Minor and trace sterols in marine invertebrates 1. General methods of analysis

699

MINOR AND TRACE STEROLS IN MARINE INVERTEBRATES 1.

GENERAL METHODS OF ANALYSIS

Simeon Popov,+ Robert M. K. Carlson, Annemarie Wegrnann and Carl Djerassi Department of Chemistry, Stanford University, Stanford, California 94305 Received:

3/l/76 ABSTRACT

SS-Hydroxy sterols occurring at a concentration of at least 0.001% of the steroi mixtures of Pseudoplexaura porosa and Plexaura homomalla have been fractionated using a series of remtechniques and subsequently analyzed using combined gas chromatography-mass spectrometry (G&MS) in the development of a procedure for examining the minor and trace components of marine sterol mixtures. A total of 49 sterols were found4 which spanned a molecular weight range of 274 to 440. In addition A 3-keto analogs of cholesterol, 24-methylcholesterol and gorgosterol Initial separation of varwere found in the extracts of P. homomalla. ious natural sterol-containing-conjugates and free sterols was found to have a number of advantages. Fractional digitonin precipitation and alumina column chromatography were found to possess greater sterol separation abilities than previously recognized. Many of the minor sterols were found to possess novel structures including a series of short side chain sterols, 19-nor sterols, 5B-stanols and 4-monomethyl sterols for which structure elucidation work is continuing. INTRODUCTION 36-Monohydroxyl sterol extracts from marine invertebrates can be extremely complex mixtures.

Over 80 sterols have so far been identified

from marine sources Cl], and evidence has been presented for the occurrence of over 30 sterols in a single marine species C21.

The discovery

of novel sterols as major components of the extracts of marine animals is rare, although there have been a number of recent exceptions: dinosterol (4a,23,24~-trimethyl-5a-cholest-22-en-36-o1) a dinoflagellate t

the major sterol in

[3], calysterol (23,24-ethylidenecholesta-5,23-dien-

Visiting investigator on leave (1975-1976) from the Institute of Organic Chemistry, Bulgarian Academy of Sciences, Sofia, Bulgaria.

VoZwne 28, Number 5

S

T=EOXDI

November, 1976

36-01) the major sterol of a marine sponge C4], aplysterol 24,26-dimethylcholest-5-en-38-01)

((24k,25S)-

and 24,28-bisdehydroaplysterol,

the

major sterols of marine sponges of the genus Verongia [5], and a series of 19-nor-5a-stanols

again from a marine sponge [6].

The bulk of marine

invertebrates for which sterol analyses have been reported Cl], however, contain major sterols with well known structures. A number of workers have recently turned to characterizing the minor sterol components of marine extracts with very interesting results. Kobayashi, -et al., have recently characterized patinosterol nor-24-methyl-5a-cholest-22E-en-38-ol) 24-methylcholesta-5,22E-dien-38-01)

[7], occelasterol((24S)-27-nor-

[8] and amurasterol

24-methyl-5a-cholesta-7,22E-dien-3$-ol)

((24S)-27-

((24S)-27-nor-

[9] as minor components of a

scallop, marine annelid, and starfish respectively.

Kobayashi points

out [7] that these 27-nor sterols are possibly biosynthetic precursors to the 24-nor C

26

sterols which themselves are minor sterol components

of a great many marine organisms. isofucostanol in a jellyfish [lo].

Ballantine, et al., have detected 28Idler, et al., have also recently

discovered new sterols as minor components of marine invertebrates, the from a scallop Cl1 1.

most recent being (E)-24-propylidenecholest-5-en-38-01

Sterols possess very similar properties in conventional thin layer chromatography

(TLC) and column chromatography systems.

severe difficulties

This leads to

in isolation and analysis of minor sterol comoonents.

A number of reports have recently appeared which describe the use of new techniques in the fractionation of sterol mixtures. umn chromatography

over a hydroxyalkoxypropyl

LH-20 [12] and high pressure reversed-phase

These include col-

derivative of Sephadex

liquid chromatography

c13].

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TEEOIDI

Such new preparative techniques when used in conjunction with refined pre-existing techniques offer new opportunities for studying minor sterol components. A study of the minor sterols of marine organisms is pertinent for a number of important reasons. It is reasonable to assume that some of the minor sterol components are active metabolites and therefore are of major relevance to the study of sterol biosynthesis in marine organisms a promising field which is only just being opened. Other minor components may be sterols carried through the marine food chain and are, therefore, of ecological interest. Analysis of minor sterols is particularly important with respect to biosynthetic studies by radioisotope labeling because a large radiolabel incorporation in an unresolved minor component could lead to ambiguous results and misleading conclusions. As part of a search for new biosynthetically important marine sterols, and as a prelude to a sterol biosynthetic study in gorgonians, a number of different methods of sterol separations were investigated by us using the extracts of two Caribbean gorgonians, Pseudoplexaura porosa and Plexaura homomalla. Our procedure represents a refinement and synthesis of several new and older methods of sterol fractionation. Particularly noteworthy is the finding that several conventional fractionation procedures including chromatography over alumina (Activity III) and digitonin fractionation possess previously unrecognized potential in separating 3-monohydroxy sterol mixtures and that organisms from which only eight sterols have been identified previously by conventional gas chromatography-massspectrometer (GC-MS) Cl41 analysis

S

TEIROXDN

can be shown to contain over 30 sterols (many of which possess novel structures) using the fractionation and analytical procedures described below.

Further structural elucidation

and syntheses of many of these

novel sterols- is underway in this laboratory and will be the subject of forthcoming communications. RESULTS AND DISCUSSION The purpose of the procedure presented below is to create fractions of a sterol mixture with enrichments of various sterols sufficient for the production of a good quality GC-MS derived mass spectrum of each sterol component.

This requires in some cases a several hundredfold

enrichment of a particular minor sterol relative to the more abundant sterols.

Ps :e

1); however, ture.

porosa can be shown to contain over 40 sterols (see Table nine

of these sterols make up 97% of the mass of the mix-

Many of the minor and trace sterols are only present in micro-

gram quantities even if gram quantities of the sterol mixture are available.

At present GC-MS is the only viable tool for obtaining strue

tural information concerning these minor sterol components.

Present

knowledge of sterol mass spectrometric behavior has advanced so that many times GC-MS data are sufficient to suggest a structure for a newly detected sterol [15], which then warrants confirmation by synthesis. When quantities of the minor sterols are sufficient, production of fractions significantly enriched in the minor sterols also allow preparative GLC or HPLC collection of these minor components for Nuclear Magnetic Resonance

(NMR) or other analysis.

A number of chromatographic proceed-

ures were used by us to effect sufficient enrichments of the various minor sterol components.

These procedures include:

S Table

stem1 Reference Number

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

i”7 18 19 20 21 22 23 24 25 26 27 28

Structure

* * * * * * * * * * * * * * 1

* 15 7 1 I z 10 Y 5 li x

29

ii 5

30 31

T z

32 33 34 35 36 37 38 39 40 41

< G * : * * G

1.

