Methyl Ether-Derivatized Sterols and Coprostanol Produced via Thermochemolysis Using Tetramethylammonium Hydroxide (TMAH)

Abstract

Sterols are widely distributed in nature from lipids in organisms to sediments. As a conventional method, extraction and derivatization with TMS have been applied for sterol analysis, requiring a long preparation time for gas chromatography–mass spectrometry analysis. In this study, for sterol analysis, thermochemolysis using tetramethylammonium hydroxide (TMAH) was applied. This method performs hydrolysis and methylation simultaneously; thus, free and ether-bonding sterols can be analyzed as sterol methyl ethers in a relatively short time period. A sediment sample from a tideland (the Yatsu tideland, Japan) was analyzed using the TMAH method, and we detected more than 10 sterols, which include cholest-5-en-3β-ol (cholesterol), 24-ethylcholest-5-en-3β-ol (sitosterol), 24-methylcholesta-5,22E-3β-ol (brassicasterol), 24-ethylcholesta-5,24(28)Z-dien-3β-ol (isofucosterol), 4α,23,24-trimethyl-5α(H)-cholest-22E-en-3β- ol (dinosterol), and 5β(H)-cholestan-3β-ol (coprostanol). The detection of the various sterols can be attributed to multiple natural and artificial sources around the Yatsu tideland. In this paper, the mass spectra of these sterols are provided together with an interpretation of their fragmentation patterns. Additionally, the fecal pollution in the Yatsu tideland is discussed in the context of the detection of coprostanol.

1. Introduction

Sterols exist in the membranes of eukaryotic organisms, including microorganisms and terrestrial plants [1,2]. Thus, sterols have been detected in various natural environments, such as in lake and ocean waters and in sediments [3,4,5,6,7]. These natural sterols can be used to estimate ecological sources because sterol structures differ depending on the types of microorganisms [1,8]. Moreover, they have been used as environmental tracers, such as in degradation processes and for the reconstruction of paleoenvironments [9,10,11].

Sterols can be extracted with organic solvents, such as chloroform and methanol. A hydrolysis step is required if necessary. For sterol analysis by gas chromatography (GC)–mass spectrometry (MS), the extracted sterols must be derivatized to obtain enough volatility to pass through the GC. By derivatization, in the case of sterols, hydroxyl functional groups will be changed to more low-polarity functional groups. Typically, derivatization techniques are categorized into three types, namely, silylation, acylation, and alkylation. Although acylation is also used, silylation is most commonly used for derivatization.

Silylation is effective for various functional groups, including carboxylic acids, amides, and alcohols, and it is performed using silylation reagents, such as N,O-bis(trimethylsilyl)trifluoroacetamide (BSTFA) and N,O-bis(trimethylsilyl)acetamide (BSA). The mass spectra of sterols determined via these conventional derivatizations have been reported in earlier studies [12,13,14,15,16]; thus, the interpretation of the mass spectra of sterols has become common, which allows us to easily identify sterols. Sterol trimethylsilyl (TMS) ether spectra can be divided into two patterns: ions that are accompanied by the loss of the side chain (SC) and ions that are not accompanied by the loss of the side chain (Figure 1). The loss of the side chain has five main variations: SC, SC1, SC2, SC + C₂H₃, and SC + C₃H₅. SC, SC1, and SC occur via the cleavage of C₁₇–C₂₀, C₂₀–C₂₂, and C₂₂–C₂₃ bonds, respectively. SC + C₂H₃ represents the cleavage of the side chain followed by the cleavage of C₁₃–C₁₇ and C₁₅–C₁₆ bonds (C₂H₃) on the D-ring. SC + C₃H₅ is from the cleavage of SC and C₁₃–C₁₇ and C₁₄–C₁₅ bonds (C₃H₅). However, the ions that are not accompanied by the loss of the SC only have two main variations: [HO–Si(CH₃)₃]⁺ and [(CH₃)₃Si–O–C₃H₄]⁺. Sterols can be identified by these fragmentations because these fragmentation patterns are different for each sterol. Meanwhile, silylation is very sensitive to water, causing degradation of the derivatized functional groups. Additionally, because Si in TMS reagents can remain in the inlet of the GC, there is a possibility that the inlet of the GC will be contaminated. Moreover, sample preparation before derivatization (e.g., the extraction step) requires a relatively long time.

