*Article* **Triterpene Glycosides from the Far Eastern Sea Cucumber** *Thyonidium (=Duasmodactyla) kurilensis* **(Levin): The Structures, Cytotoxicities, and Biogenesis of Kurilosides A3, D1, G, H, I, I1, J, K, and K1**

**Alexandra S. Silchenko, Anatoly I. Kalinovsky, Sergey A. Avilov, Pelageya V. Andrijaschenko, Roman S. Popov, Pavel S. Dmitrenok, Ekaterina A. Chingizova and Vladimir I. Kalinin \***

> G.B. Elyakov Pacific Institute of Bioorganic Chemistry, Far Eastern Branch of the Russian Academy of Sciences, Pr. 100-letya Vladivostoka 159, 690022 Vladivostok, Russia; silchenko\_alexandra\_s@piboc.dvo.ru (A.S.S.); kaaniv@piboc.dvo.ru (A.I.K.); avilov\_sa@piboc.dvo.ru (S.A.A.); andrijashchenko\_pv@piboc.dvo.ru (P.V.A.); popov\_rs@piboc.dvo.ru (R.S.P.); paveldmt@piboc.dvo.ru (P.S.D.); chingizova\_ea@piboc.dvo.ru (E.A.C.) **\*** Correspondence: kalininv@piboc.dvo.ru; Tel./Fax: +7-(423)2-31-40-50

**Citation:** Silchenko, A.S.; Kalinovsky, A.I.; Avilov, S.A.; Andrijaschenko, P.V.; Popov, R.S.; Dmitrenok, P.S.; Chingizova, E.A.; Kalinin, V.I. Triterpene Glycosides from the Far Eastern Sea Cucumber *Thyonidium (=Duasmodactyla) kurilensis* (Levin): The Structures, Cytotoxicities, and Biogenesis of Kurilosides A3, D1, G, H, I, I1, J, K, and K1. *Mar. Drugs* **2021**, *19*, 187. https://doi.org/10.3390/ md19040187

Academic Editors: Vassilios Roussis and Hitoshi Sashiwa

Received: 25 February 2021 Accepted: 24 March 2021 Published: 27 March 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

**Abstract:** Nine new mono-, di-, and trisulfated triterpene penta- and hexaosides, kurilosides A3 (**1**), D1 (**2**), G (**3**), H (**4**), I (**5**), I1 (**6**), J (**7**), K (**8**), and K1 (**9**) and two desulfated derivatives, DS-kuriloside L (**10**), having a trisaccharide branched chain, and DS-kuriloside M (**11**), having hexa-*nor*-lanostane aglycone with a 7(8)-double bond, have been isolated from the Far-Eastern deep-water sea cucumber *Thyonidium (=Duasmodactyla) kurilensis* (Levin) and their structures were elucidated based on 2D NMR spectroscopy and HR-ESI mass-spectrometry. Five earlier unknown carbohydrate chains and two aglycones (having a 16*β*,(20S)-dihydroxy-fragment and a 16*β*-acetoxy,(20S)-hydroxy fragment) were found in these glycosides. All the glycosides **1**–**9** have a sulfate group at C-6 Glc, attached to C-4 Xyl1, while the positions of the other sulfate groups vary in different groups of kurilosides. The analysis of the structural features of the aglycones and the carbohydrate chains of all the glycosides of *T. kurilensis* showed their biogenetic relationships. Cytotoxic activities of the compounds **1**–**9** against mouse neuroblastoma Neuro 2a, normal epithelial JB-6 cells, and erythrocytes were studied. The highest cytotoxicity in the series was demonstrated by trisulfated hexaoside kuriloside H (**4**), having acetoxy-groups at C(16) and C(20), the latter one obviously compensated the absence of a side chain, essential for the membranolytic action of the glycosides. Kuriloside I1 (**6**), differing from **4** in the lacking of a terminal glucose residue in the bottom semi-chain, was slightly less active. The compounds **1**–**3**, **5**, and **8** did not demonstrate cytotoxic activity due to the presence of hydroxyl groups in their aglycones.

**Keywords:** *Thyonidium kurilensis*; triterpene glycosides; kurilosides; sea cucumber; cytotoxic activity

#### **1. Introduction**

The investigations of the triterpene glycosides from different species of sea cucumbers have a range of goals. Among them are the drug discoveries based on the promising candidates, demonstrating the target bioactivity [1–6], the solving of some taxonomic problems of the class Holothuroidea based on the specificity of the glycosides having characteristic structural peculiarities for the certain systematic groups [7–10], the ascertaining of biologic and ecologic functions of these metabolites [11–15], and the discovery of novel compounds, especially minor ones, that can be the "hot metabolites" clarifying the biosynthetic pathways of triterpene glycosides [16–18].

As a continuation of our investigation of glycoside composition of the sea cucumber *Thyonidium* (=*Duasmodactuyla*) *kurilensis* (Levin), we report herein the isolation and structure elucidation of nine glycosides, kurilosides A3 (**1**), D1 (**2**), G (**3**), H (**4**), I (**5**), I1 (**6**), J (**7**), K (**8**), and K1 (**9**) as well as two desulfated derivatives, DS-kuriloside L (**10**) and DS-kuriloside

M (**11**). The animals were collected near Onekotan Island in the Sea of Okhotsk. The structures of the compounds **1**–**11** were established by the analyses of the 1H, 13C NMR, 1D TOCSY, and 2D NMR (1H,1H-COSY, HMBC, HSQC, ROESY) spectra as well as HR-ESI mass spectra. All the original spectra are presented in Figures S1–S85 in the Supplementary Materials. The hemolytic activities against mouse erythrocytes, cytotoxic activities against mouse neuroblastoma Neuro 2a, and normal epithelial JB-6 cells have been reported.

#### **2. Results and Discussion**

#### *2.1. Structural Elucidation of the Glycosides*

The concentrated ethanolic extract of the sea cucumber *Thyonidium (=Duasmodactyla) kurilensis* was chromatographed on a Polychrom-1 column (powdered Teflon, Biolar, Latvia). The glycosides were eluted with 50% EtOH and separated by repeated chromatography on Si gel columns using CHCl3/EtOH/H2O (100:100:17) and (100:125:25) as mobile phases to give five fractions (I–V). The glycosides **1**–**9** (Figure 1) were isolated as a result of subsequent HPLC of the fractions II–V on a reversed-phase semipreparative column Phenomenex Synergi Fusion RP (10 × 250 mm).

**Figure 1.** Chemical structures of glycosides isolated from *Thyonidium kurilensis:* **1**—kuriloside A3; **2**—kuriloside D1; **3**—kuriloside G; **4**—kuriloside H; **5**—kuriloside I; **6**—kuriloside I1; **7**—kuriloside J, **8**—kuriloside K, **9**—kuriloside K1.

− The molecular formula of kuriloside A3 (**1**) was determined to be C54H87O29SNa from the [MNa − Na]<sup>−</sup> ion peak at *m/z* 1231.5063 (calc. 1231.5059) in the (−)HR-ESI-MS. Kuriloside A3 (**1**) as well as the reported earlier kurilosides A, A1, and A2 [19] belong to the same group of glycosides, so these compounds have the identical monosulfated pentasaccharide chains that were confirmed by the coincidence of their 1H and 13C NMR spectra corresponding to the carbohydrate chains (Table S1). The presence of five characteristic doublets at δ<sup>H</sup> = 4.64–5.18 (*J* = 7.1–7.6 Hz), and corresponding signals of anomeric carbons at δ<sup>C</sup> = 102.3–104.7 in the 1H and 13C NMR spectra of the carbohydrate part of **1** indicate the presence of a pentasaccharide chain and *β*-configurations of the glycosidic bonds. Monosaccharide composition of **1**, established by the analysis of the 1H,1H-COSY, HSQC, and 1D TOCSY spectra, includes one xylose (Xyl1), one quinovose (Qui2), two glucoses (Glc3 and Glc4), and one 3-O-methylglucose (MeGlc5) residue. The signal of C-6 Glc4 was observed at δ<sup>C</sup> = 67.1 due to α-shifting effect of a sulfate group at this position. The positions of interglycosidic linkages were established by the ROESY and HMBC spectra (Table S1). The analysis of NMR spectra of the aglycone part of **1** (Table S2) indicated the presence of 22,23,24,25,26,27-hexa-*nor*-lanostane aglycone with a 16α-hydroxy,20-oxofragment and 9(11)-double bond due to the characteristic signals: (δ<sup>C</sup> 149.0 (C-9) and 114.2 (C-11), δ<sup>C</sup> = 71.1 (C-16) and δ<sup>H</sup> = 5.40 (brt, *J* = 7.5 Hz, H-16), δ<sup>C</sup> = 208.8 (C-20)). The ROE

correlations H-16/H-15β and H-16/H-18 indicated a 16α-OH orientation in the aglycone of kuriloside A3 (**1**). 17αH-orientation, common for the sea cucumber glycosides, was deduced from the ROE-correlation H-17/H-32. The same aglycone was found earlier in kuriloside F [19].

The (−)ESI-MS/MS of **1** demonstrated the fragmentation of [MNa − Na]<sup>−</sup> ion at *m/z* 1231.5. The peaks of fragment ions were observed at *m/z* 1069.5 [MNa – Na − C6H10O5 (Glc)]−, 1055.4 [MNa – Na − C7H12O5(MeGlc)]−, 923.4 [MNa − Na − C6H10O5(Glc) − C6H10O4(Qui)]−, 747.3 [MNa − Na − C6H10O5(Glc) − C6H10O4(Qui) − C7H12O5(MeGlc)]−, 695.1 [MNa −Na − C24H37O3(Agl) − C6H10O5 (Glc) − H]−, 565.1 [MNa − Na − C24H37O2 (Agl) − C6H10O5(Glc) − C6H10O4 (Qui) − H]−, 549.1 [MNa − Na − C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − H]−, 417.1 [MNa – Na − C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [MNa − Na − C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − C7H12O5(MeGlc) − H]−, corroborating the structure of kuriloside A3 (**1**).

All these data indicate that kuriloside A3 (**1**) is 3β-*O*-{β-D-glucopyranosyl-(1→4)-β-Dquinovopyranosyl-(1→2)-[3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23,24,25,26,27-hexa-nor-16α-hydroxy,20 oxo-lanost-9(11)-ene.

The molecular formula of kuriloside D1 (**2**) was determined to be C66H107O36SNa from the [MNa − Na]<sup>−</sup> ion peak at *m/z* 1507.6291 (calc. 1507.6268) in the (−)HR-ESI-MS. The hexasaccharide monosulfated carbohydrate chain of **2** was identical to that of previously reported kuriloside D [19] since their 1H and 13C NMR spectra corresponding to the carbohydrate moieties were coincident (Table S3). Actually, six signals of anomeric doublets at δ<sup>H</sup> = 4.70–5.28 (d, *J* = 7.5–8.2 Hz) and corresponding signals of anomeric carbons at δ<sup>C</sup> = 103.7–105.7 indicated the presence of a hexasaccharide chain in kuriloside D1 (**2**). The presence of xylose (Xyl1), quinovose (Qui2), three glucose (Glc3, Glc4, Glc5), and 3-O-methylglucose (MeGlc6) residues were deduced from the analysis of the 1H,1H-COSY, HSQC, and 1D TOCSY spectra of **2**. The positions of the interglycosidic linkages were elucidated based on the ROESY and HMBC correlations (Table S3). The presence in the 13C NMR spectrum of kuriloside D1 (**2**) of the only signal of the *O*-methyl group at δ<sup>C</sup> 60.5 and the upfield shift of the signal of C-3 Glc4 to δ<sup>C</sup> 71.5 indicated the presence of a non-methylated terminal Glc4 residue. Analysis of the 1H and 13C NMR spectra of the aglycone part of **2** indicated the presence of a lanostane aglycone (the signals of lactone ring are absent and the signals of methyl group C-18 are observed at δ<sup>C</sup> 16.9 and δ<sup>H</sup> 1.30 (s, H-18) with normal side chain (30 carbons) and 9(11)-double bond (the signals at δ<sup>C</sup> 149.0 (C-9), 114.9 (C-11), and δ<sup>H</sup> 5.35 (brd, *J* = 6.2 Hz; H-11) (Table 1). The comparison of the 13C NMR spectra of **2** and kuriloside D showed their great similarity, except for the signals of the side chain from C-23 to C-27. Two strongly deshielded signals at δ<sup>C</sup> 216.3 (C-16) and 217.6 (C-22) corresponded to carbonyl groups, whose positions were established on the base of the HMBC correlations H-15/C-16, H-21/C-22, H-23/C-22, and H-24/C-22. The signals of protons assigned to the methylene group adjacent to 22-oxo group were deshielded to δ<sup>H</sup> 3.67 (dd, *J* = 10.6; 18.2 Hz; H-23a) and 3.43 (dt, *J* = 7.8; 18.2 Hz; H-23b) and correlated in the 1H,1H-COSY spectrum of **2** with one signal only at δ<sup>H</sup> 2.27 (t, *J* = 7.8 Hz; H-24). These data, along with the deshielded signal of quaternary carbon at δ<sup>C</sup> 69.0 (C-25) and the almost coinciding signals of methyl groups C-26 and C-27 (δ<sup>C</sup> 30.0 and 29.5, δ<sup>H</sup> 1.42 and 1.41, correspondingly), indicated the attachment of the hydroxy-group to C-25. Therefore, the side chain of kuriloside D1 (**2**) is characterized by the 22-oxo-25-hydroxy-fragment (Table 1).

The (−)ESI-MS/MS of **2** demonstrated the fragmentation of [MNa − Na]<sup>−</sup> ion at *m/z* 1507.6. The peaks of fragment ions were observed at *m/z* 1349.5 [MNa − Na − C8H15O3 + H]−, corresponding to the loss of the aglycone fragment from C(20) to C(27), 1187.5 [MNa − Na − C8H15O3 − C6H10O5(Glc) + H]−, 1025.4 [MNa − Na − C8H15O3 − C6H10O5(Glc) − C6H10O5 (Glc) + H]−, 879.4 [MNa − Na − C8H15O3 − C6H10O5(Glc) − C6H10O5 (Glc) − C6H10O4(Qui) + H]−, 565.1 [MNa − Na − C30H47O4(Agl) − C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) − H]−, 417.1 [MNa − Na − C30H47O5(Agl) − C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [MNa − Na − C30H47O5 (Agl) − C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − C7H12O5(MeGlc) − H]−, corroborating the structure of kuriloside D1 (**2**).

All these data indicate that kuriloside D1 (**2**) is 3β-*O*-{β-D-glucopyranosyl-(1→3)-β-Dglucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)-[3-*O*-methyl-β-D-glucopyranosyl- (1→3)-6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-16,22-dioxo-25-hydroxylanost-9(11)-ene.

**Table 1.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of the aglycone moiety of kuriloside D1 (**1**).


<sup>a</sup> Recorded at 176.03 MHz in C5D5N/D2O (4/1). <sup>b</sup> Recorded at 700.00 MHz in C5D5N/D2O (4/1).

The molecular formula of kuriloside G (**3**) was determined to be C61H98O37S2Na2 from the [M2Na − Na]<sup>−</sup> ion peak at *m/z* 1509.5102 (calc. 1509.5132) and the [M2Na − 2Na]2<sup>−</sup> ion-peak at *m/z* 743.2624 (calc. 743.2626) in the (−)HR-ESI-MS. In the 1H and 13C NMR spectra of the carbohydrate part of kuriloside G (**3**), six characteristic doublets at δ<sup>H</sup> 4.65–5.19 (*J* = 7.0–8.1 Hz) and signals of anomeric carbons at δ<sup>C</sup> 102.1–104.8, correlated with each anomeric proton by the HSQC spectrum, were indicative of a hexasaccharide chain and *β*-configurations of glycosidic bonds (Table 2). The signals of each monosaccharide unit were found as an isolated spin system based on the 1H,1H-COSY, and 1D TOCSY spectra of **3.** Further analysis of the HSQC and ROESY spectra resulted in the assigning of the monosaccharide residues as one xylose (Xyl1), one quinovose (Qui2), two glucoses (Glc3 and Glc5), and two 3-*O*-methylglucose (MeGlc4 and MeGlc6) residues.


**Table 2.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of carbohydrate moiety of kuriloside G (**3**).

<sup>a</sup> Recorded at 176.03 MHz in C5D5N/D2O (4/1). <sup>b</sup> Bold = interglycosidic positions. <sup>c</sup> Italic = sulfate position. <sup>d</sup> Recorded at 700.00 MHz in C5D5N/D2O (4/1). Multiplicity by 1D TOCSY.

> The positions of interglycosidic linkages were established by the ROESY and HMBC spectra of **3** (Table 2) where the cross-peaks between H-1 Xyl1 and H-3 (C-3) of an aglycone,

H-1 Qui2 and H-2 (C-2) Xyl1; H-1 Glc3 and H-4 (C-4) Qui2; H-1 MeGlc4 and H-3 Glc3; H-1 Glc5 and H-4 Xyl1; H-1 MeGlc6 and H-3 (C-3) Glc5 were observed.

The signals of C-6 MeGlc4 and C-6 Glc5 in the 13C NMR spectrum of **3** were observed at δ<sup>C</sup> 67.0 and δ<sup>C</sup> 67.1, correspondingly, due to α-shifting effects of the sulfate groups at these positions. Thus, the hexasaccharide disulfated chain of kuriloside G (**3**) was first found in the sea cucumber glycosides. The NMR spectra of the aglycone part of **3** coincided with that of kuriloside A3 (**1**), indicating the identity of these aglycones (Table S2).

The (−)ESI-MS/MS of **3** demonstrated the fragmentation of [M2Na − Na]<sup>−</sup> ion at *m/z* 1509.5. The peaks of fragment ions were observed at *m/z* 1389.6 [M2Na – Na − NaHSO4] −, 1333.5 [M2Na – Na − C7H12O5(MeGlc)]−, 1231.5 [M2Na – Na − C7H11O8SNa(MeGlcSO3Na)]−, 1069.4 [M2Na – Na − C7H11O8SNa(MeGlcSO3Na) − C6H10O5(Glc)]−, 923.4 [M2Na – Na − C7H11O8SNa(MeGlcSO3Na) − C6H10O5(Glc)] − C6H10O4(Qui)]−.

All these data indicate that kuriloside G (**3**) is 3β-*O*-{6-*O*-sodium sulfate-3-*O*-methylβ-D-glucopyranosyl-(1→3)-β-D-glucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)- [3- *O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*-sodium sulfate*-*β-D-glucopyranosyl-(1→4)]-*β*-D-xylopyranosyl}-22,23,24,25,26,27-hexa-*nor*-16*α*-hydroxy,20-oxo-lanost-9(11)-ene.

The molecular formula of kuriloside H (**4**) was determined to be C64H101O42S3Na3 from the [M3Na − Na]<sup>−</sup> ion peak at *m/z* 1683.4701 (calc. 1683.4730), [M3Na − 2Na]2<sup>−</sup> ion peak at *m/z* 830.2425 (calc. 830.2419), and [M3Na − 3Na]3<sup>−</sup> ion peak at *m/z* 545.8332 (calc. 545.8315) in the (−)HR-ESI-MS. The presence of three-charged ions in the (−)HR-ESI-MS of kuriloside H (**4**) was indicative for the trisulfated glycoside.

The 1H and 13C NMR spectra corresponding to the carbohydrate chain of kuriloside H (**4**) (Table 3) demonstrated six signals of anomeric protons at δ<sup>H</sup> 4.63–5.21 (d, *J* = 7.1– 8.6 Hz) and the signals of anomeric carbons at δ<sup>C</sup> 102.8–104.7 deduced by the HSQC spectrum, indicative of hexasaccharide moiety with *β*-glycosidic bonds. The signals of each sugar residue were assigned by the analysis of the 1H,1H-COSY, 1D TOCSY, ROESY, and HSQC spectra, enabling the identification of monosaccharide units in the chain of **4** as one xylose (Xyl1), one quinovose (Qui2), three glucoses (Glc3, Glc4 and Glc5), and one 3-*O*-methylglucose (MeGlc6). Therefore, the monosaccharide composition of **4** was the same as in kuriloside D1 (**2**).

However, in the 13C NMR spectrum of **4** three signals at δ<sup>C</sup> 67.6 (C-6 Glc3), 67.4 (C-6 Glc5), and 67.0 (C-6 MeGlc6), characteristic for sulfated by C-6 hexose units, were observed instead of one signal at δ<sup>C</sup> 67.0 (C-6 Glc5) in the spectrum of **2**. The signal of the OMegroup observed at δ<sup>C</sup> 60.4 indicated one terminal monosaccharide residue was methylated. Actually, the protons of the OMe-group (δ<sup>H</sup> 3.75, s) correlated in the HMBC spectrum with C-3 MeGlc6 (δ<sup>C</sup> 86.1), which was, in turn, attached to C-3 Glc5 (ROE-correlation H-1 MeGlc6 (δ<sup>H</sup> 5.13 (d, *J* = 7.4 Hz)/H-3 Glc5 (δ<sup>H</sup> 4.13 (t, *J* = 8.6 Hz)). At the same time, the fourth (another terminal) monosaccharide unit was glucose (the signal of C-3 Glc4 was shielded to δ<sup>C</sup> 77.7 due to the absence of *O*-methylation). The positions of all interglycosidic linkages were elucidated based on the ROESY and HMBC correlations (Table 3).

Hence, kuriloside H (**4**) has a hexasaccharide chain with a non-methylated terminal Glc4 residue and three sulfate groups. This carbohydrate chain is first found in the glycosides of the sea cucumbers and kuriloside H (**4**) is the most polar glycoside discovered so far as well as two tetrasulfated pentaosides isolated from *Psolus fabricii* [20].

The analysis of the 13C NMR spectrum of the aglycone part of **4** demonstrated its identity to the aglycone of kurilosides A1 and C1, isolated earlier [19]. Therefore, kuriloside H (**4**) contains a 22,23,24,25,26,27-hexa-*nor*-lanostane aglycone with 9(11)-double bond and acetoxy-groups at C-16 and C-20. *β*-orientation of the acetoxy group at C-16 and (20*S*)-configuration were established on the base of coincidence of the coupling constants (*J*16/17 = 7.7 Hz and *J*17/20 = 10.6 Hz), observed in the 1H NMR spectra of **4** and kuriloside A1, and confirmed by the ROE-correlation H-16/H-32 in the spectrum of **4** (Table S4).


**Table 3.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of carbohydrate moiety of kuriloside H (**4**).

<sup>a</sup> Recorded at 176.04 MHz in C5D5N/D2O (4/1). <sup>b</sup> Bold = interglycosidic positions. <sup>c</sup> Italic = sulfate position. <sup>d</sup> Recorded at 700.13 MHz in C5D5N/D2O (4/1). Multiplicity by 1D TOCSY.

> The (−)ESI-MS/MS of kuriloside H (**4**) demonstrated the fragmentation of the [M3Na − Na]− ion at *m/z* 1683.5. The peaks of fragment ions were observed at *m/z* 1503.5 [M3Na – Na − CH3COOH − NaHSO4] <sup>−</sup>, 1443.5 [M3Na – Na − 2CH3COOH − NaHSO4] −, 1281.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C6H10O5(Glc)]−, 1165.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C7H11O8SNa(MeGlcOSO3)]−, and 1003.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C7H11O8SNa(MeGlcOSO3) − C6H10O5(Glc)]−, corroborating its carbohydrate chain structure.

All these data indicate that kuriloside H (**4**) is 3β-*O*-{β-D-glucopyranosyl-(1→3)-6- *O*-sodium sulfate-β-D-glucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)-[6-*O*-sodium sulfate-3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*-sodium sulfate*-*β-D-glucopyranosyl- (1→4)]-β-D-xylopyranosyl}-22,23,24,25,26,27-hexa-*nor*-16β,(20*S*)-diacetoxy-lanost-9(11)-ene.

The molecular formula of kuriloside I (**5**) was determined to be C54H87O35S3Na3 from the [M3Na − Na]<sup>−</sup> ion peak at *m/z* 1437.3952 (calc. 1437.3991), [M3Na − 2Na]2<sup>−</sup> ion peak at *m/z* 707.2049 (calc. 707.2049), and [M3Na − 3Na]3<sup>−</sup> ion peak at *m/z* 463.8076 (calc. 463.8069) in the (−)HR-ESI-MS, indicating the presence of three sulfate groups. The 1H and 13C NMR spectra corresponding to the carbohydrate part of kuriloside I (**5**) (Table 4) demonstrated five characteristic doublets at δ<sup>H</sup> 4.63–5.13 (d, *J* = 6.6–7.8 Hz) and corresponding signals of anomeric carbons at δ<sup>C</sup> 102.4–104.7 deduced by the HSQC spectrum, which indicated the presence of five monosaccharide residues in the carbohydrate chain of **5**. The signals at δ<sup>C</sup> 67.0, 67.6, and 67.7 indicated the presence of three sulfate groups as in the carbohydrate chain of kuriloside H (**4**). Indeed, the comparison of the 13C NMR spectra of kurilosides I (**5**) and H (**4**) showed that they differed by the absence in the spectrum of **5** of the signals corresponding to non-sulfated terminal glucose residue attached to C-3 Glc3 in the carbohydrate chain of **4**. The signal of C-3 Glc3 in the 13C NMR spectrum of **5** was observed at δ<sup>C</sup> 76.9 (instead of δ<sup>C</sup> 86.3 in the spectrum of **4**), demonstrating the absence of a glycosylation effect. The presence of xylose (Xyl1), quinovose (Qui2), two glucose (Glc3, Glc4), and one 3-*O*-methylglucose (MeGlc5) residue was deduced from the analysis of the 1H,1H-COSY, HSQC and 1D TOCSY spectra of **5**. The positions of interglycosidic linkages were elucidated based on the ROESY and HMBC correlations (Table 4) and indicated the presence of the branched at the C-4 Xyl1 pentasaccharide chain in **5**, with the same architecture as in the other pentaosides of *T. kurilensis*. Thus, kuriloside I (**5**) contains a new pentasaccharide branched trisulfated chain.

The analysis of the 13C and 1H NMR spectra of the aglycone part of **5** indicated the presence of 22,23,24,25,26,27-hexa-*nor*-lanostane aglycone having a 9(11)-double bond (Table 5). The signals of methine group CH-16 were observed at c δ<sup>C</sup> 72.8 (C-16) and at δ<sup>H</sup> 4.82 (dd, *J* = 7.1; 14.9 Hz, H-16) due to the attachment of the hydroxyl group to this position. The HMBC correlations H-15/C-16 and H-20/C-16 confirmed this. The signals of C-20 and H-20 were shielded to δ<sup>C</sup> 66.5 and δ<sup>H</sup> 4.38 (dd, *J* = 6.0; 9.5 Hz), correspondingly, when compared with the same signals in the spectra of kuriloside H (**4**) (δC-20 69.4, δH-20 5.46 (dd, *J* = 6.1; 10.6 Hz)), containing (20S)-acetoxy-group. Hence, it was supposed that the attachment of the hydroxyl group to C-20 was in the aglycone of kuriloside I (**5**) instead of the acetoxy group in the aglycone of kuriloside H (**4**).

The ROE-correlations H-16/H-17 and H-16/H-32 indicated a 16*β*-OH orientation in the aglycone of kuriloside I (**5**). (20*S*)-configuration in **5** was determined on the base of the closeness of the coupling constant *J*20/17 = 9.5 Hz to those in the spectra of kurilosides A1, C1 [19], and H (**4**) and corroborated by the observed ROE-correlations H-17/H-21, H-20/H-18 and biogenetic background. Hence, kuriloside I (**5**) has an aglycone with a 16*β*,(20*S*)-dihydroxy-fragment that is unique in marine glycosides.