TXIEOXDb

703

4-Demethyl' Sterols Present in ps. porosaand P. homoma.l.la M+

274 288 290 300 302 312 314 314 316 316 318 318 330 344 358 360 372 384 384 386 396 386 389 398 398 398 400 402 412 412 414 4J.4 414 416 416 424 426 426 426 426 428 428

Number of Carbon Atoms

19 20 20 21 21 21 22 22 22 21 22 21 23 24 25 25 26 27 27 27 27 27 27 28 28 28 28 28 29 29 29 29 29 29 29 30 30 30 30 30 30 30

Relative Retention Time 3% ov-17 276Y

Fraction Used in GC-MS+ Analysis

T5-6 0.13 TS-6 0.25 T5-6 0.25, b c 0.53 ' TS-6 0.23 T5-6 0.57 T5-6 0.57 T5-6 0.31 T4-lt2 0.27 TS-6,T6-12 0.57 T5-6 0.27 TS-6 0.57 TS-6 0.33 0.6-0.8 T5-6 0.6-0.8 T5-6 0.6-0.6 T5-6 0.92b T8-20t21 g 1.17 0.93 T8-19 T6-7 l.OOb 1.15 Tg8-20+21 1.21 T6-(l-4jd 1.00 TB-15t16 1.13 c,Tt?-22+23 1.2gb 1.35 T8-20+21 1.27 T6-7 1.27 T8-(l-4)d 1.36 T8-13t14 1.65 TS-3 1.54 T6-7 1.65 T8-(l-4jd 1.62 T7-5 T7-5 1.33 1.53 T8-(l-4jd 2.25 T8-22t23.c 2.20 T6-10tllj 3.38a Tgs-(1-4) 1.53a 2.15 TB-22 2.20_ T;-(l-4)d 1.53.=

Percent of Sterol Mixture Mixture II.phomomalla Ps. porosa Percent of Sterol

0.3
0.8

0.4 go.001
a.002 so.001 51 0.03 0.05
> Structure elucidation to be reported in a forthcomingpublication. t

e.g., "TS-6" refers to Table 5 (Digitonin Fractionation)Fraction number 6.

a Relative retention time measured on OV-25 3% on Gas Chrom Q, 256OC. b Relative retention time measured as the acetate relative to cholesterolacetate = 1.00. ' Silver nitrate-silicagel column chromatography. d Followed by perbenzoic acid oxidation and isolation of stanols. e Weight percent of the stem1 ester fraction. f Mass spectmmetric data not obtained. ' Preparative GC 3% OV-25 on Gas Chrom Q. 256'X.

Table 2.

4-Monomethyl Sterols Present in Ps. porosa and P. homomalla M+

Sterol Structure Reference #

43

C

Number of Carbon Atoms

Relative Retention Time 3% OV-17 27b°Ca

Fraction Used in GC-MS Analysisb

Percent of Sterol Mixture Ps.porosa -~

Percent of Sterol Mixture P. homomallg

414

29

1.49

T7-(6-10)

44

C

414

29

1.35

T7d6-10)

0.01

0.2

45

C

416

29

1.47

T7_(6-10)

0.05

3.3

46

C

424

30

1.70

T746-10)

47

C

426

30

1.55

T7d6-10)

48

-18

428

30

1.52

T645-6)

49

C

-

0.2

-

co.1

-

co.1

0.7

6

T746-10) 440

3.5-4

31

T&l-4)d

-

0.01

a. Cholesterol = 1.00; b. -e.g., "T -(S-lo)" refers to Table 7, Fraction 6 through 10; c. Structure elucida ?* Ion to be reported in a forthcoming publication; d. Followed by perbenzoic acid oxidation and isolation of stanols.

Table 3. A4-3-Keto Steroids present in P. Sterol Reference #

Structure

M+

Number of Carbon Atoms

homomallaa

Relative Retention Time 3% ov-17 276='cb

Fraction Used in GC-MS AnalysisC

Percent of Sterol Mixture

50

19

384

27

1.50

T7-3

0.46

51

20

398

28

1.91

T7-3

0.17

52

2a

424

30

3.32

T7-3

0.20

a. A4-3-Keto steroids were not found in Ps.porosa; b. Cholesterol = 1.00; c. 'IT7-3" refers to Table 7, Fraction number 3.

S

TDEOXDD

&

705

w

HO

\

0

& ’

S

706

TIIROID~

1) Initital fractionation of high molecular weight fatty acid sterol esters, naturally occurring sterol acetates, free sterols and polar sterol conjugates

(sterol sulfates, glycosides, etc.).

2) Fractionation of free sterols (either naturally occurring free sterol or saponification products) using alumina Brockmann Grade (III) column chromatography

enabling separation of A4-3 keto

steroids, 5f3-stanols, 4-methyl sterols and 4-demethyl sterols. The alumina column chromatographic procedure also yielded fractions containing significant enrichments of the various 4-demethyl sterols on the basis of carbon number and unsaturation in side chain. 3) Fractionations

over silver-nitrate

impregnated alumina or silica

gel, procedures often used in marine sterol fractionations. 4) Fractionations via

a fractional digitonin precipitation proced-

ure. 5) Fractionations using reversed-phase tography

high pressure liquid chroma-

(HPLC).

Each fractionation and enrichment procedure will be discussed in turn and the advantages and limitations of each procedure will be discussed with respect to the analysis of minor and trace components of marine sterol mixtures. INITIAL ISOLATION Numerous extraction procedures have been employed by previous researchers.

Some methods employ initial saponification of the fresh

tissues followed by extraction of the nonsaponifiable cedure should be avoided for a number of reasons.

lipids.

This pro-

Sterols in gorgonians

and other organisms occur in several easily separable forms; high molecular weighty fatty acid sterol esters, free sterols and sterols conjugated to very polar moieties (sterol sulfates, glycosides, etc.). We have found that a number of gorgonians also contain a reasonable quantity of naturally occurring sterol acetates (see Table 4) which can be initially separated from the other sterol-containingconstituents. The sterol constitution of each of these fractions need not be the same.

In fact,

significant variations in the sterol composition occur in the various sterol-containinggroups as is evident from Table 4.

Not only are the

variations in the sterol composition of the various sterol and sterol ester or glycoside fractions of biological and biochemical significance, but these variations represent fractions with naturally occurring enrichments of various sterol components. Saponification of the total lipid extract destroys these natural enrichments. The following fractionation procedures are essentially enrichment procedures and so initial saponification results in loss of the aforementioned naturally enriched fractions in addition to resulting in loss of information concerning sterol distributions among the various conjugates and destruction of interesting labile terpenes or other compounds, the analysis of which is an additional bonus of sterol conjugate fractionation. A good extraction procedure should extract all lipid material from the sample without causing lipid hydrolysis or degradation and without requiring an excessive amount of time or number of manipulations. The extraction procedures employed in past studies can be grouped into two distinct classes: 1) Continued hot extraction of dried tissues employing a Soxhlet or modified Soxhlet apparatus using either a series of solvents

Table 4.

Fraction

Major Sterol Percent Compositions of Sterols Contained in Free Sterol and Sterol Conjugate Fractions of P. homomalla.

10

lb

20

24

41

6

Sterol Reference Numbera 29 48 31 27 25 6.5

1.5

30

38 33

4.2

5

1

1

% of Tissue Dry Weight 0.02

2c

-

15

4

2

1

5

3.5

0.7

0.3 68

0.03

gd

0.8

32

11

18

8

4

0.3

0.2

2.2 14

0.002

4e

0.1

50

10

9

9

3.7

0.2

0.8

4

13

0.2

1

63

QJo.02

gf gg

-

19

3.5

26

5

4

6

13

6

2 -

-

1.5 47

0.01

a. See Table 1 of identification of sterol from sterol reference number; b. High molecular weighty fatty acid (mainly palmitic and stearic acids. (See Experimental Section for method of fatty acid analysis) containing sterol esters; c. Slightly more polar high molecular weight sterol ester fraction with fatty acid moieties of as yet unidentified structures; d. Naturally occurring sterol acetates; e. Free Sterols; f. Sterols released by KOH hydrolysis of extracted tissues; g. Sterols released by HCl hydrolysis of extracted tissues. of increasing polarity or a single relatively polar solvent. 2) Cold extraction of fresh homogenized or dried ground tissues using a series of solvents of increasing polarity, a single polar solvent or a single mixed solvent system, usually chloroform-methanol. It might be expected that the hot continuous extraction procedures would be more efficient; however, this is not true. found that room temperature chloroform-methanol

Giese cl61 has

extraction of marine ani-

mal tissues results in the extraction of up to 40% more lipid material than the corresponding Soxhlet procedure employing ethyl ether. found the same superior efficiency of cold chloroform-methanol

We have extrac-

tion procedure 1171 (Bligh and dyer method) as opposed to a continuous hot acetone Soxhlet extraction of gorgonian tissues.

The hot continuous

acetone extract of dried tissues requires a number of hours before free

sterols are no longer extracted as revealed by TLC (System 1 - see Experimental) with a cholesterol marker. Subsequent ethanol extraction of the acetone extracted tissue resulted in no further extraction of free sterols as revealed again by TLC (System 1) with a cholesterol marker. Base and acid hydrolysis of the extracted tissues, however, did result in release of further sterof material. The sterols released by base hydrolysis had a composition similar to the sterols of the sterol ester fraction #2 of Table 4.