Since it is a rapid method, thermochemolysis with tetramethylammonium hydroxide (TMAH) was applied for sterol analysis in this study. This method involves hydrolysis and methylation simultaneously, which allow the analysis of both ester and ether compounds and nonbonding compounds that have hydroxyl and carboxyl functional groups [17]. Thus, the TMAH method can provide various compounds, including lignin phenols, amino acids, sugars, and sterols, in one analysis in a relatively short time period [18,19,20,21,22,23]. Additionally, a recent study on the TMAH method extended the analysis to nucleobases [24]. Typically, thermochemolysis with TMAH includes using a pyrolyzer [25,26,27]. Several milligrams of the sample (the optimal weight is varied with the organic carbon content in the sample) was wrapped in a pyrofoil, and it was heated in a pyrolyzer that was directly connected to a GC–MS. Alternatively, thermochemolysis with TMAH can be carried out in a glass ampule [28]. Using this approach, thermochemolysis is performed on an ampule that contains a sample and the TMAH reagent that is sealed under vacuum in an oven.

Upon the TMAH reaction, sterols are provided as sterol methyl ethers. The application of the TMAH method for sterols can produce different types of sterols, including cholesterol, sitosterol, brassicasterol, and stanols [5,6,29]. The efficiency of methylation in sterols reaches >90% [29], indicating that sterols are able to be stably analyzed. Using the TMHA method, both free and bonding (e.g., cholesteryl ester) sterols can be analyzed (Figure 2). However, there are few reports on the interpretation of mass spectra of sterol methyl ethers, which could help detect them. Therefore, in this study, we provide the mass spectra of sterol ethers obtained using the TMAH method from a sediment sample and also provide an interpretation of these mass spectral patterns to show how to detect them.

2. Materials and Methods

2.1. A Tideland Surface Sediment Sample

In this study, the Yatsu tideland, Japan, was selected as the research location (Figure 3). The Yatsu tideland is located in the innermost location of Tokyo Bay, and is surrounded by industrial and residential areas that have expanded through reclamation works. In 1993, for the conservation of ecosystems and landscapes in the natural environment, the Yatsu tideland was registered in the Ramsar Convention. Various organisms, such as lugworms, gobies, clams, sea lettuces, and migratory birds inhabit the Yatsu Tideland [30,31,32,33], although the area is small (40 hectares). Therefore, various organic compound inputs are expected. A sediment sample was taken from the edge of the Yatsu tideland using an Ekman–Birge grab (Figure 3). The sediment sample was freeze-dried immediately after sampling and then powdered well. The powdered sample was stored in a freezer at −20 °C until the sterols were analyzed.

2.2. Thermochemolysis with TMAH

Several milligrams of the sample were added into a 20-mL glass ampule with 150 µL (25% in methanol) of TMAH reagent (Sigma–Aldrich Co., St. Louis, MO, USA) and with nonadecanoic-d37 acid (99.1%, CDN isotopes Co., Pointe-Claire, QC, Canada) as an internal standard. The glass ampule was placed into a desiccator to dry the solvents for approximately 15 min. To remove the solvents more sufficiently, a 40 °C hot plate with a nitrogen stream was applied for approximately 10 min. After the solvents were dried, the glass ampule was sealed under vacuum conditions and was heated in an oven at 300 °C for 30 min. After the ampule was cooled to room temperature, the reacted products were extracted with ethyl acetate thrice. The solvent was removed once, and the extracted products were re-resolved with ethyl acetate to an accurate volume of 100 µL. Several microliters within the re-resolved products were injected into the GC–MS system.

2.3. GC–MS Conditions

An Agilent GC–MS (GC, 6890N; MS, 5973 inert) was used for this study. A fused silica capillary GC column for the DB-5 ms (inner diameter, 0.25 mm; column length, 30 m; film thickness, 0.25 µm) was installed on the GC system with helium gas (purity: >99.9 vol.%) flowing as the carrier gas. The spitless injection mode was employed. The injection room temperature was set to 310 °C. The setting for the oven was as follows: 60 °C (20 min), 310 °C (6 °C/min), 310 °C (20 min). The mass ion source and quadrupole were programmed at 230 °C and 150 °C, respectively. MS data were recorded using the full scanning mode from 50 to 650 Da (Dalton). Ionization was achieved using the electron impact mode at 70 eV.

2.4. Identification of Sterols

The mass spectrum of the sterols obtained using the TMAH method was interpreted by comparing it with reported mass spectra from previous studies on methyl ether derivatization [34] and TMS derivatization [12,13,14,15]. The National Institute of Standards and Technology (NIST)/ United States Environmental Protection Agency (EPA)/ National Institutes of Health (NIH) Mass Spectral Library (NIST 05) was also employed for the interpretation of the sterols. The epicoprostanol-to-coprostanol and coprostanol-to-cholesterol ratios were calculated from their concentrations, which were obtained by comparison with the internal standard.