The (−)ESI-MS/MS of kuriloside I (**5**) demonstrated the fragmentation of the [M3Na − Na]<sup>−</sup> ion at *m/z* 1437.5. The peaks of fragment ions were observed at *m/z* 1317.4 [M3Na – Na − NaHSO4] <sup>−</sup>, 1197.4 [M3Na – Na − 2NaHSO4] <sup>−</sup>, 1173.4 [M3Na − Na − C6H9O8SNa(GlcOSO3)]−, 1039.4 [M3Na – Na − NaHSO4 − C7H11O8SNa(MeGlcOSO3)]−, 1027.3 [M3Na – Na − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui)]−, 907.3 [M3Na – Na − NaHSO4 − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui)]−, 895.4 [M3Na – Na − C6H9O8SNa (GlcOSO3) − C7H11O8SNa(MeGlcOSO3)]−, 667.4 [M3Na – Na − C24H39O2(Agl) − C6H9O8 SNa(GlcOSO3) − C6H10O4(Qui) − H]−, 519.0 [M3Na – Na − C24H39O3(Agl) − C6H9O8SNa (GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, and 417.1 [M3Na – Na − C24H39O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − NaHSO3] −, corroborating the structure of the glycoside.


**Table 4.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of carbohydrate moiety of kurilosides I (**5**) and I1 (**6**).

<sup>a</sup> Recorded at 176.04 MHz in C5D5N/D2O (4/1). <sup>b</sup> Bold = interglycosidic positions. <sup>c</sup> Italic = sulfate position. <sup>d</sup> Recorded at 700.13 MHz in C5D5N/D2O (4/1). Multiplicity by 1D TOCSY.

> All these data indicate that kuriloside I (**5**) is 3β-*O*-{6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)-[6-*O*-sodium sulfate-3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23, 24,25,26,27-hexa-nor-16β,(20*S*)-dihydroxy-lanost-9(11)-ene.

> The molecular formula of kuriloside I1 (**6**) was determined to be C58H91O37S3Na3 from the [M3Na − 2Na]2<sup>−</sup> ion peak at *m/z* 749.2148 (calc. 747.2155) and [M3Na − 3Na]3<sup>−</sup> ion peak at *m/z* 491.8146 (calc. 491.8139) in the (−)HR-ESI-MS. Kuriloside I1 (**6**) as well as kuriloside I (**5**) belong to one group because they have identical trisulfated pentasaccharide chains and, therefore, parts of the 1H and 13C NMR spectra corresponding to the carbohydrate chains are coincident (Table 4). 22,23,24,25,26,27-hexa-*nor*-lanostane aglycone of kuriloside I1 (**6**) is identical to that of kurilosides H (**4**), A1 and C1 [19] (Table S4) and characterized by the presence of 16*β*,(20*S*)-diacetoxy-fragment.


**Table 5.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of the aglycone moiety of kurilosides I (**5**) and K (**8**).

<sup>a</sup> Recorded at 176.03 MHz in C5D5N/D2O (4/1). <sup>b</sup> Recorded at 700.00 MHz in C5D5N/D2O (4/1).

The (−)ESI-MS/MS of **6** demonstrated the fragmentation of the [M3Na − Na]<sup>−</sup> ion at *m/z* 1521.4 and [M3Na − 2Na]2<sup>−</sup> ion at *m/z* 749.2. The peaks of fragment ions were observed at *m/z*: 1281.4 [M3Na – Na − 2CH3COOH − NaHSO4] <sup>−</sup>, 1197.4 [M3Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3)]−, 1137.4 [M3Na – Na − 2CH3COOH − C6H9O8SNa(GlcOSO3)]−, 859.4 [M3Na – Na − 2CH3COOH − C6H9O8SNa(GlcOSO3) − C7H11O8SNa(MeGlcOSO3)]−, 719.2 [M3Na − 2Na − CH3COOH]<sup>2</sup>−, 629.2 [M3Na − 2Na − NaHSO4] <sup>2</sup>−, and 557.2 [M3Na − 2Na − 2CH3COOH − C6H9O8SNa(GlcOSO3)]<sup>2</sup>−, which confirmed its structure, established by the NMR data.

All these data indicate that kuriloside I1 (**6**) is 3β-*O*-{6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)-*β*-D-quinovopyranosyl-(1→2)-[6-*O*-sodium sulfate-3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23, 24,25,26,27-hexa-*nor*-16β,(20*S*)-diacetoxy-lanost-9(11)-ene.

The molecular formula of kuriloside J (**7**) was determined to be C56H90O33S2Na2 from the [M2Na−Na]<sup>−</sup> ion peak at *m/z* 1377.4687 (calc. 1377.4709) and [M2Na−2Na]2<sup>−</sup> ion peak at *m/z* 677.2413 (calc. 677.2408) in the (−)HR-ESI-MS. In the 1H and 13C NMR spectra of the carbohydrate part of kuriloside J (**7**) (Table 6), five signals of anomeric protons at δ<sup>H</sup> 4.65–5.12 (d, *J* = 7.2–7.9 Hz) and corresponding five signals of anomeric carbons at δ<sup>C</sup> 102.0–104.7, deduced by the HSQC spectrum, were observed, which indicated the presence of a pentasaccharide chain similar to compounds **5** and **6**. Actually, the comparison of the 13C NMR spectra of sugar parts of kurilosides I (**5**) and J (**7**) revealed the closeness of the signals of four monosaccharide residues, except the signals of the third unit, attached to C-4 Qui2. The analysis of the signals of this residue in the 1H,1H-COSY, HSQC, 1D TOCSY, and ROESY spectra of kuriloside J (**7**) showed that it is a glucose without a sulfate group (δC-6 Glc3 61.8, δC-5 Glc3 77.7), while in the carbohydrate chain of **5**, this residue is sulfated. The other sulfate groups occupy the same positions at C-6 Glc4 (δC-6 Glc4 67.1, δC-5 Glc4 75.1) and at C-6 MeGlc5 (δC-6 MeGlc5 66.7, δC-5 MeGlc5 75.5) as in the sugar chains of kurilosides I (**5**) and I1 (**6**). The positions of interglycosidic linkages in the carbohydrate chain of **7**, elucidated by the ROESY and HMBC correlations (Table 6), were the same as in kurilosides of groups A [19] and I. Thus, kuriloside J (**7**) is a branched disulfated pentaoside with the sulfate groups bonding to C-6 Glc4 and C-6 MeGlc5 in the upper semi-chain.


**Table 6.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of carbohydrate moiety of kuriloside J (**7**).

<sup>a</sup> Recorded at 176.04 MHz in C5D5N/D2O (4/1). <sup>b</sup> Bold = interglycosidic positions. <sup>c</sup> Italic = sulfate position. <sup>d</sup> Recorded at 700.13 MHz in C5D5N/D2O (4/1). Multiplicity by 1D TOCSY.

> The analysis of the 1H and 13C NMR spectra of the aglycone part of kuriloside J (**7**) (Table 7) revealed the presence of the hexa-*nor*-lanostane aglycone having a 9(11)-double bond, similar to the majority of the other glycosides of *T. kurilensis* [19]. The signals at δ<sup>C</sup> 171.2 and 21.1 were characteristic for the acetoxy group, bonded to C-16, that was deduced from the characteristic δ<sup>C</sup> 75.1 value of C-16 and the ROE-correlation between

the signal of *O*-acetyl methyl group (δ<sup>H</sup> 2.17 (s)) and H-16 (δ<sup>H</sup> 5.76 (m). Actually, in the spectrum of **7**, the signal of C-16 was deshielded by 2.3 ppm due to the presence of the acetoxy-group when compared with the corresponding signal in the spectrum of kuriloside I (**5**), having a 16-hydroxy-group. The presence of hydroxyl group at C-20 was deduced from the characteristic signals at δ<sup>C</sup> 64.8 (C-20) and δ<sup>H</sup> 4.28 (dd, *J* = 6.4; 10.0 Hz, H-20). Hence, the hydroxyl group is attached to C-20 in the aglycones of kuriloside I (**5**) and J (**7**). The ROE-correlation H-16/H-32 indicated 16*β*-O-Ac orientation in the aglycone of kuriloside J (**7**), which was confirmed by the coupling constant *J*16/17 = 7.9 Hz, indicating both protons, H-16 and H-17, to be α [21]. (20*S*)-configuration in **7** was corroborated by the coupling constant *J*17/20 = 10.0 Hz and the ROE-correlations H-17/H-21, H-20/H-18. Hence, kuriloside J (**7**) is characterized by the new hexa-*nor*-lanostane aglycone with a 16*β*-acetoxy,(20*S*)-hydroxy-fragment.

**Table 7.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of the aglycone moiety of kuriloside J (**7**) and K1 (**9**).


<sup>a</sup> Recorded at 176.03 MHz in C5D5N/D2O (4/1). <sup>b</sup> Recorded at 700.00 MHz in C5D5N/D2O (4/1).

The (−)ESI-MS/MS of kuriloside J (**7**) demonstrated the fragmentation of [M2Na − Na]− ion at *m/z* 1377.5. The peaks of fragment ions were observed at *m/z* 1317.4 [M2Na – Na − CH3COOH]−, 1257.4 [M2Na – Na − NaHSO4] <sup>−</sup>, 1197.5 [M2Na – Na − CH3COOH − NaHSO4] <sup>−</sup>, 1155.4 [M2Na – Na − CH3COOH − C6H10O5 (Glc)]−, 1039.4 [M2Na – Na − CH3COOH − C7H11O8SNa(MeGlcOSO3)]−, 1009.4 [M2Na – Na − CH3COOH − C6H10O5(Glc) − C6H10O4(Qui)]−, 889.4 [M2Na − Na − NaHSO4 − CH3COOH − C6H10O5(Glc) − C6H10O4(Qui)]−, 667.4 [M2Na – Na − C26H41O3(Agl) − C6H10O5(Glc) −

C6H10O4(Qui) − H]−, 519.0 [M2Na – Na − C26H41O3(Agl) − C6H10O5(Glc) − C6H10O4 (Qui) − C5H8O4(Xyl) − H]−, 417.1 [M2Na – Na − C26H41O3(Agl) − C6H10O5(Glc) − C6H10O4 (Qui) − C5H8O4(Xyl) − NaHSO3] −, corroborating the structure of its aglycone and the carbohydrate chain.

All these data indicate that kuriloside J (**7**) is 3β-*O*-{β-D-glucopyranosyl-(1→4)-β-Dquinovopyranosyl-(1→2)-[6-*O*-sodium sulfate-3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6- *O*-sodium sulfate*-β*-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23,24,25,26,27-hexa*nor*-16β-acetoxy*,(*20*S*)-hydroxy-lanost-9(11)-ene.

The molecular formula of kuriloside K (**8**) was determined to be C54H88O32S2Na2 from the [M2Na − Na]<sup>−</sup> ion peak at *m/z* 1335.4573 (calc. 1335.4603) and the [M2Na − 2Na]2<sup>−</sup> ion peak at *m/z* 656.2357 (calc. 656.2356) in the (−)HR-ESI-MS. In the 1H and 13C NMR spectra of the carbohydrate part of kuriloside K (**8**) (Table 8), five signals of anomeric protons at δ<sup>H</sup> 4.62–5.19 (d, *J* = 6.5–8.5 Hz) and five signals of anomeric carbons at δ<sup>C</sup> 102.7–104.8, deduced by the HSQC spectrum, were indicative for the pentasaccharide chain with the *β*-configuration of glycosidic bonds. The comparison of the 13C NMR spectra of oligosaccharide parts of trisulfated kuriloside I (**5**) and kuriloside K (**8**) revealed the coincidence of the monosaccharide residues, except for the signals of a terminal, 3-*O*-methylglucose (MeGlc5) unit. The analysis of the signals of this residue in the 1H,1H-COSY, HSQC, 1D TOCSY, and ROESY spectra of kuriloside K (**8**) showed the absence of a sulfate group (δC-6 MeGlc5 61.6, δC-5 MeGlc5 77.5), in contrast with the carbohydrate chain of **5** (δC-6 MeGlc5 67.0, δC-5 MeGlc5 75.4). The positions of interglycosidic linkages in the carbohydrate chain of **8**, deduced by the ROESY and HMBC correlations (Table 8), showed that kuriloside K (**8**) has branching at C-4 Xyl1 in the disulfated pentasaccharide chain with the sulfate groups at C-6 Glc3 and C-6 Glc4.

The NMR spectra as well as the ROE-correlations of the aglycone part of kuriloside K (**8**) were coincident to that of kuriloside I (**5**), indicating the presence of a 22,23,24,25,26,27 hexa-*nor*-lanostane aglycone with 16β,(20*S*)-dihydroxy-fragment (Table 5).

The (−)ESI-MS/MS of **8** demonstrated the fragmentation of the [M2Na − Na]<sup>−</sup> ion at *m/z* 1335.4 resulted in the fragment ions observed at *m/z*: 1215.4 [M2Na – Na − NaHSO4] −, 1159.4 [M2Na – Na − C7H12O5(MeGlc)]−, 1071.4 [M2Na – Na − C6H9O8SNa(GlcOSO3)]−, 925.4 [M2Na – Na − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui)]−, 895.4 [M2Na – Na − C6H9O8SNa(GlcOSO3) − C7H12O5(MeGlc)]−, 713.3 [M2Na – Na − C24H39O2(Agl) − C6H9O8SNa(GlcOSO3) − H]−, 417.1 [M2Na – Na − C24H39O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [M2Na – Na − C24H39O3(Agl) − C6H9O8SNa (GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − C7H12O5(MeGlc) − H]−, which confirmed the chemical structure established by the NMR data.

All these data indicate that kuriloside K (**8**) is 3β-*O*-{6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)-[3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*sodium sulfate*-*β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23,24,25,26,27-hexa-*nor*-16*β,(*20*S*)-dihydroxy-lanost-9(11)-ene.

The molecular formula of kuriloside K1 (**9**) was determined to be C56H90O33S2Na2 from the [M2Na − Na]<sup>−</sup> ion peak at *m/z* 1377.4723 (calc. 1377.4709) and the [M2Na − 2Na]2<sup>−</sup> ion peak at *m/z* 677.2426 (calc. 677.2408) in the (−)HR-ESI-MS. The comparison of the 1H and 13C NMR spectra of the carbohydrate chains of kuriloside K1 (**9**) and kuriloside K (**8**) demonstrated their coincidence (Table 8) due to the presence of the same pentasaccharide, branched by C-4 Xyl1, sugar parts with the sulfate groups at C-6 Glc3 and C-6 Glc4. The analysis of the NMR spectra of the aglycone part of **9** indicated the presence of 22,23,24,25,26,27-hexa-*nor*-lanostane aglycone with 16*β*-acetoxy,(20*S*)-hydroxy-fragment (Table 7), identical to that of kuriloside J (**7**). Hence, kuriloside K1 (**9**) is an isomer of kuriloside J (**7**) by the position of one of the sulfate groups, that was confirmed by the presence of the ion-peaks having coincident *m/z* values in their (−)ESI-MS/MS spectra.


**Table 8.** 13C and 1H NMR chemical shifts, HMBC, and ROESY correlations of the carbohydrate moiety of kurilosides K (**8**) and K1 (**9**).

<sup>a</sup> Recorded at 176.04 MHz in C5D5N/D2O (4/1). <sup>b</sup> Bold = interglycosidic positions. <sup>c</sup> Italic = sulfate position. <sup>d</sup> Recorded at 700.13 MHz in C5D5N/D2O (4/1). Multiplicity by 1D TOCSY.

> The (−)ESI-MS/MS of **9** demonstrated the fragmentation of [M2Na − Na]<sup>−</sup> ion at *m/z* 1377.5. The peaks of fragment ions were observed at *m/z* 1317.4 [M2Na – Na − CH3COOH]−, 1197.5 [M2Na – Na − CH3COOH − NaHSO4] <sup>−</sup>, 1069.5 [M2Na – Na − C6H10O5(Glc)]−, 1053.4 [M2Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3)]−, 877.4 [M2Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3) − C7H12O5(MeGlc)]−, 731.3 [M2Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3) − C7H12O5(MeGlc) − C6H10O4(Qui)]−, 565.1 [M2Na – Na − C26H41O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − H]−, 417.1 [M2Na – Na − C26H41O4(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl)]−.

> All these data indicate that kuriloside K1 (**9**) is 3β-*O*-{6-*O*-sodium sulfate-β-D-glucopyranosyl-(1→4)-β-D-quinovopyranosyl-(1→2)-[3-*O*-methyl-β-D-glucopyranosyl-(1→3)-6-*O*sodium sulfate-β-D-glucopyranosyl-(1→4)]-β-D-xylopyranosyl}-22,23,24,25,26,27-hexa-*nor*-16β-acetoxy*,*(20*S*)-hydroxy-lanost-9(11)-ene.

When the studies on the glycosides of *T. kurilensis* were started [22], the complexity of glycosidic mixture became obvious. Therefore, the part of the glycosidic sum was subjected to solvolytic desulfation to facilitate the chromatographic separation and isolation of the glycosides. However, the obtained fraction of desulfated glycosides was separated only recently as part of the effort to discover some minor glycosides possessing interesting structural peculiarities. As a result, the compounds **10** and **11** were isolated (Figure 2). Their structures were elucidated by thorough analysis of 1D and 2D NMR spectra, similar to the natural compounds **1**–**9** and confirmed by the HR-ESI-MS.

**Figure 2.** Chemical structures of desulfated glycosides isolated from *Thyonidium kurilensis:* **10**—DS-kuriloside L; **11**—DSkuriloside M.

The molecular formula of DS-kuriloside L (**10**) was determined to be C41H64O15 from the [M − H]<sup>−</sup> ion peak at *m/z* 795.4169 (calc. 795.4172) in the (−)HR-ESI-MS. Compound **10** has a trisaccharide sugar chain (for NMR data see Tables S5 and S6, for original spectra see Figures S69–S76) and a hexa-*nor*-lanostane-type aglycone identical to that of kuriloside A2 [19].

The molecular formula of DS-kuriloside M (**11**) was determined to be C54H88O26 from the [M − H]<sup>−</sup> ion peak at *m/z* 1151.5469 (calc. 1151.5491) in the (−)HR-ESI-MS. DSkuriloside M (**11**), characterized by the 7(8)-double bond in the hexa-*nor*-lanostane nucleus and pentasaccharide chain, differed from the chains of kurilosides of the groups A, I, J, and K by the absence of sulfate groups (see Tables S7 and S8 for the NMR data, Figures S77–S85 for the original spectra). Noticeably, all of the isolated kurilosides, with the exception of **11**, contained a 9(11)-double bond in the polycyclic systems.

#### *2.2. Bioactivity of the Glycosides*

− Cytotoxic activities of compounds **1**–**9** against mouse neuroblastoma Neuro 2a, normal epithelial JB-6 cells, and erythrocytes were studied (Table 9). Known earlier cladoloside C was used as a positive control because it demonstrated a strong hemolytic effect [23]. Erythrocytes are an appropriate model for the studying of structure–activity relationships of the glycosides, since, despite many of them demonstrate hemolytic activity, the effect strongly depends on the structure of the compound. Normal epithelial JB-6 cells were used to search the compounds, not cytotoxic against this cell line, but having selective activity against other cells. Triterpene glycosides of sea cucumbers are known modulators of P2X receptors of immunocompetent cells when acting in nanomolar concentrations [24]. Neuroblastoma Neuro 2a cells are convenient model for the study of agonists/antagonists of P2X receptors—the targets in the treatment of selected nervous system diseases. Therefore, the activators, modulators, and blockers of purinergic receptors are of great interest [4] and the compounds demonstrating high cytotoxicity against Neuro 2a cells could be more deeply studied with the models of neurodegenerative diseases.


**Table 9.** The cytotoxic activities of glycosides **1**–**9** and cladoloside C (positive control) against mouse erythrocytes, neuroblastoma Neuro 2a cells, and normal epithelial JB-6 cells.

Kuriloside H (**4**), having a hexasaccharide trisulfated chain and the aglycone with acetoxy-groups at C(16) and C(20), was the most active compound in the series, demonstrating strong cytotoxicity against erythrocytes and JB-6 cells and a moderate effect against Neuro 2a cells. Kuriloside I1 (**6**), differing from **4** by the lack of a terminal glucose residue in the bottom semi-chain, was slightly less active. The effect of this glycoside is obviously explained by the presence of the acetoxy-group at C(20) in their aglycones, which compensates for the absence of a side chain, essential for the demonstration of the membranolytic action of the glycosides. Kurilosides J (**7**) and K1 (**9**), differing by the position of the second sulfate group attached to C(6) of different terminal monosaccharide residues, but having the same aglycones with 16*β*-acetoxy-group, were moderately cytotoxic against erythrocytes and JB-6 cells and had no any effect against Neuro 2a cells. However, the presence of the hydroxyl group in this position causes the loss of activity, so, the rest of compounds **1**−**3**, **5**, and **8** were not cytotoxic.

#### *2.3. Biosynthetic Pathways of the Glycosides*

The analysis of the structural peculiarities of the aglycones and carbohydrate chains of all the glycosides (kurilosides) found in the sea cucumber *T. kurilensis* allowed us to construct the metabolic network based on their biogenetic relationships. As a result, some biosynthetic pathways are taking shape (Figure 3).

Since the triterpene glycosides of sea cucumbers are the products of a mosaic type of biosynthesis [17], the carbohydrate chains and the aglycones are biosynthesized independently of each other. The main biosynthetic transformations of sugar parts of kurilosides are glycosylation and several rounds of sulfation that can be shifted in time relatively to each other (Figure 3). This has led to the formation of the set of compounds having 11 different oligosaccharide fragments. Meanwhile, there are some missing links (biosynthetic intermediates) in these biogenetic rows: biosides consisted of the glucose bonded to the xylose by *β*-(1→4)-glycosidic linkage, then triosides and tetraosides having glucose bonded to C(2) Xyl1—the precursors on kuriloside E, two types of disulfated hexaosides with a non-methylated terminal Glc4 unit that should biosynthetically appear between the carbohydrate chains of kurilosides of groups D and H; J and H; K and H, which have not so far been isolated. DS-kuriloside L (**10**) with a trisaccharide sugar chain is perfectly fit into the network as one of the initial stages of biosynthesis, illustrating the stepwise glycosylation of the synthesized chain. The structure of its sugar chain as well as the chain of kuriloside C1 [19] suggests the glycosylation of C(4) Xyl1 and initialization of the growth of the upper semi-chain precedes the glycosylation of C(2) Xyl1. There are some branchpoints of the biosynthetic pathways where the processes of sulfation and glycosylation or sulfation and methylation are alternative/concurrent. The final product of such transformations is the trisulfated hexaoside kuriloside H (**4**), the most biologically active compound in the series

(Table 9), which can be formed by different pathways, and is a characteristic feature of a mosaic type of biosynthesis. However, this glycoside is minor (0.9 mg) in the glycosidic sum of *T. kurilensis*, while the main compounds are kurilosides of group A (~150 mg), and these carbohydrate chains can be considered as the most actively metabolized and resulted in the formation of at least three different types of sugar chains (kurilosides of the groups D, J, and K). Thus, their formation is a mainstream of the biosynthesis of carbohydrate chains of the glycosides of *T. kurilensis*.

**Figure 3.** The metabolic network of the carbohydrate chains of the glycosides from *T. kurilensis.*

As for the directions of biosynthesis of the aglycone parts of kurilosides (Figure 4), the scheme presented earlier [19] was complemented by some structures found recently, representing intermediate biosynthetic stages. DS-kuriloside M (**11**) is the only glycoside from *T. kurilensis* characterized by the 7(8)-double bond in the lanostane nucleus, when all the other kurilosides contain a 9(11)-double bond in the polycyclic systems. This finding indicates the existence of two oxidosqualene cyclases (OSCs)—enzymes converted 2,3-oxidosqualene into different triterpene alcohols giving rise various skeletons of the aglycones—in this species of sea cucumbers. These data are in good agreement with the results of the investigations of the genes coding OSCs in the other species of the sea cucumbers—*Eupentacta fraudatrix* [25], *Stichopus horrens* [26], and *Apostichopus japonicus* [27], demonstrating that even when the glycosides preferably contain the aglycones with one certain position of intra-nucleus double bond (Δ7(8)-aglycones in *E. fraudatrix* [13,18] and *S. horrens* [28,29], and Δ9(11)-aglycones in *A. japonicus* [30,31]), the genes of at least two OSCs, producing aglycone precursors with different double bond positions, are expressed, albeit with different efficiency.

**Figure 4.** The biosynthetic pathways to aglycones of glycosides from *T. kurilensis.*

The constituent hexa-*nor*-lanostane aglycones of kurilosides are biosynthesized via the oxidative cleavage of the side chain from the precursors having normal side chains (for example, kurilosides D [19] and D1 (**2**)) and oxygen-containing substituents at C-20 and C-22 (Figure 4). As result, the aglycone of kuriloside E [19] was formed. The subsequent biosynthetic transformations of the aglycones can occur in two directions. The first one started from the reduction of the C-20-oxo-group to the hydroxy-group, followed by the oxidation of C-16 to the hydroxy-group with the formation of the aglycones of kurilosides I (**5**) and K (**8**). It is important that the latter reaction is carried out by the cytochrome P450 monooxygenase selectively bonding to the *β*-hydroxy-group to C-16 in the derivatives containing the hydroxy-group at C-20. The next steps lead to the acetylation of hydroxyl group at C-16 (as in the aglycones of kurilosides J (**7**) and K1 (**9**)) followed by the acetylation of the hydroxyl group at C-20 (the aglycones of kurilosides A1, C1, H (**4**), and I1 (**6**) correspond to this conversion). Obviously, the oxidation of C-16 precedes the acetylation of C-20 since no aglycones with a 16-hydroxy,20-acetoxy-fragment have been found.

The second direction of the aglycone biosynthesis occurs through the introduction of the α-hydroxyl group to C-16, resulting in the formation of aglycone of kurilosides A3 (**1**), G (**3**), and F [19]. Moreover, the transformation leading to hexa-*nor*-lanostane aglycones having a 16α-hydroxy,20-oxo-fragment is the same in the biosynthetic precursors with 7(8)- and 9(11)-double bonds, which is confirmed by the aglycone structure of **11**. Subsequent acetylation of the 16*α-*OH-group leads to the aglycone of kuriloside A, while intramolecular dehydration to the aglycone of kuriloside A2 and DS-kuriloside L (**10**). Therefore, an *α*-hydroxy-group was selectively introduced to C-16 of the 20-oxo-lanostane precursors.

#### **3. Materials and Methods**

#### *3.1. General Experimental Procedures*

Specific rotation, Perkin-Elmer 343 Polarimeter (Perkin-Elmer, Waltham, MA, USA); NMR, Bruker Avance III 700 Bruker FT-NMR (Bruker BioSpin GmbH, Rheinstetten, Germany) (700.00/176.03 MHz) (1H/13C) spectrometer; ESI MS (positive and negative ion modes), Agilent 6510 Q-TOF apparatus (Agilent Technology, Santa Clara, CA, USA), sample concentration 0.01 mg/mL; HPLC, Agilent 1260 Infinity II with a differential refractometer

(Agilent Technology, Santa Clara, CA, USA); column Phenomenex Synergi Fusion RP (10 × 250 mm, 5 μm) (Phenomenex, Torrance, CA, USA).

