Energy & Fuels 1990,4,675-688
675
Preparation of Bacteriopetroporphyrins by Partial Synthesis from the Chlorobium Chlorophylls Norman W. Smith and Kevin M. Smith* Department of Chemistry, University of California, Davis, California 95616 Received April 27, 1990. Revised Manuscript Received July 12, 1990 Chemical degradation of the natural bacteriochlorophyll d homologous mixture to give the corresponding bacteriopetroporphyrin methyl esters is described. Reduction of the correspondingmethyl bacteriopheophorbide d homologue with excess of sodium borohydride, followed by oxidation with 2,3-dichloro-5,6-dicyanobenzoquinone, affords the corresponding bacteriopetroporphyrin. 'H NMR spectroscopyof the nickel(I1) complexes is used to establish that the pigments are identical with those isolated from immature Messel oil shale. Similar degradative transformations of the bacteriochlorophyll c homologues (and also of the bacteriochlorophyll e mixture) to give "bacteriopetroporphyrins C" (which have not yet been observed in oil shales) are reported, and the proton NMR signatures of their nickel(I1) complexes are obtained and assigned.
Introduction Alfred Treibs was the first to discover petroporphyrins in petroleum oil shales, coals, etc. In the 1 9 3 0 ~he , ~iso~~ lated and identified the major metalloporphyrin present in petroleum sources, vanadyl deoxophylloerythroetioporphyrin (DPEP) (1) and vanadyl etioporphyrin 111. In 2b
CH,
1
1936 it was proposed3 that these compounds were degradation products of the chlorophylls and hemins and a scheme was suggested to account for their origin. New petroporphyrins continue to be isolated and identified, and it has recently been stated4 that there are now over 40 confirmed structures. The petroporphyrins are now generally accepted to be derived by geochemical degradation of chlorophyll over and there is significant recent evidence in support of There has been much speculation about the origin of petroporphyrins containing more than 32 carbons. One possible source for higher molecular weight petroporphyrins would be the Chlorobium chlorophylls (such as the bacteriochlorophylls c , d , and e , which have homologated side chains at positions 4 and 5 and, for the bacteriochlorophylls c and e, also at the &meso position). However, early analyses of maleimides from degradation of high molecular weight geoporphyrins showed that not only is (1) Treibs, A. Liebigs Ann. Chem. 1934, 510, 42. (2) Treibs, A. Liebigs Ann. Chem. 1934, 509, 103. (3) Treibs, A. Angew. Chem. 1936,49,682. (4) See other papers in this issue of Energy Fuels. (5) Quirke, J. M. E.; Eglinton, G.; Maxwell, J. R. J . Am. Chem. SOC. 1970, 101, 7693. (6) Quirke, J. M. E.; Maxwell, J. R. Tetrahedron 1980,36, 3453. (7) Quirke, J. M. E.; Maxwell, J. R.; Eglington, G.; Sanders, J. K. M. Tetrahedron Lett. 1980,21, 2897. (8)Fookes, C. J. R. J. Chem. Soc., Chem. Commun. 1983, 1472. (9) Fookes, C. J. R. J . Chem. SOC.,Chem. Commun. 1983, 1474. (10) Verne-Mismer, J.; Ocampo, R.; Callot, H. J.; Albrecht, P. Tetrahedron Lett. 1986,27, 5257. (11) Fookes, C. J. R. J . Chem. SOC.,Chem. Commun. 1985, 706.
0887-0624/90/2504-0675$02.50/0
the 4-isobutyl substituent on pyrrole subunit B not f o ~ n d , ~but ~ - Quirke l~ and c o - ~ o r k e r s found '~ alkyl substituents containing up to 11carbons that could not have arisen from simple degradation of Chlorobium chlorophylls. This led Baker and PalmerI2 to conclude that the likelihood of the Chlorobium chlorophylls being precursors to certain petroporphyrins was small because transalkylation mechanisms can satisfactorily explain higher molecular weight porphyrins. Recently, however, Ocampo and colleague^^^^^ have isolated a series of petroporphyrins from Messel oil shale that have homologous side chains at positions 4 and 5 showing a clear origin from photosynthetic (Chlorobium) bacteria. In a preliminary publication, we have reported their syntheses.18 Thus it is apparent that while the Chlorobium-type chlorophylls are contributors to the wide variety of petroporphyrins, they are probably not a major source for higher molecular weight porphyrins found in mature sediments or petr01eum.l~
Results and Discussion Although there are over 70 known petr~porphyrins,~ many of them have been structurally characterized only by NMR [with appropriate 2D and nuclear Overhauser enhancement (NOE) studies]. Although this method is an excellent characterization technique, for an organic chemist the ultimate proof of structure is total or partial synthesis using rational synthetic approaches; it is always reassuring to verify the NMR data by conducting a synthesis of the molecule in question and then comparing the data for the synthetic material against that of the natural product. (12) Baker, E. W.; Palmer, S. E. In The Porphyrins; Dolphin, D., Ed.; Academic Press, New York, 1978; Vol. 1, pp 486-552. (13) Quirke, J. M. E.; Shaw, G. J.; Soper, P. D.; Maxwell, J. R. Tetrahedron 1980, 36, 3621. (14) Barwise, A. J. G.; Whitehead, E. V. In Adoances in Organic Geochemistry; Maxwell, J. R., Douglas, A. G., Eds.; Pergamon Press:
Oxford, 1980; pp 181-92. (15) Ocampo, R.; Callot, H. J.; Albrecht, P. J . Chem. Soc., Chem. Commun. 1985, 198. (16) Ocampo, R.; Callot, H. J.; Albrecht, P. J . Chem. Soc., Chem. Commun. 1985, 200. (17) Ocampo, R.; Callot, H.; Albrecht, P. In Metal Complexes in Fossil
Fuels; Filby, R. H., Branthaver, J. F., Eds.; American Chemical Society: Washington, DC, 1987; pp 68-72. (18) Smith, N. W.; Smith, K. M. J. Chem. SOC., Perkin Trans. 1 1989, 188. (19) Filby, R. H.; Van Berkel, G. In Metal Complexes in Fossil Fuels; Filby, R. H., Branthaver, J. F., Eds.; American Chemical Society: Washington, DC, 1987; pp 2-39.