3. Results and Discussion

3.1. Identified Sterols in the Samples

From the Yatsu sediment, seven types of sterols from three Δ⁵-sterols, four 5α(H)-stanols, two 5β(H)-stanols, one Δ^5,22-sterol, two Δ²²-sterols, two Δ^5,24(28)-sterols, and one 4α-Me-Δ²²-sterol were determined (Table 1 and Figure 4). These multiple differences indicate that there are various input sources present for the Yatsu tideland, which can result from the rich ecosystem and human-living area being adjacent to each other.

3.2. Δ⁵-Sterols

The compounds of cholest-5-en-3β-ol (cholesterol), 24-methyl-cholest-5-en-3β-ol (campesterol), and 24-ethylcholest-5-en-3β-ol (sitosterol) were detected from the tidal sediment, and they were grouped as Δ⁵-sterols (Figure 4 and Figure 5). C₂₇ and C₂₈ sterols, including cholesterol and campesterol, are widely present in plankton [1,8]. However, C₂₉ sterols, such as sitosterol, are present in terrestrial plants [35,36]. Therefore, the distributions of C₂₇, C₂₈, and C₂₉ sterols have been used for the estimation of sources in sedimentary sterols [8]. However, the results must be interpreted carefully, because C₂₉ sterols are also found in phytoplankton [1].

A peak at m/z 71, a characteristic ion for Δ⁵-sterol, was observed for all Δ⁵-sterols, representing [CH₃–O–C₃H₄]⁺. Peaks at m/z 255, 229, and 213 were due to an ion accompanying the loss of the SC (Figure 5). These peaks (m/z 71, 255, 229, and 213) did not change with the type of derivatization because these peaks appear together with the loss of a derivatized functional group at the C₃ position. Therefore, these peaks have also been found in Δ⁵-sterol TMS derivatives [12,13,14]. The peak at m/z 255 is derived from the losses of the side chain and HO–CH₃ from the Δ⁵-sterols ([M − (SC + 32)]⁺). In addition, the peak at m/z 229 results from the loss of C₂H₃ (without a hydrogen atom) from the D-ring ([M − (SC + 32 + C₂H₃ − H)]⁺). The peak at m/z 213 is from the loss of m/z 255 from the molecular ion of Δ⁵-sterols followed by the loss of C₃H₅ (with a hydrogen atom) on the D-ring ([M − (SC + 32 + C₃H₅ + H)]⁺). The peaks at m/z 255 and m/z 213 are clear in all the Δ⁵-sterols detected, although the peak at m/z 229 is weak for campesterol.

[M]⁺, [M − 15]⁺, [M − 32]⁺, [M − (32 + 15)]⁺, [M − 71]⁺, [M − (32 + 67)]⁺, [M − (71 + 54)]⁺, and [M − (71 + 82)]⁺ are all ions that were not accompanied by the loss of the SC (Figure 5). These ions were also confirmed from the fragmentation of sterol TMS derivatives, although their intensities in methyl ethers are relatively high than in TMS ethers. [M − 15]⁺ is produced by the loss of a methyl group from sterols. [M − 32]⁺ represents the loss of HO–CH₃. The additional loss of a methyl group and C₅H₇ from [M − 32]⁺ yields [M − (32 + 15)]⁺ and [M − (32 + 67)]⁺, respectively. [M – 71]⁺ arises from the loss of CH₃–O–C₃H₄. Additionally, [M − (71 + 54)]⁺ and [M − (71 + 82)]⁺ are generated by an additional loss of C₄H₆ on the A-ring or C₆H₁₀ on the A- and B-rings, respectively. [M − (32 + 43 − H)]⁺ can result from the loss of HO–CH₃ from Δ⁵-sterols, followed by the loss of C₃H₇ (without a hydrogen atom) at the end of the SC (Figure 5).

3.3. 5α(H)-Stanols

Four 5α(H)-stanols, 5α(H)-cholestan-3α-ol (epi-cholestanol), 5α(H)-cholestan-3β-ol (cholestanol), 24-methyl-5α(H)-cholestan-3β-ol (campestanol), and 24-ethyl-5α(H)-cholestan-3β-ol (sitostanol) were determined (Figure 4 and Figure 6). Typically, 5α(H)-stanols are produced under anoxic conditions via bacterial conversion from sterols [7,37,38]. Thus, the ratio of 5α(H)-stanol/Δ⁵-sterol can be applied as a tracer for redox conditions in the field of organic geochemistry [5,6,39]. However, the 5α(H)-stanol/Δ⁵-sterol ratio could be large due to the preferential degradation of sterols in terrestrial organic matters, even under oxic conditions [35].