Whether the sterols released upon acid and base

hydrolysis are from occluded sterols or sterol esters or from the

hy-

drolysis of very polar or high molecular weight sterol conjugates (sulfates, glycosides, etc.) not released by acetone is under investigation. The Bligh and Dyer chloroform/methanol/waterprocedure [17] can be completed in as little as five minutes for small tissue samples and is therefore also more effective with respect to time than hot continuous extractions. The reasons for the remarkable efficiency of the chloroform/methanol procedure have been the subject of a number of publications 1187. The chloroform/methanolprocedure does have a few disadvantages [19], however, and other cold multicomponent solvent extraction systems have been described C20]. From the standpoint of lipid oxidation and degradation, an efficient low temperature extraction of fresh undried tissues is clearly preferable. This is particularly important in the isolation of sterol fatty acid esters, the fatty acid moiety of which may be quite susceptible to oxidation. Free sterols are particularly stable and therefore worries regarding auto-oxidation are not as great as in the investigation of other lipid classes. Several marine sterols, however, have

S

710

TIOROIDS

been demonstrated, and still others are believed to be products of autooxidation [21]. Smith, -et _*y al of 24-nor-chol-5-en-36-01,

have recently shown [22] by theirdetection

androst-5-en-36-01

and preg-5-en-36-01,

that

sterols with short side chains can be minor products of cholesterol auto-oxidation.

This problem is clearly most critical in the analysis

of minor sterol components.

Studies concerning the possible sterol

auto-oxidation origin of several of the minor sterols indicated in Table 1 are underway in this laboratory.

Analysis of the scope of this prob-

lem with regard to tissue storage and workup will be considered in a forthcoming communication concerning structure elucidation of the low molecular weight minor sterols presented in Table 1. DIGITONIN FRACTIONAL PRECIPITATION Digitonin has been used for the isolation of 36-hydroxy sterols since the initial discovery [23] of sterol-digitonin Windaus in 1909.

precipitation by

Digitonin was used extensively for the isolation and

quantitative analysis of sterols (usually cholesterol) before more modern chromatographic methods of separation and instrumental methods of analysis became available. of the digitonin-sterol

Recent studies [24] have shown that part

complex always remains in solution which lessens Digitonin has

the value of digitonin in quantitative sterol analysis.

also been found to precipitate steroidal ketones (A4-3-ones, cholestanone, A5-cholestenone,

5a-pregnan-20-one,

5-pregnene-20-one,

and other ster-

oical ketones), some 3B-acetoxy sterols, some 3a-hydroxy sterols and 4monomethyl and 4,4-dimethyl sterols [24,25,26].

Digitonin precipita-

tion is, therefore, not specific for 3&hydroxy-4-demethyl

sterols, al-

though it is known that 3t3-hydroxy sterols form digitonides more quickly

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TBEOXDl

711

and completely than the aforementioned steroids. Among the 36-hydroxy sterols, 56-stanols and A 537-sterols form digitonin complexes least rapidly [26,27]. Ease of precipitation with digitonin has been found to be a function of sterol configuration at C-5 and C-17, the length of the side chain, and the presence or absence of D-ring substituents [26]. Variation in double bond positions within the steroid nucleus has not been found C261 to be a significant factor in sterol-digitonin complex formation. Although the above variations in sterol-digitonin complex formation have been noted in the literature, the use of these properties in the separation of complex sterol mixtures doesnot appear to have been documented. Utilizing the differences in the ease of sterol-digitonincomplex formation and solubility, we have developed a fractional precipitation procedure which can create fractions significantly enriched in certain of the minor sterols of the gorgonian sterol extract so that the GC-MS analysis of these minor sterols was possible. The results demonstrate (Table 5) that higher molecular weight sterols form digitonides more readily than do low molecular weight sterols (C sterols > C sterols 2 C sterols). Double bond position 29 28 27 within the sterol side chains does not influence ease of digitonide for22 YA2'+(26)I mation (A , although sterols with unsaturated side chains precipitate slightly less readily than their saturated analogs. A major exception to the finding of increased ease of precipitation of high molecularweight sterols is gorgosterol (2) and 23-demethylgorgosterol(8). The presence of the 22,23-cyclopropanering in gorgosterol and demethylgorgosterol significantly reduces the ease of precipitation of these

sterols by digitonin. 01,

It should be pointed out, however, that gorgoster-

a C 30 sterol, precipitates more easily than 23-demethylgorgosterol,

a c29

sterol, this fact perhaps being a reflection of the earlier men-

tioned finding of increased ease of precipitation

of sterols as their mo-

lecular weight increases. The real power of this digitonin fractional precipitation procedure lies in its ability to create remarkable enrichments of the short side chain or low molecular weight sterols (Table 1 compounds l-16).

The

third soluble digitonin fraction (see Table 5) had nearly a 50 fold enrichment of many of the short side chain sterols (Table 1, compounds l16) which allowed the production of good quality GC-MS derived mass spectra and even preparative HPLC or GLC collection of some

samples

for NMR and further analysis. Table 5.

Major Sterols Present in Digitonin Fractionally Precipitated Ps. porosa Sterol Mixture: Percent Compositions.

Fractionsb

10

20

Sterol Reference Numbera 24 27 25 29 31

l-Dig. Precip. I

0.03

10

13

15

4

8

2.5

4

40

2-Dig. Precip. II

0.6

10.5

12.5

15

4

1.5

1.2

12

45

3-Dig. Precip. III

2.5

7

12

12

4

0.4

0.5

17

42

4-Soluble Dig. I

2

20

12

18

6

6

1

6

30

5-Soluble Dig. II

1.8

10

11.5

18

6

0.5

0.5

14

41

5

25

6-Soluble Dig. III 34

2


30

37

a. See Table 1 for identification of sterol from sterol reference number; b. See Experimental section for method of production of each fraction. ALUMINA CHROMATOGRAPHY Neutral alumina (Brockmann grade II or III) column chromatography has been employed by past researchers for the separation of 4-monomethyl and 4,4-dimethyl sterols from 4-demethyl sterols [2,6].

We have found

that neutral alumina (Brockmann grade III) column chromatography employing a gentle gradient of diethyl ether in hexane has significantly more utility in that A4-3-one steroids, 58-stanols, 4-methyl sterols and 4-demethyl sterols (see Tables 6 and 7) can be separated. In addition different 4-demethyl sterols are enriched in various fractions. A ratio of alumina (III) to sterol mixture of at least 3OO:l is required to achieve separations of 4-demethyl sterols. Higher alumina (III) to sterol mixture ratios improve the separations. Neutral alumina (III) provides separation better than alumina of greater or less Brockmann activity. Sterols were eluted in the following order as can be seen in Table 5: 1) A5-C27 sterols, 2) A5-C28 sterols, 3) A5y22-C28 sterols, 4) A5-C28 stercls, 5) A5y22-C28 sterols, 6) gorgosterol, together with a short side chain sterol of M+ 316. The latter is a C21 sterol possessing an additional (non-hydroxy) oxygen atom which may explain the polar nature of this sterol. Although the alumina column chromatographicprocedure described here does not afford clean separations of the 4-demethyl sterols, it does create fractions with various enrichments of a number of the minor sterols sufficient for production of good quality GC-MS derived mass spectra as indicated in Table 1.

Recently an important separation of 24& and 2

isomers of steranes corresponding to the naturally occurring marine or plant sterols has been achieved using alumina (I) [28]. These reported sterane separations, together with the 4-demethyl sterol separations on alumina activity III reported here suggest that the alumina column chromatographic separation of sterols might be improved still further. Investigations along these lines using microparticulate alumina (5 micron diameter) and high pressure liquid chromatographic techniques are now underway in this laboratory.

S Table 6.

TRBOXD6

Major Sterol Compositions of Al ~_~ 0 (Act. III) Column Chromatographic Fractions of Ps.porosa t&o1 Mixture.

Sterol Reference Numbera Fraction 10 5

-

6

--

7

20

24

2%

4%

3%

1%

6%

3%

1%

13

35

2

13

22

31

5

0.05 25 12

27

8

-

9

-

510

10

-

2

4.52 1.2

11

2

0.3

12

9

-_

13

17

25/29

5.2

-

4

48

Eluent Hex:Et 0 (125 m? for each 37 Fraction)

Total Wt. mgs.