#### *3.2. Animals and Cells*

Specimens of the sea cucumber *Thyonidium* (*=Duasmodactyla*) *kurilensis* (Levin) (family Cucumariidae; order Dendrochirotida) were collected in August 1990 using an industrial rake-type dredge in the waters of Onekotan Island (Kurile Islands, the Sea of Okhotsk) at a depth of 100 m by the medium fishing refrigerator trawler "Breeze" with a rear scheme of trawling during scallop harvesting. The sea cucumbers were identified by Prof. V.S. Levin; voucher specimens are preserved at the A.V. Zhirmunsky National Scientific Center of Marine Biology, Vladivostok, Russia.

CD-1 mice, weighing 18–20 g, were purchased from RAMS 'Stolbovaya' nursery (Stolbovaya, Moscow District, Russia) and kept at the animal facility in standard conditions. All experiments were performed following the protocol for animal study approved by the Ethics Committee of the Pacific Institute of Bioorganic Chemistry No. 0085.19.10.2020. All experiments were conducted in compliance with all of the rules and international recommendations of the European Convention for the Protection of Vertebrate Animals Used for Experimental Studies.

Mouse epithelial JB-6 cells Cl 41-5a and mouse neuroblastoma cell line Neuro 2a (ATCC ® CCL-131) were purchased from ATCC (Manassas, VA, USA).

#### *3.3. Extraction and Isolation*

The extract of the glycosides, obtained by the standard procedure, and the initial stages of their separation were discussed in a previous paper [19]. As result of the chromatography on Si gel columns using CHCl3/EtOH/H2O (100:125:25) as the mobile phase, the fractions II–V were obtained, which were subjected to HPLC on a Phenomenex Synergi Fusion RP (10 × 250 mm) column. The separation of fraction II with MeOH/H2O/NH4OAc (1 M water solution) (63/35/2) as the mobile phase resulted in the isolation of individual kuriloside A3 (**1**) (79.2 mg). HPLC of fraction III with MeOH/H2O/NH4OAc (1 M water solution) (60/38/2) as the mobile phase gave 3.1 mg of kuriloside K1 (**9**) and 0.9 mg of kuriloside K (**8**). Fraction IV was the result of the HPLC using MeOH/H2O/NH4OAc (1 M water solution) (68/31/1) as the mobile phase was separated to the subfractions 1–7. Further rechromatography of subfraction 7 with MeOH/H2O/NH4OAc (1 M water solution) (63/34/3) followed by (60/37/3) as the mobile phases gave 3.1 mg of kuriloside I1 (**6**). The use of the ratio of MeOH/H2O/NH4OAc (1 M water solution) (62/35/3) for subfraction 4 gave 2.3 mg of kuriloside J (**7**) and the ratio (58/39/3) for subfraction 3 gave 7 mg of kuriloside D1 (**2**). For the HPLC of the most polar fraction V, obtained after Si gel chromatography, the ratio of the same solvents (60/39/1) was applied, which led to the isolation of 10 subfractions. Some of them were minor, thus only the main ones were submitted for further separation. For subfraction 10, the ratio (64/34/2) was applied to give 0.9 mg of kuriloside H (**4**). The ratio (54/43/3) used for HPLC of subfraction 4 gave 1.9 mg of kuriloside G (**3**) and 2.3 mg of kuriloside I (**5**).

The fraction of desulfated derivatives obtained earlier by the standard methodology (~350 mg) was submitted to column chromatography on Si gel using CHCl3/EtOH/H2O (100:50:4) and CHCl3/MeOH/H2O (250:75:3) as mobile phases to give subfractions DS-1−DS-8, which were subsequently subjected to HPLC on the same column as compounds **1**–**9**. Individual DS-kuriloside M (**11**) (3.8 mg) was isolated as a result of separating the subfraction DS-6 with 66% MeOH as the mobile phase which gave several fractions, followed by the HPLC of one of them with 32% CH3CN as the mobile phase. HPLC of subfraction DS-2 with 50% CH3CN as the mobile phase, followed by 46% CH3CN as the mobile phase, gave 4.0 mg of DS-kuriloside L (**10**).

#### 3.3.1. Kuriloside A3 (**1**)

Colorless powder; [*α*] 20 <sup>D</sup> −1◦ (*c* 0.1, 50% MeOH). NMR: See Tables S1 and S2, Figures S1–S6. (−)HR-ESI-MS *m/z*: 1231.5063 (calc. 1231.5059) [MNa − Na]−; (−)ESI-MS/MS *m/z*: 1069.5 [MNa – Na − C6H10O5(Glc)]−, 1055.4 [MNa − Na–C7H12O5(MeGlc)]−, 923.4 [MNa − Na–C6H10O5(Glc) − C6H10O4(Qui)]−, 747.3 [MNa − Na–C6H10O5(Glc) − C6H10O4(Qui) − C7H12O5(MeGlc)]−, 695.1 [MNa – Na − C24H37O3(Agl) − C6H10O5(Glc) − H]−, 565.1 [MNa − Na–C24H37O2(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − H]−, 549.1 [MNa − Na– C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − H]−, 417.1 [MNa − Na–C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [MNa − Na–C24H37O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) – C7H12O5(MeGlc) − H]−.

#### 3.3.2. Kuriloside D1 (**2**)

Colorless powder; [*α*] 20 <sup>D</sup> –39◦ (*c* 0.1, 50% MeOH). NMR: See Table 1 and Table S3, Figures S7–S13. (−)HR-ESI-MS *m/z*: 1507.6291 (calc. 1507.6268) [MNa − Na]−; (−)ESI-MS/MS *m/z*: 1349.5 [MNa – Na − C8H15O3 + H]−, 1187.5 [MNa – Na − C8H15O3 −C6H10O5 (Glc) + H]−, 1025.4 [MNa – Na − C8H15O3 − C6H10O5(Glc) − C6H10O5(Glc) + H]−, 879.4 [MNa – Na − C8H15O3 −C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) + H]−, 565.1 [MNa – Na − C30H47O4(Agl) − C6H10O5(Glc) − C6H10O5(Glc) −C6H10O4(Qui) − H]−, 417.1 [MNa – Na − C30H47O5(Agl) − C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [MNa – Na − C30H47O5(Agl) − C6H10O5(Glc) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − C7H12O5(MeGlc) − H]−.

#### 3.3.3. Kuriloside G (**3**)

Colorless powder; [*α*] 20 <sup>D</sup> −2◦ (*c* 0.1, 50% MeOH). NMR: See Table 2 and Table S2, Figures S14–S22. (−)HR-ESI-MS *m/z*: 1509.5102 (calc. 1509.5132) [M2Na − Na]−; 743.2624 (calc. 743.2626) [M2Na − 2Na]2−, (−)ESI-MS/MS *m/z*: 1389.6 [M2Na – Na − NaHSO4] −, 1333.5 [M2Na – Na − C7H12O5(MeGlc)]−, 1231.5 [M2Na – Na − C7H11O8SNa(MeGlcSO3Na)]−, 1069.4 [M2Na – Na − C7H11O8SNa(MeGlcSO3Na) − C6H10O5(Glc)]−, 923.4 [M2Na –Na − C7H11O8SNa(MeGlcSO3Na) − C6H10O5(Glc)] − C6H10O4(Qui)]−.

#### 3.3.4. Kuriloside H (**4**)

Colorless powder; [*α*] 20 <sup>D</sup> −3◦ (*c* 0.1, 50% MeOH). NMR: See Table 3 and Table S4, Figures S23–S31. (−)HR-ESI-MS *m/z*: 1683.4701 (calc. 1683.4730) [M3Na − Na]−, 830.2425 (calc. 830.2419) [M3Na − 2Na]2−, 545.8332 (calc. 545.8315) [M3Na − 3Na]3−; (−)ESI-MS/MS *m/z*: 1503.5 [M3Na – Na − CH3COOH − NaHSO4] <sup>−</sup>, 1443.5 [M3Na – Na − 2CH3COOH − NaHSO4] <sup>−</sup>, 1281.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C6H10O5(Glc)]−, 1165.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C7H11O8SNa(MeGlcOSO3)]−, 1003.4 [M3Na – Na − 2CH3COOH − NaHSO4 − C7H11O8SNa(MeGlcOSO3) − C6H10O5 (Glc)]−.

#### 3.3.5. Kuriloside I (**5**)

Colorless powder; [*α*] 20 <sup>D</sup> −9◦ (*c* 0.1, 50% MeOH). NMR: See Tables 4 and 5, Figures S32–S40. (−)HR-ESI-MS *m/z*: 1437.3952 (calc. 1437.3991) [M3Na − Na]−, 707.2049 (calc. 707.2049) [M3Na − 2Na]2−, 463.8076 (calc. 463.8069) [M3Na − 3Na]3−; (−)ESI-MS/MS *m/z*: 1317.4 [M3Na – Na − NaHSO4] <sup>−</sup>, 1197.4 [M3Na – Na − 2NaHSO4] −, 1173.4 [M3Na – Na − C6H9O8SNa(GlcOSO3)]−, 1039.4 [M3Na – Na − NaHSO4 − C7H11O8SNa(MeGlcOSO3)]−, 1027.3 [M3Na – Na − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui)]−, 907.3 [M3Na – Na − NaHSO4 − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui)]−, 895.4 [M3Na – Na − C6H9O8SNa (GlcOSO3) − C7H11O8SNa (MeGlcOSO3)]−, 667.4 [M3Na – Na − C24H39O2(Agl) − C6H9O8 SNa(GlcOSO3) − C6H10O4(Qui) − H]−, 519.0 [M3Na – Na − C24H39O3(Agl) − C6H9O8SNa (GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 417.1 [M3Na – Na − C24H39O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − NaHSO3] −.

#### 3.3.6. Kuriloside I1 (**6**)

Colorless powder; [*α*] 20 <sup>D</sup> −5◦ (*c* 0.1, 50% MeOH). NMR: See Table 4 and Table S4, Figures S41–S47. (−)HR-ESI-MS *m/z*: 1749.2148 (calc. 747.2155) [M3Na − 2Na]2−, 491.8146 (calc. 491.8139) [M3Na − 3Na]3−; (−)ESI-MS/MS *m/z*: 1281.4 [M3Na – Na − 2CH3COOH − NaHSO4] <sup>−</sup>, 1197.4 [M3Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3)]−, 1137.4 [M3Na – Na − 2CH3COOH − C6H9O8SNa (GlcOSO3)]−, 859.4 [M3Na – Na − 2CH3COOH − C6H9O8SNa(GlcOSO3) − C7H11O8SNa(MeGlcOSO3)]−, 719.2 [M3Na − 2Na − CH3COOH]2<sup>−</sup>, 629.2 [M3Na − 2Na − NaHSO4] <sup>2</sup>−, 557.2 [M3Na − 2Na − 2CH3COOH − C6H9O8SNa (GlcOSO3)]<sup>2</sup>−.

#### 3.3.7. Kuriloside J (**7**)

Colorless powder; [*α*] 20 <sup>D</sup> −10◦ (*c* 0.1, 50% MeOH). NMR: See Tables 6 and 7, Figures S48–S56. (−)HR-ESI-MS *m/z*: 1377.4687 (calc. 1377.4709) [M2Na − Na]−, 677.2413 (calc. 677.2408) [M2Na − 2Na]2<sup>−</sup>; (−)ESI-MS/MS *m/z*: 1317.4 [M2Na – Na − CH3COOH]−, 1257.4 [M2Na – Na − NaHSO4] <sup>−</sup>, 1197.5 [M2Na – Na − CH3COOH − NaHSO4] −, 1155.4 [M2Na – Na − CH3COOH − C6H10O5(Glc)]−, 1039.4 [M2Na – Na − CH3COOH − C7H11O8 SNa(MeGlcOSO3)]−, 1009.4 [M2Na – Na − CH3COOH − C6H10O5(Glc) − C6H10O4(Qui)]−, 889.4 [M2Na – Na − NaHSO4 − CH3COOH − C6H10O5(Glc) − C6H10O4(Qui)]−, 667.4 [M2Na – Na − C26H41O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − H]−, 519.0 [M2Na – Na − C26H41O3(Agl) − C6H10O5(Glc) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 417.1 [M2Na – Na − C26H41O3(Agl) − C6H10O5 (Glc) − C6H10O4 (Qui) − C5H8O4 (Xyl) − NaHSO3] −.

#### 3.3.8. Kuriloside K (**8**)

Colorless powder; [*α*] 20 <sup>D</sup> −7◦ (*c* 0.1, 50% MeOH). NMR: See Tables 5 and 8, Figures S57–S65. (−)HR-ESI-MS *m/z*: 1335.4573 (calc. 1335.4603) [M2Na − Na]−, 656.2357 (calc. 656.2356) [M2Na − 2Na]2<sup>−</sup>; (−)ESI-MS/MS *m/z*: 1215.4 [M2Na – Na − NaHSO4] −, 1159.4 [M2Na – Na − C7H12O5(MeGlc)]−, 1071.4 [M2Na – Na − C6H9O8SNa(GlcOSO3)]−, 925.4 [M2Na – Na − C6H9O8SNa (GlcOSO3) − C6H10O4 (Qui)]−, 895.4 [M2Na – Na − C6H9O8SNa(GlcOSO3) − C7H12O5(MeGlc)]−, 713.3 [M2Na – Na − C24H39O2(Agl) − C6H9O8SNa(GlcOSO3) − H]−, 417.1 [M2Na – Na − C24H39O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − H]−, 241.0 [MNa – Na − C24H39O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl) − C7H12O5(MeGlc) − H]−.

#### 3.3.9. Kuriloside K1 (**9**)

Colorless powder; [*α*] 20 <sup>D</sup> −4◦ (*c* 0.1, 50% MeOH). NMR: See Tables 7 and 8, Figures S66–S68. (−)HR-ESI-MS *m/z*: 1377.4723 (calc. 1377.4709) [M2Na − Na]−, 677.2426 (calc. 677.2408) [M2Na − 2Na]2−; (−)ESI-MS/MS *m/z*: 1317.4 [M2Na – Na − CH3COOH]−, 1197.5 [M2Na – Na − CH3COOH − NaHSO4] <sup>−</sup>, 1069.5 [M2Na – Na − C6H10O5 (Glc)]−, 1053.4 [M2Na – Na − CH3COOH − C6H9O8SNa(GlcOSO3)]−, 877.4 [M2Na – Na − CH3COOH −C6H9O8SNa(GlcOSO3) − C7H12O5(MeGlc)]−, 731.3 [M2Na – Na − CH3COOH − C6H9O8 SNa(GlcOSO3) − C7H12O5(MeGlc) − C6H10O4(Qui)]−, 565.1 [M2Na – Na − C26H41O3(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − H]−, 417.1 [M2Na – Na − C26H41O4(Agl) − C6H9O8SNa(GlcOSO3) − C6H10O4(Qui) − C5H8O4(Xyl)]−.

#### *3.4. Cytotoxic Activity (MTT Assay)*

All compounds (including cladoloside C used as the positive control) were tested in concentrations from 1.5 μM to 100 μM using two-fold dilution in dH2O. The solutions (20 μL) of tested substances in different concentrations and cell suspension (180 μL) were added in wells of 96-well plates (1 × 104 cells/well) and incubated 24 h at 37 ◦C and 5% CO2. After incubation, the medium with tested substances was replaced by 100 μL of fresh medium. Then, 10 μL of MTT (thiazoyl blue tertrazolium bromide) stock solution (5 mg/mL) was added to each well and the microplate was incubated for 4 h. After that, 100 μL of SDS-HCl solution (1 g SDS/10 mL dH2O/17 μL 6 N HCl) was added to each well followed by incubation for 4–18 h. The absorbance of the converted dye formazan

was measured using a Multiskan FC microplate photometer (Thermo Fisher Scientific, Waltham, MA, USA) at a wavelength of 570 nm. Cytotoxic activity of the substances was calculated as the concentration that caused 50% metabolic cell activity inhibition (IC50). All the experiments were made in triplicate, *p* < 0.01.

#### *3.5. Hemolytic Activity*

Blood was taken from CD-1 mice (18–20 g). Erythrocytes were isolated from the blood of albino CD-1 mice by centrifugation with phosphate-buffered saline (pH 7.4) for 5 min at 4 ◦C by 450× *g* on a LABOFUGE 400R (Heraeus, Hanau, Germany) centrifuge for three times. Then, the residue of erythrocytes was resuspended in ice cold phosphate saline buffer (pH 7.4) to a final optical density of 1.5 at 700 nm, and kept on ice. For the hemolytic assay, 180 μL of erythrocyte suspension was mixed with 20 μL of test compound solution (including cladoloside C used as the positive control) in V-bottom 96-well plates. After 1 h of incubation at 37 ◦C, plates were exposed to centrifugation for 10 min at 900× *g* on a LMC-3000 (Biosan, Riga, Latvia) laboratory centrifuge. Then, we carefully selected 100 μL of supernatant and transferred it to new flat-plates respectively. Lysis of erythrocytes was determined by measuring the concentration of hemoglobin in the supernatant with a microplate photometer Multiskan FC (Thermo Fisher Scientific, Waltham, MA, USA), λ = 570 nm. The effective dose causing 50% hemolysis of erythrocytes (ED50) was calculated using the computer program SigmaPlot 10.0. All experiments were made in triplicate, *p* < 0.01.

#### *3.6. Solvolytic Desulfation*

A part of the glycosidic sum (350 mg) was dissolved in a mixture of pyridine/dioxane (1/1) and refluxed for 1 h. The obtained mixture was concentrated in vacuo and subsequently purified by using Si gel column chromatography (as depicted in the Section 3.3).

#### **4. Conclusions**

Thus, nine unknown earlier triterpene glycosides were isolated from the sea cucumber *Thyonidium (=Duasmodactyla) kurilensis* in addition to the series of kurilosides found recently [19]. Five new types of the carbohydrate chains (kurilosides of the groups G–K) were discovered. There were trisulfated penta- (kurilosides of the group I (**5**, **6**)) and hexaosides (kuriloside H (**4**)) among them. Kuriloside H (**4**) is the second example of the most polar triterpene glycosides, along with tetrasulfated pentaosides found earlier in the sea cucumber *Psolus fabricii* [20]. The structures of disulfated hexa- and pentasaccharide chains of kurilosides of the groups G (**3**), J (**7**), and K (**8**, **9**) clearly illustrate a combinatorial (mosaic) type of biosynthesis of the glycosides, namely, the positions of the sulfate group attachment. At the same time, the position of one of the sulfate groups (at C(6) Glc, attached to C(4) Xyl1) remained the same in all glycosides found in this species. Three new non-holostane aglycones lacking a lactone ring, two of them being the 22,23,24,25,26,27-hexa-*nor*-lanostane type and one having a normal side chain, were found in glycosides **1**–**9**. The majority of the aglycones of *T. kurilensis* glycosides differed from each other in the substituents at C-16 (*α*- and *β*-oriented hydroxy- or acetoxy groups, or keto-group) and C-20 (hydroxy-, acetoxy-, or keto-groups), representing the biogenetically related rows of the compounds. As mentioned in a previous paper [19], the glycosides with 16*α*-substituents were isolated from *T. kurilensis* only. The finding of 16*β*-hydroxylated aglycones is also for the first time. Such compounds can be considered as "hot metabolites", biosynthetic intermediates or precursors of the aglycones with the 16*β*-acetoxy-group.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/md19040187/s1. Table S1. NMR spectrometric data of the carbohydrate moiety of kuriloside A3 (**1**); Table S2. NMR spectrometric data of the aglycone moiety of kurilosides A3 (**1**) and G (**3**); Table S3. NMR spectrometric data of the carbohydrate moiety of kuriloside D1 (**2**); Table S4. NMR spectrometric data of the aglycone moiety of kurilosides H (**4**) and I1 (**6**); Table S5. NMR spectrometric data of the carbohydrate moiety of DS-kuriloside L (**10**); Table S6. NMR spectrometric data of the

aglycone moiety of **10**; Table S7. NMR spectrometric data of the carbohydrate moiety of DS-kuriloside M (**11**); Table S8. NMR spectrometric data of the aglycone moiety of **11**; Figure S1. 1H NMR and 13C NMR spectra of **1**; Figure S2. COSY spectrum of **1**; Figure S3. HSQC spectrum of **1**; Figure S4. ROESY spectrum of **1**; Figure S5. HMBC spectrum of **1**; Figure S6. HR-ESI-MS and ESI-MS/MS spectra of **1**; Figure S7. 13C NMR spectrum of **2**; Figure S8. 1H NMR spectrum of **2**; Figure S9. COSY spectrum of **2**; Figure S10. HSQC spectrum of **2**; Figure S11. HMBC spectrum of **2**; Figure S12. ROESY spectrum of **2**; Figure S13. HR-ESI-MS and ESI-MS/MS spectra of **2**; Figure S14. 13C NMR spectrum of **3**; Figure S15. 1H NMR spectrum of **3**; Figure S16. COSY spectrum of **3**; Figure S17. HSQC spectrum of **3**; Figure S18. HMBC spectrum of **3**; Figure S19. ROESY spectrum of **3**; Figure S20. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **3**; Figure S21. 1 D TOCSY spectra of the MeGlc4, Glc5 and MeGlc6 of **3**; Figure S22. HR-ESI-MS and ESI-MS/MS spectra of **3**; Figure S23. 13C NMR spectrum of **4**; Figure S24. 1H NMR spectrum of **4**; Figure S25. COSY spectrum of **4**; Figure S26. HSQC spectrum of **4**; Figure S27. ROESY spectrum of **4**; Figure S28. HMBC spectrum of **4**; Figure S29. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **4**; Figure S30. 1 D TOCSY spectra of Glc4, Glc5 and MeGlc6 of **4**; Figure S31. HR-ESI-MS and ESI-MS/MS spectra of **4**; Figure S32. 13C NMR spectrum of kuriloside I (**5**); Figure S33. 1H NMR spectrum of **5**; Figure S34. COSY spectrum of **5**; Figure S35. HSQC spectrum of **5**; Figure S36. HMBC spectrum of **5**; Figure S37. ROESY spectrum of **5**; Figure S38. 1D TOCSY spectra of Xyl1, Qui2 and Glc3 of **5**; Figure S39. 1D TOCSY spectra of Glc4 and MeGlc5 of **5**; Figure S40. HR-ESI-MS and ESI-MS/MS spectra of **5**; Figure S41. 13C NMR spectrum of **6**; Figure S42. 1H NMR spectrum of **6**; Figure S43. COSY spectrum of **6**; Figure S44. HSQC spectrum of **6**; Figure S45. ROESY spectrum of **6**; Figure S46. HMBC spectrum of **6**; Figure S47. HR-ESI-MS and ESI-MS/MS spectra of **6**; Figure S48. 13C NMR spectrum of kuriloside J (**7**); Figure S49. 1H NMR spectrum of **7**; Figure S50. COSY spectrum of **7**; Figure S51. HSQC spectrum of **7**; Figure S52. HMBC spectrum of **7**; Figure S53. ROESY spectrum of **7**; Figure S54. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **7**; Figure S55. 1 D TOCSY spectra of Glc4 and MeGlc5 of **7**; Figure S56. HR-ESI-MS and ESI-MS/MS spectra of **7**; Figure S57. 13C NMR spectrum of kuriloside K (**8**); Figure S58. 1H NMR spectrum of **8**; Figure S59. COSY spectrum of **8**; Figure S60. HSQC spectrum of **8**; Figure S61. HMBC spectrum of **8**; Figure S62. ROESY spectrum of **8**; Figure S63. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **8**; Figure S64. 1 D TOCSY spectra of Glc4 and MeGlc5 of **8**; Figure S65. HR-ESI-MS and ESI-MS/MS spectra of **8**; Figure S66. 13C NMR spectrum of **9**; Figure S67. 1H NMR spectrum of **9** in; Figure S68. HR-ESI-MS and ESI-MS/MS spectra of **9**; Figure S69. 13C NMR spectrum of DS-kuriloside L (**10**); Figure S70. 1H NMR spectrum of **10**; Figure S71. COSY spectrum of **10**; Figure S72. HSQC spectrum of **10**; Figure S73. HMBC spectrum of **10**; Figure S74. ROESY spectrum of **10**; Figure S75. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **10**; Figure S76. HR-ESI-MS (−) and ESI-MS/MS spectra of **10**; Figure S77. 13C NMR spectrum of DS-kuriloside M (**11**); Figure S78. 1H NMR spectrum of **11**; Figure S79. COSY spectrum of **11**; Figure S80. HSQC spectrum of **11**; Figure S81. HMBC spectrum of **11**; Figure S82. ROESY spectrum of **11**; Figure S83. 1 D TOCSY spectra of Xyl1, Qui2 and Glc3 of **11**; Figure S84. 1 D TOCSY spectra of Glc4 and MeGlc5 of **11**; Figure S85. HR-ESI-MS (−), ESI-MS/MS spectra of **11**.

**Author Contributions:** Conceptualization, A.S.S. and V.I.K.; Methodology, A.S.S. and S.A.A.; Investigation, A.S.S., A.I.K., S.A.A., R.S.P., P.S.D., E.A.C. and P.V.A.; Writing—original draft preparation, A.S.S. and V.I.K.; Writing—review and editing, A.S.S. and V.I.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** The investigation was carried out with the financial support of a grant from the Ministry of Science and Education, Russian Federation 13.1902.21.0012 (075-15-2020-796) (isolation of individual triterpene glycosides) and a grant from the Russian Foundation for Basic Research No. 19-04-000-14 (elucidation of structures of the glycosides and their biotesting).

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki, and approved by the Ethics Committee of the Pacific Institute of Bioorganic Chemistry (Protocol No. 0085.19.10.2020).

**Acknowledgments:** The study was carried out on the equipment of the Collective Facilities Center "The Far Eastern Center for Structural Molecular Research (NMR/MS) PIBOC FEB RAS". The authors are very appreciative to Professor Valentin A. Stonik (PIBOC FEB RAS, Vladivostok, Russia) for reading and discussion of the manuscript.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Review* **Application of MS-Based Metabolomic Approaches in Analysis of Starfish and Sea Cucumber Bioactive Compounds**

**Roman S. Popov \*, Natalia V. Ivanchina and Pavel S. Dmitrenok \***

G.B. Elyakov Pacific Institute of Bioorganic Chemistry, Far Eastern Branch of Russian Academy of Sciences, 159 Prospect 100-let Vladivostoku, Vladivostok 690022, Russia; ivanchina@piboc.dvo.ru

**\*** Correspondence: popov\_rs@piboc.dvo.ru (R.S.P.); paveldmt@piboc.dvo.ru (P.S.D.);

Tel.: +7-423-231-1132 (P.S.D.)

**Abstract:** Today, marine natural products are considered one of the main sources of compounds for drug development. Starfish and sea cucumbers are potential sources of natural products of pharmaceutical interest. Among their metabolites, polar steroids, triterpene glycosides, and polar lipids have attracted a great deal of attention; however, studying these compounds by conventional methods is challenging. The application of modern MS-based approaches can help to obtain valuable information about such compounds. This review provides an up-to-date overview of MS-based applications for starfish and sea cucumber bioactive compounds analysis. While describing most characteristic features of MS-based approaches in the context of starfish and sea cucumber metabolites, including sample preparation and MS analysis steps, the present paper mainly focuses on the application of MS-based metabolic profiling of polar steroid compounds, triterpene glycosides, and lipids. The application of MS in metabolomics studies is also outlined.

**Keywords:** starfish; sea cucumber; polyhydroxysteroids; triterpene glycosides; steroid glycosides; lipids; mass spectrometry; metabolomics; metabolomic profiling

#### **1. Introduction**

The emergence of novel diseases and resistant forms of known diseases in recent years and the emergence of multidrug-resistant pathogens has led to a renewed interest in the exploration of new sources of bioactive compounds. The sea environment possesses extraordinary ecological variety, and its inhabitants exhibit enormous biochemical diversity. At present, over 29,000 marine natural products have been discovered [1–4]. To date, several dozen marine natural products or their derivatives have been approved as therapeutic agents or are undergoing Phase III, II, or I drug development [5]. As the biodiversity of marine organisms is higher than that of terrestrial plants and animals, and as only a minor portion of metabolites present in marine species has been studied, it can be assumed that the number of new marine compounds will continue to increase, providing new therapeutic alternatives.