The peaks at m/z 257, 248, 230, and 215 for 5α(H)-stanols are due to an ion accompanying the loss of the SC in 5α(H)-stanols (Figure 6). The peak at m/z 248 is derived from the losses from the side chain and the D-ring from 5α(H)-stanols ([M – (SC + C₃H₅)]⁺). The ion at m/z 257 represents the losses of HO–CH₃ and the side chain ([M – (SC + 32)]⁺), and the ion at m/z 230 is produced by the additional loss of the D-ring from the ion at m/z 257 ([M – (SC + 32 + C₂H₃)]⁺). The peak at m/z 215 is a base peak in 5α(H)-stanols and is due to the further loss of the D-ring (with a hydrogen atom) from the m/z 257 ion ([M – (SC + 32 + C₃H₅ + H)]⁺). As an ion that is not accompanied by the loss of the SC, for example, in cholestanol, peaks at m/z 402 ([M]⁺), 387 ([M − 15]⁺), 370 ([M − 32]⁺), and 355 ([M − (32 + 15)]⁺) were confirmed (Figure 6). The peaks appearing at m/z 345 and 262 in the spectrum of cholestanol were not found in the spectra of the TMS ethers (Figure 6). The peak at m/z 345 may originate from the loss of C₃H₇ from the SC and a methyl group ([M – (42 + 15)]⁺). Additionally, the peak at m/z 262 can be from the loss of CH₃–O–C₃H₄ followed by the loss of C₅H₉ at the A- and B-rings ([M − (71 + 69)]⁺). Although campestanol was also confirmed, the spectrum is not shown in Figure 6, since it was ambiguous.

3.4. 5β(H)-Stanols (Coprostanol and Epicoprostanol)

From the sediment sample, 5β(H)-cholestan-3β-ol (coprostanol) and 5β(H)-cholestan-3α-ol (epicoprostanol) were determined (Figure 4 and Figure 6). Coprostanol can be produced through the reduction of cholesterol by bacteria within the gut of higher animals including humans [40,41], which is ubiquitously present in their fecal matter [42,43]. Therefore, coprostanol has been used as an indicator of fecal pollution and has been widely applied to environments [30,44,45,46,47,48,49].

The mass fragments that appeared for coprostanol and epicoprostanol were similar to those of the 5α(H)-stanols, implying that the same fragmentation as in 5α(H)-stanols occurs in 5β(H)-stanols (Figure 6). However, these intensities are considerably different from those of 5α(H)-stanols. As with 5α(H)-stanols, the peaks at m/z 257, 248, 230, and 215 were confirmed from coprostanol and epicoprostanol as an ion accompanying the cleavage of the SC (Figure 6). An intense peak at m/z 248 represents C₃H₅ in the D-ring according to the molecular weight followed by the loss of the side chain ([M – (SC + C₃H₅)]⁺). The peak at m/z 257 is derived from the loss of HO–CH₃ and the side chain ([M – (32 + SC)]⁺), and the peak at m/z 230 is from an additional loss of C₂H₃ in the D-ring from the m/z 257 ion ([M – (SC + 32 + C₂H₃)]⁺). The peak at m/z 215 represents the loss of C₃H₅ (with molecular hydrogen) in the D-ring from the m/z 257 ion ([M − (SC + 32 + C₃H₅ + H)]⁺), which is a characteristic ion for 5α(H)- and 5β(H)-stanols [50,51,52]. Additionally, in the spectrum of coprostanol and epicoprostanol, the peaks at m/z 402 ([M]⁺), 387 ([M − 15]⁺), 370 ([M − 32]⁺), and 355 ([M − (32 + 15)]⁺) were confirmed to be due to ions that are not accompanied by the loss of the SC (Figure 6).