31

30

90%

-

-

- (94:6)

0.3

90%

-

-

- (94:6)

1.2

7

5

- (94:6)

2.2

2

19

9.5 (91:9)

8 8

-

0.5

5

_-

-

-

-

-

-

_

103

0.1

7

65 (91:9jb

73

-

5

79 (91:9)b

58

2.7

89 (91:9)

33

_

-

91 (91:9)

3

-

-

79 (91:9)

13

a. See Table 1; b. 50 ml of eluent used instead of 125 ml.

pble

7.

Major Sterol Percent Compositions of Al,& Fractions of P. homomalla Sterol Hixtws

(Act. III) Column Chromatographic

EluelIt 7/10

20

Fraction 3 4 5 6 7 6 9 10 11 12 13 14 15

0.3 0.7 6 16 45

_ ___ 1 2 -__ __ -__ 32 13 7 11 10

Sterol Reference Numbera 24 27 25/29 4% 31 49 -

_

_-

_

-

_ -

6 6 3 4 _

12 3.5 2.5 3 _

_ _ _ _ _ _ 11 18 7.5 8 _

10 70 90 90 70 50 -

30

37

5052

_ _ -60 _ -60 15 -_-__ __-___-_-4 92l0.5 5 50 0.2 .l 67 0.1 45 _ -40 -

a See Table 1 for identification of sterol from sterol reference number.

Hex-Et 0 33/34 (100 9/ frac.) 5 50 10 -

94:6 94:6 94:6 94:6 94:6 94:6 94:6 94:6 91:9 91:9 91:9 91:9 91:9

Total Wt. mgs. 2.5 2.1 9.4 16.2 9.7 4.6 2.0 4.6 105 75.5 6.3 2.9 1.1

s

715

WDEOXDrn

CHROMATOGRAPHY USING SILVER NITRATE IMPREGNATED ADSORDENTS Column chromatography using silver nitrate impregnated silica gel or alumina has been widely used in marine sterol isolations. In fact this technique coupled with argentic TLC [29] has been the most often used technique in the resolution of marine sterol mixtures. The selectivities of these techniques have been graphically illustrated in a number of articles [2] and the mechanism of separation via. silver ion double bond complexation has been discussed [29]. We have found that argentic chromatographyusing alumina as the support not only results in better separations of quickly eluting sterols but also results in some contamination of more slowly running sterols (such as those possessing =CH2 groups) with polyoxygenated compounds. Argentic silica gel chromatography does not result in this contamination problem and can therefore be used in the fractionation of the most slowly running sterols. The first three fractions presented in Table 8 contained enrichments of the stanols which were easily recognized by the intense molecular ions of the stanol acetates in the mass spectra of these fractions as contrasted with intense M-HOAc peaks and no molecular ions for A5steno1 acetates. The stanols were subsequently isolated using m-chloroperbenzoic acid oxidation and TLC system 1.

The next fractions (Prac-

tions 4-9, Table 8) contained good enrichments of dinosterol [3] (Sterol 48, Table 2) probably because the only olefinic linkage in dinosterol is a 22-23 trisubstituted double bond which, it is reasonable to assume, had decreased interaction with silver ions due to steric effects.

S

716

TIlROIDCl

The most polar fractions contained significant enrichments of a series of A5-19-nor-sterols

[30] (Sterols 17,21, and 26, Table 1).

These sterols have an increased exposure of the A5 double bond due to the absence of the C-19 angular methyl group and therefore have the possibility of increased silver ion - double bond interactions which explains the greater retention of these sterols.

The slowly eluting nature

of certain high molecular weight sterols along with limited mass spectral information suggests that these sterols possess =CH2 groups in the side chain; work is currently in progress on their structure.

Table

Fraction *

1*

8.

Major Stem1 Compositions of Al 0 /AgNO of Ps porosa Sterol Mixtw+3--3

10

19

-

_

:* 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23

-

_

0.1 0.1 __ -_ 0.1 0.1 0.1 0.1 2 15 22 -_ -

-

1.2 8 25 22 48 23 _

20

Column Chmmatographic

Sterol Reference Nun$era 24 27 25/29 4S 31

_

_

15 10 9 9 9 8 8 10 6 2.5 0.1 -78 -70 -73 ----

0.2 1 7.2 SO

34 30

37 35 20 16 10.5 9.5 9 12.5 11 5 0.1 -20 -30 -30

4.5 5.5 6 8 7.5 12 13 23 22 3 2

___13 4 3 5 10 5 1.8 __ ___ __ _ _ ___-

_ 2 2 2 1 2 2 0.4 1.5 0.6 0.5 0.1 __ -

30

~7

_

-

5 10 13 12.5 11.5 10.5 15 14 20 19 11 2.2 -

_ __ _

_ 23 39 45 50 48 47 55 51 50 46 54 15 14 3 1.5 0.5 -

Fractions

Elulmt Hex:Et20

Weight mgs.

l:Ob l:oc 100:ld 100:ld 100:zd 100:zd 100:3d 100:3d 100:4d 100:4d 100:sd 100:5d 100:7d 100:7d 100:lOd 100:lOd 100:lSd 100:15d 100:20d 100:20d 100:30d 100:30d BinzeneC

4 0.7 0.6 0.6 2.8 1.6 1.5 2 3.4 13.5 18.6 8.8 3 1.6 3.8 4.5 3.8 1.1 1 2.6 4.3 24.1 19.1

a. See Table 1 for identification of sterol from sterol reference number; b. 150 ml eluent/fraction; C. 200 ml eluent/fraction; d. 100 ml eluent/fraction; stStanolcontaining fractions; e. dinosterol plus an unidentified component.

REVERSED-PHASE HIGH PRESSURE LIQUID CHROMATOGRAPHY In a recent and initial application of reversed-phase HPLC to the separation of sterol acetate mixtures, Rees, et al. [13], have demonstrated that the technique can separate C27, C28, and C29 sterol mixtures

with some selectivity for position of unsaturation (although the

method is not as selective as argentic TLC). Working with free sterols for mass spectrometric reasons [35], we have applied this technique to the complex sterol mixtures of Es.porosaandP. homomalla. A solvent system of methanol-H20 (92:8) was found to give optimum resolution (see Fig. 1) for the column employed (see Experimental), It was also found necessary to inject the sterol sample in solvent of the exact composition of the elution solvent because use of a solvent of higher polarity was impossible for solubility reasons and use of a solvent of lowerpolarity resulted in much of the sterol mixture eluting with the solvent peak

and a further quantity slowly bleeding from the column and contaminating later fractions. This

could

only be detected by GLC analysis of the frac-

tions since the chromatogram recorded from the differential refractometer showed little sign ofcontaminationand indeed looked identical to Fig. 1 even when the various peaks were significantly contaminated with unresolved sterol mixture as revealed by GLC.

The order of elution

5,24(28) (see Fig. 1) was A5y22-C27, A5,22-C28, A5-C27, A -'28' A5-C28, A5(22-23 cyclopropyl)_c 5(22-23 cyclopropyl)_C 29 and A Clean prepa30: rations of 24S-methylcholesta-5,22-dien-3-01, ergosta-5,24(28)-dien-301, 246-methylcholest-5-en-3B-01,23-demethylgorgosteroland gorgosterol (see Nos. 24, 25, 4, 12 and 37 in Table 1) were found in various fractions of the gs.porosa sterol mixture as revealed by GLC.

High Pressure Liquid Chromatogram of 2. porosa I-beeSterol Mixture. column: u Bondapak C 8, 2 columns in series tsee Experimental) sample: 1.0 mgr in 0.5 ml of elution solvent. elution solvent: Methanol-H20 (92:8) pressure: 21OO/psi flow rate: 1.6 ml/min * Numbers abcve peaks refer to sterol reference numbers (see Table 1).

Application

of more than a milligram of sterol mixture unfortunately

results in a significant decrease in resolution of the various components. The technique, therefore, is not a viable tool for the initial examination of very minor or trace components of marine sterol mixtures, but is a valuable tool for the resolution of enriched fractions obtained by the previously described procedures. STRUCTURAL ELUCIDATION Using the GC-MS system described below (see Experimental),

each com-

ponent needs to be present at a concentration of at least 5% w/w of the sterol mixture with at least 1 pg of the component present in the injected mixture for the production of a good quality mass spectrum.