0 1990 American Chemical Society
Smith and Smith
676 Energy & Fuels, Val. 4, No. 6,1990 Chart I. Structures of BacteriopetroporphyrinsIsolated from Messel Oil Shale (as Methyl Esters) and Characterized by Ocampo et al.16
Chart 11. Structures of Bacteriochlorophyll d Homologues (2) and of Corresponding Methyl Bacteriopheophorbides d (4)
I "
I
CO*CH,
3 R4
RS
Total synthesis (especially of an unsymmetrical porphyrin) requires a large number of steps and therefore requires much time and effort in order to carry through the procedure. On the other hand, performing a partial synthesis can greatly simplify matters depending on the starting material, target molecule, and the routes available to achieve conversion. In the present case, the petroporphyrins that were recently discovered in Messel oil shale by Ocampo et (Chart I) were a particularly convenient system to study because of the readily available bacteriochlorophylls d (Chart 11)to which those particular petroporphyrins are so clearly related. In a certain sense, conversion of the bacteriochlorophylls d into the corresponding Messel oil bacteriopetroporphyrins would constitute a biomimetic synthesis. The skeletal relationship between structures %a,e,f,g and 3a-d is clear. The methyl bacteriopheophorbidesd (Bmph-d) (4) are readily obtained from the bacteriochlorophylls d, and the steps necessary to achieve the desired conversion of pheophorbides into the required porphyrins were envisioned to be possible, so the partial syntheses described below were undertaken. The green and brown sulfur bacteria are found in stagnant ponds, lakes, and estuarine habitats having vertical gradients of light (from above) and hydrogen sulfide (from be lo^).^ The main light-harvesting antenna of these sulfur bacteria are the homologous series of bacteriochlorophylls c, d, and e known as the Chlorobium chlorophylls.21 The bacteriochlorophylls d were first isolated from Chlorobium thiosulfatophilum and identified by Holtn-24and MacDonald and co-workers.25 The main esterifying alcohol at the 7-propionic position was determined to be farnesol.21 Smith and co-workers subsequently verified the stereochemistry of the 2-(l-hydroxyethyl) group for the homologues by NMR, HPLC,and X-ray analysis.26 For the bacteriochlorophylls d, cultures of Chlorobium vibrioforme forma thiosulfatophilum were grown on the (20) Simpson, D. J.; Smith, K. M. J.Am. Chem. SOC.1988,110,1753. (21) Holt, A. S. In The Chlorophylls; Vernon, L. P., Seely, G. R., Eds.; Academic Press: New York, 1966; p 111. (22) Holt, A. S.; Purdie, J. W. Can. J. Chem. 1965, 43, 3347. (23) Holt, A. S.; Hughes, D. W. J. Am. Chem. SOC.1961, 83, 499. (24) Holt, A. S.; Hughes, D. W.; Kende, H. J.; Purdie, J. W. J.Am. Chem. SOC.1962,84, 2835. (25) Archibald, J. L.; Walker, D. M.; Shaw, K. B.; Markovac, A.; MacDonald, S. F. Can. J . Chem. 1966,44, 345. (26) Smith, K. M.; Goff, D. A. J. Chem. Soc., Perkin Trans. I 1985, 1099.
R4
RS
2 M = Mg, R = Famesyl (Bacteriochlorophylls-d) 4 M = 2H,R = Methyl (Methyl Bacteriopheophorbides-d)
4.5- or 20-L scale for several days as described elsewhere.% The cells were collected by centrifugation and extracted with methanol, and a crude mixture of carotenoids and bacteriochlorophylls d was obtained. Treatment of the extract with sulfuric acid in methanol served to demetalate and transesterify the 7-propionic ester to give the homologous series of Bmph-d (4). The 5-ethyl series of homologues was separated from the 5-methyl series by preparative normal-phase HPLC, and the homologues within each subset were then separated by preparative reverse-phase HPLC. On paper, the necessary steps to achieve conversion of the Bmph-d (4) to the desired petroporphyrins 3 included (i) reduction of the 2-hydroxyethyl group to ethyl, (ii) reduction of the 9-keto to methylene, and (iii) oxidation of the chlorin to the porphyrin macrocycle. The porphyrin could then be chelated with nickel for proton NMR investigation. Procedures chosen to accomplish these steps had to leave the 7-methyl propionate substituent intact (Le., no reduction). Initial considerations included the following: Removal of the hydroxy group from the 2hydroxyethyl substituent is fairly straightforward. Dehydration of hydroxyethyl groups by treatment with a catalytic amount of p-toluenesulfonic acid in hot 1,2-dichlorobenzenez7has been done before on porphyrin compounds. This could then be followed by catalytic hydrogenation of the resulting vinyl group to give the ethyl group. Oxidations of chlorins to porphyrins can be effected by treatment with high potential quinones such as 2,3dichloro-5,6-dicyano-1,4-benzoquinone (DDQ).28 Attention, therefore, was focused first to determining an efficient way of removing the 9-keto group. Baker et al.29reported that treatment of pheophytins with hydrazine hydrate and strong base in triethylene glycol at high temperature (Wolff-Kishner conditions) effected reduction of the 9-keto group to methylene and also accomplished oxidation of the chlorin to the corre~~~~-
(27) Smith, K. M. Porphyrins and Metalloporphyrins;Smith, K.M., Ed.; Elsevier: Amsterdam, 1975. (28) Kenner, G. W.; McCombie, S. W.; Smith, K. M. J. Chem. Soc., Perkrn Trans. 1 1973, 2517. (29) Baker, E. W.; Corwin, A. H.; Klesper, E.; Wei, P. E. J. Org. Chem. 1968,33,3144.