The detection of coprostanol from sediment suggests that the Yatsu tideland is affected by some fecal pollution. This might be because industrial and residential areas were expanded by reclamation works, and they have enhanced the environmental load on the Yatsu tideland. A biplot of the coprostanol/cholesterol ratio with the epicoprostanol/coprostanol ratio in the sediment is shown in Figure 7. Epicoprostanol is typically converted from coprostanol via a bacterial reaction during sewage treatment processes, and therefore is observed from treated or old sewage samples [53]. Thus, a high epicoprostanol/coprostanol ratio is observed from “old or treated” sewage samples [54]. In our Yatsu tideland sediment sample, the epicoprostanol/coprostanol ratio shows approximately 0.4, indicating that the source is a mixture of new and old. The ratio of coprostanol/cholesterol can estimate the degree of fecal contamination [55,56,57]. According to a proposal by Grimalt and Albaigés [56], when the coprostanol/cholesterol ratio is greater than 0.2 in an environmental sample, the environment can be considered to have fecal contamination. The coprostanol/cholesterol ratio in our sediment sample was approximately 0.5 (Figure 7), which is over the value of the definition. Therefore, the Yatsu tideland is an environment that is affected by fecal pollution according to this definition. However, the value is not higher as with large contaminated areas (Figure 7). Additionally, our data is limited to one site. Thus, a more accurate investigation of the polluted area of the Yatsu tideland requires that different sample areas be analyzed.

Of note, coprostanol analysis via the TMAH method has methodological advantages because the process can be performed quickly with a small amount of solvent. These advantages can result in the treatment of a large number of samples and the reduction of solvents during analysis.

3.5. Δ^5,22-Sterol (Brassicasterol)

A Δ^5,22-sterol, 24-methylcholesta-5,22E-dien-3β-ol (brassicasterol), was determined from the sediment (Figure 4 and Figure 8). Brassicasterol is present in diatoms as a major sterol [58,59,60]. In particular, pennate diatoms have a high relative abundance of brassicasterol (>60% of sterols) [61]. However, this sterol cannot be a specific biomarker for diatoms because the sterol is present in other algal groups [61].

The peaks at m/z 314 ([M − (SC1 + H)]⁺), 299 ([M − (SC1 + H + 15)]⁺), 285 ([M − (SC + 2H)]⁺), 255 ([M − (SC + 32)]⁺), 229 ([M − (SC + 32 + C₂H₃ − H)]⁺), 213 ([M − (SC + 32 + C₃H₅ + H)]^+,), and 199 ([M − (SC + 32 + C₄H₇ + H)]⁺) in the brassicasterol are due to ions that accompany the cleavage of the SC (Figure 8). These ions are present in the spectra of Δ⁵-sterols and Δ²²-sterols as well. Additionally, the peaks at m/z 255 and 213 were also found in the TMS derivative, because these ions do not include a derivatized functional group [12,62]. However, peaks at m/z 412 ([M]⁺), 397 ([M − 15]⁺), 380 ([M − 32]⁺), 365 ([M − (32 + 15)]⁺), 341 ([M − 71]⁺), and 337 ([M − (32 + 43)]⁺) were found to result not by the loss of the SC, and they are also observed for Δ⁵-sterols and Δ²²-sterols (Figure 8). The peak at m/z 125 represents the molecular weight of its SC. These ions are important for the determination of Δ^5,22-sterols.

3.6. Δ²²-Stanols

Two Δ²²-stanols, 24-methyl-5α(H)-cholest-22E-en-3β-ol (brassicastanol) and 23,24-dimethyl-5α(H)-cholest-22E-en-3β-ol, were determined from the sediment (Figure 4 and Figure 9). As with 5α(H)-sterols, Δ²²-stanols can be produced from the bacterial conversion from Δ^5,22-sterols. Indeed, a sterol analysis of particle matter at the oxic–anoxic boundary in the Black Sea implied that the brassicastanol/brassicasterol ratio increases at the anoxic water layer [7]. Additionally, a continuous record of the brassicastanol/brassicasterol ratio from California late Quaternary sediments has shown historical changes in redox conditions over 40 kyr [5].

The peaks at m/z 316, 301, 287, 257, 229, 215, and 201 are due to ions that accompany the cleavage of the SC (Figure 9). The peak at m/z 316 results from the loss of the side chain (SC1) with a hydrogen atom from Δ²²-stanols ([M – (SC1 + H)]⁺), and the additional loss of a methyl group yields the ion at m/z 301 ([M – (SC1 + H + 15]⁺). The peak at m/z 287 originates from the loss of the side chain (with two hydrogen atoms) from Δ²²-stanols ([M – (SC + 2H)]⁺). An intense peak at m/z 257 is derived from the loss of HO–CH₃ and the side chain from Δ²²-stanols ([M – (32 + SC)]⁺). This ion, m/z 257, is a characteristic ion that is used for the determination of Δ²²-sterols, as in the case of TMS derivatization [51,63,64], and it shows that the sterol skeleton is saturated (no double bond on the C₅–C₆ position) as with 5α(H)-stanols [51]. The peaks at m/z 229, 215, and 201 can arise from the loss of C₂H₃ (with a hydrogen atom) at the D-ring ([M – (32 + SC + C₂H₃ + H)]⁺), the loss of C₃H₅ (with a hydrogen atom) at the D-ring ([M – (32 + SC + C₃H₅ + H)]⁺), and C₄H₇ (with a hydrogen atom) at the D- and C-rings from the ion at m/z 257 ([M – (32 + SC + C₄H₇ + H)]⁺). Ions [M]⁺, [M − 15]⁺, [M − 32]⁺, [M − (32 + 15)]⁺, and [M − (32 + 43)]⁺ were confirmed to be not accompanied by the loss of the SC (Figure 9). [M − (32 + 43)]⁺ can arise from the loss of HO–CH₃ from Δ²²-stanols, which is followed by the loss of C₃H₇ on the end of the SC.