This con-

centration limit varies somewhat with the GC resolution and relative concentration of accompanying components. fractions of the preceding enrichment

Tables 1, 2, and 3 indicate which procedures met the above criteria

and therefore resulted in good quality mass spectrometric

information

(from the standpoint of purity and diagnostic peak intensities) for the sterols and A4-3-keto steroids considered. Sterols with reference numbers 19, 20, 23, 24, 25, 27, 28, 29, 30, 31, 37, and 41 (see Table 1) have well known structures which were identified by GC coinjection with an authentic sample and comparison of their identical mass spectra. Structural elucidation of the A5-19-nor-sterols (sterol No. 17, 21, 26 - Table 1, structures 15, 16 and II, respectively) is the subject of 30 another communication. 5a-23-Demethylgorgostanol(Sterol No. 32, Table 1, Structure -12). The stanol containing fractions (first fractions eluted by Hex-Et20 (1:O and 1OO:l)) from argentic alumina column chromatography of the p. homomalla sterol mixture was subjected to m-chloroperbenzoicacid oxidation (see Experimental) and the stanols isolated by preparative TLC (System 1).

The GC of the isolated stanols exhibited a peak with retention

time (1.65, Table 1) identical to 23-demethylgorgosterol(8_)(M+=412). The mass spectrum of this component possessed a molecular ion = 414 with a base peak of -m/e 316 characteristic of sterols possessing a 22-23 cyclopropane ring31 since the method of isolation excludes the presence of double bonds in the side-chain. A sample of the hitherto undescribed 5a-23-demethylgorgostanol(12) was synthesized from an authentic sample of 23-demethylgorgosterol(see Experimental) and was found to have a retention time and mass spectrum identical to those of the naturally occurring p. homomalla stanol. 56,246-Ethylcholestan-36-01(Number 34, Table 1, Structure -14). Column chromatography on A1203 (Act. III) of the total sterols of

S

720

-P. homomalla

TDEOIDrn

(Table 7, Fraction 4-6) resulted in the isolation of two

components eluted immediately after the A4-3-keto steroids (see Table 3). Preparative TLC (System 4) resulted in the purification of these components from contaminating 4-monomethyl sterols and A4-3-keto steroids. of this fraction revealed two closely running components.

Each component

was enriched by preparative GLC (3% OV-17, see Experimental) spectrum was run.

The major component

The GLC

and its mass

(Retention time=1.33, OV-17, see

Table 1) had a molecular ion = 416 and fragmentation typical for 245ethylcholestan-

36-01: -m/e 401, 398, 383, 257, 233, 214, 215, 216, 217.

32

The retention time however was somewhat less than that of authentic 5e,, 24c-ethylcholestan-3B-01.

Furthermore, the mass spectrum of this com-

ponent also showed an intense -m/e 149 peak characteristic of 56-stanols.

32

It is also known that 56-steranes elute faster than 5a-steranes during alumina chromatography during

28

and 5$-stanols elute faster than 5a-stanols

florisil chromatography.

33

The mass spectrum did not exhibit

an intense M-H20 characteristic of 3a-hydroxy-5@stanols, well with 36-hydroxy-56-stanol

M-H20 intensities.

32

that this component was 58,245-ethylcholestan-36-01 and MS comparisons with an authentic sample.

34

but correlated

The indications was confirmed by GC

The structure of the

second component (M+=414, Retention time=1.62) is under investigation. Dinosterol

(Sterol No. 48, Table 2, Structure -18).

Fractions 4 and 5 (Table 6) from alumina chromatography

of the -Ps.

porosa sterol mixture consisted to the extent of over 90% of a sterol with a GC retention time of 1.52 (3% OV-17) and with Mt=428 (corresponding to a C3D sterol).

The base peak in the mass spectrum was found to

be at -m/e 287 with intense peaks at -m/e 271 and 316.

Peaks of lesser in-

S

TDEOSDI

721

tensity occurred at 413(M-15), 410(M-18), 385(M-43) and 367(M-61). The mass spectrum of the acetate exhibited all intense peaks mentioned above but shifted by 42 mass units (428 to 470, 316 to 358, and 287 to 329) thus indicating the absence of a A5-double bond (which would have resulted in 2 mass unit shifts due to the loss of HOAc). Furthermore, the peaks at -m/e 271 and 287 in the spectrum of the free sterol correspond to ions of masses 257 and 273

(14 mass shifts)

respectively in the mass

spectrum of cholestanol which indicates a saturated ring system containside ing an additional methyl group and possessing an unsaturated C 10 chain. The double bond must be located at positions 22 and 23 as indicated by the -m/e 385 (M-C3H7) and 367 (M-(H20 + C3H7) peaks with the C3H7 group originating from carbons 25, 26, and 27, and especially by the intense m/e 316 peak which is due to the characteristic vinylic 20-22 fission -(with hydrogen transfer) reported for A22-unsaturated steroids.35 The NMR spectrum exhibited only a single olefinic proton (6 = 4.875) indicat3 ing alkylation at positions 22 or 23.

The retention time, mass spectrum

and NMR spectrum of an authentic sample of the C-23 alkylated sterol, dinosterol (E), isolated from an Atlantic dinoflagellate3 and supplied by Prof. Y. Shimizu proved to be identical with our C30 sterol. This report constitutes the first identification of dinosterol in a marine animal. A4-3-Ketones (Steroid Nos. 51, 51, 52, Table 3). The first fractions (3-5, Table 7) from alumina column chromatography of the P. homomalla sterol mixture were found to contain components with TLC (System 1) mobility equivalent to cholest-4-en-3-oneand with an W maximum at 238 nm.

The GLC of the fraction displayed peaks of retention

S

722

TIIROIDII

times 1.50, 1.91 and 3.32 in a ratio of 4:1.7:2.

The GC-MS of this frac-

tion showed that the components had molecular weights of 384, 398 and 424 respectively.

Furthermore, the mass spectrum of each component exhibited

no loss of 18 mass units (H20), an intense loss of 42 mass units (CH2= C=O), a peak at -m/e 229 diagnostic of a diunsaturated nucleus, and a very 36 (except compointense peak at -m/e 124 characteristic of A4-3-ketones, nent with M

+

= 424,

see below).

The component with M=384 was found to

have a retention time and mass spectrum equal to that of an authentic sample of cholest-4-en-3-one. The second component (ret. time = 1.91) had a mass spectrum displaying peaks at -m/e 398 CM+), 383 (M-CH3) and 356 (M-42) which are equivalent to the corresponding cholest-4-en-3-one plus an additional methyl group.

fragments 384, 369 and 342

The mass spectrum also exhibited a peak

at -m/e 275 which is equivalent to the mass 261 ring C and D fragment of cholest-4-en-3-one

plus an additional side chain methyl group.

ogy to the many naturally occurring 24-methylated marine sterols

By anal1

we

assume that this new A4-3-keto sterol analog is 245-methylcholest-4-en3-one. The final component (M=424) had a mass spectral fragmentation different from the other two components: small loss of CH 3, no loss of CH2=C=0 but exhibiting peaks corresponding to (M-43) and (M-79) fragments.

The

mass spectrum did exhibit an -m/e 312 peak corresponding to the -m/e 314 peak in the mass spectrum of gorgosterol and indicating a second unsaturation in the nucleus.

The mass spectrum also displayed peaks at -m/e 124,

269, 270, and 298 indicating a diunsaturated nucleus and either a 22-23 cyclopropane-containing

side chain or a side chain possessing a 22-23

S 31 double bond.