Marine invertebrates have long been considered an inexhaustible source of novel natural products. Although Porifera and Cnidaria are the two major sources of new marine natural products, Echinodermata is viewed as another abundant source of new bioactive compounds. Over the past five years, just over two hundred new compounds have been isolated from echinoderms [1–4,6]. The phylum Echinodermata includes about 7500 species found in all seas at every depth, from intertidal to abyssal, and in all ecosystems, from coral reefs to shallow shores. Echinoderms are divided into five different taxonomic classes, including Asteroidea (starfish) and Holothuroidea (sea cucumbers).

Starfish and sea cucumbers are extensively employed in traditional medicine, being a rich source of bioactive compounds. Several starfish species are used to treat rheumatism or as tonics in traditional Chinese medicine [7,8]. Sea cucumbers are one of the most valuable aquaculture species in China, Korea, and Japan, as well as other countries, where they are

**Citation:** Popov, R.S.; Ivanchina, N.V.; Dmitrenok, P.S. Application of MS-Based Metabolomic Approaches in Analysis of Starfish and Sea Cucumber Bioactive Compounds. *Mar. Drugs* **2022**, *20*, 320. https:// doi.org/10.3390/md20050320

Academic Editor: Espen Hansen

Received: 19 April 2022 Accepted: 11 May 2022 Published: 12 May 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

used as functional foods. Traditional medicine in China and other countries in Asia and the Middle East uses sea cucumbers widely to treat a broad range of diseases, including asthma, arthritis, hypertension, and kidney disease [9,10].

Unique pharmacological properties, including anticancer, antioxidant, antithrombotic, and immunostimulating activities, among others, are associated with bioactive starfish and sea cucumber compounds [11–15]. Moreover, the secondary metabolites of sea cucumbers affect the biological clock and circadian rhythm of lipid metabolism [16], reduce fat accumulation [17], and protect against high fat diet-induced metabolic disorders in mice [18]. Extracts of certain specimens may accelerate wound healing and tissue regeneration [19].

The distinctive chemical composition of starfish and sea cucumbers seems to be the main reason for these beneficial properties. Starfish and sea cucumbers are high in valuable nutrients such as vitamins, minerals, and metabolites such as peptides, sterols, phenolics, sphingolipids, glycosaminoglycans, sulfated polysaccharides, and lectins [10]. Among these compounds the most exciting are unique polar steroid compounds and triterpene glycosides, which are characteristic of starfish and sea cucumbers. These compounds have unusual chemical structures and demonstrate a variety of biological effects, such as cytotoxic, antifungal, antiviral, antibacterial, anti-inflammatory, analgesic, ichthyotoxic, hemolytic, anti-biofouling, anticancer, immunomodulating, and neuritogenic actions [12,20–33].

Secondary metabolites are usually present in the extracts of starfish and sea cucumbers as complex mixtures of very similar compounds. Conventional methods for the structural study of bioactive compounds are usually time-consuming and labor-intensive procedures that include the isolation of individual compounds by a combination of chromatographic techniques and structure elucidation through a combination of different methods [20]. The final structure confirmation of a new compound is always performed with a set of independent methods, such as nuclear magnetic resonance (NMR) spectroscopy, mass spectrometry (MS), or other analytical methods and chemical transformation. Despite the instrumentation developments of recent years, the analysis of bioactive natural compounds using conventional approaches remains a challenging task due to the difficulty of isolating individual compounds from fractions consisting of many components and with high chemical diversity covering a broad concentration range. As a result, only certain compounds can be described; overall, the entire metabolite pool remains poorly studied.

At present, modern mass spectrometry techniques are widely employed for the identification and structural analysis of novel natural compounds [34–36]. Hyphenated techniques combining various separation methods with mass spectrometry are applied for metabolomic and target profiling and allow for the characterization of compounds in complex mixtures extracted from biological material. Different MS-imaging techniques precisely localize and quantify the metabolites in tissues [37]. Recently developed ion mobility (IM) methods add dimension to conventional chromatography separation, allowing the stereoisomers that cannot be separated by liquid chromatography (LC) to be identified [38].

Mass spectrometry is used in various fields of marine sciences today, including marine proteomics [39], metabolomics [40,41], lipidomics [42], marine toxicology [43], ecology studies [44], and others. The introduction of modern mass-spectrometric approaches has greatly contributed to the development of metabolomics as a transdisciplinary science that aims at the qualitative and quantitative determination of the whole metabolite pool of organisms. Although no single analytical method exists that can determine all members of the metabolome simultaneously, MS-based metabolomics has been successfully applied to analyze a wide range of compounds from various sources. In recent years, MS-based metabolomics has emerged as a useful tool in natural product research. In addition to metabolic fingerprinting, two approaches used in metabolomics studies can be distinguished [45]. The first, metabolic profiling, focuses on the analysis of structure-related metabolites or metabolites related to a specific metabolic pathway. Such an approach provides information on the chemical composition of extracts or fractions and allows for the dereplication of known bioactive compounds and detection of new compounds as well as the evaluation of the their isolation possibility [34]. The metabolome-oriented approach

aims to detect differences in metabolic profiles that occur in response to stress, disease, changing environmental conditions, or other influences in comparative experiments.

Metabolomic studies of marine organisms is a field that uses a variety of modern approaches, including MS, NMR methods, and hyphenated techniques [40,41,44]. This review focuses on the use of MS-based metabolomics techniques applied in studies of extracts and fractions of starfish and sea cucumber bioactive compounds. The following section provides a general overview of the characteristic features in workflows, including sample preparation and analytical approaches. Section 3 includes illustrative examples of MS-based applications in the analysis of bioactive compounds such as polar steroids, triterpene glycosides, cerebroside, and ganglioside, as well as examples of the multiclass approach application. Instrumental and methodological details are highlighted and summarized in tables (Table 1, Table 2, Tables S1 and S2). Section 4 reviews the recently published advances in the field of metabolome-oriented studies of starfish and sea cucumbers.

#### **2. Overview of MS-Based Metabolomic Workflows in the Analysis of Starfish and Sea Cucumber Bioactive Compounds**

In terms of workflow, a typical MS-based metabolomic study involves the stages of sample collection, extraction, fractionation and/or purification, measurement, identification, and analysis of the results (Figure 1). The analytical protocols used in MS-based marine metabolomics have several important differences from those used to analyze the metabolites of terrestrial animals and plants [41]. This section discusses the characteristic features of MS-based metabolomic approaches in the context of starfish and sea cucumber bioactive compounds, including steps of sample preparation, acquisition, and analysis.

**Figure 1.** Main research stages in MS-based metabolomics studies of starfish and sea cucumber bioactive compounds.

#### *2.1. Sample Preparation*

The sample preparation stage, which comprises sample collection and extraction, is the most important in metabolomics research. Most of the studied starfish and sea cucumbers are collected from the coastal area manually or by SCUBA divers, or, if the depth exceeds 30 m, by bottom trawling. The main difficulties encountered in the collection of echinoderms are related to accessibility and their limited quantity. Many starfish and sea cucumbers are common species found in coastal areas where collection is unproblematic, while others occur in restricted or inaccessible geographic areas or in limited populations. Certain sea cucumber species, such as *Apostichopus japonicus* and *Holothuria scabra*, are aquaculture species, making their collection much simpler than the collection of wild specimens. In contrast to terrestrial ecosystems, when collecting marine samples the depth, salinity, and oxygen concentration of the water must be considered in addition to general factors such as temperature and light. A specimen's location, physiological state, sex, and

season can have a great metabolic influence. Difficulties are often caused by significant distances between the collection site and the laboratory, which requires more complicated logistics and specific sample preparation protocols.

Stress caused by handling induces responses in animals at the biochemical level [46] and can cause sea cucumber evisceration, the expulsion of the internal organs from the body [47]. To avoid such changes, as well as metabolomic changes resulting from enzymatic turnover during transportation or sample processing, it is highly recommended to quench the metabolism rapidly [48]. There are protocols designed for quenching, including flashfreezing using liquid nitrogen or dry ice, lyophilization, and freeze-drying; however, some of these are difficult to implement when animals are collected in the wild. Therefore, in most cases researchers use alternative protocols such as freezing or direct extraction with organic solvents [41].

The collected sample material must be processed to extract the metabolites of interest and remove salts and impurities. Extractions with organic solvents are commonly used for this purpose. Due to the high chemical diversity of metabolites, there is no single solvent capable of capturing all the required compounds without related impurities and contaminants. Generally, polar and semi-polar metabolites such as triterpene glycosides, asterosaponins, and gangliosides are preferentially extracted with hydro-alcoholic solutions, while lipid, sterol, terpene, and other non-polar compound extraction can be achieved with hydrophobic solvents (chloroform, hexane) or liquid–liquid extraction (LLE) by Folch's [49] and Bligh and Dyer's [50] methods. Extraction with methyl tert-butyl ether (MTBE) [51] can be used for the recovery of both polar and non-polar metabolites into separate fractions. In addition, the selected extraction protocols and solvents must be related to the analytical methods used.

It should be noted that most starfish and sea cucumber extracts contain significant amounts of salts, even if a non-polar solvent is used for extraction. Such samples are incompatible with analytical techniques such as mass spectrometry and NMR because of the effect of salts on analytical performance. For example, the presence of a small concentration of NaCl can cause the appearance of unexpected adducts at ESI MS, while larger concentrations can suppress analyte ionization and lead to salt crystal deposits in the ion source and the capillary, which can cause the instrument malfunction. MALDI MS is more tolerant to salt impurities and can be used for preliminary screening of extracts without additional purification. Desalting of marine extracts typically involves column chromatography (CC), liquid–liquid extraction, and solid-phase extraction (SPE).

Another problem can be the presence of lipid impurities and/or proteins in samples of polar secondary metabolites. For example, when extracting starfish polar steroid compounds or sea cucumber triterpene glycosides the crude hydro-alcoholic extracts may contain a large concentration of phospholipids, which can complicate chromatographic separations and suppress the ionization of the target analytes. If lipid compounds are not included in the target pool, additional purification of the extract can improve both LC separation and MS identification of target analytes. In order to remove such interfering compounds, column chromatography with Amberlite XAD-4, Sephadex LH-60, or other sorbents, LLE or SPE is usually used. In order to simplify the analysis of extremely complex extracts and obtain a mixture containing only structure-related metabolites of interest, fractionation using column chromatography, flash chromatography, or HPLC is used.

The choice of extraction solvent and purification methods affects the efficiency of the sample preparation stage. The use of unsuitable solvents and extraction methods can result in quantitatively and qualitatively incomplete extraction, while the use of suboptimal purification or fractionation procedures can lead to loss of the target metabolites. To the best of our knowledge, there are no published studies comparing the effectiveness of the most commonly used sample preparation protocols for the analysis of starfish and sea cucumber bioactive compounds.

#### *2.2. Data Acquisition*

Structural elucidation of starfish and sea cucumber bioactive compounds remains a difficult task due to the great diversity of these compounds and the complexity of the analyzed mixtures. Usually, these compounds form very complicated mixtures which are difficult to separate into pure compounds by chromatography. In the past, the application of chemical methods was required in order to identify the structure of such compounds. In particular, acid hydrolysis was used to recognize steroid and triterpene glycoside structures. While this approach allowed the partial characterization of aglycon structures and the determination of qualitative and quantitative monosaccharide composition, the destruction of native aglycon was frequent.

For a long time, electron ionization (EI) was the only possible mass spectrometry technique. Rashkes et al. carried out mass spectrometry research on six polyhydroxysteroid compounds and glycosides isolated from the Far Eastern starfish *Patiria pectinifera* and determined the characteristic fragmentation pattern of starfish polyhydroxysteroid under EI conditions [52]. EI and GC-EI MS were widely used for the determination of structures of aglycones and oligosaccharide chains of asterosaponins and triterpene glycosides after hydrolysis of glycosides and chemical derivatization of monosaccharides [20]. GC coupled with EI MS remains one of the most suitable metabolomic techniques for analyzing the wide range of volatile, semi-volatile non-polar compounds and derivatized polar metabolites. Electron impact ionization results in highly reproducible fragmentation patterns that can be used for identification by database search along with retention times indexes.

The application of fast atom bombardment (FAB) MS allows for analysis of the more polar and unstable compounds. Introduced in 1983, FAB has been successfully used for the determination of the structures of starfish steroid glycosides and sea cucumber triterpene glycosides as well as cerebrosides and gangliosides, which could not be analyzed by EI MS [53]. FAB mass spectra of starfish and sea cucumber glycosides can show molecular ions as well as fragmentation products, providing information about molecular formulae, the presence and location of sulfate groups, the structures of carbohydrate chains, and aglycon. Collision-induced dissociation (CID) experiments can provide additional structural information on the structural features of aglycon, the quantity and type of monosaccharides attached to aglycon, and their location.

Electrospray ionization (ESI) and Matrix-Assisted Laser Desorption/Ionization (MALDI) have significantly expanded the possibilities of mass spectrometry for the analysis of natural products. ESI has had an enormous impact on the analysis of polar and nonvolatile molecules as well as large biomolecules. In contrast to electron ionization, in-source fragmentation under ESI conditions is practically unrealized; tandem MS methods are used to initiate the fragmentation of these ions. ESI mass spectrometry is currently the most common ionization technique; it has been widely used for the characterization of natural compounds, including steroid and triterpene glycosides, polar lipids, and other compounds from purified starfish and sea cucumber extracts. MALDI MS is another efficient method for the analysis of natural compounds. The necessity of using matrices and the presence of matrix ion peaks at spectra in the low mass range are drawbacks; however, due to its high sensitivity, high speed of analysis, and tolerance to inorganic salts impurities, MALDI MS is widely used for rapid screening and chemical characterization of complex mixtures. Recent advances in analytical techniques, including high-resolution time-of-flight (TOF), Fourier transform (FT), and Orbitrap mass analyzers have high scan speeds along with extended dynamic range and sensitivity, allowing for the development of hybrid instruments and new ionization interfaces such as nanoelectrospray (nanoESI) and heated electrospray ionization (HESI) and leading to the establishment of high-throughput protocols for the analysis of the most complex mixtures of natural compounds. The development of hyphenated techniques combining liquid chromatography or gas chromatography with mass spectrometry (LC-MS or GC-MS) makes allows for straightforward analysis of the compounds present in complicated extracts.

#### *2.3. Data Analysis*

Data analysis is the next important stage of MS-based research. Generally, the processing of data obtained using chromatography-MS methods has included the steps of identifying *m*/*z* signals, chromatographic peak detection, filtering, alignment, and identification [54]. Many freely available (XCMS [55], MZmine 2 [56], OpenMS [57], and MS-DIAL [58]) and commercial software tools are currently available for the processing of LC-MS and GC-MS data. While the processing of GC-MS data is well-established and relatively simple, the results are usually limited to known compounds presented in databases. LC-MS is a more versatile method, covering broad chemistries and sensitivity ranges, although it produces more complex data. Due to the lower resolution and reproducibility of LC separation and the presence of adduct, isotope, fragment, and contamination peaks in ESI spectra, LC-MS data processing is much more difficult.

In certain cases, special approaches are useful for data processing. In order to process MS profiling data, methods based on scanning neutral losses, characteristic fragments, and an in-house library for rapid screening of the compounds of interest are often used. For example, the construction of ion chromatograms for negative fragment ions at *m*/*z* 96.96 can be used for detecting sulfated compounds like asterosaponins and sulfated triterpene glycosides, and cerebrosides can be detected according to the neutral loss fragments of 180 Da [59].

Similar to common metabolomic studies, metabolite identification is a current bottleneck in the analysis of starfish and sea cucumber metabolites. Chromatography-MS-based analysis can result in a huge number of peaks that are extremely difficult to identify. Even when analyzing well-studied organisms, only small percentages of the data collected in a typical LC-MS experiment can be matched to known molecules [60]. The chemical composition of starfish and sea cucumbers remains poorly investigated and the percentage of identified compounds can be extremely low. According to Metabolomics Standards Initiative recommendations, high identification confidence can be obtained by comparing an accurate high-resolution monoisotopic mass, MS/MS spectra, and retention times with data from an authentic chemical standard [61]. However, the available libraries of certified standards do not cover the entire scope of biochemical diversity, especially in the area of marine bioactive compounds. Although only putative annotation is possible without matching experimental data to data for authentic chemical standards [61], the availability of comprehensive open-access databases is extremely important for the successful application of mass spectrometry to the analysis of complex mixtures of natural compounds. Existing databases cover various natural compounds [62], and several databases, such as the GNPS database [63], MassBank [64], Metlin [65], the Human Metabolome Database [66], and MassBank of North America (https://mona.fiehnlab.ucdavis.edu, accessed on 1 April 2022) include MS/MS spectra of natural compounds from different sources. Databases such as the Dictionary of Marine Natural Products (https://dmnp.chemnetbase.com, accessed on 1 April 2022) and MarinLit (https://marinlit.rsc.org, accessed on 1 April 2022) include structural information, and the NMR and UV spectra of marine-derived compounds, although MS and MS/MS data on marine natural compounds in all existing databases is extremely limited. There are currently no databases covering taxonomic, structural, and experimental mass spectrometry data on bioactive metabolites of marine echinoderms.

Moreover, unlike peptides, oligosaccharides, and lipids, the MS fragmentation of most secondary metabolites is less studied due to the vast structural variability, and the de novo identification of metabolites by MS/MS spectra is very difficult. Several computational approaches based on machine learning or quantum chemistry calculations have been proposed for the in silico generation of MS/MS spectra or the prediction of structural features of compounds based on the experimental MS/MS spectra [67,68]. The molecular networking approach is based on the clustering of detected compounds by the similarity of their MS/MS spectra and allows for the annotation of related metabolites [63]. Using models for the in silico prediction of LC retention times can help to improve the reliability of identification in metabolomics analysis [69]. However, despite the recent advances in computational approaches, the currently used algorithms need to be significantly improved before effective identification and structural elucidation of marine bioactive compounds is possible. Along with the huge degree of structural variability, these issues limit the application of MS for annotation, dereplication, and structural elucidation in studies of metabolites from marine organisms.

In the case of research aimed at discovering metabolomic alterations between distinct biological groups of organisms, the statistical analysis is applied to the peak lists obtained after the processing of MS data. The choice of statistical methods is often determined by the study design, and can be divided into univariate (*t*-test, analysis of variance (ANOVA), fold-change analysis) and multivariate (unsupervised Principal Component Analysis (PCA) and supervised Partial Least Squares Discriminant Analysis (PLS-DA)) methods of analysis. Generally, the statistical methods used in the analysis of starfish and sea cucumber metabolites are the same as those used for conventional metabolomics studies, which are thoroughly discussed in [70]. Tables S1–S3 provide a general overview of the statistical approaches used for the treatment of analytical data in research on starfish and sea cucumber bioactive compounds.

#### **3. MS-Based Metabolomic Profiling Approaches to the Study of Starfish and Sea Cucumber Bioactive Compounds**

#### *3.1. Starfish Polar Steroid Compounds*

Unlike other echinoderms, starfish are characterized by a wide variety of steroid compounds, both non-polar sterols and polar steroid compounds. The latter form a large group of biologically active compounds, including polyhydroxysteroids, related glycosides, and steroid oligoglycosides (asterosaponins) (Figure 2). Starfish polyhydroxysteroids are steroid compounds, and usually contain from four to nine hydroxy groups in a steroidal nucleus and side chain. Polyhydroxylated glycosides have one, two, or rarely three monosaccharides attached to a steroid moiety, either to side chains or to the steroid nucleus and side chain simultaneously. The most common sugar residues in these compounds are xylose or its derivatives and arabinose. Polyhydroxylated glycosides have been found in both sulfated and non-sulfated forms. A characteristic feature of the asterosaponins is the 3β,6α-dihydroxysteroid aglycon with a 9(11)-double bond and a sulfate group at C-3. The asterosaponin carbohydrate chain consists of four to six sugars and is attached to C-6. The oligosaccharide chain of pentaosides contains one branching at the second monosaccharide, while hexaosides can have one or two branches at the second and third monosaccharide residues of the chain. Hexoses (glucose, galactose), pentoses (arabinose, xylose), and deoxyhexoses (fucose, quinovose) are the most common sugar residues in asterosaponins. Monosaccharides in asterosaponins are always in pyranose forms and are connected, as a rule, by β-glycosidic bonds (Figure 2).

**Figure 2.** The structures of typical starfish polyhydroxysteroid (5α-cholestane-3β,6α,8,15α,16β,26 hexaol (**1**) from the starfish *Protoreaster nodosus* [71]), a glycoside of polyhydroxysteroid (linckoside

#### A (**2**) from the starfish *Linckia laevigata* [72]), and asterosaponin (thornasteroside A (**3**) from the starfish *Acanthaster planci* [73]).

Demonstrating significant structural diversity, individual representatives of starfish polar steroids show a variety of biological effects including cytotoxic, neuritogenic hemolytic, antibacterial, antiviral, and anti-inflammatory effects [12,20,27–31]. Several starfish polar steroids are promising antitumor and cancer-preventing agents [14]. A recent study has reported that starfish polar steroids in combination with X-ray radiation affect colony formation and apoptosis induction in human colorectal carcinoma cells [74]. Starfish polar steroids have shown a combined anticancer effect with alga polysaccharides on human cell lines in models of 2D and 3D cultures [75,76].

ESI and MALDI are currently the most suitable and widely used ionization techniques for analyzing starfish polar steroids (Table 1). Typically, starfish sulfated steroid glycosides are detected as [M + Na]+ and [M − Na]<sup>−</sup> ions in the positive and negative ion modes of ESI MS, respectively. Non-sulfated polyhydroxysteroids and related glycosides are usually revealed as [M − H]<sup>−</sup> and [M + Cl]<sup>−</sup> ions in the negative ion mode and as [M + Na]+ ions in the positive ion mode. Although both positive and negative ion mass spectra are good enough to characterize all types of glycosides, the negative ion mode contains peaks of higher intensities and is more suitable for the analysis of sulfated compounds, while non-sulfated glycosides are analyzed by the positive ion mode [77–79].

Preliminary asterosaponin structures can be predicted from experimental tandem MS data because typical fragment ions and neutral losses provides valuable information about aglycon structures and sequences of monosaccharide units in carbohydrate chains. Tandem mass spectra show intensive characteristic fragment peaks for all asterosaponins and sulfated glycosides, indicating the sulfate group (peak at *m*/*z* 96.9 in negative ion mode and neutral loss of 120 Da and peak at *m*/*z* 142.9 in positive ion mode) (Figure 3). In the negative product ion spectra, an intense Y-type ion series (nomenclature by Domon and Costello [80]) associated with the cleavages of glycosidic bonds and corresponding sequential losses of sugar units has been observed. The analogous Y-type product ion series corresponding to losses of monosaccharide units as well as B- and C-type product ion series can be observed in the positive ion spectra of asterosaponins. The fragmentation of certain asterosaponins under CID conditions produces a very intense characteristic product ion series corresponding to the loss of side chain neutral fragments. For example, the spectra of many asterosaponins display neutral loss of a fragment of 100 Da as well as the Y–100 product ion series. This fragmentation corresponds to the loss of the C6H12O molecule associated with the C-20–C-22 bond cleavage and 1H transfer, which is characteristic of asterosaponins containing an aglycon with a 20-hydroxy-cholestan-23-one side chain (Figure 3) [20]. The similar product ion series Y–114 and Y–128 indicate aglycons with 20-hydroxy-24-methyl-cholestan-23-one and 20-hydroxy-24-ethyl-cholestan-23-one side chains.

The structural characterization of polyhydroxysteroids and glycosides of polyhydroxysteroids is challenging due to the great diversity of compounds in this class. Spectra of polyhydroxysteroid compounds and related glycosides contain fragmentation patterns indicate a number of hydroxy groups and structures of the side chains and steroid nuclei. Tandem spectra of polyhydroxysteroid glycosides with sulfated monosaccharide unit usually show diagnostic ion B0 at *m*/*z* 241.0 [C6H9O8S]−, 225.0 [C6H9O7S]− or 210.9 [C5H7O7S]−, which are characteristic of sulfated hexose, methylated pentose, or pentose units, respectively, whereas the presence of intense Y-type ions is associated with non-sulfated compounds or sulfated aglycon. In certain cases the A- and X-type product ions formed by cross-ring cleavages of sulfated monosaccharides can be detected, potentially allowing isomeric monosaccharides with different positions of the sulfate group to be distinguished [81,82]. In other cases, the structure of the polyhydroxysteroidal aglycon can be proposed from both obtained MS data and from biosynthetic considerations [77,83].

**Figure 3.** ESI MS/MS spectrum of [M − Na]<sup>−</sup> precursor ion at *m*/*z* 1213 identified as ophidianoside F (modified from [77]).

Table 1 provides a general overview of the approaches used for metabolomic profiling of starfish polar steroids and triterpene glycosides of sea cucumbers, including extractions, purification methods, and analytical techniques. Table S1 provides more expansive technical details of these particular approaches, including sample preparation protocols and the instrumental setups of the ESI MS, MALDI MS, and hyphenated techniques.

**Table 1.** Selected examples illustrating MS-based approaches for the analysis of starfish polar steroids and sea cucumber triterpene glycosides \*.



**Table 1.** *Cont.*


**Table 1.** *Cont.*


**Table 1.** *Cont.*

\* Abbreviations: CC, column chromatography; CSI, captive spray ionization; ESI, electrospray ionization; FTICR, Fourier-transform ion cyclotron resonance; HPLC, high-performance liquid chromatography; IM, ion mobility; IT, ion trap; LC, liquid chromatography; LLE, liquid-liquid extraction; NMR, nuclear magnetic resonance; MALDI, matrix-assisted laser desorption/ionization; MS, mass spectrometry; MSI, mass spectrometry imaging; MSPD, matrix solid-phase dispersion; nanoESI, nanoelectrospray; QOrbitrap, quadrupole-Orbitrap; QTOF, quadrupole time-of-flight; QQQ, triple-quadrupole; RPLC, reverse-phase liquid chromatography; SPE, solid-phase extraction; TOF, time-of-flight.

Polar steroid compounds are usually extracted from animal materials using methanol, ethanol, or hydro-alcoholic solutions. As crude extracts contain a large concentration of impurities and inorganic salt, there is a need for additional purification procedures before analysis. SPE with reverse-phase (RP) sorbents is the most fast and versatile way of obtaining the purified total fraction of polar steroids [77,78]. Regarding asterosaponin fraction extraction, LLE is preferred. The most commonly used protocol is adapted from that in [108], and includes the dilution of the dry extract in 90% methanol followed by successive partitioning against *n*-hexane, dichloromethane, and chloroform. After column chromatography with an Amberlite XAD-4 column, the eluate is extracted against isobutanol. The resulting fraction contained purified asterosaponins [85,90].

Although LC coupled to MS through electrospray ionization interface is the most popular combination for profiling due to its efficiency, versatility, and capability in analyzing isomeric compounds, MALDI is commonly used as the primary technique for the rapid screening of extracts [85]. All of the researchers reviewed here used columns with RP C18 sorbents for analytical separation; however, it should be noted that the high complexity of the extracts and fractions places high demands on both chromatography and detection. The use of high-resolution MS analyzers (TOF and Orbitrap) enables calculation of the elemental composition of the analytes via high accuracy measurements and true isotopic patterns, while using tandem techniques allows for structural characterization of the detected compound.