3.7. Δ^5,24(28)-Sterols

Two Δ^5,24(28)-sterols, 24-methylcholesta-5,24(28)-dien-3β-ol (24-methylenecholesterol) and 24-ethyl-cholesta-5,24(28)Z-dien-3β-ol (isofucosterol), were determined from the sediment (Figure 3 and Figure 9). 24-methylenecholesterol is present in some microalgae, such as diatoms [1], which are found in various aquatic environments including in the Arctic and Antarctic oceans [65,66,67]. Isofucosterol has been observed in green microalgae as a major sterol [1].

The spectra of both Δ^5,24(28)-sterols show peaks at m/z 328, 313, 296, 285, 281, 255, 243, 229, and 213 (Figure 10). Of these ions, the peaks at m/z 328, 313, 285, and 243 were also confirmed in the spectra of the Δ²⁴⁽²⁸⁾-sterols. The ion at m/z 328 arises from the loss of the side chain (SC2) with a hydrogen atom and is also a characteristic ion for Δ^5,24(28)-sterols. The peaks at m/z 255, 253, 229, and 213 are also visible in the fragmentation pattern of other sterols, such as Δ²⁴⁽²⁸⁾-sterols and Δ⁵-sterols. The peak at m/z 296 results from the losses of the side chain (SC1) and HO–CH₃ from Δ^5,24(28)-sterols ([M − (SC1 + 32)]⁺). The additional elimination of a methyl group yields m/z 281 ([M − (SC1 + 32 + 15)]⁺). Ions [M]⁺, [M − 15]⁺, [M − 32]⁺, and [M − (32 + 15)]⁺ were confirmed to be ions that are not accompanied by the loss of the SC, which are also present for other sterols, including Δ⁵-sterols (Figure 10).

3.8. 4α-Me-Δ²² Sterol

A 4α-Me-Δ²² sterol, 4α,23,24-trimethyl-5α(H)-cholest-22E-en-3β-ol (dinosterol), was identified from the sediment (Figure 4 and Figure 11). Sterols having a methyl functional group at the C₄ position, including dinosterol, are abundant in dinoflagellates [68,69,70]. Therefore, dinosterol has been used as a unique biomarker for dinoflagellates [71,72].

In the spectrum of dinosterol, [M]⁺, [M − 15]⁺, [M − 32]⁺, m/z 330, 315, 301, 271, 245, and 229 were confirmed, which can be interpreted by the 14 Da (CH₃) higher shift of the corresponding fragmentation patterns in the spectrum of 23,24-dimethyl-5α(H)-cholest-22E-en-3β-ol methyl ether (Figure 11).

3.9. Identification of Sterols from the Extracted Ion Chromatograms

Determining each sterol from the GC–MS results can be effectively achieved from the extracted ion chromatograms of each sterol. The ion at m/z 255 is useful to determine Δ⁵-, Δ^5,22-, and Δ^5,24(28)-sterols. Ion chromatograms with m/z 215, 257, 328, and 330 represent 5α- and 5β-stanols, Δ²²-stanols, Δ^5,24(28)-sterols, and 4α-Me-Δ²²-sterols, respectively (Figure 4).

The retention time in the GC–MS results is also important for determining sterols. For example, coprostanol and cholestanol, isomers of the H atom at the C₅ position (i.e., 5β(H) or 5α(H)), elute at substantially different times even though the structures are similar (Figure 4). The retention time of coprostanol is earlier than that of cholestanol. Additionally, 5α(H)-stanols and Δ²²-stanols were obtained immediately after their corresponding Δ⁵-sterols and Δ^5,22-stanols, which is useful for the determination of these compounds (Figure 4). Additionally, the number of double bonds and their positions (C₂₂ and/or C₅ positions) appear at different retention times; for example, retention times become longer in the order of 28(24)Δ^5,22E (brassicasterol), 28(24)Δ^22E (brassicastanol), and 28(24)Δ⁵ (campesterol) (Figure 4).