Preparation of Bacteriopetroporphyrins
Energy & Fuels, Vol. 4, No. 6,1990 677
Scheme I. Wolff-Kischner Degradation of Bmph-d
Scheme I1 Et
M
e
M
R
Me
4
Me' '
W
H
7
+
C0,Me
13%
sponding porphyrin. When [n-Pr,Et]-Bmph-d (4f) was subjected to the same conditions, the desired compound was obtained, but in rather disappointing yields. Only 13% of the porphyrin and 4% of the corresponding chlorin were obtained (Scheme I). Since reduction of the 9-keto group can readily be accomplished with NaBH, in virtually quantitative yield, methods for conversion of the 9-keto group into the corresponding hydroxy group, followed by reductive deoxygenation, were next investigated. Also at this point, it was decided to try model reductive studies on methyl mesopyropheophorbide a (5), because of its more ready availability (compared with Bmph-d) from Spirulina maxima alga. Although reduction of aliphatic alcohols can seldom be accomplished by hydrogenolysis,3° reduction of benzylictype alcohols can often be performed in this way. Techniques to accomplish this objective have been reviewed by R ~ l a n d e r . ~As~ a model system, 5 was reduced to the 9-hydroxy compound 6 with NaBH, and then this was hydrogenated over 10% palladized charcoal in an attempt to obtain the 9-deoxo compound 7; however, only starting
&NH Me,"'
W
N 3 M
e R2
A
Me02C
Me
Et
R'
5 R',' = 0 6 R' = H, R2 = OH 7R''=H
material 6 was recovered. The attempted hydrogenation of the same compound in the presence of trifluoroacetic acid (TFA)32was also unsuccessful, this causing decomposition of the starting material. Attempts to deoxygenate with P21t3or with triethylsilane in the presence of TFA% gave very small amounts of product as determined by thin-layer chromatography (TLC). Raney nickel has been used to reduce nickel(I1) complexes of methyl pyropheophorbide a to provide mixtures
Me0,C
M
e
Me" "
-
I a
W
M
e
ti CHO CHO Me0,C
9
of its corresponding9-deoxo compound and di-, tetra-, and hexahydr~porphyrins.~~~~ Although the conditions have been worked out to give the 9-deoxo compound 7 in yields as high as 52%, similar treatment of Bmph-d (4) gave a mixture of several compounds, all in disappointingly minor amounts. An attempt to dehydrate the 9-hydroxy group of 6 by acid catalysis to give the 9,lO-dehydrocompound followed by catalytic hydrogenation gave similar results as obtained p r e v i o u ~ l y . ~The ~ product isolated was identified as a 9,lO-diformylchlorin. Apparently the dehydration proceeds to give the exocyclic alkeno ring 8 which is then attacked by dioxygen in air to give the ring-opened compound 9 (Scheme 11). Other methods to accomplish these reductive deoxygenations are abundant; they include the Clemmensen hydride reagents in combination with Lewis acids such as LiA1H4/A1C13,39s40 and many others.30 Less direct methods, such as conversion of the hydroxy group to the tosylate or another sulfonate ester, followed by displacement with LiA1H4,41-43with NaBH, in a dipolar aprotic solvent,44with LiEt3BH, and with Bu3SnH-Na16 are also available. Use of LiAlH, was not an option in our case, however, because that would have also reduced the propionic ester. The following method proved to be quite successful. It is a modification of work developed earlier involving the use of hydrides in conjunction with Lewis acids. Gribble and co-workersa reported that reduction of aryl carbonyl and hydroxy groups proceeded in high yield upon treatment with NaBH, in the presence of TFA. This method seemed particularly well suited for use on the Bmph since they contain both aryl alcohol and aryl ketone functionalities. It was hoped that this NaBH,/TFA method could provide both reduction of the 2-hydroxyethylto ethyl and also of the 9-keto to 9-CH2. Model studies using methyl pyropheophorbide a ( 5 ) gave encouraging results (75% yield of 7), so it was applied to the Bmph-d. Small-scale treatment of the homologous mixture 4e-h also gave very encouraging results, so conversion of a single homologue into its corresponding deoxo compound was attempted next. Because Ocampo et al.16 reported full NOE data for the Goff, D. A. Ph.D. Dissertation, University of California, Davis, Mengler, C.-D. Dissertation, Braunschweig, 1966. Lai, J.-J. PbD. Diasertation, University of Califomia, Davis, 1983. Martin, E. L.Org. React. 1942, 1 , 155. Brown, H. C.; Subba Rao, B. C. J. Am. Chem. SOC. 1956,78,2582. Blackwell, J.; Hickinbottom, W. J. J. Chem. SOC.1961, 1405. Dimitriadis. E.: Massy-Westrop. R. A. Aust. J. Chem. 1982.35. 1895.
(30) March, J. Advanced Organic Chemistry;Wiley: New York, 1985. over Platinum Metals: (31) Rylander, P. N. Catalytic Hydroaenation - -
Academic Press:. New York,-1967. (32) Dar'eva; Miklukhin, J. Gen. Chem. USSR 1959,29, 620. (33) Suzuki, H.; Tani, H.; Kubota, H.; Sato, N. Chem. Lett. 1983,247. (34) Adlington, M. G.; Orfanopoulos, M.; Fry, J. L.Tetrahedron Lett. 1976, 2955.
(42) Eschenmoser, A.; Frey, A. Helv. Chim. Acta 1952, 35, 1660. (43) Rapoport, H.; Bonner, R. M. J. Am. Chem. SOC.1951, 73,2872. (44) Hutchins, R. 0.;Hoke, D.; Keogh, J.; Koharski, D. Tetrahedron Lett. 1969, 3495. (45) Bell, H. M.; Vanderslice, C. W.; Spehar, A. J. Org. Chem. 1969, 34, 3923. (46) Gribble, G. W.; Leese, R. M.; Evans, B. E. Synthesis 1977, 172.
Smith and Smith
678 Energy & Fuels, Vol. 4, No. 6, 1990 Scheme 111
;
;
$
&
/
-
e
MH
Me'"
-N
Me I
HN
.
\
L
'H
4f
0 C02Me
NaBHJFA
14
OH
H---CMe
Figure 1. Structureof the by-product 14 from the DDQ oxidation of the correspondingchlorin, 12, and network of NOE connec-
Me
tivities.
J
C0,Me P
- @ &
-
-
10
C02Me
e M
Me;&-
11 H2/Pd-C
-
6O2Me
DDQ
-N
HN
b02Me
13 H
-N
Me"'
.