LC-NMR-MS was used for the profiling and characterization of *Asterias rubens* asterosaponins [84,109]. The on-flow LC-NMR-MS screening showed novel asterosaponins in *A. rubens*, and their tentative structures were proposed by MS and NMR data [109]. Using a similar experimental setup, the authors replaced time-consuming classical extraction with matrix solid-phase dispersion (MSPD) extraction, which combines both sample homogenization and extraction in a single step starting from the intact sample material. The structures of seventeen asterosaponins were established based on complementary structural information from both MS and NMR detection, including 1H-NMR spectra obtained in on-flow mode, 2D WET-TOCSY spectra from the MS-triggered stopped-flow mode, information about molecular mass before and after H-D exchange, and fragmentation patterns and characteristic neutral losses [84].

Further studies on *A. rubens* using a combination of MALDI MS, MALDI imaging (MALDI MSI), and LC-MS have focused on the diversity, body distribution, and localization of asterosaponins [85,86]. Asterosaponins from the body walls, stomach, pyloric caeca, and gonads were extracted and analyzed by MALDI-TOF MS and LC-ESI MS [85]. As a result, seventeen known and nine novel asterosaponins were detected. It was found that each organ was characterized by a specific mixture of asterosaponins, and that their concentration varies considerably among individuals. MALDI MSI was used to clarify the inter- and intraorgan distribution of asterosaponins [86]. Sample preparation is a particularly important step in MALDI imaging. Because the starfish body wall contains calcareous ossicles, the researchers used carboxymethyl cellulose as an embedding medium to facilitate the cryosectioning procedure. The results confirmed that asterosaponin distributions are not homogeneous, and revealed that certain asterosaponins are located both inside the body wall and within the outer mucus layer, where they probably protect the animal.

As a part of starfish polar steroid exploration, metabolite profiling of polar steroids in the Far Eastern starfishes *Aphelasterias japonica* and *Patiria pectinifera* was performed [77,78]. A detailed LC-MS analysis of the complicated mixture of polar steroids from *A. japonica* revealed 68 polar steroid metabolites, including 33 asterosaponins, 28 polyhydroxysteroid glycosides, and seven polyhydroxysteroids [77]. Fragmentation analysis indicated asterosaponins with rare and atypical units in their oligosaccharide chains that have thus far not been identified from marine sources. The profiling of polar steroid compounds of *P. pectinifera* using the LC-MS allowed many different polar steroid compounds to be discovered [78]. LC-ESI MS analysis revealed 72 components (35 asterosaponins, 15 sulfated glycosides of polyhydroxysteroids, and 22 polyhydroxylated steroids). Annotation was based on MS data obtained in both negative and positive ion modes. Liquid chromatography coupled with atmospheric pressure photoionization (LC-APPI) MS was applied for non-sulfated polyhydroxysteroid compounds. APPI MS/MS exhibited extensive fragmentation, with sequential neutral losses of H2O molecules and cleavages in side chains and tetracyclic nucleus. The comparison of the steroid constituents of *P. pectinifera* and *A. japonica* revealed significant differences associated with details of the biosynthesis of starfish polar steroids.

A combination of SPE and ultra-high performance liquid chromatography (UPLC) coupled with ion trap (IT) mass spectrometry was used to profile asterosaponins from the Brazilian starfish *Luidia senegalensis* [88]. Seven components were detected as a result, and five of which were characterized as asterosaponins. ESI MS was used together with NMR to detect and characterize asterosaponins and sulfated polyhydroxysteroid glycosides in bioactive fractions obtained by LLE and chromatography purification of the ethanolic extract from the starfish *Heliaster helianthus* [91].

The profiling of polar steroids from the starfish *Lethasterias fusca* was carried out by nanoflow liquid chromatography coupled with captive spray ionization (CSI) mass spectrometry [79]. As a result, the structure of the largest number of polar steroid metabolites was discovered, and the MS fragmentation of a large series of starfish polar steroids was studied. A total of 207 compounds, including 106 asterosaponins, 81 glycosides of polyhydroxysteroids, and 14 polyhydroxylated steroids, were detected and characterized. Further study of the distribution of the detected compounds in *L. fusca* body components showed that the polar steroid compositions in the body walls, coelomic fluid, gonads, stomach, and pyloric caeca were qualitatively and quantitatively different [89]. Research on the distribution of asterosaponins from *Echinaster sepositus* revealed eleven compounds, and found significant variability in asterosaponin composition depending on the organ, sex, and season [90].

In summarizing the aforementioned results it is necessary to note the huge variety of starfish polar steroids. Each studied species contained dozens, and in several cases, hundreds of polar steroids, most of which had not been previously described [77–79]. Additional studies of the polar steroids in the studied species often lead to the identification of both known compounds and new previously-undescribed metabolites. For example, the study of asterosaponins of *A. rubens* by LC-NMR-MS revealed seventeen asterosaponins [84]. Subsequent studies led to the detection of both previously discovered compounds as well as the discovery of new asterosaponins [85,86]. This is related both to the instrumental advancements involving increasing sensitivity and selectivity of analysis and to the great variability in the polar steroid composition of starfish, even among representatives of the same species. The observed large structural variability together with the huge number of structures in each species studied and the small number of species studied (a total of six species have been studied using metabolomic methods) may indicate a potentially huge chemical space for starfish polar steroids.

Most studies have focused on both the description of the structural diversity and on the study of the localization of the discovered compounds. It has been determined that each organ is characterized by a certain composition of polar steroids. The comparison of the content of individual steroids in different starfish organs probably suggests the different biological roles of these metabolites in the starfish. Asterosaponins, which are the most toxic starfish compounds, have been found in all organs of the starfish [85,86,89]. However, the body walls often show the highest content of asterosaponins. In addition, these compounds have been found in the outer layer of mucus [86]. This may be due to the toxic, protective, or antimicrobial properties of these compounds. The main potion of polyhydroxysteroid glycosides is located in the pyloric caeca, which confirms the digestive function of these steroids in starfish [89,110]. At the same time, the levels of polar steroids can vary greatly depending on the individual, season, and sex [85,90,111]. This high interindividual variability may be associated with different physiological statuses of the animals, and partly with the biogenesis of certain compounds from dietary steroids.

#### *3.2. Sea Cucumber Triterpene Glycosides*

Triterpene glycosides are the characteristic secondary metabolites of sea cucumbers. Their chemical structures are characterized by the large variability of certain structural features, although the general structure of these compounds is rather conservative. Most sea cucumber triterpene glycosides have a lanostane-type aglycon with an 18(20)-lactone. Usually, aglycon has a polycyclic nucleus with a 7(8)- or 9(11)-double bond and oxygencontaining substituents, which may be bonded to C-12, C-17, or C-16. Structures of the side chains of aglycons demonstrate significant natural diversity and may have one or more double bonds, hydroxyl or acetate groups, and other substituents. Certain glycosides have aglycons with shortened side chains. The carbohydrate chains of sea cucumber glycosides may include up to six sugar units and be attached to C-3 of the aglycon. Xylose, glucose, quinovose, 3-*O*-methylglucose, and rarely 3-*O*-methylxylose are the most common sugar residues in triterpene glycosides. The first monosaccharide unit is always xylose, and monosaccharides with the 3-*O*-methyl group are always terminal ones. Many glycosides have up to four sulfate groups in the first xylose, glucose, and 3-*O*-methylglucose units. The oligosaccharide chains that have up to four monosaccharide units usually represent a linear structure, while the penta- and hexaosides contain a branching at the first or second monosaccharide unit (Figure 4) [21,23,24,32,33].

**Figure 4.** The structures of typical holothurian triterpene glycosides with holostane aglycon (okhotoside B1 (**4**) from the sea cucumber *Cucumaria okhotensis* [112]) and rare non-holostane aglycon (kurilosides A1 (**5**) from the sea cucumber *Thyonidium* (*=Duasmodactyla*) *kurilensis* [113]), demonstrating different carbohydrate chain architecture.

Triterpene glycosides demonstrate both biological and pharmacological effects, including cytotoxic, antifungal, bactericidal, antiviral, and antiparasitic effects [21,22,32,33]. The most interesting of these is the ability of certain glycosides to induce apoptosis and inhibit tumor cell growth; thus, sea cucumbers have become a promising source for the discovery of new drugs [114]. Additionally, certain triterpene glycosides have been reported to exhibit immunomodulatory properties [115]. Several species of sea cucumber are an important aquaculture resource and are used as functional foods [116]. It has been suggested that dietary triterpene glycoside supplements can improve lipid metabolism, significantly suppress adipose accumulation, and reduce serum and hepatic lipids [117].

Most triterpene glycosides have been detected within *m*/*z* range from 1000 to 1600 as [M + Na]<sup>+</sup> ions. In most cases, sulfated and disulfated triterpene glycosides are detected in negative ion mode as [M − Na]<sup>−</sup> and [M − 2Na]2<sup>−</sup> ions, respectively, whereas nonsulfated compounds are detected as [M − H]<sup>−</sup> ions. The tandem mass spectra of triterpene glycosides usually reveal B- and C-type product ion series arising from the cleavage of glycosidic bonds. These product ion series are characteristic, and they provide information about the sequence of monosaccharide residues in carbohydrate chains (Figure 5).

**Figure 5.** ESI MS/MS spectrum of [M <sup>−</sup> Na]– precursor ion at *<sup>m</sup>*/*<sup>z</sup>* 1277, identified as cucumarioside F2 (modified from [118]).

Tandem mass spectra of many triterpene glycosides show typical mass losses related to aglycon fragmentation, and can provide information about the structure of the nucleus and the side chain. In the MS/MS spectra of certain glycosides a mass loss of 60 Da between the precursor and the intense fragment ion has been detected, which corresponds to the loss of the C2H4O2 molecule and is a characteristic of glycosides containing an acetoxy group. An intense fragment ion with a mass loss of 104 Da from the precursor is related to the loss of a [C2H4O2 + CO2] fragment, which is characteristic of compounds with an acetoxy group and an 18(20)-lactone cycle. Certain triterpene glycosides tend to lose the neutral fragments of the side chain under CID conditions.

Regarding the extraction of triterpene glycosides from animal tissue, the most preferred solvents are 70% ethanol and methanol, although ethanol and ethyl acetate:methanol mixtures have been used (Tables 1 and S1). Crude extracts must be purified before analysis to remove inorganic salts, lipid, and protein contaminants. For these purposes, most researchers use successive liquid–liquid partitioning against *n*-hexane, dichloromethane, and chloroform, followed by column chromatography with an Amberlite XAD-4 column and extraction against butanol or similar LLE-based protocols. Another purification approach includes SPE with C18 cartridges [99,100]. Omran et al. used LLE with a solvent combination of MTBE/MeOH/H2O for the purification of crude ethanol extract to obtain the polar fraction of the triterpene glycosides and non-polar fraction of lipids in a single extraction step [105]. The purified extracts can be fractionated using HPLC [96,106] or flash chromatography [107].

MALDI MS is often used as the primary technique for rapid screening of triterpene glycoside mixtures. As sea cucumber triterpene glycosides are characterized by the presence of isomeric compounds, the LC-MS technique can be used as a tool for discriminating different isomers and structure confirmation. In order to separate glycosides, most LC-MS applications use analytical columns with C18 sorbents. Ion mobility technology provides additional orthogonal separation for the discrimination and structural characterization of isomeric compounds [119].

A combination of MALDI MS and ESI MS was used to annotate the triterpene glycosides in purified fractions of the Australian sea cucumber *Holothuria lessoni* [120–123]. As a result, a series of known and novel triterpene glycosides were annotated by extensive MS fragmentation. It should be noted that the authors determined the structures of novel glycosides based only on MS data. However, it is known that different epimeric monosaccharides as well as types of bonds between sugars and absolute configuration of asymmetric atoms cannot be strictly distinguished by MS [124]. The proposed structures are therefore tentative, and must be verified by additional approaches such as NMR spectroscopy.

MS-based approaches were used for screening, characterization, and study of the bodily distribution of the triterpene glycosides of the sea cucumber *Holothuria forskali* [92,93,95]. Triterpene glycosides were extracted from two different body components, the body wall and the Cuvierian tubules (a defensive organ that can be ejected in response to predator attacks), and analyzed by a combination of MALDI MS, MALDI MSI, and LC-ESI MS. The analysis revealed at least 26 triterpene glycosides, including twelve glycosides in the body wall and twenty-six in the Cuvierian tubules. The glycosides detected in the body wall were found in the Cuvierian tubules, with the latter containing fourteen other specific glycosides as well. A more detailed study of triterpene glycoside localization in the body wall revealed that the glycosides were mainly localized in the epidermis and mesothelium [95]. A combination of MALDI, LC-ESI MS, and LC-ESI-IM MS was used in order to better tackle the structural complexity of *H. forskali* glycosides [101]. As a result, at least 10, 16, and 22 different triterpene glycosides within the body wall, gonads, and Cuvierian tubules, respectively, were detected. Glycosides with pentasaccharide chains were dominant within the extracts from the gonads and the Cuvierian tubules, whereas the body wall extract exhibited equally abundant tetra-, penta- and hexaosides. In addition, the authors described the interaction of branched triterpene glycosides with sodium ions, and proposed a new schematic for data representation using sector diagrams constructed from MS data.

The diversity of triterpene glycosides was studied in five tropical sea cucumber species [94]. Triterpene glycosides from the body wall and the Cuvierian tubules were extracted and analyzed with a combination of MALDI MS, ESI MS, and LC-MS. The researchers indicated that the smallest number of glycosides was observed in *Holothuria atra*, which contained a total of four compounds, followed by *Holothuria leucospilota*, *Pearsonothuria graeffei*, and *Actinopyga echinites* with six, eight, and ten compounds, respectively. *Bohadschia subrubra* showed the highest triterpene glycoside diversity. Differences between the glycoside composition in the body walls and the Cuvierian tubules were highlighted.

The profiles of three tropical sea cucumber species were determined and compared to examine their chemical diversity with phylogenetic data [96]. Semi-purified extracts from the body wall of *Holothuria scabra*, *H. impatiens*, and *H. fuscocinerea* were first analyzed by MALDI-FT MS and chip-HPLC-ESI MS. The obtained data showed holothurines common for three species (for example, holothurin A) as well as glycosides specific for certain species (for example, impatienside A in *H. impatiens*). Glycosidic fractions of three species contained approximately the same number of compounds (32 glycosides in *H. scabra* and *H. impatiens* and 33 glycosides in *H. fuscocinerea*); however, the glycoside profiles were both quantitatively and qualitatively different from each other. Moreover, the authors demonstrated a relationship between metabolomic and phylogenetic data. Their obtained results show the possibility of effectively using MS-based metabolomic profiling for chemotaxonomy purposes in sea cucumbers.

The MALDI MS analysis of triterpene glycosides of *Holothuria scabra* revealed six major compounds saved during the processing of the body wall [97]. Mitu et al. showed that *H. scabra* releases triterpene glycosides into the surrounding seawater [100]. The characterization of these compounds by LC-multimode source MS led to the annotation of sixteen new and known compounds. A recent study used MALDI MS and LC-IM MS to characterize the triterpene glycoside composition of the viscera of *H. scabra* [107]. A combined analysis revealed 26 sulfated triterpene glycosides.

MALDI MS was used for the rapid structural characterization of triterpene glycosides in the body wall and Cuvierian tubules of the sea cucumber *Holothuria sanctori* [98]. Mass spectrometry analysis revealed eighteen triterpene glycosides, including eight novel compounds. Body wall triterpene glycosides showed higher diversity than those from the Cuvierian tubules.

LC-MS profiling of the sea cucumber *Eupentacta fraudatrix* revealed 26 sulfated, 18 nonsulfated, and 10 disulfated triterpene glycosides [99]. Many novel compounds were characterized by tandem MS, including those with previously unknown oligosaccharide chain

types. Two new glycosides were isolated and their tentative structures were confirmed by NMR. Based on the literature data and obtained results, a biosynthetic pathway for oligosaccharide fragments of *E. fraudatrix* glycosides was proposed. LC-MS analysis of extracts from the respiratory trees, body walls, gonad tubules, guts, and aquapharyngeal bulbs indicated triterpene glycosides in all body components, although quantitative variability for certain triterpene glycosides was observed.

A combined technique involving microscopic analysis, MALDI MS, and MALDI MSI was used to study triterpene glycoside localization in the body wall of *Holothuria leucospilota* [102]. MALDI MS analysis of the body wall and the epidermal tissue extracts revealed twelve triterpene glycosides. The following MALDI MSI analysis showed the presence of detected glycosides in the epidermis of *H. leucospilota*, whereas in the dermis the circular and longitudinal muscle bands had no glycosides.

The composition of triterpene glycoside in the sea cucumber *Holothuria atra*, collected in the Persian Gulf, was studied using a modern analytical approach combining LC-MS, molecular networking, pure compound isolation, and NMR spectroscopy. In order to evaluate the entire pool of triterpene glycosides, the purified extract was subjected to LC-MS analysis using a column with pentafluorophenyl phase. The obtained MS data were used to create a molecular network using the GNPS Molecular Networking website. As a result, twelve triterpene glycosides were found, including three novel glycosides. Four major triterpene glycosides were isolated by HPLC, and their structures were confirmed by NMR [103].

The metabolic profiling of the polar fractions obtained from MTBE-based LLE of six sea cucumbers allowed for the identification of two sulfated glycosides found in all species and two species-specific nonsulfated glycosides [105]. Metabolic profiling was used for the screening of crude extracts of nine tropical sea cucumber species related to anti-fouling activities [106]. LC-MS analysis detected glycosides in all extracts; in total 102 triterpene glycosides were detected. The extract of *Bohadschia argus* represents one of the most active anti-fouling extracts, and includes 23 glycosides. The obtained results demonstrate that anti-fouling activities in sea cucumber extracts are species-specific and related to both triterpene glycoside total concentration and structures of presented triterpene glycosides. In another study, the relation between anti-fouling activities and the presence of triterpene glycosides was observed [125]. The UPLC-MS approach was used to evaluate the glycoside composition of *Apostichopus japonicus* [104]. As a result, five triterpene glycosides were detected and the variability of content of identified triterpene glycosides among the different types of *A. japonicus* was described.

The results of the aforementioned metabolomics studies indicate a huge structural variability of triterpene glycosides, as in the case of starfish polar steroids. Interestingly, certain species contain only a few triterpene glycosides, while dozens of glycosides have been found in the extracts of others. The maximum number of triterpene glycosides was detected in the extract of *E. fraudatrix*, with 54 compounds [99]. It should be noted that certain researchers used only MALDI MS methods, which do not distinguish between isomeric compounds in mixtures; thus, the real number of structures may be somewhat higher. Several studies have demonstrated the strict taxonomic specificity of the chemical composition of studied systematic groups of sea cucumbers [96,105]. Thus, MS-based metabolomic profiling can easily be used for chemotaxonomy purposes, both to clarify the species identity of unknown specimens and to confirm or revise the status of taxa.

The application of MALDI MSI and LC-MS profiling of extracts from various organs provides a better understanding of the distribution and localization of triterpene glycosides in animal tissues. Although glycosides are present in all organs, their distribution is heterogeneous [99]. Maximum total concentrations and number of structures can be found for the body walls and Cuvierian tubules [94,98,101]. By using MS-based methods, it has been found that sea cucumbers secrete toxic triterpene glycosides into the surrounding water [95]. These facts confirm the suggestion that triterpene glycosides have multiple defensive roles, including defense against predators and protection from parasites and microorganisms.

#### *3.3. Starfish and Sea Cucumber Lipids*

Marine invertebrates are known to be a valuable source of bioactive and dietary lipids, which are connected to the prevention of diseases and have applications in nutrition, cosmetics, pharmacy, and other fields. In this respect, an essential part of modern lipidomic studies is focused on fatty acids, glycerophospholipids, and sphingolipids in marine organisms such as sponges, cnidarians, worms, molluscs, and arthropods. Regarding starfish and sea cucumbers, research interest is largely linked to studying fatty acid composition and bioactive sphingolipids, including unique cerebrosides and gangliosides [126].

Sea cucumbers and starfish contain fatty acids in relatively small amounts [127]. Sea cucumber fatty acids usually account for less than 8% of their total weight, enriched in unsaturated fatty acids that may account for up to 70% [128]. The main fatty acids of sea cucumbers are 20:4 (n-6), 20:1, 20:5 (n-3), 16:0 and 18:0 [129]. Starfish are characterized by a high level of polyunsaturated acids, among which 20:5 (n-3) and 20:4 (n-6) are dominant [130]. MS-based analysis of fatty acids can be considered a well-established approach in view of both sample preparation and analysis protocols. Nowadays, GC-MS with chemical derivatization is routinely used to obtain an exhaustive view of the composition and metabolism of fatty acids. Analytical approaches and scientific results on fatty acid composition in marine invertebrates, including sea cucumbers and starfish, are discussed in detail in [13,131–133]. Fatty acid and sterol compositions of numerous sea cucumber [134] and starfish species [135] have been analyzed using GS-MS-based approaches. In addition, the variability in fatty acid content studied by GC-MS is used to analyze the effects of diet [136] and geographical origin [137,138] and to distinguish between wild and cultured animals [139].

The major phospholipids in Echinodermata include phosphatidylcholine, phosphatidylethanolamine, and phosphatidylserine. Sea cucumbers are characterized by high phosphatidylinositol content, while their lysophosphatidylcholine, lysophosphatidylethanolamine, diphosphatidylglycerol, phosphatidic acid, and phosphatidylinositol-4-phosphate contents are low [140].

Sample preparation for phospholipid determination in sea cucumbers and starfish is commonly based on extractions from the fresh whole body, body walls, or viscera via the classic Folch method [141], the Bligh and Dyer method [142], or extraction using a solvent combination of MTBE/MeOH/H2O [105] (Table 2 and Table S2). Different phospholipid headgroups and alterations in chain length, amount and position of double bonds in fatty acids, and ester bond types lead to an extremely complex composition of phospholipid mixtures. Traditionally, protocols that use deacylation and derivatization followed by GC or GC-MS identification were used to establish the fatty acid composition of phospholipids. However, such approaches are associated with the destruction of the original structures. In contrast, LC-MS methods allow for the detailed investigation of complex lipid mixtures in a high-throughput manner without prior purification and chemical modification. However, good chromatographic separation is critical for the accurate identification of lipids in such complex mixtures. Most frequently used reverse-phase sorbents, such as C8 or C18, allow for the separation of the molecules of phospholipids according to chain length and degree of saturation of acyl fatty acids. The use of HILIC or normal-phase (NP) columns, on the other hand, allows for the separation of molecules based on the structure of the polar head groups.

**Table 2.** Selected examples illustrating MS-based approaches for the analysis of starfish and sea cucumber lipids \*.



**Table 2.** *Cont.*

\* Abbreviations: CC, column chromatography; ESI, electrospray ionization; HESI, heated electrospray ionization; HILIC, hydrophilic interaction chromatography; IT, ion trap; LC, liquid chromatography; LLE, liquid-liquid extraction; LPC, lysophosphatidylcholine; LPE, lysophosphatidylethanolamine; NPLC, normal-phase liquid chromatography; MS, mass spectrometry; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; QOrbitrap, quadrupole-Orbitrap; QTOF, quadrupole time-of-flight; RPLC, reverse-phase liquid chromatography; SPE, solid-phase extraction; TOF, time-of-flight.

Although phospholipids have drawn intense interest in recent years as substances with the potential for human health benefits, and although modern LC-MS lipidomic methods have been used extensively, only limited studies are available on the phospholipid content of starfish and sea cucumbers [148]. The investigation of the composition of phospholipids in several echinoderm species by LC-ESI MS revealed the predomination of alkylacyl-PC and alkenylacyl-PE forms in starfish and sea cucumbers [141]. Structural characterization was achieved by comparing the retention times and in-source fragmentation patterns obtained in negative and positive ion modes with single-stage MS. The NPLC-ESI TripleTOF MS was employed to study the phospholipid composition of the dried body walls of six sea cucumber species [142]. The application of normal-phase liquid chromatography as a separation technique allowed for the division of complex mixtures into subclasses with reproducible retention times as well as the separation of isobaric molecules in the same subclass. As a result, between 295 to 445 molecular species belonging to eleven phospholipid subclasses (phosphatidylcholines, phosphatidylserines, phosphatidylethanolamines, phosphatidylinositol, phosphatidic acids, phosphatidylglycerols, lysophosphatidylcholines, lysophosphatidylethanolamine, lysophosphatidylserine, lysophosphatidylinositol, and rare phosphonoethanolamine) were detected in each species. Identification and structure elucidation were based on retention times, ion forms, and specific fragmentation patterns obtained in negative ion mode using the LIPIDMAPS database. Semiquantitation followed by statistical analysis demonstrated differences in the phospholipid profiles of the studied species. Analysis of the nonpolar fraction obtained by extraction of six Egyptian sea cucumber species was performed using UPLC-Orbitrap MS in positive and negative ion modes [105]. For metabolite identification, the obtained data (*m*/*z*, retention time, isotope and fragmentation patterns) were searched against public databases. A total of fifteen free fatty acids and 45 triacyl glycerols were detected. Statistical analysis showed that quantitative variations in lipid content between the studied species were associated with habitat or food changes, not with taxonomical relationships.

Sphingolipids constitute an extremely diverse class of bioactive polar lipids. As components of cell membranes and intracellular mediators, sphingolipids are involved in cell recognition and signal transduction processes [149,150]. Sphingolipids from marine organisms exhibit various activities, including antitumor, immunomodulatory, antiviral, neuritogenic, and other activities [126,151].

Sphingolipids, particularly cerebrosides and gangliosides, are important components of echinoderm lipids that have drawn attention because of their structure and bioactivity. Compared to those present in mammals and plants, the sphingolipids present in echinoderms have notable structural differences. Variations in chain length and in the degree of saturation and/or hydroxylation of the sphingoid backbone and fatty acids lead to the extensive variety of cerebrosides structures. Starfish and sea cucumber gangliosides remain little studied. These compounds have a specific sugar core, including sialic acids within carbohydrate chains, as well as additional monosaccharide residues and unusual types of glycosidic bonds between them (Figure 6) [126]. The determination of their diverse structures and variety of sphingoid backbones are very important for understanding the functional and nutritional significance of dietary sphingolipids. To date, about 150 sphingolipids from fifteen starfishes and nine sea cucumbers have been studied, several of which have demonstrated biological activity [126]. Cerebrosides from sea cucumbers and starfish show activity against nonalcoholic fatty liver disease [152] and an inhibitory effect on cell proliferation through the induction of apoptosis in cancer cells [153]. Sea cucumber sphingosine has strong cytotoxicity against colon cancer cells [154]. Gangliosides of starfish and sea cucumbers show slight neuritogenic activity. However, the bioactivity of these compounds remains poorly investigated [126].

Regarding cerebroside extraction from starfish and sea cucumbers, LLE with methanol: chloroform mixtures is preferred, although extraction following the Bligh and Dyer method or modifications of it is applicable. Subsequent purification of crude extracts by LLE, SPE, or silica gel column chromatography provides pure fractions containing cerebrosides. Highspeed counter-current chromatography (HSCCC) has recently been pointed out as a way to improve the resulting cerebroside fractions without requiring extraction procedures or additional purification [145]. Gangliosides, as the more polar compounds, can be extracted from sea cucumber body walls by homogenization in water followed by purification using LLE with a solvent combination of CHCl3/MeOH/ H2O followed by SPE purification using C8 sorbent [147] (Tables 2 and S2).

**Figure 6.** The structures of typical echinoderm cerebrosides (luidiacerebroside A (**6**) from the starfish *Luidia maculata* [155] and glucocerebroside HPC-3-A (**7**) from the sea cucumber *Holothuria pervicax* [156]) and gangliosides (ganglioside molecular species from the sea cucumber *H. pervicax* (**8**) [157] and acanthaganglioside I (**9**) from the starfish *Acanthaster planci* [158]).