4. Conclusions

A tideland surface sediment from the Yatsu tideland, Japan, was analyzed using thermochemolysis with tetramethylammonium hydroxide (TMAH) for sterols. From the sediment, 10 sterols, including Δ⁵-sterols, 5α(H)-sterols, Δ^5,22-sterols, Δ²²-sterols, Δ^5,24(28)-sterols, 4α-Me-Δ²²-sterols, and coprostanol, were determined. The Yatsu tideland sediment sample enabled the determination of these various sterols, because the Yatsu tideland is located where rich ecosystems and human-living areas are adjacent to each other. The MS results of these sterols helped determine them. Characteristic mass fragments at m/z 255, m/z 215, m/z 257, and m/z 330 were useful to detect Δ⁵-type sterols (i.e., Δ⁵-sterols, Δ^5,22-sterols, and Δ^5,24(28)-sterols), 5α(H)-sterols, 5β(H)-sterols, Δ²²-sterols, and 4α-Me-Δ²²-sterols from the extracted mass chromatograms. Additionally, some fecal pollution in the Yatsu tideland was suggested by the detection of coprostanol. The TMAH method can treat samples in a relatively short time with a small amount of sample and then provide sterol methyl ethers. Thus, this method is suitable for studies that analyze many samples. The provided sterol spectra in this study, including other such studies, will help determine the kinds of sterols present in a sample using the GC–MS result.

Author Contributions

Conceptualization, M.N. and S.Y.; Investigation, M.N., Y.Y., N.Y., and K.T.; Resources, M.N., N.Y., and K.T.; Data Curation, M.N., Y.Y., and S.Y.; Writing – Original Draft Preparation, M.N. and S.Y.; Writing – Review and Editing, Y.Y., N.Y., K.T., and S.Y.; Visualization, M.N.; Supervision, M.N. and S.Y.; Funding Acquisition, M.N.

Founding

The research was financially supported by the Sasakawa Scientific Research Grant (2018) from The Japan Science Society.

Acknowledgments

The authors thank the laboratory members of our university, and would like to thank MARUZEN-YUSHODO Co., Ltd. (https://kw.maruzen.co.jp/kousei-honyaku/) for the English language editing.

Conflicts of Interest

The authors declare no conflict of interest.

Figures and Table

Figure 1. Diagnostic fragmentation of the sterols. Each number represents the position of the carbons. X varied with derivatization, for example, –Si(CH3)3 in trimethylsilyl, –COCH3 in acetyl, and –CH3 in methyl ethers. The X of the original sterols is H (i.e., –OH). SC refers to the side chain.

Figure 2. Generation processes for sterol methyl ether from cholesteryl stearate via the tetramethylammonium hydroxide (TMAH) reaction. This figure was drawn based on Asperger et al. [29] and Challinor [17].

Figure 3. Pictures of the sampling point (Yatsu Tideland, Japan). (a) Picture of the sample. (b) State of the sampling place. The sample was taken from a bridge, as shown in the picture, using an Ekman–Birge grab.

Figure 4. Partial total ion chromatogram and extracted ion chromatograms (m/z 255, 215, 257, 328, and 330) focused on sterols (38.00–41.88 min) via thermochemolysis using tetramethylammonium hydroxide (TMAH). The symbols are listed in Table 1.

Figure 5. Electron ionization (70 eV) mass spectrum of Δ5-sterol. Bold numbers indicate an ion that accompanies the cleavage of the side chain. (a)–(h) Ions that are not accompanied by the loss of the side chain or other groups.

Figure 6. Electron ionization mass spectra of 5α(H)-stanol and 5β(H)-stanol (coprostanol). Bold numbers indicate an ion that accompanies the cleavage of the side chain. (a)–(f) Ions that not accompanied by the loss of the side chain or other groups.

Figure 7. A biplot of the coprostanol/cholesterol ratio with the epicoprostanol/coprostanol ratio for the Yatsu sediment. Our data is plotted on a redrawn figure from Mudge and Seguel [54] and Mudge and Ball [55].

Figure 8. Electron ionization mass spectrum of 24-methylcholesta-5,22E-dien-3β-ol methyl ether (brassicasterol). Bold numbers indicate an ion accompanying the cleavage of the side chain. (a)–(f) Ions that are not accompanied by the loss of the side chain or other groups.