HN
''
3c
[4-n-Pr,5-Et] homologue (condensed t o [n-Pr,Et] throughout), we selected the same Bmph-d for conversion into the corresponding porphyrin. Thus, treatment of the [n-Pr,Et] homologue of Bmpd-d 4f with 10 equiv of NaBH, in dry TFA provided varying amounts of 9deoxo-2-(1-hydroxyethy1)-Bmph-d (10) and 9-deoxo-2vinyl-Bmph-d (11)(Scheme 111). No material was obtained that had not been completely reduced to methylene at the 9-position. Gribble found that this method was limited to compounds that were capable of forming relatively stable carbocations in acidic media, since benzyl alcohol and a series of aliphatic alcohols afforded little or no reduction product. The 9,lO-dehydro product is probably not formed in major amounts because of its instability; quenching of the carbocation with hydride to give the desired compound must be a more favored process than forming the strained exocyclic alkeno ring. In later experiments it was determined that use of a larger excess of NaBH, (in the form of pellets to allow for slower dissolution in TFA), and extension of the reaction time, gave the 9-deoxo-2-ethyl compound 12 directly. Catalytic hydrogenation of the vinyl group in 1 1 proceeded smoothly to give the ethyl compound 12 in quantitative yield. Treatment of 12 with DDQ gave the corresponding porphyrin 13 in good yield. Complexation with nickel was accomplished by dissolving in chloroform and treating with excess saturated nickel(I1)acetate in methanol47providing 3c in quantitative yield (Scheme IV). A by-product in the above DDQ reaction that was observed when the material was purified on silica gel was
material (14)which had been oxidized a t the 10-position to give the corresponding hydroxy compound. The hydroxylation was established to be a t the 10-position by proton NMR NOE difference experiments. Irradiation of the proton geminal to the hydroxyl group gave NOES a t the other ring position (position 9, as expected) and to the 7a-CH2and 7b-CH2positions. If the hydroxyl substituent were at position 9, it would have given an NOE to the 5-ethyl group (not observed) and not to the propionate substituent. Substitution a t any position other than on the exocyclic ring was clearly ruled out because all other signals in the NMR spectrum were unchanged. Figure 1 shows the NOE connectivities established by the experiments carried out. Benzylic oxidation involving DDQ has been observed previously. Lee and Harvey48 were able to oxidize the benzylic positions of a variety of polycyclic arylalkanes using DDQ. Further oxidation of the hydroxy to the corresponding aryl ketone was also observed. Apparently DDQ selectively oxidized our product a t the 10-position because that is where the most stable cation would be formed. There are a number of secondary benzylic positions around the macrocycle, but the 10-position is near a meso carbon and therefore is different from the others. Ponomarev and S h ~ l ' g anoticed ~ ~ that the 10-position of a number of cyclopentanoporphyrins was easily oxidized on the surface of silica gel. They obtained hydroxyporphyrins such as 14 upon drying preparative silica gel plates after development. I t is possible, then, that the formation of hydroxyporphyrins observed in our work is a combination of the two above factors since they both give the same result. Having established the pathway t o bacteriopetroporphyrin, the complete mixture of methyl 5-ethylbacteriopheophorbides d (HPLC trace, Figure 2) was
(47) Fuhrhop, J.-H.; Smith, K. M. In Porphyrins and Metalloporphyrins; Smith, K. M., Ed.; Elsevier: Amsterdam, 1975; p 798.
(48)Lee, H.; Harvey, R. G. J . Org. Chem. 1983, 48, 749. (49) Ponomarev, G . V.; Shul'ga, A. M. Khim. Geterotsikl. Soedin. (Eng. Trans[.)1984, 19, 389.
Me
\
\
\
Et
'H COZMe
CO@e
12
~
Energy & Fuels, Vol. 4, No. 6, 1990 679
Preparation of Bacteriopetroporphyrins
Table I. Proton NMR Chemical Shifts (a, ppm) and Assignments of the Nickel(I1) Bacteriopetroporphyrinsdin C6Da -IEt.Et1 . ~.(3b) . .In-Pr.Et1 . _ (3c) - . .li-Bu.Et1 . I3d) ref 16 this work ref 16 this work ref 16 this work 9.99 10.00 10.02 10.01 a-meso H 10.03 9.99 -
p-meso H d-meso H
10.03 9.89
;:E } 8-Me 2a-CHz 2b-CH3 7a-CH2 7b-CHz
7-OMe 9XHz 10-CHz 4a-CHz 4b-CHZlCH3 4c-CHs 5a-CH2 5b-CH3
10.06 9.93
3.39-3.40
3.40-3.41
3.9113.94 1.84 4.18 2.96 3.54 3.65 4.82 3.9113.94
1.84
3.9213.94 1.84 4.24 2.98 3.54 3.69 4.87 3.9213.94 1.84
4.00 1.93
4.03 1.94
3b
3a
iqj
0
10
20
30
40
0
B 10
3d 20
30
Retention Volume (mL)
Figure 2. Reversed-phase HPLC traces, at a flow rate of 1.0 mL/min with a Waters Associates Z Module, a 4-pm C-18 cartridge, and a variable-wavelengthdetector set at 405 nm, of (A) natural mixture of methyl 5-ethylbacteriopheophorbidesd using 10% H20/90% methanol and (B) synthetic nickel(I1)bacteriopetroporphyrin methyl esters (3a-d) using 45% H20/55%tetrahydrofuran. Assignments of methyl bacteriopheophorbides d (trace A) are, from left to right, [4-Et,5-Et],[4-n-Pr,5-Et],[4-iBu,8Et], and [4-neoPn,5-Et],respectively. subjected to the same series of transformations. Results and yields were comparable throughout the series except it was shown that treatment of 4e-h with a larger excess of NaBH4 for an extended time gave directly the 2ethyl-9-deoxopheophorbide series in 63 % yield. After oxidation to porphyrin and chelation with nickel as before, the homologous series of porphyrins was obtained and the components were separated by semipreparative HPLC. Milligram quantities of each of the three homologues were obtained in this way. The proton NMR data for the synthetic materials show very good agreement when compared with those obtained for the natural materials obtained by Ocampo et a1.16 (see Table I). Analysis of N M R NOE Spectra. Although Ocampo et al. performed a complete NOE study, we decided that only sufficient NMR work need be completed in order to confirm their assignments. Irradiation of the 10-CH2 multiplet at 4.79 ppm gave NOES at 2.95 (corresponding
10.07 9.91 3.40 3.44 3.40 3.93 1.84 4.21 2.97 3.54 3.66 4.84 3.95 2.34 1.31 4.02 1.93
10.05 9.88 3.40 3.44 3.37 3.92 1.85 4.16 2.95 3.54 3.65 4.79 3.94 2.34 1.32 4.02 1.93
10.07 9.91 3.40 3.45 3.40 3.94 1.85 4.21 2.97 3.54 3.68 4.85 3.87 2.70 1.35 4.03 1.94
.