'EEEOIDI

723

The known gorgost-4-en-3-onewas synthesized (see Experi-

mental) and was found to have a mass spectrum and retention time identical with that of the third component. CONCLUSION Many questions arise concerning the origin, mode of biosynthesis, and function of the large number of minor sterols found in Pseudoplexaura porosa and Plexaura homomalla. Questions of this nature are particularly difficult to answer for gorgonians because gorgonians are suspension feeders and, therefore, have an extremely heterogeneous diet; furthermore many gorgonians, including the species considered here, contain large quantities of the endosymbiotic zooxanthellae, Gymnodinium microadriaticum, in their tissues. Thepreciseorigin of any minor sterol be it from dietary sources, zooxanthallae or coelenterate tissues is therefore very difficult to demonstrate. Reasonable hypotheses can, however, be advanced for several of the minor sterol classes. For example, the presence of a 5S-stanol (34, Table 1, Structure 14) is not surprising since these are typical fecal sterols of higher animals and may therefore be excretory products in gorgonians although it is troubling that no other 5B-stanols were encountered. The only other reported isolation of a 5S-stanol from a marine organism is coprostanol from 38 whale oi1.37 A4-3-Keto steroids have been found in both marine animals 39 This is the first reported occurrence of 24S-methylcholestand plants. 4-en-3-one and gorgost-4-er-3-onefrom marine sources. Saturated 3-keto steroid analogs of sterols have been shown to be precursors of the steroid41 40 of higher animals. The al hormones and also to occur in the feces resolution of the role of the 3-keto steroidsin gorgonian fore awaits further investigations.

physiology there-

S

724

TDICOIDE

4-Monomethyl sterols can be easily demonstrated in gorgonian tissues using the procedures described above.

Such sterols have been demonstrated

to be present in a number of marine organisms and are generally believed to be 4-demethyl sterol precursors which may in some instances be accumulated from dietary sources.

2

Dinosterol (18)3 is a particularly -

interesting 4-monomethyl sterol.

We encountered it in reasonably high concentrations - 6% of the sterol mixture of P. homomalla and also in a somewhat lower concentration in -Ps. porosa.

Sterols alkylated at the 23 position are very rare, the prime ex-

amples being gorgosterol demethyl-A5-homologue.

44

42

and its isomers

43

and dinostero13 and its 4-

Since dinosterol is the major sterol of a plank-

tonic dinoflagellate and therefore a primary oceanic food source, and dinosterol also occurs in gorgonians which contain gorgosterol as their major sterol, the biosynthetic or dietary origin of dinosterol in gorgonians is an important problem awaiting resolution. This is the first report of the isolation of naturally occurring sterol 36-monoacetates

in marine invertebrates.

The sterol composition

of the sterol acetate fraction was quite similar to that of the freester01 fraction.

It is interesting that sterols Nos. 9 and 34 are found only

in the sterol ester fractions.

It is also interesting to note that there

is a greater proportion of the side-chain alkylated sterols in the higher sterol ester fractions (Fractions lt2, Table 4) than in the free sterol and sterol acetate fractions (Fractions 3t4, Table 4) which could lead to some interesting speculations concerning the role and origin of sterol esters.

These few findings suggest that sterol esters may be tied into

sterol biosynthetic processes in gorgonians in an interesting manner,

S

TElEOIDI

725

although more study is needed before the nature of these ties zan be elucidated. 24-N0r-C2~ sterols were not detected in the extracts of either -* Ps porosa

or p. homomalla which was disturbing to us because of the almost

45 ubiquitous reported occurrence of these sterols in marine animals. A subsequent analysis of a sample of -P, homomalla collected at a different location and time of year, however, revealed a sterol with a GLC retention time corresponding to that of 24-nor-cholest-5,22-dien-38-01at a concentration of approximately 0.01 percent of the sterol mixture. This finding is perhaps a reflection of a dietary variation. Perhaps most interesting is the occurrence of a series of A5-19-norsterols as minor components of the 2.porosa

30 sterol mixture.

Several

preliminary findings have suggested that 19-nor sterols may have a wider distribution in the minor sterols of marine animals. It is our feeling that g.porosa and P.

homomalla are neither unique, nor even unusual in

containing such a plethora of interesting minor steroid components. Further, we feel that reexamination of the minor and trace sterol components of even the most unpromising marine extracts using techniques similar to those reported here, will result in the identification of many new sterols which will either fill in gaps in suggested marine sterol biosynthetic pathways or suggest new or alternate pathways. It can safely be predicted that analyzing organisms along the food chain of a marine animal whose novel minor sterols are demonstrated not to be biosynthesized by that animal will lead to the discovery of organisms containing high concentration of these interesting sterofs.

726

S

T=ICOXD=

EXPERIMENTAL Analytical GLC was performed using a Hewlett Packard 402A chromatograph equipped with 4mm I.D. x 6' "U" shaped column containing either 3% OV-17 on Gas Chrom Q (GCQ), 3% OV-25 on GCQ or 3% Poly S-179 on GCQ (all from Applied Sci. Inc.) or 1% OV-25 on GCQ prepared in this laboratory. Oven temperatures were 276O, 256O, 260°, and 252O respectively. Helium was used as the carrier at a flow rate of 100 ml/min. Preparative GLC was performed using the same instrument equipped with 8mm I.D. x 6' "U" shaped columns containing 10% SE-30 on GCQ, 3% OV-17 on GCQ or 3% OV-25 on GCQ with oven temperatures of 270°, 276O, and 256' respectively. Effluent splitting was effected with a Hewlett Packard annular "T" shaped splitter operating at a split ratio of less than 1 to 10 (detector-collector). Helium was again the carrier and the flow rate was varied for optimum speed and resolution for each preparative separation. The standard 402A flame ionization detector was used throughout the work. Combined GC-MS analysis was performed using a Hewlett Packard 7610A gas chromatograph equipped with 2mm I.D. x 10' "U" shaped columns (3% OV-17 on GCQ or 3% Poly S-179 on GCQ (Applied Sci. Inc.) at a column temperature of 260°) and interfaced with a Varian Mat 711 double focussing mass spectrometer (equipped with an all glass Watson-Biemann dual stage separator and a PDP-11/45 computer for data acquisition and reduction)operating at a resolution of 1000 or 5000. Conventional mass spectrometry was performed using an A.E.I. MS-9 Ultraviolet spectra were recorded using a Cary 14 spectroinstrument. photometer, 1 cm path length matched quartz cells and methanol as the solvent. NMR spectra were recorded using a Varian XL-100 spectrometer (100 MHz, CDC13 solvent, and TMS internal reference). HIGH PRESSURE LIQUID CHROMATOGRAPHY (HPLC) HPLC was performed using a Haskel Model 28303 pump connected to a Valco 7000 psi loop injection valve through a series of Whitey 5000 psi rated valves, l/16" O.D. swage lock connections, 0.020" I.D. 303 stainless steel tubing and incorporating a O-5000 psi Ashcroft gauge. A Waters Associates 30 cm x 4mm I.D. micron Bondapak C in conjunction with a Waters Associates dual cell dl'+~e~~i;~lwZfZ~~ometer. The system had a maximum operating pressure of 4500 psi. All separations, however, were performed at 1000-2000 psi which resulted in a flow rate of 0.8-2.0 ml/min. for the mobile phase (methanol-water, 92:8) and column employed. EXTRACTION Air dried gorgonians (P. homomalla) were crushed in a mortar and pestle and subsequently ground in a Waring blender in pure acetone. The ground material was then transferred to a Soxhlet apparatus and continuously extracted for 36 hrs using a tissue to acetone (w/v) ratio of 1:6 Kg/l. The extract was reduced in volume under reduced pressure at 30400. The residue was subjected to initial Si02 column chromatography (see below). A portion of the extracted tissues was then subjected to base hydrolysis by refluxing in 1N KOH in ethanol for 3-4 hrs. The

