ (2), 460 [M − H₃CSC₇H₁₄]⁺ (24), 400 [M − H₃CSC₇H₁₄ − CH₃COOH]⁺ (100), 358 (10), 340 (11), 145 [H₃CSC₇H₁₄]⁺ (36), 102 [AcOCH₂CH(NHAc) − СН₂СO]⁺ (36), 84 [AcOCH₂CH(NHAc) − AсOН]⁺ (55). The 2-acetyloxy acids from FA subfraction were methylated using N-nitroso-N-methylurea. The resulting methyl esters were purified by column chromatography (SiO₂, 3.5 cm × 1.2 cm), eluting with 44 mL of hexane-ethyl acetate (10:1, v/v). The elution gave a mixture, containing the methyl esters of major mono(methylthio) and minor tris(methylthio) derivatives of 2-acetyloxy FAs (0.2 mg): ¹H-NMR (CDCl₃, 700 MHz): 5.40 (dt, J = 6.9, 15.2 Hz, –HC=CH–CH(SCH₃)–), 5.175 (dd, J = 9.0, 15.2 Hz, –HC=CH–CH(SCH₃)–), 4.99 (dd, J = 6.2, 6.9 Hz, H-2), 3.74 (s, H₃CO– at C-1), 2.99 (m, –HC=CH–CH(SCH₃)–), 2.135 (s, −OCOCH₃ at C-2), 2.04 (m, allylic CH₂), 1.965 (s, –HC=CH–CH(SCH₃)–), 1.82 (m, H₂C-3), 1.45–1.15 (m, –(CH₂)_n–), 0.88–0.89 (a series of triplets, J ≈ 7.0 Hz), the methyl esters of the mono(methylthio) derivatives of 2-acetyloxy C₂₃ acid MS: Figures S21–S24, the methyl esters of the mono(methylthio) derivatives of 2-acetyloxy C₂₂ acid MS: Figures S25–S28, the methyl esters of the tris(methylthio) derivatives of 2-acetyloxy C₂₃ acid MS: Figures S29 and S30, the methyl esters of the tris(methylthio) derivatives of 2-acetyloxy C₂₂ acid MS: Figures S31 and S32. For the methyl esters of 2-acetyloxy C₂₃ acids containing an allylic –SMe group, the order of elution was 15-methylthio-16-ene→17-methylthio-15-ene→16-methylthio-17-ene→18-methylthio-16-ene. The elution order for the methyl esters of 2-acetyloxy C₂₂ acids containing an allylic –SMe group was 14-methylthio-15-ene→16-methylthio-14-ene→15-methylthio-16-ene→17-methylthio-15-ene. The mixture of these methylthio derivatives was hydrogenated over Adams’ catalyst in AcOH at room temperature (12 h). The hydrogenation gave methyl 2-(acetyloxy)tricosanoate and methyl 2-(acetyloxy)docosanoate. The straight-chain structures of these compounds were clarified, using the GC-MS data on the methyl esters of straight-chain 2-acetyloxy tricosanoic and docosanoic acids, released from the non-oxidized cerebrosides of Aulosaccus sp. Then, methyl 2-(acetyloxy)tricosanoate and methyl 2-(acetyloxy)docosanoate, obtained by hydrodesulfurization, were de-esterified in MeCN (0.45 mL) and 5N HCl (0.05 mL) for 4 h at 73−75 °C to give free FAs. These FAs were converted into (2S)-oct-2-yl esters of 2-hydroxy FAs by treatment with 2% H₂SO₄ in (2S)-octan-2-ol (0.2 mL) for 4 h at 75 °C in a capped vial [8]. In GC analyses, the retention times of the resulting (2S)-oct-2-yl ester of 2-hydroxy tricosanoic and docosanoic acids were identical with those of the standard (2S)-oct-2-yl esters of (2R)-2-hydroxy tricosanoic and docosanoic acids, respectively.

The absolute configuration of glucose, released from oxidized cerebrosides, was determined by the GC analyses of per-acetylated (2R)-oct-2-yl glycosides according to the method of Leontein et al. [41]. The sugar (1.2 mg), (2R)-octan-2-ol (0.4 mL), and one drop of trifluoroacetic acid in a capped vial were kept for 7 h at 120 °C with stirring. Then, the mixture was concentrated in vacuo and acetylated with Ac₂O (0.4 mL) in pyridine (0.4 mL), overnight. The acetylated material was purified by column chromatography (SiO₂, 3.0 cm × 1.2 cm), eluting with a mixture of hexane/ethylacetate (5:1, v/v). The eluate was evaporated, yielding per-acetylated (2R)-oct-2-yl glucoside. Standards, D-Glc (1.0 mg) and L-Glc (1.0 mg), were treated and derivatized under the same conditions that were applied to the sugar subfractions, liberated from parts 1 and 2. The GC profiles (the retention times and intensities of GC peaks) of the derivatives of D-Glc and sugar, obtained from cerebrosides, were proven to be identical.