I
9.99 9.81 3.38 3.45 3.31 3.94 1.86 4.17 2.92 3.54 3.58 4.68 3.86 2.87 1.36 3.97 1.91
to the 7b-CH2 of the propionate group), 4.16 (corresponding to the 7a-CH2of the propionate group), and 3.65 (corresponding to the 9-CH2group of the exocyclic ring). Irradiation of the d-meso proton at 9.88 ppm gave NOEs at 3.37 and 3.40 ppm, peaks corresponding to the 1-Me and 8-Me. The methyl group that does not show an NOE is the 3-Me, the chemical shift of which is 3.44 ppm. Differentiation between the 8-Me and the 1-Me is possible by irradiation of the 7a-CH2a t 4.16 ppm. This gave rise to an NOE at 3.37 ppm that can only correspond to &Me. By elimination, therefore, the peak at 3.40 belongs to the 1-Me. Also observed were NOEs at 2.95 ppm, corresponding to 7b-CH2, and at 4.79 ppm, corresponding to the 10-CH2. Determination of which triplet is assigned to 2b and which to 5b (1.93 and 1.85 ppm) was accomplished by irradiation of the 9-CH2 multiplet at 3.65 ppm. This gave NOEs at 4.79 (corresponding to the 10-CH2)and a t 1.93 ppm corresponding to the 5b-CH3; by elimination, therefore, the peak at 1.85 ppm belongs to the 2b-CH,. The rest of the peaks could then be assigned unequivocally by decoupling except the for the a-and 0-meso peaks at 10.01 and 10.05 ppm. Since there is no methyl group near the 0-meso proton, irradiation of the 3-Me could distinguish between them; it would show an NOE to the a-meso proton. Irradiation of the 3-Me peak at 3.44 ppm showed an NOE to 10.01 ppm, which could only be the a-meso proton. Therefore the resonance at 10.05 ppm is assigned to the P-meso proton. The NOE experiments confirmed the chemical shifts of the CY-, 0-, and 6-meso protons, the 1,3,8 ring methyls, the 7-propionate substituent, the 9- and 10-CH2groups, and the 2b- and 5b-methyls. The 4b-CH2and the 4c-CH3can be assigned by inspection. The only remaining resonances that needed to be characterized were the 2a-, 4a-, and 5a-CH2 groups which overlap each other in the region 3.9-4.0 ppm, and these were distinguished by decoupling experiments. The decoupling experiments also confirmed the assignments of Ocampo et al. Decoupling of the 5bCH, at 1.93 ppm collapsed the quartet at 4.02 ppm (5aCHJ. Decoupling of the 2b-Me at 1.85 ppm collapsed the quartet (partially obscured by the 4a triplet) at 3.92 ppm (2a-CH2). Decoupling the 4b-CH2 multiplet at 2.34 ppm collapsed the triplet at 1.32 ppm (4c-CH3)and the triplet at 3.94 ppm that corresponds to the 4a-CH2. Decoupling the 4c-CH3triplet at 1.32 ppm collapsed the multiplet of the 4b-CH2 at 2.34 ppm. Figure 3 summarizes these NOE connectivities and decoupling data. The proton NMR spectrum is shown in Figure 4.
Smith and Smith
680 Energy & Fuels, Vol. 4, No. 6, 1990 Chart 111
A '
1
Me Me@:;
'Id"
H
-N
Me'"
. 'H
N \
'' 0
R02C
R.'
&O,Me
R5
Figure 3. Proton NMR (300 MHz) NOE and decoupling cond nectivities for nickel(I1)[%n-Pr,5-Et]-bacteriopetroporphyrin methyl ester (312). Arrows with no asterisk indicate NOE only; arrows with one asterisk indicate decoupling only; arrows with two asterisks indicate NOE and decoupling.
15 R = Famesyl, M = Mg (Bacteriochlorophylls-c)
16 R = Me, M = 2H (Methyl Bacteriopheophorbides-c: Bmph-c)
Chart IV
Me
El
(a) R4 = Et
(b) R4 = n-Pr
10
8
(c) R4 = i-Bu
17 R = Famesyl, M = Mg (Bacteriochlorophyll\-e) I
"
'
6
l
"
' 4
I
18 R = Me, M = 2H (Methyl Bactenopheophorbidew
"
2
PPm
Figure 4. Proton NMR spectrum (300 MHz in C&) of nickel(I1) [4-n-Pr,5-Et]-bacteriopetroporphyrin d methyl ester (3c).
Preparation of "Bacteriopetroporphyrinsc and e *. Having thus further verified the structures of the petroporphyrins isolated by Ocampo et a1.,16 we decided to convert the bacteriochlorophylls c (15) (Chart 111) and e (17) (Chart IV) into their corresponding nickel(I1) bacteriopetroporphyrins. Although petroporphyrins with a methyl group a t the 6-meso position have not yet been isolated, there is no reason why they should not be at some point in the future. If and when these compounds are eventually found in petroleum, their recognition will be facilitated by the availability of synthetic samples. These two Chlorobium chlorophyll series were also readily available from existing bacteria cultures maintained in our laboratory, so they presented a convenient starting point for preparation of the target bacterioporphyrins. Bacteriochlorophyll c (15), the main chlorophyll component of the green sulfur bacteria Prosthecochloris aestuarii, was first reported in 1953 and characterized by H ~ l int 1961. ~ ~ The &methyl group was identified by extensive synthetic s t ~ d i e and s ~spectroscopic ~ ~ ~ ~ ~stud(50)Morley, H.V.;Holt, A. S. Can. J . Chem. 1961, 39, 755.
Bmph-e)
ies.24p53*" The complete structures were finally unequivocally determined (including stereochemistries at position 2) for these compounds by Smith et al.55 Like the bacteriochlorophylls d, the bacteriochlorophylls c (15) and consequently the Bmph-c (16) occur as homologous mixtures of compounds in which the substituents a t position 4 can be ethyl, n-propyl, or isobutyl, and at position 5 can be methyl or ethyl. Bacterial cultures of P. aestuarii were grown in 20-L batches according to procedures already described in the l i t e r a t ~ r e .Harvesting ~~ of the cells was done by a simple organic solvent extraction of the media with acetonef ether. This was more convenient than other methods such as centrifugation or filtration of the bacterial cells on Celite followed by organic solvent extraction of the collected cells. Not only was this method faster, but more of the chlorophyll was actually extracted rather than left in the su(51) Holt, A. S.; Hughes, D. W.; Kende, H. J.; Purdie, J. W. J . Plant Cell Physiol. (Tokyo) 1963,41, 49. (52)Kenner, G. W.; Rimmer, J.; Smith, K. M.; Unsworth, J. F. Philos. Trans. R. SOC.London, B 1976,273, 367. (53)Holt, A. S.;Purdie, J. W.; Wasley, J. W. F. Can. J . Chem. 1966, 44, 88.