hydrolysate was extracted with Et 0, washed, dried (Na2S04) and the sterols isolated via TLC system 3 using a cholesterol marker. A second portion of the ex=cted gorgonian tissues was subjected to acidic hydrolysis: refluxing in 5% HCl for 3-4 hrs followed by Et20 extraction, neutralization (NaHCO ) and drying (Na2SO ) of the extract, and isolation via TLC system 1 ?see Table 4, Fractions 5 and 5). of the sterols presen9 THIN LAYER CHROMATOGRAPHIC (TLC) SYSTEMS Silica gel used was E. Merck HF-254+366 Type 60 for both analytical and preparatyve applications. Aluminum oxide used was E. Merck GF-254 Type E. The visualizing agent was ceric sulfate in H2S04. System 1: Silica gel, hexane-Et20 (1:l) System 2: Silica gel, hexane-benzene (3:2) System 3: Silica gel, hexane-benzene (1:l) System 4: Alumina, hexane-Et 0 (1O:l) 29 Argentic TLC was performed using ?he technique of Idler. System 5: AgNO impregnated silica gel, hexane-benzene (5:2) with two developments. INITIAL SILICA GEL COLUMN CHROMATOGRAPHY OF FREE STEROLS AND STEROL CONJUGATES The residue containing sterol esters, free sterols, etc., was applied to a silica gel column (E. Merck 60, 0.063 - 0.2 mm dia. particles) at a w/w ratio of 1:lOO residue to adsorbent. Elution of the column was stepwise using pure hexane, hexane-benzene (3:2) and hexane-Et20 (1:l). The column was subsequently eluted with Et20-methanol mixtures yielding several polyhydroxy sterols which are not considered here. Fractions (50 ml) were collected (for a 100 g silica gel column) and each fraction was examined by TLC beginning with cholesterolstearatemarker and TLC system 2. The elution solvent system was changed after no further compounds were eluted by the solvent in use. A cholesterol acetate and cholesterol marker were used in the TLC (System 3 and System 1 respectively) examination of more polar fractions. High sterol esters, sterol acetates and free sterols were all purified using preparative TLC with the solvent specified above. DIGITONIN FRACTIONAL PRECIPITATION 20 The sterol precipitation procedure is basically that of Windaus, although the sterol/digitoninproportions are4garied. The digitonide regeneration procedure is that of Schoenheimer. Digitonin (20 mg) (J. T. Baker Chem. Co) dissolved in 10 ml of 90% ethanol-water was added to 40 mgs of P. porosa sterol mixture dissolved in 10 ml of 95% ethanol-water. After 2-hrs at room temperature (20-25O) the precipitate was recovered by filtration, and both the filtrate (Filtr. I) and the precipitate were retained. The precipitate was then dissolved in a minimal volume of dry pyridine (anal. reagent) with heating as necessary. A large volume of ether, 50-60 times the volume of the pyridine solution was then added to the sterol-digitonin-pyridinesolution. After one hour the ether-pyridine solution was filtered. The filtrate corresponds to the free sterols (Dig. Precip. I) in Table 5. The filtrate from the original precipitation (Filtr I) was then evaporated to dryness (under reduced pressure) and extracted twice with lo-15 ml of dry diethyl ether to isolate uncomplexed

free sterols , 31.5 mg (uncomplex. I). The insoluble residue consisted of soluble (in ethanol-water) digitonin-sterol complexes and so was dissociated with pyridine and free sterols recovered with diethyl ether as described above. This sterol fraction recovered from the soluble digitonides is represented by "Soluble Dig. I" in Table 5. The uncomplexed sterols from the first precipitation (Uncomplex If were then precipitated with a larger quantity of digitonin, 40 mg, with the precipitation and work-up the same as described above for the initial free E. porosa sterol mixture. This second precipitation step resulted in the formation of a second precipitated digitonin sterol complex fraction (Dig. Precip. II), a second soluble digitonin sterol fraction (Soluble Dig. II) and the recovery of 18 mgs of noncomplexed sterols. The mixture of noncomplexed sterols was subjected to a final precipitation with digitonin using the same procedure as for Step 1 and 2, but with 45 mgs of digitonin. This final precipitation step yielded a third precipitated digitonin-sterol complex fraction (Precip. Dig. III) and 2 mg of an uncomplexed material which is non-steroidal. All sterol fractions were analyzed via GLC as described below, and certain fractions were analyzed by GC-MS= indicated in column 6 of Table 1. Saponification of sterol esters was performed by boiling in 1N KOH in methanol for 2 hrs followed bv the standard work-up. Sterol acetates were formed-from 3 -hydroxy sterbls by standard methods (25O, 24 hrs) with pyridine-acetic anhydride.

Al,0

(BrockmannAct. III) Column Chromatography 3Naturallv occurring free sterols or sterols derived from saponification of sterol esters w&e chromatographed over a column of neutral alumina (BrockmannActivity III)(E. Merck Aluminum oside 90) at an adsorbent to sterol mixture (w/w) ratio of at least 300/l. Elution solvents, the total weight and major sterol compositions of all fractions of an initial 300 mg sample of R.porosa and 300 mg sample of P. homomalla sterols are given in Table 6and 7 respectively. All fractyons were analyzed by GLC and by GC-MS as indicated in Table 1. 58-Stanols (fraction 4-7, Table 7) were purified by preparative TLC system 4.

ARGENTIC CHROMATOGRAPHY Sterol acetates (formed using the above procedure) were chromatographed using 30% (w/w) AgNO - Alumina (E. Merck Aluminum Oxide 90). The AgN03 was dissolved in tfl. e smallest possible volume of acetonitrile and the solution was added to the alumina in a foil-covered round bottom flask. The acetonitrile was removed under reduced pressure at 50° for 5-7 hrs (it is very important to remove all traces of the acetonitrile). The argentic adsorbent was packed into a foil-covered column and washed with the starting solvent (hexane). Elution solvents, the total weight and sterol compositions of all fractions of a 300 mg sample of _R.porosa sterols acetates are presented in Table 8. m-CHLOROPERBENZOIC ACID.OXIDATION: ISOLATION OF STANOLS The procedure is a slight modification of Goad's.L The first frac-

tions from argentic alumina column chromatography (see Table 8, Fractions l-4) contaifing significant enrichments of stanol acetates contaminated with some A sterol acetates were combined and 5 mg was dissolved in 5 ml of CHCl and mixed with 5 ml of CHC13 containing m-chloroperbenzoicacid (Aldricil). After stirring for 4 hrs at R.T. the solution was dried over anhydrous Na SO4 and evaporated under reduced pressure. The product was chromatograpZed (TLC System 1) with a cholesterol acetate marker, and the band with Rf equal to cholesterol acetate consisted of 56-stanol acetates (56-stanol acetates and monomethyl stanol acetates have slightly higher Rf's) as evidenced by subsequent mass spectra. The isolated stanol acetates amounted to 3.6 mg, i.e., 60% of the weight of the starting combined argentic alumina chromatographicfractions. FATTY ACID ANALYSIS The high fatty acid sterol esters isolated by SiO2 column chromatography (Table 4, Fraction 1) and TLC System 2 were saponified as described above except that the basic solution left after ether extraction of sterols and other unsaponifiable lipids was washed, acidified with dilute HCl to a pH 1 and subsequently extracted with ether. The ether extract was washed with water to neutral pH, dried and evaporated under reduced pressure to a white solid. The fatty acid methyl esters were then formed by refluxing,lO mg of fatty acid extract with 0.5 ml of a 14% w/v solution of boron trifluoride in methanol (Supelco, Inc.) for 2 minutes. Water was added to the reaction mixture and the fatty acid methyl esters were extracted into Et20. The fatty acid methyl esters were subsequently analyzed using temperature programmed GLC (80° to 260° on 3% OV-17 on GCQ program rate 5O/min) and comparison with authentic samples by GC-MS (see footnotes in Table 4). Gorgost-4-en-3-on? Gorgosterol (3 mgs) was refluxed with 0.5 ml cyclohexanone ( dry) and 10 mgs of aluminiwn isoproperoxide (Eastman) in 2.0 ml of dry toluene for 5 hrs. The solution was evaporated under reduced pressure and the residue chromatographed (TLC system 1) using a cholest-4-en-3-one marker. The band with Rf = cholest-4-en-3-one (~1 mg) was found to have only a single peak (retention time+= 3.32) in the GC and displayed the expected UV maximum (239 nm) and M = 424. 5a-23-Demethylgorgostanol 23-Demethylgorgosterol(1.5 mg) was dissolved in ethyl acetatemethanol (1:l) containing 1.0 mg of platinum oxide, hydrogenated for 24 hrs at atmospheric pressure and worked up by standard methods. Acknowledgements. We are grateful to Prof. L. J. Goad and Prof. J. T. Baker for copies of their reviews in advance of publication and to Robert G. Ross for mass spectra obtained on the MS-9 spectrometer. We thank Prof. A. J. Weinheimer (University of Oklahoma) for a generous sample of xs.porosa sterols and Dr. F. S. Alvarez (Syntex Research, Palo Alto) for generous samples of 2. homomalla and several sterol containing fractions from that animal. We also thank Prof. Y. Shimizu (University

730

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TIIEOXD=

of Rhode Island) for a sample of dinosterol, Dr. Leif Aringer (Karolinska Siukhuset, Stockholm) for a sample of 5$,24-ethylcholestan-38-01, Prof. F. J. Schmitz (University of Oklahoma) for 23-demethylgorgosterol, and Prof. P. J. Scheuer (University of Hawaii) for gorgosterol. Thanks also go to M. H. Kohrman for preparation of several GC packings and columns, L. Dunham for the 10% SE-30 preparative GC column, Dr. D. A. Schooley (Zoecon Corp.) for suggesting the reversed-phase HPLC technique and for invaluable help in the design of our HPLC instrument and W. C. Dow for the construction and operation of that instrument. This research was supported by grants from the National Institutes of Health (GM 06840 and RR 00612). REFERENCES 1.