Because of their structural complexity, sphingolipids are difficult to analyze using one method. Formerly, sphingolipids were quantified by thin-layer chromatography (TLC); structure elucidation comprised many stages of chemical decomposition and derivatization followed by GC, GC-MS, HPLC, MS, and NMR analyses [159]. Today, LC-MS with electrospray ionization is the main tool for the detection and annotation of sphingolipids, including both known and novel molecular species. RPLC with isocratic elution with 95% MeOH with 5 mM ammonium acetate and 0.05% acetic acid or elution with 95% ACN is commonly used to separate sphingolipids. HILIC-LC on the GOLD-amino column has been used in the separation of gangliosides from sea cucumbers [147]. In this approach, the ganglioside subclasses are eluted in a specific time range based on their amounts of sialic acid residues.

The ESI in positive ion mode is the most extensively used ionization technique for the analysis of cerebrosides (Table 2). Although the locations of the double bonds in the fatty acyl chain often cannot be unequivocally identified by CID, the specific fragment ions from fatty acid, sphingosine, and sugar units allow putative structures to be proposed [59,143–145]. Gangliosides detected in the negative ion mode mainly form deprotonated or double deprotonated ions. Characteristic fragment ions and neutral losses formed by oligosaccharide chain fragmentation reveal a monosaccharide composition and different sialic acid types [147].

The investigation of cerebrosides from various sources by LC-MS showed that the cerebroside composition of sea cucumbers differs from those of plants (maize, rice) and mushrooms (maitake). [160]. Research into the cerebroside composition of the sea cucumbers *Apostichopus japonicus*, *Thelenota ananas*, *Acaudina molpadioides*, *Bohadschia marmorata*, *Cucumaria frondosa*, and *Pearsonothria graeffei* using LC-MS led to the discovery of a large series of compounds and showed that sea cucumber sphingolipids are much more diverse than was conventionally thought [59,143–145] (Table 2). Each studied species contained several dozen cerebroside molecular species, with the most complicated composition belonging to *T. ananas*. Several sea cucumber species were found to have similar sphingolipid compositions, while the profiles of others were dramatically distinctive. The analysis of many structures allowed identification of the characteristic structural features of sea cucumber cerebrosides. A sphingoid base (d17:1) is typically predominant in sea cucumber cerebrosides and is not widely found in plants, mammals, or fungi. In addition, the occurrence of C23:1h is characteristic of sea cucumber cerebrosides and is rarely found in

plants, mammals, or fungi. The FA contained in cerebrosides from sea cucumbers is similar to those of common mammals, although it has more double bonds and hydroxylation. The study of the cerebroside composition of the gonads, viscera, and whole body of the starfish *Asterias amurensis* revealed a characteristic structure distribution that can be divided into three major structural groups [146].

HILIC-ESI MS was used to identify gangliosides in six sea cucumber species. Seventeen ganglioside subclasses were detected, and their oligosaccharide chains were characterized by tandem MS [147]. The results indicated that sea cucumber gangliosides differ from mammalian gangliosides in monosaccharide composition, number, and types of sialic acids. Moreover, gangliosides with phosphoinositidyled sialic acid and tetrasialogangliosides were identified in sea cucumbers for the first time.

#### *3.4. Multi-Class Profiling Studies*

In contrast to the single-class approach, the multi-class approach attempts to simultaneously detect the qualitative and quantitative characterization of various metabolites of different chemical groups in a single analysis run. Essentially, research that uses a multiclass approach aims to investigate crude extracts or study the composition of bioactive fractions. Due to the complicated composition of marine invertebrates, such extracts or fractions contain a wide diversity of compounds. The chemical pool of such extracts depends primarily on the solvent used and/or on the extraction protocol. Methanol, ethanol, dichloromethane, chloroform, ethyl acetate, and their mixtures are often used for extraction. Non-polar solvents extract complex mixtures of fatty acids, lipids, carotenoids, triterpenes, and sterols, while the use of polar solvents leads to extracts containing mixtures of polar steroid compounds or triterpene glycosides, polar lipids, and other compounds. The results depend on the analytical method and platform. The widely used multi-class GC-MS approach is used for analysis and accurate identification via a database search of sterols, triterpenes, and fatty acids such as methyl esters. LC-MS is used to characterize more polar compounds.

The LC-MS metabolic profiling of sea cucumber *Holothuria spinifera* extract indicated secondary metabolites of several classes [161]. Gradient elution on the C18 column with ESI-Orbitrap MS operating in positive and negative ion modes was used for the investigation of methanol:dichloromethane extract. It is worth noting that only 4% of metabolites were detected in either mode. The molecular formula was predicted using the MZmine algorithm and identification was achieved using MarinLit and the Dictionary of Natural Products databases. As a result, thirteen secondary metabolites belonging to the fatty acids, phenolic diterpenes, and triterpenes were identified.

Investigation of antifouling and antibacterial activities of three extracts from different organs of the sea cucumber *Holothuria leucospilota* showed that ethyl acetate extract of the body wall possessed the most pronounced activity [162]. In order to determine its bioactive compounds, GC-MS was used. Using the NIST GC-MS library, seventeen metabolites, including five terpenes and terpenoids and six fatty acids, were identified.

The lipid, fatty acid, and sterol compositions of four sea cucumbers were analyzed to assess their feeding habits [163]. The extracts were obtained using a modified Bligh and Dyer protocol and analyzed using a thin-layer chromatography-flame ionization detector (TLC-FID) analyzer to quantify lipid classes and GC-MS for the identification of individual metabolites. The sea cucumbers were found to be rich in phytosterols and algal-derived fatty acids, suggesting tight trophic coupling to phytodetritus, while the relatively large proportions of stanols were probably the result of enteric bacteria. GC-MS was used for targeted profiling of fatty acids as methyl esters and amino acids after sample derivatization with N-methyl-N-tert-butyldimethylsilyltrifluoroacetamide (BSTFA) and trimethylchlorosilane (TMCS) of three sea cucumber species [127].

Dichloromethane, methanol, and aqueous extracts of *Linckia laevigata*, *Fromia indica*, *Cryptasterina pentagona*, and *Archaster typicus* were tested in order to identify surface-bound metabolites that protect the starfish from fouling; the most biologically active fractions

were analyzed by GC-MS [164]. Several fatty acids and sterols were identified using the NIST and Wiley GC-MS databases. GC-MS analyses of the surface-extracted metabolites of each starfish specimen identified hexadecanoic acid, cholesterol, lathosterol, and sitosterol as the compounds responsible for the antifouling effects.

Pereira et al. proposed the GC-MS method for simultaneous analysis of fatty acids, amino acids, sterols, and lupanes in marine animals and applied this method to the characterization of the starfish *Marthasterias glacialis* extract [165]. Using ethanol as the extraction solvent at 40 ◦C and N-methyl-N-(trimethylsilyl)-trifluoroacetamide (MSTFA) as the derivatization reagent, forty compounds (including fifteen amino acids, sixteen fatty acids, six sterols, and three lupanes were detected and quantified.

The LC-MS approach was used for the profiling and identification of the active compounds of dichloromethane:methanol extract of the holothurian *Pseudocolochirus violaceus*, which exhibited strong antiproliferative effects [166]. The mixture was separated on a C18 column and metabolites were detected using ESI MS in positive and negative ionization modes. Compound identification was performed by accurate mass measurement and comparison of the obtained data with previously reported information. As a result, 24 compounds, belonging mainly to the terpenes, steroids, and fatty acids, were detected using both positive and negative ionization modes, several of which have previously been reported to exhibit antiproliferative capacity in cancer cells.

A combination of the GC-MS and LC-MS methods was applied to obtain a comprehensive view of the composition of ethyl acetate extract of the sea cucumber *Holothuria forskali* [167]. The GC-MS analysis revealed 25 major components identified by the NIST GC-MS library, while LC-MS showed eight molecules, including fat-soluble vitamins, phytosterols, and phenolic acids, identified by comparison of retention times and MS data obtained for pure standards.

Zakharenko et al. proposed an alternative method for the extraction of bioactive compounds from sea cucumbers using a two-step process with carbon dioxide extraction followed by extraction with CO2 and ethanol as a co-solvent [168]. The obtained extracts were tested by LC-IT MS in negative and positive ion modes and tandem MS. The identification of the metabolites was achieved using the Bruker library database and literature data, and revealed the presence of fifteen triterpene glycosides, eighteen styrene compounds, and fourteen carotenoids.

Variability in the chemical composition of the viscera and body walls of the sea cucumber *A. japonicus* extracted by methanol was investigated by the LC-MS approach, and 85 metabolites were determined [169]. Multivariate data analysis using PCA and PLS-DA revealed significant differences between the viscera and body walls. To identify the main characteristic compounds of viscera, several sphingoid-based nucleoside analogs were isolated and their structures were confirmed by MS/MS and NMR methods.

The investigation of sea cucumber metabolites is not restricted to metabolite profiling of extract, and can be applied to the profiling of volatile compounds as well. Volatile compounds of eight dried sea cucumber species with different geographical origins were analyzed using a combination of headspace solid-phase microextraction and GC-MS [170]. Metabolite identification was achieved by matching data with NIST and Wiley GC-MS databases and confirmed by comparing retention times and mass spectra with standard compounds. As a result, 42 volatile compounds, including aldehydes, alcohols, aromatic compounds, and furans, were identified in the dried sea cucumbers, several of which were determined as odour-active compounds.

#### **4. Applications of Metabolome-Oriented Approaches in Studies of Starfish and Sea Cucumbers**

MS-based targeted metabolomics were applied to investigate the influence of different environmental factors on the polar steroids of the starfish *P. pectinifera* [87]. Extracts of control starfishes and starfishes exposed to water heating, oxygen deficiency, feeding, injury, and different water salinity levels were purified by SPE and analyzed using LC-ESI-QTOF

MS. An in-house library of retention times and MS data on previously characterized polar steroid metabolites of *P. pectinifera* were used for metabolite identification. Univariate and multivariate statistical analyses revealed variations in the steroid metabolome between the control and treatment groups. In order to further evaluate stress-induced differences, PCA and PLS-DA analyses were carried out on each group of starfish individually with the control group. The results revealed that differences caused by feeding, injury, and heating were greater than in the other starfish groups. These states had similarities in their effects on the steroid metabolome of starfish. Most asterosaponins were reduced, and most polyhydroxysteroids and related glycosides were increased. These differences in steroid metabolite profiles may relate to the biological multifunctionality of these compounds.

MALDI MSI was applied to study the precise localization of sea cucumber *H. forskali* triterpene glycosides in the Cuvierian tubules of control and stressed sea cucumbers [93]. Stressed animals were mechanically disturbed for 4 h by repetitive hitting using a specific device. Statistical multivariate tests using PCA showed statistical differences in triterpene glycoside composition between the control and stressed groups. Triterpene glycosides with corresponding ions at *m*/*z* 1287 and 1303 were mainly localized in the connective tissue of the tubules of both control and stressed sea cucumbers. Glycoside ions at *m*/*z* 1125 and 1141 were present in relaxed animals, while ions at *m*/*z* 1433, 1449, 1463, and 1479 were observed in the Cuvierian tubules of stressed animals in the outer part of the connective tissue. The authors proposed that the latest glycosides are stress-specific compounds formed by modifications of the glycosides with shortened oligosaccharide chains. Another study revealed that *H. forskali* releases glycosides into the surrounding seawater. Among these secreted glycosides, holothurinoside G was detected in the seawater surrounding relaxed sea cucumbers, while holothurinosides C, F, M, L, and desholothurin A were secreted when the animals were stressed [95].

The most extensive metabolomics research has been performed on the commercially important sea cucumber *Apostichopus japonicus* (Table S3). Most of these studies used a non-targeted UPLC-QTOF MS approach and focused on primary metabolites such as amino acids, sugars, fatty acids, and common metabolites, and did not involve sea cucumberspecific triterpene glycosides and gangliosides.

High temperature and low oxygen concentration are the common environmental stress factors for marine invertebrates, and their impact on *A. japonicus* has been studied using MS-based metabolomics [171]. Changes in the concentrations of 84, 68, and 417 metabolites related to the responses to heat, hypoxia, and combined stress, respectively, were detected by LC-MS and multivariate statistical analysis. Among the detected metabolites, compounds atypical for echinoderms such as the plant glycoside tokoronin, the synthetic drug tirofiban, and others were found, which may be due to identification errors (the authors did not provide information on how metabolite identification was performed). Another investigation of acute hypoxia in *A. japonicus* showed that levels of most lipids increased with the elongation of hypoxia. These results imply that the homeostasis of synthesis and degradation of lipids and their derivatives are strongly affected by hypoxic stress [172]. Liu et al., used GC-MS to compare the metabolic profiles of a thermotolerant strain of *A. japonicus* with a control group, and found significant differences in the concentrations of 52 metabolites [173]. Evisceration is a well-known stress response of sea cucumbers, although the biochemistry of this process is unclear. Metabolomic analysis of coelomic fluids ejected during *A. japonicus* evisceration using LC-MS followed by univariate and multivariate analysis revealed five significantly changed signaling pathways [47]. In response to high temperatures, sea cucumbers can enter a state characterized by inactivity, cessation of feeding, gut degeneration, and decreased metabolic rate. This physiological state is called aestivation. Yang et al. used transcriptomic and metabolomic approaches to explore alterations in *A. japonicus* during the aestivation stage [174]. LC-MS analysis revealed that downregulated metabolites were associated with fatty acid metabolism, carbohydrate metabolism, and the TCA cycle. UPLC-QTOF MS was used to describe the metabolic changes induced by skin ulceration syndrome, the main disease affecting the development of *Apostichopus japonicus* in the aquaculture industry [175]. As a result, variations in metabolites mainly related to amino acid metabolism, energy metabolism, immunity, osmoregulation, and neuroactive ligand-receptor interaction has been discovered.

Another research focus has been directed towards the study of metabolomic changes induced by the impact of various factors. LC-MS has been used to highlight metabolomic differences between cage-cultured, pond-cultured, and bottom-sowed *A. japonicus* [176]. Multivariate analysis and enrichment of metabolic pathway analyses revealed differential metabolites participating in lipid, amino acid, carbohydrate, and nucleotide metabolism. The investigation of *A. japonicus* coelomic fluids in different sexes and reproductive states by UPLC-QTOF MS and multivariate statistical analysis revealed variations in phenylalanine metabolism and unsaturated fatty acid synthesis [177]. LC-MS highlighted significant metabolic differences in the muscle tissue of animals between the nonbreeding and growth stages [178]. The metabolite profiles obtained using UPLC-QTOF-MS of four *A. japonicus* varieties (green, white, purple, and spiny) were compared, and differences were identified using multivariate analysis [179]. Differential metabolites included fatty acids, amino acids, phospholipids, and sugars. In another study, a similar approach was applied to reveal the metabolic changes in white, green, and purple *A. japonicus* body walls during the pigmentation process [180]. Statistical analysis differentiated the body wall chemical composition among the three color morphs, and thirteen annotated metabolites showed significant differences in white, green, and purple sea cucumbers. UPLC-QTOF MS metabolomic profiling was applied to distinguish *A. japonicus* from different geographical origins [181]. Data analysis using OPLS-DA showed that differential metabolites mainly included amino acids and lipids.

Melatonin-induced metabolomic changes in the muscle tissues of *A. japonicus* have been tested using UPLC-QTOF MS [182]. Statistical analysis with PCA, PLS-DA, foldchange analysis, and *t*-test showed alterations in the levels of 22 different metabolites, including serotonin, retinoic acids, and fatty acids, which can explain the observed sedative effect of melatonin on this species. The LC-MS metabolomic analysis revealed that pedal peptide-type neuropeptides involved in the regulation of locomotor behavior in *A. japonicus* induce changes in the levels of certain phospholipids [183].

Most of the mentioned metabolomics studies of *A. japonicus* used a similar workflow involving the extraction of tissue samples with methanol, methanol:water, or methanol: acetonitrile mixtures, homogenization, centrifugation, and LC-MS analysis mainly using RP separation and ESI-QTOF mass spectrometers for the detection of metabolites operating in the mass ranges from 50 to 1200 Da in positive and negative ion modes. Although a huge number of metabolites (from several dozen to 4435 metabolites) are found in almost every work, the compounds that are mainly responsible for the bioactive properties of sea cucumbers (triterpene glycosides, cerebrosides, and gangliosides) remain undetected, and their variations under the studied conditions remain unclear. At the same time, the results of the other works [87,93] indicate statistically significant changes in the levels of specific metabolites, namely, starfish polar steroids and sea cucumber triterpene glycosides, in response to stresses and environmental factors. Thus, the influence of many factors and physiological phenomena, such as aestivation and evisceration, on large groups of bioactive metabolites remains unexplored. It is well known that starfish and sea cucumbers have extraordinary regenerative potential, however, the features of this process remain poorly explored at the metabolome level.

#### **5. Conclusions and Perspectives**

The unique features of biosynthesis and metabolism and the diversity of their metabolites explain the researchers' interest in these organisms. Starting in the middle of the last century, chemical research on echinoderm metabolites has resulted in hundreds of compounds, many of which have demonstrated biological and pharmacological effects. The structural elucidation of the bioactive compounds of starfish and sea cucumbers is a difficult task, combining isolation of pure compounds with modern MS and NMR techniques to unambiguously determine the structures of new compounds.

MS-based metabolomics approaches have proven to be a powerful research tool in the natural product area. The application of MS-based techniques has made it possible to study chemical compounds without the laborious process of isolating individual compounds. Using modern metabolomics methods in the marine sciences allows evaluation of the biochemical diversity of marine systems and expands our understanding of the chemical space of marine compounds.

To date, only a few dozen species of starfish and sea cucumbers have been studied using MS-based metabolomics approaches, a very small fraction of the more than 3600 known species. Most of the studied species are aquaculture species or readily available and widely distributed species, and most deep-sea and rare species remain almost unexplored.

In summarizing MS-based metabolomics studies on the bioactive secondary metabolites of starfish and sea cucumbers, it was found that this approach allows for the ready detection and annotation of polar steroids, triterpene glycosides, and lipids in producer organisms. Obtained data allow for their exact or preliminary structures to be proposed. The data obtained thus far make it possible to assess the prospects for the search for new bioactive molecules as well as to draw conclusions about their taxonomic distribution, biogenesis, and biological functions. These methods are used to compare metabolomic profiles of different echinoderm species and populations in ecological, dietary, and biosynthesis studies.

The main difficulties of applying MS-based metabolomics approaches are related to the extreme complexity of the mixtures being analyzed. However, the use of modern chromatography and mass spectrometry methods allows these methods to be successfully applied. UPLC with analytical columns packed with sub-2-μm sorbents allows metabolites with closely related structures to be separated. The introduction of cutting-edge massspectrometry techniques such as ion mobility, improved ion dissociation techniques, and ultra-high resolution mass analyzers can provide high-performance MS analyses. While identification of detected peaks remain the main problem in MS data analysis, constructing specialized databases of echinoderm metabolites and improving in silico computational algorithms may improve the obtained results considerably.

**Supplementary Materials:** The following supporting information can be downloaded at: https: //www.mdpi.com/article/10.3390/md20050320/s1, Table S1: Instrumental and methodological details of MS-based applications for the analysis of starfish polar steroids and sea cucumber triterpene glycosides; Table S2: Instrumental and methodological details of MS-based applications for the analysis of starfish and sea cucumbers lipids; Table S3: Instrumental and methodological details in metabolomics studies of starfish and sea cucumbers.

**Author Contributions:** Writing—original draft preparation and writing—review and editing, R.S.P. and N.V.I.; conceptualization and validation, P.S.D. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was carried out with the support of the Russian Science Foundation (RSF) Grant Number 21-73-00180.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


## *Review* **Sphingolipids of Asteroidea and Holothuroidea: Structures and Biological Activities**

**Timofey V. Malyarenko 1,2,\*, Alla A. Kicha 1, Valentin A. Stonik 1,2 and Natalia V. Ivanchina 1,\***


**Abstract:** Sphingolipids are complex lipids widespread in nature as structural components of biomembranes. Commonly, the sphingolipids of marine organisms differ from those of terrestrial animals and plants. The gangliosides are the most complex sphingolipids characteristic of vertebrates that have been found in only the Echinodermata (echinoderms) phylum of invertebrates. Sphingolipids of the representatives of the Asteroidea and Holothuroidea classes are the most studied among all echinoderms. In this review, we have summarized the data on sphingolipids of these two classes of marine invertebrates over the past two decades. Recently established structures, properties, and peculiarities of biogenesis of ceramides, cerebrosides, and gangliosides from starfishes and holothurians are discussed. The purpose of this review is to provide the most complete information on the chemical structures, structural features, and biological activities of sphingolipids of the Asteroidea and Holothuroidea classes.

**Keywords:** sphingolipids; ceramides; cerebrosides; gangliosides; sialic acid; Asteroidea; Holothuroidea; biological activity; neuritogenic activity

#### **1. Introduction**

Being the second-largest clade in a superphylum Deuterostomia after chordates, Echinodermata (echinoderms) is a phylum of exclusively marine invertebrates, inhabiting all the oceans in all the depths. These animals are characterized by radial symmetry, a particular water vascular system, and calcareous particles (ossicles) embedded in the dermis of their body walls. In some habitats, echinoderms are the dominant species in marine communities. There are five living classes of Echinodermata: Holothuroidea (sea cucumbers), Asteroidea (starfish), Ophiuroidea (brittle stars), Echinoidea (sea urchins), and Crinoidea (sea lilies and feather stars). These invertebrates present a rich source of diverse low molecular biologically active metabolites, including triterpene glycosides, polar steroids, and their glycosides, peptides, fatty acids, carotenoids, quinoid pigments, and different lipids, including sphingolipids. Our group is carrying out long-term studies on natural products from echinoderms [1–6], but sphingolipids from these invertebrates [7] so far were not in our main spotlight. However, our recent metabolomic studies on secondary metabolites from echinoderms, showing their extremal diversity [8–13], and successful attempts of application of some compounds as chemotaxonomic markers required the examination of perspectives of similar use of sphingolipids.

Sphingolipids, a group of heterogeneous lipids known as constituents of the plant, fungal, and animal cellular membranes, play a fundamental role in important phenomena such as cell-cell recognition and antigenic specificity [14,15]. Sphingolipids include ceramides, the hydrophobic molecules, involving a long-chain base (LCB) and an amide-

**Citation:** Malyarenko, T.V.; Kicha, A.A.; Stonik, V.A.; Ivanchina, N.V. Sphingolipids of Asteroidea and Holothuroidea: Structures and Biological Activities. *Mar. Drugs* **2021**, *19*, 330. https://doi.org/10.3390/ md19060330

Academic Editor: Vassilios Roussis

Received: 14 May 2021 Accepted: 2 June 2021 Published: 8 June 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

linked fatty acid residue (FAR) and their glycoconjugated derivatives. Glycosylated ceramides are named cerebrosides, except for the corresponding oligoglycosides with carbohydrate chains, comprising one, two, three, or more sialic acid residues, which are known as gangliosides [16]. Sphingolipids were isolated from a number of biological sources, including marine invertebrates such as sea anemones [17], sponges [18–20], octocorals [21], ascidians [22], and representatives of other taxa. Various biological activities of ceramides, cerebrosides, and gangliosides, including plant growth stimulatory action [23], anti-inflammatory effects [24], the improving of the barrier function of the skin [25], cancerprotective action [26], proangiogenic action [27] have been reported.

In their majority, reviews about sphingolipids from marine organisms [28], including those concerning the corresponding natural products from echinoderms, were published from 12 to 20 years ago [29–31]. The present review includes data concerning chemical structures of sphingolipids from two classes of the phylum Echinodermata and their biological activities and covers the literature from 2000 to March 2021. We have focused our attention on the structures of these compounds, modern methods of analyses of complicated fractions of these lipids, and their bioactivities. Current problems of these studies are also discussed.

#### **2. Ceramides**

Ceramides are biosynthesized at the reaction of S-acyl-coenzyme A (usually C16-CoA) with serine, catalyzed by serine palmitoyl transcriptase or related enzymes, followed by reduction of carbonyl group by ketosphinganine reductase and the N-acylation by ceramide synthase. Surprisingly, hydroxylation of long-chain bases (LCBs) that leads to so-called phytosphinganine derivatives, takes a place in plants and in many echinoderms. When hydroxylases act on fatty acid residues (FARs) in these invertebrates, an additional hydroxyl is introduced also into α–position of FARs [32]. As result, four main types of ceramides are known from different organisms including echinoderms, namely, A—containing sphinganine bases and nonhydroxylated fatty acid residues, B—consisting of sphinganine bases and α–hydroxylated fatty acids, C—containing phytosphinganine bases and nonhydroxylated fatty acids, and D—consisting of phytosphinganine bases and α–hydroxylated fatty acids (Figure 1). Both bases and fatty acids moieties in this type of natural products may contain normal chains, as well as those with *iso*- and/or *anteiso*-branching. Therefore, ceramides have great structural variety.

**Figure 1.** Scheme of biosynthesis and structures of main types of ceramides in echinoderms.

#### *Class Asteroidea*

Three ceramides **1**–**3** were isolated from the starfish *Distolasterias nipon* collected off the coast of the East Sea, Republic of Korea [33]. Structures of **1**–**3** were established by spectroscopic techniques and chemical transformations as (2*S*,3*R*,4*E*,8*E*,10*E*)-2-[(2*R*)-2 hydroxyhexadecanoylamino]-9-methyl-4,8,10-octadecatriene-1,3-diol (**1**), (2*S*,3*S*,4*R*,7*Z*)-2- [(2*R*)-2-hydroxyhexadecanoylamino]-7-docosene-1,3,4-triol (**2**), and (2*S*,3*R*,4*E*,7*E*)-2-[(2*R*)-2 hydroxyhexadecanoylamino]-7-docosene-1,3,4-triol (**4**) (Figure 2). Later ceramides **2** and **3** (iteratively) and **4**–**11** (additionally) were extracted from the same species of starfish and purified by silica gel column chromatography and reversed-phase high-performance liquid chromatography [34]. The high-energy collision-induced dissociation (CID) spectra of ceramides with various structures, differing from each other in the number and positions of double bonds on both the *N*-acyl and sphingoid chains as well as in the presence of hydroxy groups or a double bond at the C-4 position of the sphingoid chains as well as an additional α-hydroxy group in *N*-acyl chains, were established. The CID mass spectrum of the monosodiated ion [M + Na]+ of each ceramide molecular species provided structural data concerning fatty acyl chains and sphingoid long-chain bases. This technique allowed determining complete structures of ceramides and cerebrosides in a mixture of sphingoid lipids and showed great potential for analysis of other sphingolipids isolated from various biological sources [34].

**Figure 2.** Ceramides from the starfish *Distolasterias nipon* and *Luidia maculata*.

A sphingosine-type ceramide LMCer-1-1 (**12**) and three phytosphingosine-type ceramides, LMCer-2-1 (**13**), LMCer-2-6 (**14**), and LMCer-2-7 (**15**), were isolated from the ceramide molecular species LMCer-1 (**16**) and LMCer-2 (**17**), obtained from the chloroform– methanol extract of the whole bodies of *Luidia maculata* [35]. The structures of ceramides **12**–**15** were determined on the basis of spectroscopic and chemical evidence as (2*S*,3*R*,4*E*)- 2-[(2*R*)-2-hydroxyhexadecanoylamino]-16-methyl-4-octadecene-1,3-diol (**12**), (2*S*,3*S*,4*R*)- 2-[(2*R*)-2-hydroxyhexadecanoylamino]-16-methyl-octadecane-1,3,4-triol (**13**), (2*S*,3*S*,4*R*)- 2-[(2*R*)-2-hydroxydocosanoylamino]-hexadecane-1,3,4-triol (**14**), and (2*S*,3*S*,4*R*)-2-[(2*R*)-2 hydroxydocosanoylamino]-14-methyl-hexadecane-1,3,4-triol (**15**) (Figure 2).