(54) Smith, K. M.; Unsworth, J. F. Tetrahedron 1975, 31, 367. (55) Smith, K. M.; Craig, G. W.; Kehres, L. A.; Pfennig, N. J . Chromatogr. 1983,281, 209.
Preparation of Bacteriopetroporphyrins
Energy & Fuels, Vol. 4, No. 6, 1990 681
Meg:: Scheme V
Chart V. Bacteriopetroporphyrins c Resulting from Diagenesis of Bacteriochlorophylls c (15) and e (17)
OH
';""
H+Me
MeH
-N
Me"
.
NaBH4 TFA
'H
Me
''
0
20
+
l6
Me0&
C02CH3
22 M
e
m
R
[ol
Me M
*R5e
4
\
\
M Me
e
m
W
R'
R'
\ U
I
Me02C
19
pernatant liquid or filtrate. Evaporation of the extract was foliowed by transesterification and demetalation with sulfuric acid/methanol. Removal of the carotenoids and other unwanted materials by column chromatography on alumina gave the Bmph-c. Since the NaBH,/TFA method for deoxygenation worked so well for the Bmph-d series, it was decided to follow the same strategy for the Bmph-c series. The reactions were performed on the entire homologous mixture, with the ultimate goal of separating the products by HPLC a t the end of the reaction sequence. Thus, treatment of the homologous mixture of Bmph-c (16) with an excess of NaBH, in TFA provided the 9-deoxo-2-ethyl mixture 19 in good yield (Scheme V). Quite often, the NMR spectra of the product showed small amounts of the 9-deoxo-2vinyl compound 20 to be present, so the crude material after workup was hydrogenated over 10% palladized charcoal for several hours. Removal of the catalyst by filtration on Celite and purification on an alumina column gave 19 in 73% yield. The next step in the reaction sequence was oxidation of 19 into the corresponding porphyrin 21. Whereas in the case of the deoxygenated Bmph-d, the oxidation to porphyrin with DDQ could be done virtually by titration from a buret, in this case it was considerably more difficult. Treatment of 19 with DDQ was initially done exactly the same way as with the deoxy-Bmph-d compounds. By dissolving the chlorin in CH2C12and adding a solution of DDQ in benzene dropwise, it was hoped the oxidation should give the desired porphyrin. Instead, TLC monitoring showed the reaction quickly produced large amounts of chromatographically polar material and there was as much starting chlorin as porphyrin. It was thought most likely that DDQ was over-oxidizing the chlorin; after production of the porphyrin, it was in turn oxidized to the corresponding cation radical. Attempts to quench the proposed cation radical with NaBH, were unsuccessful. Using less than 1 equiv of DDQ gave the same results. The reaction products were separated by alumina chromatography. The starting chlorin and the desired porphyrin ran very close together and in many instances they were not completely separated, even through the use of a very long alumina column and eluting with 25% cyclohexane/75% CH2ClP.Use of a silica gel column or preparative silica gel TLC plates was avoided for fear of causing the same benzylic oxidation at position 10 that was previously observed. Since it appeared that over-oxidation was taking place, it was decided to try a weaker oxidizing agent. Treatment
of 19 with 1,Cbenzoquinone under a variety of conditions did not oxidize the chlorin at all and quantitative recovery of starting material was obtained. Intermediate in oxidizing strength between DDQ and 1,4-benzoquinone is p-chloranil. This, too, was tried without much success on the free base chlorin; mostly starting material was recovered even though the reaction mixture was refluxed overnight in CHC13. I t is knownMl5' that the oxidation potential of porphyrins and chlorins depends on the metal chelated within. The presence of zinc(I1) imparts a higher electron density onto the periphery of the macrocycle, and this facilitates oxidation. Therefore, oxidation with p chloranil was attempted on the zinc(I1) complex of 19. This succeeded but the reaction was stopped before all the starting material was consumed because TLC indicated that a large quantity of chromatographicallypolar material was being formed. After demetalation of the zinc(I1) complex with TFA, the best yield of the free base porphyrin among several attempts was 1590, along with 27% of unreacted starting material. This was unfortunately about the same yield as obtained by treating the free base with DDQ, so there was still a disadvantage here in having to insert and then remove the chelated zinc. DDQ oxidation of nickel(I1) complexes gave mostly decomposition products. When 21 (obtained from the DDQ reaction) was itself treated with DDQ, the same polar compounds were obtained. It seemed, therefore, that the oxidation to porphyrin occurs quickly, but the resulting porphyrin can be oxidized almost as readily as the starting chlorin. The relative amounts of starting material and desired porphyrin were always about the same. Since, on the basis of numerous other attempts to control the oxidation, there seemed to be no better alternative to using DDQ on the free base chlorin, further preparations of the porphyrin were carried out in that way. After sufficient quantity of porphyrin was collected, nickel(I1) chelation was accomplished by refluxing 21 in chloroform containing saturated Ni(OAc)2/methanolto give the homologous mixture 22 (Chart V). The HPLC trace is shown in Figure 5. Separation of the homologues was performed by reversed-phase semipreparative HPLC, which gave baseline separation. Because of the relatively small amount of the [i-Bu,Et] homologue, it was necessary (56) Fuhrhop, J.-H. J . Am. Chem. SOC.1973,95, 5140. (57) Fuhrhop, J.-H. In Porphyrins and Metalloporphyrins; Smith, K. M., Ed.; Elsevier, Amsterdam, 1975; p 629.