Figure 9. Electron ionization mass spectrum of Δ22-stanol. Bold numbers indicate an ion that accompanies the cleavage of the side chain. (a)–(e) Ions that are not accompanied by the loss of the side chain or other groups.

Figure 10. Electron ionization mass spectrum of Δ5,24(28)-sterol. Bold numbers indicate an ion that accompanies the cleavage of the side chain. (a)–(d) Ions that are not accompanied by the loss of the side chain.

Figure 11. Electron ionization mass spectrum of 4α,23,24-trimethyl-5α(H)- cholest-22E-en-3β-ol methyl ether (dinosterol). Bold numbers indicate an ion that accompanies the cleavage of the side chain. (a)–(c) Ions that are not accompanied by the loss of the side chain.

Table 1

Sterols identified from the Yatsu surface sediment.

Retention Time (min)	Symbol *	Systematic Name	Trivial Name	Formula(Methylated Ether)	Molecular Weight (Methylated Ether)	m/z **
38.358	Coprostanol	5β(H)-Cholestan-3β-ol methyl ether	Coprostanol	C28H50O	402	215, 230, 248, 257, 345, 355, 370, 387, 402
38.671	Epicoprostanol	5β(H)-Cholestan-3α-ol methyl ether	Epicoprostanol	C28H50O	402	215, 230, 248, 257, 345, 355, 370, 387, 402
38.777	Epicholestanol	5α(H)-Cholestan-3α-ol methyl ether	Epicholestanol	C28H50O	402	215, 230, 248, 257, 345, 355, 370, 387, 402
39.096	27Δ5	Cholest-5-en-3β-ol methyl ether	Cholesterol	C28H48O	400	213, 229, 255, 275, 301, 326, 329, 353, 368, 385, 400
39.228	27Δ0	5α(H)-Cholestan-3β-ol methyl ether	Cholestanol	C28H50O	402	215, 230, 248, 257, 345, 355, 370, 387, 402
39.578	28(24)Δ5,22E	24-Methylcholesta-5,22E-dien-3β-ol methyl ether	Brassicasterol	C29H48O	412	199, 213, 229, 255, 285, 299, 314, 337, 341, 365, 380, 397, 412
39.722	28(24)Δ22E	24-Methyl-5α(H)-cholest-22E-en-3β-ol methyl ether	Brassicastanol	C29H50O	414	201, 215, 229, 257, 287, 301, 316, 339, 367, 382, 399, 414
40.166	28(24)Δ5,24(28)	24-Methylcholesta-5,24(28)-dien-3β-ol methyl ether	24-Methylenecholesterol	C29H48O	412	213, 229, 243, 255, 281, 285, 296, 313, 328, 365, 380, 397, 412
40.229	28(24)Δ5	24-Methylcholest-5-en-3β-ol methyl ether	Campesterol	C29H50O	414	213, 255, 261, 289, 301, 315, 340, 343, 367, 382, 399, 414
40.360	28(24)Δ0	24-Methyl-5α(H)-cholestan-3β-ol methyl ether	Campestanol	C29H52O	416	215, 230, 248, 257, 359, 369, 384, 401, 416
40.616	29(23,24)Δ22E	23,24-Dimethyl-5α(H)-cholest-22E-en-3β-ol methyl ether		C30H52O	428	201, 215, 229, 257, 287, 301, 316, 353, 385, 396, 413, 428
41.161	29(24)Δ5	24-Ethylcholest-5-en-3β-ol methyl ether	Sitosterol	C30H52O	428	213, 229, 255, 275, 303, 329, 354, 357, 381, 396, 413, 428
41.217	30(4,23,24)Δ22E	4α,23,24-Trimethyl-5α(H)-cholest-22E-en-3β-ol methyl ether	Dinosterol	C31H54O	442	271, 301, 315, 330, 410, 427, 442
41.298	29(24)Δ0	24-Ethyl-5α(H)-cholestan-3β-ol methyl ether	Sitostanol	C30H54O	430	215, 230, 248, 257, 290, 373, 383, 398, 415, 430
41.336	29(24)Δ5,24(28)Z	24-Ethylcholesta-5,24(28)Z-dien-3β-ol methyl ether	Isofucosterol	C30H50O	426	213, 229, 243, 255, 281, 285, 296, 313, 328, 379, 394, 411, 426

* The symbols are given as n(m)Δ^p, where n is the carbon number of a sterol, m is the position of the methyl or ethyl group in the side chain, and p is the position of a double bond (s). ** m/z: bold and italic denote the base peak and molecular ion, respectively.