The phytosphingosine-type ceramide asteriaceramide A was isolated from the whole bodies of the Northern Pacific starfish *Asterias amurensis* [23]. The structure of this compound was determined as identical to compound **2**. Asteriaceramide A (**2**) showed a stimulatory activity toward root growth of *Brassica campestris*. The plant growth activity of the ceramide was reported for the first time.

Since 2000, there were no data on the isolation, structure elucidation, and determination of biological activities of sea cucumber ceramides.

Thus, representatives of all the above-mentioned structural groups of ceramides (Figure 1A–D) were found from starfish. The structural diversity of these metabolites is connected with the presence of many variants of both sphingoid and fatty acid moieties. It should be noted that generally ceramides from starfish were studied worse than other groups of sphingolipids. Perhaps this is due to the difficulty of isolation of individual ceramides or their molecular species.

#### **3. Cerebrosides**

Cerebrosides are glycosylceramides that contain glucose, galactose, or other monosaccharide residues in their carbohydrate moieties. These compounds are synthesized by enzymes: UDP-glucose:ceramide β-D-glucosyl-transferase, UDP-galactose:ceramide β-Dgalactose-transferase, and other glycosyl-transferases [32]. Cerebrosides can be divided into three classes: monoglycosides, biglycosides (mainly lactosides), and oligoglycosides. This class of complex lipids can contain an aminosugar residue (globosides) in their carbohydrate moieties or be sulfated at a terminal monosaccharide residue [30]. Cerebrosides, such as ceramides, are part of the plasmatic membranes of cells and perform a number of important biological functions: they take part in the formation of new membranes, such as phospholipids, sterols, and cellular membrane proteins, and also participate in the transmission of cellular signals [14]. Moreover, the cerebrosides in cellular membranes act as cell surface antigens and receptors. Interest in sphingolipids and their derivatives mainly is associated with their high biological significance. Some studies have shown that sphingolipids can inhibit the growth of microalgae, fungi, and bacteria. The presumptive mechanism of this action is associated with the ability of this type of compound to perforate cell membrane, in addition, in the presence of sphingolipids, the ability of bacterial cells to adhere is reduced [36–38]. The ability of sphingolipids to stimulate plant growth [23], demonstrate an anti-inflammatory effect [24], and improve the barrier function of the skin [25] has also been shown.

An even larger variety of cerebrosides containing one or more monosaccharide residues, in comparison with ceramides, was isolated from starfish and sea cucumbers.

#### *3.1. Class Asteroidea*

From the chloroform–methanol–water extract of gonads and body walls of the Patagonian starfish *Allostichaster inaequalis,* glucosylceramides were isolated [39], along with the previously known phalluside-1 and two glucosylceramides earlier isolated from the starfish *Cosmasterias lurida* [40]. Compounds were described as (2*S*,3*R*,4*E*,8*E*,10*E*)-1-*O*-(β-Dglucopyranosyl)-2-[(2*R*)-2-hydroxy-15-tetracosenoylamino]-4,8,10-octadecatrien-3-ol (**18**) and (2*S*,3*R*,4*E*,15*Z*)-1-*O*-(β-D-glucopyranosyl)-2-[(2*R*)-2-hydroxyhexadecanoylamino]-4,15 docosadien-3-ol (**19**) using spectroscopic and chemical methods (Figure 3).

**Figure 3.** Cerebrosides from the starfish *Allostichaster inaequalis* and *Luidia maculata*.

Glucosylcerebrosides, luidiacerebrosides A (**20**) and B (**21**), were isolated from the cerebroside fraction obtained from the extract of the starfish *Luidia maculata* using HPLC [41]. The structures of cerebrosides were determined as 1-*O*-β-D-glucopyranoside of (2*S*,3*S*,4*R*)-2- [(2*R*)-2-hydroxyhexadecanoylamino]-16-methyl-octadecane-1,3,4-triol (**20**) and (2*S*,3*S*,4*R*)-2- [(2*R*)-2-hydroxytetracosanoylamino]-16-methyl-octadecane-1,3,4-triol (**21**), respectively [41]. In continuation of the studies on sphingolipids from the same starfish four cerebrosides, luidialactosides A–D (**22**–**25**), were isolated from its the water-insoluble lipid fraction [42]. They were proved to contain lactosyl carbohydrate chains attached to C-1 of 2-[(2*R*)-2-hydroxytetracosanoylamino]-16-methyl-4-octadecene-1,3-diol (**22**), 2-[(2*R*)-2 hydroxydocosanoylamino]-14-methyl-1,3,4-hexadecanetriol (**23**), 2-[(2*R*)-2-hydroxyhexade canoylamino]-9-docosene-1,3,4-triol (**24**), and 2-[(2*R*)-2-hydroxydocosanoylamino]-1,3,4 hexadecanetriol (**25**) (Figure 3).

Eight glucosylceramides were found in the Patagonian starfish *Anasterias minuta* [43]. One of these constituents, anasterocerebroside A (**26**), was identified as a new glucosylceramide, while the earlier known glucosylceramide (**27**) was isolated and characterized for the first time as a pure compound. It was earlier isolated in a mixture with related glucosylceramides from the Patagonian starfish *Cosmasterias lurida* [40]. The structures of these sphingolipids were established by different spectroscopic and chemical methods as (2*S*,3*R*,4*E*,8*E*,10*E*)-1-*O*-(β-D-glucopyranosyl)-2-[(2*R*)-2-hydroxy-14-tricosenoylamino]- 4,8,10-octadecatrien-3-ol (**26**) and (2*S*,3*R*,4*E*,8*E*,10*E*)-1-*O*-(β-D-glucopyranosyl)-2-[(2*R*)-2 hydroxy-15-tetracosenoylamino]-9-methyl-4,8,10-octadecatrien-3-ol (**27**) (Figure 4).

The glucocerebroside, linckiacerebroside A (**28**), and known glucocerebroside S-2a-3 were isolated from the chloroform–methanol extract of the starfish *Linckia laevigata*, together with three pseudo homogeneous glucocerebrosides [44]. The structures of this cerebroside were determined as (2*S*,3*S*,4*R*)-1-*O*-(β-D-glucopyranosyl)-2-[(2*R*)-2 hydroxyhexadecanoylamino]-16-methyl-heptadecane-1,3,4-triol (**28**) (Figure 4).

A galactocerebroside molecular species CNC-2 (**29**) were isolated from the extract of the tropical starfish *Culcita novaeguineae* [45] as a phytosphingosine type galactocerebroside with nonhydroxylated and hydroxylated fatty acyl moieties (Figure 4).

**Figure 4.** Structures of cerebrosides from the starfish *Anasterias minuta*, *Cosmasterias lurida*, *Linckia laevigata*, *Culcita novaeguineae*, *Oreaster reticulatus*, and *Narcissia canariensis*.

The glucocerebroside, asteriacerebroside G (**30**), and two known cerebrosides, asteriacerebrosides A and B, were isolated from the chloroform–methanol extract of the whole bodies of the Northern Pacific starfish *Asterias amurensis* [23]. The structure of **30** was determined on the basis of chemical and spectroscopic evidence as (2*S*,3*R*,4*E*,13*Z*)-1-*O*-(β-Dglucopyranosyl)-2-[(2*R*)-2-hydroxytetradecanoylamino]-4,13-docosadiene-1,3-diol (Figure 4). Asteriacerebrosides A, B, and G exhibited growth-promoting activity for the whole body of *Brassica campestris*.

The starfish *Oreaster reticulatus* contains nine glycosphingolipids named oreacerebrosides A–I (**31**–**39**) (along with earlier known ophidiacerebrosides C–E [46], Figure 4). All these compounds have a 4,8,10-triunsaturated sphingoid base. Oreacerebrosides A–C (**31**–**33**) are β-glucosylceramides in contrast with oreacerebrosides D–I (**34**–**39**), all these compounds were the first examples of β-galactosylceramides containing this unusual sphingoid base. Four representative glycosphingolipids were tested for cytotoxic activity on rat glioma C6 cells and shown to be mildly cytotoxic. Previously, it was established that the glucosylceramides were more active than the galactosylceramides. In addition, oreacerebroside I (**39**) was shown to exert proangiogenic activity and was able to increase VEGF-induced human endothelial cell proliferation.

Mixtures of three known glucocerebrosides (F13-3), ophidiacerebrosides B–D (**40**–**42**), were isolated from the starfish *Narcissia canariensis* collected off the coasts of Dakar, Senegal [47]. This fraction included three homologous cerebrosides identified as peracetylated derivatives on the basis of spectroscopic and chemical data (Figure 4). These compounds contain a β-glucopyranose as a sugar unit, 9-methyl-branched 4,8,10-triunsaturated longchain aminoalcohols as sphingoid bases, and amide-linked 2-hydroxy fatty acid chains. The major component (63%) has an amide-linked 2-hydroxydocosanoic acid chain and was identified as ophidiacerebroside C (**41**), isolated from the starfish *Ophidiaster ophidianus* for the first time [48]. The minor components of F13-3 had one more or one less methylene group and were identified as ophidiacerebrosides B (**40**) and D (**42**). The cytotoxic activity of F13-3 was detected using KB cells. It was shown that three human cancerous cell lines, KMS-11 (adherent plasma cells obtained from patients with multiple myeloma) were inhibited by these cerebrosides with IC50 = 15.2 ± 4 μM, GBM (astrocytoma cells obtained after tumor resection of patients with glioblastoma multiforme-primary culture) with IC50 = 34.6 ± 5.1 μM, and HCT-116 (colorectal adenocarcinoma cells derived from a patient with Lynch's syndrome) with IC50 = 18 ± 3.9 μM.

In total, 21 galactocerebrosides, including 16 new compounds (**43**–**58**) (Figure 5), were identified as cerebroside molecular species obtained from the chloroform–methanol extract of pyloric caeca cut out from the starfish *Protoreaster nodosus* [49]. These compounds were phytosphingosine-type galactocerebrosides with hydroxylated fatty acyl moieties. It is important, that GC–MS analysis, followed by methanolysis and periodate oxidation of these metabolites, gave reliable structural information of ceramide moiety rapidly in minute amounts. The structures of earlier known compounds were the same as those of galactosylcerebrosides previously found from other starfish and even mammalians.

Six glucocerebrosides (**59**–**64**) were isolated from the eggs of the starfish *Asterias amurensis* by extraction and different type of column chromatography, including HPLC [50]. It was shown that the structures of cerebrosides could be completely characterized, based on their sodium-adducted molecules, using FAB tandem mass spectrometry. The lipid part of the glucocerebrosides **59**–**64** consisted of saturated and monounsaturated α-hydroxy fatty acids and sphinganine type of the long-chain base (Figure 5).

Glucosyl ceramides (GlcCers) were later isolated from the viscera of the starfish *Asterias amurensis* [51]. Degraded GlcCers generated *A. amurensis* sphingoid bases (ASBs) that mainly consisted of the triene-type bases d18:3 and 9-methyl-d18:3. Actions of these bases on ceramide synthesis and content were analyzed using normal human epidermal keratinocytes (NHEKs). The bases significantly raised the de novo ceramide synthesis in NHEKs and expression of genes, encoding enzymes such as serinepalmitoyltransferase and ceramide synthase. Total ceramide (GlcCers) and sphingomyelin contents increased highly upon ASB treatment. In particular, GlcCers bearing fatty acids with large carbon atoms (≥ C28) exhibited a significant content increasing. These ASB-induced enhancements on de novo ceramide synthesis were only observed in undifferentiated NHEKs. This stimulation of de novo sphingolipid synthesis may improve skin barrier functions.

Four cerebrosides (**65**–**68**) were isolated from the starfish *Distolasterias nipon* by extraction and different type of column chromatography, including reverse-phase HPLC [34]. Structural elucidation was conducted using tandem mass spectrometry of monosodiated ions desorbed by fast atom bombardment. Fatty acids in glucocerebrosides **65**–**68** were identified as saturated and monounsaturated α-hydroxylated derivatives. The glucocerebroside long-chain bases were found to be of di- and triunsaturated sphingenine types (Figure 5).

**Figure 5.** Cerebrosides from the starfish *Protoreaster nodosus*, *Asterias amurensis*, and *Distolasterias nipon*.

#### *3.2. Class Holothuroidea*

Overall, 18 glucocerebrosides (**69**–**86**) were detected in admixture from the sea cucumber *Holothuria coronopertusa* [52]. Their structures were established on the basis of liquid-secondary ion mass spectrometry (LSIMS) experiments. The CID mass spectrum of the lithiated molecules ([M + Li]+) led to diagnostic fragment ions, which were further identified by tandem mass spectrometry (MS/MS). Fatty acids in glucocerebrosides **69**–**86** were indicated as saturated and monounsaturated α-hydroxyl fatty acids. The glucocerebroside long-chain bases were of sphingosine type (Figure 6).

Moreover, 10 glucocerebrosides, HPC-3-A–HPC-3-J (**87**–**96**), were isolated from the extract of the sea cucumber *Holothuria pervicax* [53]. All these compounds were mixtures of regio-isomers for terminal methyl groups in the LCB moiety, namely, mixtures of *iso*- and *anteiso*-isomers (Figure 6).

**Figure 6.** Cerebrosides from the sea cucumbers *Holothuria coronopertusa* and *Holothuria pervicax*.

Five glucocerebroside molecular species (SJC-1-SJC-5, **97**–**101**) were isolated from the extract of the sea cucumber *Stichopus japonicus* [54]. Cerebrosides **97**–**99** were sphingosineand phytosphingosine-type derivatives with nonhydroxylated and hydroxylated fatty acyl moieties. At the same time, cerebroside molecular species **100** and **101** were also sphingosine-type glucocerebroside molecular species with hydroxylated fatty acid moieties, although they were new compounds with unique sphingosine bases containing additional two hydroxy groups (Figure 7).

**Figure 7.** Cerebrosides molecular species from the sea cucumber *Stichopus japonicus*.

Later, the content and components of cerebrosides from the sea cucumber *Stichopus japonicus* were analyzed by Duan et al. [55]. The absorption of cerebrosides from *S. japonicus* was studied with an in vivo lipid absorption assay. The result revealed that *S. japonicus* was a rich source of cerebrosides that contained considerable amounts of odd carbon chain sphingoid bases. The cumulative recoveries of d17:1 and d19:2 consisting cerebrosides were 0.31 ± 0.16% and 0.32 ± 0.10%, respectively, for 24 h after administration. In addition, dietary supplementation with sea cucumber cerebrosides to hairless mouse improved the skin barrier function and increased the short-chain fatty acid content in caecal fraction, which demonstrated its effects on host.

An *anteiso*-type regio-isomer on the LCB moiety HLC-2-A (**102**) from the extract of the sea cucumber *Holothuria leucospilota* were isolated from its glucocerebroside molecular species HLC-2 (**104**), composed of *iso*- and *anteiso*-isomers [56]. Other glucocerebroside molecular species HLC-1(**103**) and HLC-3 (**105**) were indicated together with HLC-2 (Figure 8).

**Figure 8.** Cerebrosides from the sea cucumber *Holothuria leucospilota*.

Sugavara et al. reported the sphingoid base composition of cerebrosides from sea cucumber (species was not identified) and their cytotoxicity against human colon cancer cell lines [57]. The composition of sphingoid bases obtained from a sea cucumber was different from that of mammals, and the major constituents were supposed from mass spectra as containing branched C-17–C-19 alkyl chains with 1–3 double bounds. The viability of DLD-1, WiDr, and Caco-2 cells treated with sea cucumber sphingoid bases was reduced in a dose-dependent manner and was similar to that of cells treated with sphingosine. The sphingoid bases induced such a morphological change as condensed chromatin fragments and increased caspase-3 activity, indicating that these sphingoid bases reduced the cell viability by causing apoptosis in the above-mentioned cells.

The galactocerebroside BAC-4-4a (**106**) was isolated from its parent galactocerebroside molecular species BAC-4 (**107**), which was obtained from the extract of the sea cucumber *Bohadschia argus* [58]. BAC-4 was obtained together with earlier known glucocerebroside molecular species [53,54,56]. The structure of **106** was determined as (2*S*,3*R*,4*E*)-1-*O*-(β-D-

galactopyranosyl)-2[(2*R*,15*Z*)-2-hydroxytetracosenoylamino]-4-heptadecene-1,3-diol (Figure 9). Before this study, galactocerebrosides were not found in sea cucumbers.

**Figure 9.** Cerebrosides from the sea cucumbers *Bohadschia argus* and *Acaudina molpadioides*.

The cerebroside molecular species AMC-2 (**108**) was isolated from the extract of the sea cucumber *Acaudina molpadioides* [59]. The amide-linked fatty acid units were established to contain four saturated and monounsaturated α-hydroxy fatty acids, the long-chain dihydroxy sphingoid base, having one double bond, and the glucose residue (Figure 9). It was shown the anti-fatty liver activity of 108 in rats with fatty liver, induced by orotic acid. AMC-2 (**108**) significantly reduced hepatic triglyceride and total cholesterol levels at a diet supplement of 0.03% and 0.006%. The indexes of stearoyl–CoA desaturase activity and mRNA expression were significantly decreased by **108**. This indicated that AMC-2 (**108**) ameliorated nonalcoholic fatty liver disease through suppression of stearoyl–CoA desaturase activity and impaired the biosynthesis of monounsaturated fatty acids in the livers of the rats.

Glucocerebrosides from three specimens of sea cucumbers, specifically, *Acaudina molpadioides*, *Cucumaria frondosa*, and *Apostichopus japonicus*, were rapidly identified by liquid chromatography–ion trap–time-of-flight mass spectrometry [60]. Various long-chain bases of glucosylcerebrosides were detected in these sea cucumbers. Two of the most common LCBs were identified as 2-amino-1,3-dihydroxy-4-heptadecene (d17:1) and 4,8 sphingadienine (d18:2), which were acylated to form saturated and monounsaturated nonhydroxylated and monohydroxylated fatty acids with 18–25 carbon atoms. The glucocerebroside fractions were the most complicated in the sea cucumber *C. frondosa* and were the simplest in the sea cucumber *A. molpadioides*.

It was found that a continuous oral administration of cerebrosides obtained from the sea cucumber *Acaudina molpadioides* at the dose of 50 mg/kg body mass per day suppressed body weight loss through alleviating adipose atrophy in cancer-associated cachexia mice [61]. The long-chain base, hydrolyzed from the cerebroside, contains 2-amino-1,3-dihydroxy-4-heptadecene (d17:1), which is a typical predominant sphingoid base in sea cucumbers. The possible mechanism by which dietary cerebrosides prevent adipose atrophy in cancer-associated cachexia mice was related to reducing serum inflammatory cytokine levels, regulating over lipolysis, enhancing the function of lipogenesis, and decreasing the lipid over-utilization. To elucidate the structure–activity relationships of cerebrosides and their long-chain base, the antitumor activities were compared between them. The results indicated that LCBs exhibited a more prominent antitumor effect both in vivo and in vitro.

In addition, sea cucumber cerebrosides and their main structural units, long-chain bases, were obtained from *Acaudina molpadioides* and then administered to high fat dietinduced obese C57BL/6J mice at a diet supplement dosage of 0.025% for 5 weeks to evaluate their effects on obesity-related metabolic disorders [62]. Cerebrosides and long-chain bases significantly decreased epididymal adipose tissue weights, lowered hepatic triacylglycerol levels, and reduced serum glucose, insulin levels, and insulin resistance HOMA-IR index in mice. The activities of hepatic lipogenetic proteins including FAS, ME, and the mRNA levels encoding proteins SREBP-1c and FAS were reduced by cerebrosides and long-chain bases treatment. However, cerebrosides and LCBs showed no effect on the hepatic lipolysis pathway. Moreover, cerebrosides and LCBs efficiently upregulated the gene expression of SREBP-1c, FAS, ACC, ATGL, and HSL, and downregulated the gene expression of LPL and VLDL-r in the adipose tissue. These results demonstrated that cerebrosides and LCBs were effective in suppressing hepatic SREBP-1c mediated lipogenesis, inhibiting lipid uptake, and increasing TG catabolism in the adipose tissue. The ameliorative degree and regulatory mechanisms of these two groups of natural products were basically the same, suggesting that long-chain bases are the key active structural units of cerebrosides [62].

Three glucocerebrosides, CF-3-1, CF-3-2, and CF-3-3 (**109**–**111**), were isolated from the cerebroside fraction, which was obtained from the chloroform–methanol extract of the sea cucumber *Cucumaria frondosa* by La et al. [63]. The structures of these cerebrosides were determined as 1-*O*-β-D-glucopyranosides of (2*S*,3*S*,4*R*)-2-[(2*R*,15*Z*)-2-hydroxy-15 tetracosenoylamino]-14-methylhexadecane-1,3,4-triol (**109**), (2*S*,3*R*,4*E*)-2-[(2*R*,15*Z*)-2-hydroxy-15-tetracosenoylamino]-15-methyl-4-hexadecene-1,3-diol (**110**), and (**111**) (2*S*,3*R*, 4*E*,8*Z*)-2-[(2*R*,15*Z*)-2-hydroxy-15-tetracosenoylamino]-4,8-octadecadiene-1,3-diol (Figure 10). Compounds **110** and **111** were obtained as pure compounds for the first time.

**Figure 10.** Cerebrosides from the sea cucumber *Cucumaria frondosa*.

Three glucocerebroside molecular species (CFC-1, CFC-2, and CFC-3, **112**–**114**) were isolated from total cerebrosides from the sea cucumber *Cucumaria frondosa* by Xu et al. (Figure 10) [64]. The structures of these substances were elucidated on the basis of spectroscopic and chemical evidence: fatty acids were identified mainly as saturated (C22:0 and C18:0), monounsaturated (C24:1 and C20:1), and α-hydroxylated derivatives (C24:1h, C23:0h, C23:1h, and C22:0h), the LCB were identified as dihydroxy (d17:1, d18:2, and d18:1) and trihydroxy (t17:0 and t16:0) compounds. The composition analysis of long-chain bases showed that the ratio of d18:2 and d17:1 was approximately 2:1. Four glucocerebrosides and long-chain bases from sea cucumber *Cucumaria frondosa* were evaluated for their cytotoxic activities against Caco-2 colon cancer cells in in vitro assays. The obtained results indicated that both glucocerebrosides and LCB demonstrated an inhibitory effect on cell proliferation. Moreover, **114** was the most effective substance from these four glucocerebrosides in the Caco-2 cell viability test. The inhibitory effects of long-chain bases were much stronger than glucocerebrosides.

Glucocerebrosides, isolated from the sea cucumber *Cucumaria frondosa* (CFC), were investigated on their antiadipogenic activity in vitro [65]. These glucocerebrosides inhibited the lipid accumulation of 3T3-L1 cells and suppressed PPARγ and C/EBPα expressions, which confirmed their antiadipogenic effect. Furthermore, CFCs suppressed lipogenesis in mature adipocytes. Glucocerebrosides enhanced β-catenin expression, promoted its nuclear translocation, and upregulated the expression of CCND1 and c-myc, two target genes of β-catenin. Moreover, after cells were treated with the β-catenin inhibitor 21H7, β-catenin nuclear translocation and transcription activity can be recovered by CFC. These findings suggested that glucocerebrosides from *Cucumaria frondosa* promoted the activation of the WNT/β-catenin pathway. Additionally, CFCs enhanced the expressions of Wnt-receptor frizzled-like protein variant 1(FZ1), low-density lipoprotein receptor-related proteins LRP5, and LRP6, while they had no effect on the expressions of Wnt10b and GSK3β proteins. These findings also confirmed that glucocerebrosides exhibit their antiadipogenic activity through enhancing the activation of the WNT/β-catenin pathway, which was mediated by FZs and LRPs.

Over the past two decades, about a hundred individual cerebrosides and their molecular species were isolated from starfish and sea cucumbers. The isolated compounds contain both sphingosine and phytosphingosine bases of *normal*-, *iso*- and *anteiso*-types. In most cases, long-chain bases include from 16 to 19 carbon atoms, but there were also longer ones, up to C-22. In addition, many LCBs were unsaturated and contained one or two double bonds. In particular, (4*E*,8*E*,10*E*)-sphinga-4,8,10-trienine; (4*E*,8*E*,10*E*)-9-methylsphinga-4,8,10-trienine; (4*E*,13*Z*)-sphinga-4,13-dienine; (4*E*,15*Z*)-sphinga-4,15-dienine; and (9*Z*)-4-hydroxy-9-sphingenine long-chain bases were often found. At the same time, unique oxidized LCB (4*E*,9*E*)-9-methyl-8,11-dihydroxy-sphinga-4,9-dienine, and (4*E*,10*E*)- 9-methyl-8,9-dihydroxy-sphinga-4,10-dienine were found in the sea cucumber *Stichopus japonicus*.

In most cases, the fatty acids in the cerebrosides were long-chain C-22–C-24 (2*R*)- 2-hydroxy acids of *normal*-, *iso*-, and *anteiso*-types. However, shorter FAs such as C-18, C-16, and even C-14 were also found. Some fatty acids in the isolated cerebrosides were unsaturated and most of them had the (15*Z*)-double bond. In contrast to cerebrosides from starfish, cerebrosides from sea cucumbers contained non-α-hydroxylated FA with different long polymethylene chains.

The carbohydrates in cerebrosides of starfish and sea cucumbers were represented by the β-D-glucopyranose and, more rarely, the β-D-galactopyranose. Thus far, no other types of monosaccharide residues have been found in cerebrosides of starfish and sea cucumbers. In addition, cerebrosides lactosides (with Gal-(1→4)-Glc-(1→1)-Cer moieties) were isolated from the starfish *Luidia maculata*. Other variants of cerebroside biglycosides or oligoglycosides in starfish and sea cucumbers have not been found.

The following types of biological activity of cerebrosides from starfish and sea cucumbers were established: *i.* growth-promoting activity on *Brassica campestris*, *ii.* cytotoxic activity against epidermal carcinoma of the mouth KB cells and rat glioma C6 cells; and *iii.*proangiogenic activity. More detailed data are given in Table 1. The conducted studies showed the promising prospects of the practical use of cerebrosides of starfish and sea cucumbers. Accordantly, further expansion of the studies on the biological activity of this class of glycolipids is required, as well as additional data concerning the molecular mechanisms of their action.


#### **Table 1.** Composition and biological activity of starfish and sea cucumber sphingolipids mentioned in this review.


**Table 1.** *Cont.*

#### **4. Gangliosides**

Gangliosides are known as additionally hydroxylated derivatives of cerebrosides with one or more sialic acid residues in their carbohydrate chains. Sialic acids are a group of higher carbohydrates with nine carbon atoms, which includes several dozens of derivatives of neuraminic acid (NeuAc) [87]. Gangliosides were so named for the first time because they were isolated from brain ganglion cells. It is considered that gangliosides are metabolites of vertebrates; however, they were also found in all classes of Echinoderms and may indicate a high organization of their nervous system. To designate gangliosides, they most often use abbreviated names according to Svennerholm's nomenclature, in which gangliosides are divided into so-called series, indicated by the number of sialic acid units and their position in the carbohydrate chain. Gangliosides are biosynthesized from the corresponding cerebrosides by sialyltransferases on the inner plasma membrane or in the Golgi apparatus, and then they are incorporated into the plasmatic membrane, where these glycosphingolipids perform their biological functions [88]. Gangliosides play an important role in binding to some lectins and affect the activity of receptor protein kinases, taking part in the transmission of cellular signals. In addition, gangliosides, similar to other sphingolipids and cholesterol, play an important role in stabilizing plasma membranes with positive curvature and also affect the surface charge of the membrane. Finally, gangliosides can act as receptors for viruses, bacteria, and toxins, thus being part of the immune system [88].