682 Energy & Fuels, Vol. 4, No. 6, 1990 22b
Smith and Smith Table 11. Proton NMR Chemical Shifts (6, ppm) and Assignments of the Nickel(I1) Bacterioporphyrins c in CaDa" - [Et,Et] [n-Pr,Et] [i-Bu,Et] [Et,Me] (228) (22b) (22c) (22d) 1-Me 3.12 (s) 3.12 (s) 3.12 (9) 3.13 (9) 2a-CH2 3.72 (4) 3.72 (q) 3.72 (q) 3.73 (4) 2b-CH3 1.66 (t) 1.67 (t) 1.66 (t) 1.66 (t) 3-Me 3.20 (s) 3.19 (9) 3.23 (9) 3.24 (9) 4a-CH2 3.72 (4) 3.72 (4) 3.73 (t) 3.66 (d) 4b-CH2/CH3 1.67 (t) 1.67 (4) 2.17 (m) 2.56 (m) 4c-CH3 1.16 (t) 1.19 (d) 5a-CH2/CH3 3.22 (s) 3.80 (q) 3.81 (4) 3.84 (q) 5b-CH3 1.77 (t) 1.78 (t) 1.79 (t) 7a-CHz 4.09 (t) 4.08 (t) 4.09 (t) 4.12 (t) 7b-CH2 2.81 (t) 2.81 (t) 2.81 (t) 2.83 (t) 7-OMe 3.41 (s) 3.41 (s) 3.41 (s) 3.41 (s) 8-Me 3.15 (9) 3.14 (6) 3.14 (s) 3.16 (9) &Me 3.51 (9) 3.49 (8) 3.50 (9) 3.52 (s) 9-CH2 3.45 (m) 3.53 (m) 3.54 (m) 3.55 (m) 10-CH2 4.71 (m) 4.70 (m) 4.71 (m) 4.74 (m) a-meso 9.57 (9) 9.56 (s) 9.58 (s) 9.60 (s) P-meso 9.61 (s) 9.63 (s) 9.65 (s) 9.66 (s) "s = singlet; q = quartet; t = triplet; m = multiplet.
I
1
0
10
I
20
I
30
40
Retention Volume (mL) Figure 5. Reversed-phase HPLC traces, at a flow rate of 1.0 mL/min with a Waters Associates RCM 8 X 10 Module, a 4-pm (2-18 cartridge,and a variable-wavelengthdetector set at 405 nm, of nickel(I1) bacteriopetroporphyrin c methyl ester homologues (22a-d) using 45% H20/55%tetrahydrofuran. to do numerous injections in order to obtain a significant quantity of this compound. Analysis of NMR NOE Spectra. As with the porphyrins obtained from the bacteriochlorophyll d series, it was necessary to record NOE difference spectra to establish the peak assignments (particularly for the a- and @-mesopeaks and the methyl peaks). It is most convenient to begin analysis of the NOE data by examining the data for compounds other than the [Et,Me] homologue 22a; the a- and P-meso protons in the [Et,Me] homologue both have neighboring methyl and ethyl groups (the ethyls being completely indistinguishable since they have the same chemical shift), so the observed NOEs do not give very much information on their own. For that reason some of the signals for these compounds (particularly [Et,Me]) have been assigned by analogy with the rest of the homologues. Thus we decided to begin the analysis with the [i-Bu,Et] homologue 22d and proceed with the other three in order of descending molecular weight. The NMR peak assignments for these compounds are listed in Table 11. [i-Bu,Et] Homologue (22d). Irradiation of the 4c-CH3 doublet at 1.19 ppm gave an NOE to the 4b-CHzmultiplet at 2.56 ppm. The methyl signal at 3.41 ppm gave no NOE anywhere, making its assignment the 7-OMe. Irradiation of the methyl singlet a t 3.52 ppm gave NOEs at both 3.13 and 3.16 ppm. Conversely, independent irradiation of the 3.13 and 3.16 ppm singlets both gave an NOE to the 3.52 ppm singlet. Therefore, the peak at 3.52 ppm is the 6methyl and those a t 3.13 and 3.16 ppm are the 1- and 3-methyls. Distinguishing between these two methyls, however, had to be accomplished in the next experiment. Irradiation of the 7a-CH, at 4.12 ppm gave NOEs to the multiplet at 4.74 ppm (10-CH,), the triplet a t 2.83 ppm (7b-CHz),and the methyl singlet at 3.16 ppm. Therefore, the 3.16 ppm resonance is the 8-Me, and by elimination, that a t 3.13 ppm is the 1-Me. Irradiation of the signal at 2.83 ppm (7b-CHz)gave NOEs to 4.12 (7a-CH2)and 4.74
ppm (10-CH,). The 10-CH, multiplet a t 4.74 ppm gave NO& to 4.12 (7a-CH2),2.83 (7b-CHz),and 3.55 ppm. The 3.55 ppm signal is the 9-CHz group; it is partially overlapped with the b-methyl singlet at 3.52 ppm and so when the 3.52 ppm line (b-methyl) was irradiated that a t 3.55 ppm (9-CH2)was also irradiated (unintentionally) giving an NOE to the 10-CH, group. When the meso proton signal at 9.60 ppm was irradiated, it gave rise to NOEs a t 3.73 ppm and the methyl group at 3.24 ppm. Conversely, irradiation of the 3.24 ppm peak gave an NOE a t 9.60 ppm. Since the P-meso proton does not have a neighboring methyl group, it cannot give this observed NOE. Therefore the 9.60 ppm line is the a-meso proton, and the 3.24 ppm methyl group has to be the 3-Me. Also, the 2a quartet can be assigned to 3.73 ppm. By elimination, the @-protonis a t 9.66 ppm. Irradiation of the P-meso proton at 9.66 ppm gave NOESat 3.66 and 3.84 ppm (4a- and 5a-CH2s). The position of the 4a-CH, quartet was established by simply decoupling the 4b multiplet a t 2.56 ppm and noticing the collapse of the pattern at 3.66 ppm. Since the 3.66 ppm resonance is the 4a-CH,, that a t 3.84 ppm must be the 5a-CH,. Irradiation of the triplets at 1.66 and 1.79 ppm gave rise to NOEs a t 3.73 and 3.84 ppm, respectively. Decoupling of the two triplets also caused the collapse of the same signals, and the reverse decoupling experiments were also consistent with the observed NOEs. Therefore, the 1.66 ppm resonance is the 2b-Me and that a t 1.79 ppm is the 5b-Me. Figure 6 shows the NOE and decoupling connectivities established by these experiments. [n-Pr,Et] Homologue (22c). The proton NMR spectrum of 22c is shown in Figure 7. The only difference between this homologue and 22d is that the substituent at position 4 is n-propyl instead of isobutyl. The NOE experiments were done very similarly to those above. Irradiation of the methyl singlet at 3.41 ppm gave no NOE anywhere, indicating it to be the 7-OMe. Irradiation of the meso peaks provided the same information as did the previous homologue. Again, since CY and @ have differing neighboring groups ( a has an ethyl and a methyl; /3 has an n-propyl and an ethyl), they are easily distinguishable by NOE difference. Irradiation of the meso proton at 9.58 ppm gave an NOE to the methyl singlet at 3.23 ppm and to the quartet at 3.72 ppm, therefore assigning the line at 9.58 ppm as the a-meso proton, that a t 3.23 ppm as the 3-Me, and the 3.72 ppm resonance as the 2a-CHz. (An
Energy & Fuels, Vol. 4 , No. 6,1990 683
Preparation of Bacteriopetroporphyrins
..
n
C02Me
Figure 6. Proton NMR spectrum (300 MHz) NOE and decoupling connectivities for nickel(I1) [4-n-Pr,S-Et]-bacteriopetroporphyrin c methyl ester (2212). Arrows with no asterisk indicate NOE only; arrows with one asterisk indicate decoupling only; arrows with two asterisks indicate NOE and decoupling.