(a) Goad, L. J., "The Steroids of Marine Algae and Invertebrate Animals", to be published (1976). (b) Baker, J. T. and Murphy, V., "Handbook of Marine Science Compounds from Marine Organisms, Vol. 1, CRC Press, Cleveland, Ohio, 1976.

2.

Kobayashi, M. and Mitsuhashi, H., STEROIDS, 26, 605 (1975); also see Smith, A. G., Rubinstein, I. and Goad, L. J., BIOCHEM. J., 135, 443 (1973).

3.

Shimizu, Y., Alam, M. and Kobayashi, A., J. AMER. CHEM. SOC., 98, 1059 (1976).

4.

Fattorusso, E., Magno, S., Mayol, L., Santacroce, C. and Sicz, D., TETRAHEDRON, 2, 1715 (1975).

5.

deluca, P., deRosa, M., Minale, L., Puliti, R. and Sodano, G., J. CHEM. SOC., CHEM. COMM., 825 (1973).

6.

Minale, L. and Sodano, G., J. CHEM. SOC. PERKIN I, 1888 (1974).

7.

Kobayashi, M. and Mitsuhashi, H., STEROIDS, 26, 605 (1975).

8.

Kobayashi, M. and Mitsuhashi, H., STEROIDS, 24, 399 (1974).

9.

Kobayashi, M. and Mitsuhashi, H., TETRAHEDRON, 30, 2147 (1974).

10.

Ballantine, J. A. and Roberts, J. C., TETRAHEDRON LETT., 2_, 105 (1975).

11.

Idler, D. R., Khalil, M. W., Gilbert, J. D. and Brooks, C. J. W., STEROIDS, 27, 155 (1976).

12.

Patterson, G. W., Khalil, M. W. and Idler, D. R., J. CHROMAT., 115, 153 (1975).

13.

Rees, H. H., Donnahey, P. L. and Goodwin, T. W., J. CHROMAT., 116, 281 (1976).

s

731

TBEOXDI

14.

(a) Hetzel, C. A., Chang, T. and Buhle, E., "Identificationof Some Sterols in a Gorgonian Coral", 22nd Annual Conf. on Mass Spectrometry and Allied Topics (1974). (b) Bowen, J. L., M.S. Thesis, Stanford University, 1970. (c) Ciereszko, L. S., Johnson, M. A. Schmidt, R. W. and Koons, C. B., COMP. BIOCHEM. PHYSIOL., 2, 899 (1968).

15.

For leading references see (a) Djerassi, C., PURE APPL. CHEM., 21, 205 (1970). (b) Waller, G., "Biochemical Applications of Mass Spectrometry", Wiley-Interscience,New York, N.Y., 1972. (c) Zaretskii, Z. V., "Mass Spectrometry of Steroids", Israel Universities Press, Jerusalem, 1976.

16.

Giese, A. C., Krishnaswamy, S., Vasu, B. S. and Lawrence, J., COMP. BIOCHEM. PHYSIOL., 13, 367 (1964).

17.

Bligh, E. G. and Dyer, W. J., CAN. J. BIOCHEM. PHYSIOL, 37, 911 (1959).

18.

Schmid, P. and Hunter E., PHYSIOL. CHEM. 5 141 (1973). S&mid, P., ibid., _,

19.

Schmid, P., Hunter E. and Calvert, J., PHYSIOL. CHEM. & PHYSICS, 2, 151 (1973).

20.

Schmid, P., Calvert, J. and Steiner, R., PHYSIOL. CHEM. & PHYSICS, 2, 157 (1973).

21.

Knights, B. A., PHYTOCHEM., 2, 903 (1970) and Goodwin, T. W., "Algal Physiology and Biochemistry", ed. W. D. P. Stewart, Botanical Monograph, Vol. 10, University of California Press, Berkeley (1974).

E

PHYSICS, 3_, 98 (197l)and

22. van Lier, J. E. and Smith, L. L., STEROIDS, 15, 485 (1970). 23. Windaus, A., CHEM. BER., z,

238 (1909).

24.

Homberg, E. and Seher, A., Z. LEBENSM. UNTER. FORSCH., 149, 129 (1972).

25.

Haslewood, G., BIOCHEM J., 5,

26.

Haslam, R. and Klyne, W., BIOCHEM. J., 55, 340 (1953).

27.

Haust, H., Kuksis, A. and Beveridge, J., CAN. J. BIOCHEM., 2, (1966).

28.

Mulheirn, L. J. and Ryback, G., NATURE, 256, 301 (1975) and Ryback, G ., J. CHROMAT., 116, 207 (1976).

29.

Idler, D. R. and Safe, L. M., STEROIDS, 19, 315 (1972) and Johnson, P., Rees, H. H. and Goodwin, T. W., BIOCHEM. SOC. TRANS., 2_, 1062 (1974).

639 (1947).

119

S

TDEOXDII

30.

Popov, S., Carlson, R. M. K., Wegmann, A. and Djerassi, C., TETRAHEDRON LETT., in press.

31.

Hale, R. L., Leclerq, J., Tursch, B., Djerassi, C., Gross, R. A. J., Weinheimer, A. J., Gupta, K. C. and Scheuer, P. J., J. AMER. CHEM. sot., 92, 2179 (1970) and Schmitz, F. J. and Pattabirman, T. J., J. AMER. CHEM. Sot., 92, 6073 (1970).

32.

Klein, H. and Djerassi, C., CHEM. BER., 106, 1897 (1973) and references cited therein.

33.

Miettinen, T. A., Ahrens, E. H. and Grundy, S. M., J. LIPID RES., 6_, 411 (1965).

34.

Eneroth, P., Hellstrom, K. and Ryhage, Ii., STEROIDS, 5, 707 (1965).

35.

Wyllie, S. G. and Djerassi, C., J. ORG. CHEM., 28,

36.

Shapiro, R. H. and Djerassi, C., J. AMER. CHEM. SOC., 86, 2825 (1964).

37.

Gupta, K. S. , HOPPE-SEYLER'S Z. PHYSIOL. CHEM., 348, 1688 (1967).

38.

Sheikh, Y. and Djerassi, C., TETRAHEDRON, 30, 4095 (1974).

39.

Kanazawa, A. and Yoshioka, M., BULL. JAP. SOC. SCIENT. FISH., 37, 397 (1971).

40.

Lommer, D., Dorfman, (1970).

41.

Miettinen, T. A., Arhrens, E. H. and Grundy, S. M., J. LIPID RESEARCH, 6_, 411 (1965).

42.

Ling, N. C., Hale, R. L. and Djerassi, C., J. AMER. CHEM. SOC., 92, 5281 (1970).

43.

Sheikh, Y. and Djerassi, C., CHEM. COMMUN., 217 (1971).

44.

Kanazawa, A., Teshima, S., Ando, T. and Tomita, S., BULL. JAP. SOC. SCIENT. FISH., '0, 729 (1974).

45.

Silberberg, M. S., Ph.D. Dissertation, University of Oklahoma, 1971.

46.

Schoenheimer, R. and Dan, H., Z. PHYSIOL. CHEM., 215, 59 (1933).

305 (1968).

R. I. and Forchielli, E., STEROIDOLOGIA, 1, 257