It is known that gangliosides play an extremely important role in the development of various neurodegenerative diseases, as well as in the regulation of proliferation and energy metabolism of tumor cells [89–91].

Thus, the search for new structural types of gangliosides in echinoderms, as well as a comprehensive study of their biological activity, is an actual scientific task.

#### *4.1. Class Asteroidea*

The ganglioside molecular species, AG-1, were obtained from the whole body of the starfish *Acanthaster planci* [70]. Enzymatic hydrolysis by endoglycoceramidase gave an oligosaccharide and ceramides, quantitatively. The oligosaccharide moiety was determined mainly by 2D-NMR experiments as β-Fucf-(1→4)-α-Galp-(1→4)-α-NeuAc-(2→3)-β-Galp- (1→4)-Glcp. The sphingoid moiety was elucidated as the mixture of (2*S*,2'*S*,3*S*,4*R*)-2- ((2*R*)-2-hydroxydocosanoylamino)-1,3,4-trihydroxyhexadecane and (2*S*,2'*S*,3*S*,4*R*)-2-((2*R*)- 2-hydroxytetracosanoylamino)-1,3,4-trihydroxyhexadecane. Reversed-phase HPLC of AG-1 gave two kinds of gangliosides named acanthagangliosides I (**115**) and J (**116**). It is clear that the oligosaccharide moiety of AG-1 is different in its terminal monosaccharide when compared with AG-2 and AG-3, which were isolated from *A. planci* earlier [71,72]. The terminal β-Galf of AG-2 and AG-3 is linked to C-3 of α-Galp, while the terminal β-Fucf of AG-1 is linked to C-4 of α-Galp. This interesting difference in terminal sugar linkages seems to be derived from the coexistence of different glycosyltransferases, namely, β-1,3-galactofuranosyl transferase and β-1,4-fucofuranosyl transferase. The gangliosides of *A. planci* characteristically have a terminal furanose-type sugar unit (Figure 11).

It was found by performing 1H NMR and saturation transfer difference (STD) NMR experiments that AG2 pentasaccharide (structure not shown) binds to human Siglec-2 (a mammalian sialic acid-binding protein expressed on B-cell surfaces, which involved in the modulation of B-cell mediated immune response [73]. STD NMR experiments indicated that the C-7–C-9 carbohydrate-chain and the acetamide moiety of the central sialic acid residue were located in the binding face of human Siglec-2. The binding epitope of AG2 pentasaccharide to human Siglec-2 was determined as the α-Galp(1→4)-α-NeuAc-(2→3)- Galp unit. The information concerning the binding epitope of AG2 pentasaccharide is of value toward the development of potent Siglec-2 inhibitors.

**Figure 11.** Gangliosides from the starfish *Acanthaster planci* and *Evasterias echinosoma*.

Gangliosides molecular species were isolated from the starfish *Evasterias echinosoma*, and their structures were elucidated [66]. Two major sphingolipids (**117**, **118**) were found to be disialogangliosides, whose carbohydrate chain is based on the trisaccharide β-*N*-acylgalactopyranosaminyl-(1→3)-β-galactopyranosyl-(1→4)-β-glucopyranose (acyl is formyl or acetyl). Both residues of 8-*O*-methyl-*N*-acetylneuraminic acid are attached to the *N*-acylgalactosamine residue at positions C-3 and C-6. Compound **118** is the first example of when an *N*-formyl derivative of an amino sugar was found in gangliosides. The lipid part of the gangliosides molecular species consists of monounsaturated sphingoid base and nonhydroxylated fatty acids (mainly, palmitic and stearic acids) (Figure 11).

The ganglioside (**119**) was isolated from the starfish *Linckia laevigata*, and its structure was determined by spectroscopic and chemical methods [75]. The carbohydrate part was proved to be 8-*O*-Me-(*N*-glycolyl-α-D-neuraminosyl)-(2→3)-β-D-galactopyranosyl-(1→4) β-D-glucopyranoside. The lipid moiety of this ganglioside consists of nonhydroxylated fatty acids (the major component is palmitic acid) and *iso*-C18:1-sphingenine. Based on the structure of the carbohydrate moiety, ganglioside **119** belongs to the hematoside type, characteristic of erythrocytes of vertebrates. It differs from the other known hematosides in the nature of the sialic acid. A hematoside with 8-*O*-methyl-*N*-glycolylneuraminic acid unit was found for the first time (Figure 12).

Continuing research on gangliosides of the starfish *Linckia laevigata,* ganglioside molecular species LLG-5 (**120**) were obtained from the water-soluble portion of its lipid fraction [76]. On the basis of spectroscopic and chemical data, the structure of **120** was elucidated as 8-*O*methyl-(*N*-glycolyl-α-D-neuraminosyl)-(2→11)-(*N*-glycolyl-α-D-neuraminosyl)-(2→11)-(*N*glycolyl-α-D-neuraminosyl)-(2→3)-β-D-galactopyranosyl-(1→4)-β-D-glucopyranoside of a ceramide composed of phytosphingosines and 2-hydroxy *n*-fatty acids. The major components of the fatty acids and long-chain bases moieties of 120 were identified as (2*R*)- 2-hydroxy *n*-docosanoic acid and (2*S*,3*S*,4*R*)-2-amino-1,3,4-octadecanetriol, respectively. This was the first isolation and characterization of a trisialo-ganglioside from Asteroidea (Figure 12). Furthermore, **120** is a new ganglioside molecular species containing a 2→11 linked trisialosyl moiety. The ganglioside molecular species LLG-5 (**120**) exhibited neuritogenic activity in rat pheochromocytoma PC12 cells in the presence of nerve growth factor (NGF). The proportion of cells with neurites longer than the diameter of the cell body at a concentration of 10 μM or 120 was 59.3% when compared with the control (NGF, 5 ng/mL: 20.6%). Furthermore, their effect was greater than that of the mammalian ganglioside GM1 (47.0%).

**Figure 12.** Gangliosides from the starfish *Linckia laevigata*.

In addition, the hematoside-type ganglioside LLG-1 (**121**) was obtained from the polar lipid fraction of the starfish *Linckia laevigata* [77]. The structure of LLG was elucidated on the basis of spectroscopic and chemical evidence as 1-*O*-[(*N*-glycolyl-α-D-neuraminosyl)- (2→3)-β-D-galactopyranosyl-(1→4)-β-D-glucopyranosyl]-ceramide. The ceramide moiety was composed of 2-hydroxy fatty acids and phytosphingosine units (*normal*- and *iso*-type long-chain bases). This was the first report on the isolation and structure elucidation of naked hematoside-type ganglioside from echinoderms (Figure 12).

Two monomethylated GM3-type ganglioside molecular species (**122** and **123**) were isolated from the extract of the starfish *Luidia maculata* [68]. The structures of these gangliosides were determined as 1-*O*-[8-*O*-methyl-(*N*-acetyl-α-D-neuraminosyl)-(2→3)-β-Dgalactopyranosyl-(1→4)-β-D-glucopyranosyl]-ceramide (**122**) and 1-*O*-[8-*O*-methyl-(*N*glycolyl-α-D-neuraminosyl)-(2→3)-β-D-galactopyranosyl-(1→4)-β-D-glucopyranosyl] ceramide (**123**). The ceramide moieties were composed of heterogeneous nonhydroxylated fatty acid, 2-hydroxy fatty acid, sphingosine, and phytosphingosine units. Compound **122**, designated as LMG-3, represented new ganglioside molecular species. Compound **123** was identified as a known ganglioside molecular species (Figure 13).

**Figure 13.** Ganglioside molecular species from the starfish *Luidia maculata*.

In addition, the GD3-type ganglioside molecular species LMG-4 (124) was obtained from the extract of the starfish *L. maculata* [69]. The structure of this compound was determined on the basis of spectroscopic and chemical evidence to be 1-*O*-[(*N*-acetyl-α-Dneuraminosyl)-(2→8)-(*N*-acetyl-α-D-neuraminosyl)-(2→3)-β-D-galactopyranosyl-(1→4)-β-D-glucopyranosyl]-ceramide. The ceramide moiety was composed of 2-hydroxy fatty acid and phytosphingosine moieties. GD3-type ganglioside was isolated and its particular structure elucidated for the first time from echinoderms (Figure 13). LMG-4 (**124**) exhibited neuritogenic activity toward the rat pheochromocytoma PC12 cells in the presence of NGF. The proportion of the neurite-bearing cells of **124** at a concentration of 10 μM was 47.7%, in comparison with the control (NGF, 5 ng/mL: 20.6%). The effect of **124** was the same as that of the mammalian ganglioside GM1 (47.0%).

Mono- and disialogangliosides (**125**, **126**) were isolated from gonads of the starfish *Evasterias retifera* [67]. Their structures were elucidated by spectroscopic and chemical evidence, including enzymatic hydrolysis with neuraminidase. The monosialoganglioside has the structure α-8-*O*-Me-NeuGc-(2→3)-β-GalNAc-(1→3)-β-Gal-(1→4)-β-Glc-(1→1)- Cer, while the disialoganglioside contains an additional NeuAc residue, which glycosylates GalNAc in position C-6. The lipid moieties of both gangliosides contain phytosphingosine bases (mainly C18:0) and two types of fatty acids, nonhydroxylated (mainly C16:0 and C18:0) and α-hydroxylated (mainly α-hydroxy-C16:0) (Figure 14).

The molecular species GP-3 (**127**) was obtained from the starfish *Patiria (=Asterina) pectinifera* [74]. The structure of the ganglioside was determined as 1-*O*-α-L-arabinofuranosyl-(1→3) α-D-galactopyranosyl-(1→4)-(*N*-acetyl-α-D-neuraminosyl)-(2→6)-β-D-galactofuranosyl-(1→3)-[α-L-arabinofuranosyl-(1→4)]-α-D-galactopyranosyl-(1→4)-(*N*-acetyl-α-D-neuraminosyl)-(2→3)-β-D-galactopyranosyl-(1→4)-β-D-glucopyranoside of ceramide composed of heterogeneous (2*S*,3*S*,4*R*)-phytosphingosine (*iso*-C-17-phytosphingosine as the major component) and (2*R*)-2-hydroxy fatty acid units (docosanoic acid as the major component) (Figure 14). Compound **127** represents new ganglioside molecular species possessing two residues of sialic acids at the inner part of the sugar moiety. A ganglioside molecular species GP-3

(**127**) exhibits neuritogenic activity toward the rat pheochromocytoma cell line PC12, in the presence of NGF. The proportion of the cells with neurite longer than the diameter of the cell body at the use of **127** at a concentration of 10 μM was 38.2% when compared with the control (NGF, 5 ng/mL: 20.6%). The effect of **127** was lower than that of the mammalian ganglioside GM1 (47.0%).

**Figure 14.** Gangliosides from the starfish *Evasterias retifera* and *Patiria (*=*Asterina) pectinifera*.

Three ganglioside molecular species PNG-1 (**128**), PNG-2A (**129**), and PNG-2B (**130**) were isolated from pyloric caeca of the starfish *Protoreaster nodosus* [78]. Their structures as 1-*O*-[8-*O*-methyl-(*N*-acetyl-α-neuraminosyl)-(2→3)-β-galactopyranosyl]-ceramide (**128**), 1-*O*-[β-galactofuranosyl-(1→3)-α-galactopyranosyl-(1→4)-8-*O*-methyl-(*N*-acetyl-αneuraminosyl)-(2→3)-β-galactopyranosyl]-ceramide (**129**), and 1-*O*-[β-galactofuranosyl- (1→3)-α-galactopyranosyl-(1→9)-(*N*-acetyl-α-neuraminosyl)-(2→3)-β-galactopyranosyl] ceramide (**130**) were elucidated by a combination of spectroscopic and chemical methods. The ceramide moieties of ganglioside molecular species consisted of (2*S*,3*S*,4*R*) phytosphingosines (*iso*-C-18-phytosphingosine as the major component) and (2*R*)-2-hydroxy fatty acid units (docosanoic acid as the major component). PNG-2A (**129**) and PNG-2B (**130**) represent the first GM4 elongation products in nature (Figure 15).

**Figure 15.** Gangliosides molecular species from the starfish *Protoreaster nodosus*.

#### *4.2. Class Holothuroidea*

The ganglioside molecular species HPG-7 (**131**) was isolated from the chloroform– methanol extract of the sea cucumber *Holothuria pervicax* [79]. On the basis of the spectroscopic and chemical evidence, the structure of the major component of **131** was determined as 1-*O*-[α-L-fucopyranosyl-(1→4)-(*N*-acetyl-α-D-neuraminosyl)-(2→11)-(*N*-glycolyl-α-Dneuraminosyl)-(2→4)-(*N*-acetyl-α-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-(2*S*,3*S*,4*R*)- [(2*R*)-2-hydroxytetracosanoylamino]-14-methyl-hexadecane-1,3,4-triol (Figure 16). The trisialo-ganglioside was isolated for the first time from sea cucumbers. HPG-7 (**131**) was studied for neuritogenic action toward the PC12 rat pheochromocytoma cell line. It was shown that **131** does not have neuritogenic activity, in comparison with control, at a concentration of above 10 μg/mL, similar to three other ganglioside molecular species (HPG-1, HPG-3, and HPG-8) [80].

**Figure 16.** Ganglioside molecular species from the sea cucumber *Holothuria pervicax*.

Three ganglioside molecular species, HLG-1 (**132**), HLG-2 (**133**), and HLG-3 (**134**), were isolated from the extract of the sea cucumber *Holothuria leucospilota* [81]. Structures of these gangliosides were determined as 1-*O*-[(*N*-glycolyl-α-D-neuraminosyl)-(2→6)-β-Dglucopyranosyl]-ceramide (**132**), 1-*O*-[(*N*-glycolyl-α-D-neuraminosyl)-(2→4)-(*N*-acetyl-α-

D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramide (**133**), and 1-*O*-[α-L-fucopyranosyl- (1→11)-(*N*-glycolyl-α-D-neuraminosyl)-(2→4)-(*N*-acetyl-α-D-neuraminosyl)-(2→6)-β-Dglucopyranosyl]-ceramide (**134**), respectively. The ceramide moieties were composed of phytosphingosines or sphingosines and 2-hydroxy fatty acids (Figure 17). Compounds **133** and **134** represent new ganglioside molecular species. These three substances showed slight neuritogenic activity toward the rat pheochromocytoma cell line PC12 cell in the presence of NGF.

**Figure 17.** Ganglioside molecular species from the sea cucumber *Holothuria leucospilota*.

The ganglioside molecular species SJG-2 (**135**) was obtained from the extract of the sea cucumber *Stichopus japonicus* [82]. On the basis of spectroscopic and chemical studies, the structure of SJG-2 (**135**) was determined as α-NeuAc-(2→4)-α-NeuAc-(2→3)-β-Gal- (1→8)-α-NeuAc-(2→3)-β-GalNAc-(1→3)-β-Gal-(1→4)-β-Glc-(1→1)-Cer. The ganglioside **135**, possessing a unique carbohydrate moiety, is the first corresponding substance with a branched sugar chain moiety and *N*-acetylgalactosamine residue isolated from sea cucumbers (Figure 18). Ganglioside SJG-2 (**135**) exhibited neuritogenic activity toward the rat pheochromocytoma cell line PC12 cells in the presence of NGF. The proportion of neurite-bearing cells at the use of SJG-2 (64.8 ± 7.6%) was larger than that induced by the previously isolated SJG-1 [83], (35.4 ± 4.0%) when compared with the control (NGF, 5 ng/mL: 20.6 ± 2.2%). Furthermore, the effect of SJG-2 (**135**) was more considerable than that of the mammalian ganglioside GM1 (47.0 ± 2.5%).

**Figure 18.** Ganglioside molecular species from the sea cucumber *Stichopus japonicus*.

Three ganglioside molecular species, SCG-1 (**136**), SCG-2 (**137**), and SCG-3 (**138**), were isolated from the extract of the sea cucumber *Stichopus chloronotus* [84]. On the basis of spectroscopic and chemical evidence, the structures of these gangliosides were determined to be 1-*O*-[(*N*-glycolyl-α-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramide (**136**), 1-*O*-[8-*O*sulfo-(*N*-acetyl-α-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramide (**137**), and 1-*O*- [α-L-fucopyranosyl-(1→11)-(*N*-glycolyl-α-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]- ceramide (**138**). The ceramide moieties were composed of isomeric long-chain bases and fatty acid units. The molecular species **138** is the first representative of gangliosides containing fucopyranose in the sialosyl trisaccharide moiety (Figure 19). Gangliosides **136**–**138** exhibited neuritogenic activity toward the rat pheochromocytoma PC12 cells in the presence of NGF. The proportions of the neurite-bearing cells at a concentration of **136**–**138** of 3.3 μg/mL were 34.1%, 24.4%, and 24.5%, respectively. These effects were compared with that of the mammalian ganglioside GM1 (22.1% at a concentration of 3.3 mg/mL).

Three monosialo-gangliosides, CEG-3 (**139**), CEG-4 (**140**), and CEG-5 (**141**), were obtained, together with two previously known gangliosides, SJG-1 (**142**, structure not shown, [83]) and CG-1 (**143**, structure not shown, [92]), from the extract of the sea cucumber *Cucumaria echinata* [85]. In addition, three disialo- or trisialo-gangliosides, CEG-6 (**144**), CEG-8 (**145**), and CEG-9 (**146**)**,** were also obtained along with the known ganglioside, HLG-3 (**134**, [81]) from this species of sea cucumbers [86]. Structures of these gangliosides were determined as 1-*O*-[(4-*O*-acetyl-α-L-fucopyranosyl)-(1→11)-(*N*-glycolylα-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramide (**139**), 1-*O*-[α-L-fucopyranosyl- (1→11)-(*N*-glycolyl-α-D-neuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramides (**140**, **141**), 1-*O*-[α-L-fucopyranosyl-(1→11)-(*N*-glycolyl-α-D-neuraminosyl)-(2→4)-(*N*-acetyl-α-Dneuraminosyl)-(2→6)-β-D-glucopyranosyl]-ceramide (**144**), and homologous to each other 1-*O*-[(*N*-glycolyl-D-neuraminosyl)-(2→11)-(*N*-glycolyl-D-neuraminosyl)-(2→4)-(*N*-acetyl-D-neuraminosyl)-(2→6)-D-glucopyranosyl]-ceramides (**145**, **146**). The ceramide moieties of each compound were composed of sphingosine or phytosphingosine bases and 2-hydroxyor nonhydroxylated fatty acid units (Figure 20). Gangliosides **134**, **139**–**146** demonstrated neuritogenic activity toward the rat pheochromocytoma cell line PC12 in the presence of NGF. The proportions of cells with neurites longer than the diameter of the cell body after the treatment with compounds **134**, **139**–**146** at concentration of 10 μM were of 40.2%, 50.8%, 34.0%, 35.7%, 39.1%, 43.0%, 43.0%, 40.2%, and 35.1%, respectively, in comparison with the control experiments (NGF, 5 ng/mL: 7.5%). The effects of **134**, **139**, and **142**–**145** were stronger than that of the mammalian ganglioside GM1 (35.6%). Compound **139** with an acetyl group at the terminal fucopyranosyl unit showed the most potent activity.

**Figure 19.** Ganglioside molecular species from the sea cucumber *Stichopus chloronotus*.

Enantiomeric pairs of sialic acids (D- and L-NeuAc) were converted to D- and Larabinose, respectively, by chemical degradation [93]. Using this approach, the absolute configurations of the sialic acid residues NeuAc and NeuGc as D-forms were determined in the gangliosides from the sea cucumber *Cucumaria echinata*. Although naturally occurring sialic acids have been believed to have D-configurations on the basis of biosynthetic evidence, this is the first report describing the determination of the absolute configuration of the sialic acid residues in gangliosides using chemical methods.

Starfish and sea cucumbers gangliosides remain to be less studied, in comparison with cerebrosides. At the same time, about 30 new compounds and/or molecular species have been isolated since 2000. The carbohydrate chains of the starfish and sea cucumber gangliosides differ markedly from the carbohydrate chains of mammals as well as from each other. Generally, besides sialic acid residues, these compounds contain lactoside fragment (Gal-(1→4)-Glc-(1→1)-Cer) and analogous fragment additionally glycosylated with galactosamine (GalNAc-(1→3)-Gal-(1→4)-Glc-(1→1)-Cer). Part of them are derivatives of galactosylceramides having (Gal-(1→1)-Cer) moiety.

In the sea cucumbers gangliosides, containing fragments of only two cerebrosides were found: lactosides glycosylated with galactosamine (GalNAc-(1→3)-Gal-(1→4)-Glc- (1→1)-Cer), and glucosylceramides (Glc-(1→1)-Cer).

Both starfish and sea cucumber gangliosides contain unusual sialic acid residues, including sialic acids within carbohydrate chains as well as additional monosaccharide residues and unusual types of glycosidic bonds between them. For example, terminal β-D-Fuc*<sup>f</sup>* was found in the gangliosides from the starfish *Acanthaster planci*, 8-*O*-Me-NeuAc and 8-*O*-Me-NeuGc were found in the gangliosides from the starfish *Linckia laevigata* as well as the glycosidic bond 2→11 between sialic acid residues. The ganglioside from the starfish *Evasterias echinosoma* contains an unusual β-D-*N*-formyl-galactosamine residue, while the carbohydrate chains from gangliosides of the starfish *Patiria (=Asterina) pectinifera* bears the terminal α-L-arabinofuranose residue and has three forms of galactose (β-D-Galp, β-D-Galf, and α-D-Galp). These gangliosides contain the maximum number of monosaccharide residues (up to nine), in comparison with other echinoderm gangliosides.

Gangliosides with the terminal α-L-Fuc*<sup>p</sup>* were identified in several species of sea cucumbers along with NeuAc and NeuGc residues within carbohydrate chains. A unique 8- *O*-sulfo-NeuAc residue was found in the corresponding substances from the sea cucumber *Stichopus chloronotus*. The maximum length of the carbohydrate chain in the sea cucumbers gangliosides was found in the ganglioside from *Stichopus japonicus*, which contained seven monosaccharide residues.

Lipid parts of gangliosides from both starfish and sea cucumbers were similar and contained both sphingosine and phytosphingosine bases of *normal*-, *iso*- and *anteiso*-types. Predominantly (2*R*)-2-hydroxy fatty acids of the normal type were found in these substances. For gangliosides of starfish and sea cucumbers, only one type of biological activity was studied, neuritogenic activity toward the rat pheochromocytoma cell line PC12 in the presence of NGF. In a number of cases, starfish and sea cucumbers gangliosides showed a higher neuritogenic effect at concentration 10 μM than the mammalian ganglioside GM1, while some gangliosides exhibited slighter action at the same concentration.

#### **5. Conclusions**

To the best of our knowledge, sphingolipids of 15 starfish and 9 sea cucumbers, mainly common Pacific Ocean inhabitants, have been studied (Table 1). In total, these 24 echinoderm species were used for the isolation and identification of about 150 sphingolipids. This indicates that echinoderms and, in particular, starfish and sea cucumbers are a rich source of sphingolipids, structures of which may differ markedly from the corresponding metabolites of plants and terrestrial animals.

Ceramides are the least studied group of echinoderms sphingolipids. Moreover, since 2000, only studies on starfish ceramides have been carried out. Nevertheless, a big variety of structural types of the isolated ceramides was detected, for instance, sphingosine and phytosphingosine LCBs of various lengths, *normal*-, *iso*-, and *anteiso*-types, often having one or two additional double bonds, were found in starfish ceramides. Fatty acid residues in starfish ceramides were most often identified as (2*R*)-2-hydroxy derivatives of various lengths (usually from C-18 to C-22) with normal hydrocarbon chains, which can also contain one additional double bonds. The "gray spot" in the study of starfish ceramides is the lack of data on biological activity, with the exception of the stimulating root growth of *Brassica campestris* activity by ceramides from *Asterias amurensis*.

Cerebrosides are the most studied class of starfish and sea cucumbers sphingolipids. Generally, about one hundred individual cerebrosides and their molecular species have been isolated from these animals. As in ceramides, sphingosine and phytosphingosine LCBs of various lengths with *normal*-, *iso*-, and *anteiso*-structures were found in starfish and sea cucumber cerebrosides. Unique oxidized sphingosine LCBs with additional hydroxy groups at either C-8 and C-9 or C-8 and C-11 were indicated in the sea cucumber *Stichopus japonicus*. Mainly saturated and monounsaturated (2*R*)-2-hydroxy fatty acids with normal hydrocarbon chains having various lengths were identified as constituents of these cerebrosides, but nonhydroxylated FAs were sometimes also detected. Almost all the isolated cerebrosides were monoglycosides and contained glucose or galactose residues. Cerebroside lactosides were isolated from the starfish *Luidia maculata*.

The following types of biological activities of starfish and sea cucumbers cerebrosides were studied: growth-promoting activity of *Brassica campestris*, anti-fatty liver activity in rats treated by orotic acid, alleviating adipose atrophy action in cancer-associated cachexia mice, effects on obesity-related metabolic disorders in mice, cytotoxic activities against KB, rat glioma C6 cells, and colon cancer Caco-2cells, and proangiogenic action. As result, it was shown that starfish and sea cucumbers cerebrosides possess various types of biological activities that are important for their practical application in the human diet and in the composition of food supplements (Table 1).

Starfish and sea cucumber gangliosides were also studied for some species, and their structural diversity was proved to be great. Carbohydrate chains of starfish and sea cucumbers gangliosides have interesting structural features and differ from gangliosides of terrestrial animals. Really, the residues of β-D-Fuc*f*, 8-*O*-Me-NeuAc, and 8-*O*-Me-NeuGc, β-D-*N*-formyl-galactosamine, as well as terminal α-L-Ara*<sup>f</sup>* were recently found in the starfish gangliosides. In gangliosides from holothurians (sea cucumbers), the terminal α-L-Fuc*p*, α-L-FucAc*p*, and 8-*O*-sulfo-NeuAc were detected.

For starfish and sea cucumbers gangliosides, only one type of biological activity was studied, namely, neuritogenic activity toward the rat pheochromocytoma cell line PC12 in the presence of NGF. Therefore, further research of other types of biological activities including antitumor and anti-inflammatory properties might be of interest. It is noteworthy that the starfish and sea cucumber gangliosides, as a rule, are species specific. Therefore, they could be taxonomic markers, such as some unusual starfish polar steroidal compounds [94,95] and sea cucumber triterpene glycosides [96]. However, the structures of gangliosides were less studied than those of other secondary metabolites of starfish and sea cucumbers and require further research.

Previously, we studied the metabolic profile of polar steroid compounds of three species of starfish and their changes under stress conditions, as well as the metabolic profile of triterpene glycosides from the sea cucumber *Eupentacta fraudatrix* [8–13]. The study of the metabolomic profiles of sphingolipids and their changes under various environmental conditions can also be one of the directions of metabolomics research. However, first of all, it is necessary to systematize the literature data on the structures of all types of sphingolipids, including ceramides, cerebrosides, and gangliosides, in these animals. We believe this review can help meet this challenge.

**Author Contributions:** Writing—original draft preparation and writing—review and editing, T.V.M., A.A.K. and N.V.I.; conceptualization and validation, V.A.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was carried out with the support of the Russian Science Foundation (RSF) Grant Number 20-14-00040.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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