I( 10
8
4
6
2
PPm
Figure 7. Proton NMFt nickel(I1) n . spectrum (300MHz in C a s )of/an.\ I 1 . ri t 1+n-rr,a-ac]-DacLeriopetroporpnyrin c memyi ewer I.
r m . 3
I
1
I
[LAC).
NOE is also observed at 9.58 ppm when the peak a t 3.23 ppm is irradiated.) Irradiation of the @-mesoproton at 9.65 ppm gave NOEs to the triplet a t 3.73 ppm (4a-CHJ and to the quartet a t 3.81 ppm (5a-CH2). Irradiation of the most downfield methyl singlet (3.50 ppm) gave NOEs to the 2-methyl peaks at 3.12 and 3.14 ppm. Irradiation of these latter two peaks gave NOEs to the 3.50 ppm singlet. Therefore, the line a t 3.50 ppm is the &methyl and the latter two correspond to the 1- and 8-methyls. A small NOE was observed at 3.72 ppm when the 3.12 ppm methyl group was irradiated, indicating the 3.12 ppm peak to be 1-Me, since 8-Me cannot give an NOE to a CH, group. This is not an unexpected observation, but the enhancement may be too small to be significant; one would expect that irradiation of the 1-Me could produce an NOE to the 2a-CH2. Irradiation of the other methyl in question did not show the same small NOE, providing further evidence. Also, the assignment of the 1-Me as the peak a t 3.12 ppm and the 8-Me as that at 3.14 ppm is completely analogous to the assignments for the corresponding methyl groups in the [i-Bu,Et] homologue. In fact, all of the assignments made thus far for this compound are not unexpected; they
are all completely analogous with the assignments made for the [i-Bu,Et] homologue. No crossing over of peaks was observed and the chemical shifts changed only very slightly when moving from the previous homologue to this one. The rest of the assignments, then, were made by simple decoupling experiments and analogy to the previous homologue. Decoupling the 4a-CH, (whose position was established above by NOE) collapsed the multiplet at 2.17 ppm. Therefore the 2.17 ppm resonance is the 4b-CH2. Decoupling the upfield triplet at 1.16 ppm collapsed the multiplet at 2.17 ppm to a triplet, thus proving the position of the 4c-CH3. Decoupling the triplet at 1.78 ppm collapsed the quartet at 3.81 ppm. Since the peak a t 3.81 ppm was shown (by NOE above) to be the 5a quartet, that at 1.78 ppm must belong to the 5b-Me. The triplet corresponding to 4a-CH, a t 3.73 ppm is partially obscured under the 7-OMe and &methyl peaks, and it disappears when the multiplet at 2.17 ppm is decoupled. [Et,Et] Homologue (22b). The only difference here is again at the 4-position. However, this only serves to simplify the spectrum. NOE and decoupling experiments were performed exactly as for the previous homologue, and there were no unexpected observations. There is again no crossing over of peaks. Although the chemical shifts change very slightly, the relative positions of the methyl and meso peaks does not. [Et,Me] Homologue (22a). Distinguishing between some of the signals is difficult for this homologue, even with the use of NOE difference data. Attempts to distinguish the meso protons, for example, by irradiating them and comparing the NOEs to the neighboring groups is uneffective because they give almost the same NOE difference spectra. Both meso protons neighbor a methyl group and an ethyl group. Since the ethyl signals fall on top of each other, they cannot be used to assign the neighboring meso protons. The NOEs to the methyls are different, but unfortunately without a definite assignment of the 3-Me and/or the &Me, they too, are of little value. The only other way to distinguish these methyl groups would be to establish a connectivity between the 9-CHz group and the 5-Me. Although the methyl peaks are well resolved, thus making the experiment easy to perform, the expected connectivity between the 9-CH2and the 5-methyl was not observed. There is, however, another way the 5-methyl peak could be tentatively assigned. Due to less than complete separation of the homologues by HPLC there is a little [Et,Et] homologue present as an impurity. This makes the signal corresponding to the 5-Me of the [Et,Me] homologue integrate to less than three protons relative to the other methyl peaks. This smaller peak appears a t 3.22 ppm. Further proof of this tentative assignment is that this shorter peak at this position relative to the other methyl peaks is not observed for the other homologues which have an ethyl substituent a t position 5 instead of a methyl group. If the 5-Me is a t 3.22 ppm, then the 3-Me is at 3.20 ppm, which agrees very well with the observed chemical shift of the 3-Me in the other homologues. Since the 3- and 5-methyl groups are now distinguished on the basis of the preceding argument, the CY- and p-meso protons and the methyl groups can be distinguished in exactly the same way as described above for the other homologues. The remaining signals are easily assigned by NOEs, decoupling, and analogy. Two trends were apparent during this analysis which helped in making the assignments of the peaks in this series of proton NMR spectra. (i) In each of the homo-
Smith and Smith
684 Energy & Fuels, Vol. 4, No. 6, 1990 Scheme VI HfMe
YHO
Me
M#
P H Me02C
Me0,C
\
I
eft‘
19
NaBH4
THF
$HdTFA
Me@ Me
H
-N
\:’
Et
Me’ ‘H Me02C
23
R4
10
NaBH4
cat H 2
Me
19
mum growth of bacteria. The harvesting of the bacteriochlorophylls e, conversion to the corresponding Bmph-e, and isolation of the homologous mixture of Bmph-e procedures were exactly the same as were done for the Bmph-c. The amount of Bmph-e mixture obtained from a 20-L carboy ranged from 184 to 500 mg (i.e., 9.2-25 mg/L of culture). Several attempts to treat the Bmph-e (18) with NaBH, in TFA under the same conditions as described earlier gave a minute amount of a compound that coeluted on TLC with an authentic sample of 19, along with a large amount of polar baseline material. Since the only difference between Bmph-c and Bmph-e is the 3-formyl group, it was thought possible that reduction of Bmph-e to the 3hydroxymethyl compound followed by treatment with NaBH,/TFA might give better results. B r o ~ k m a n n ~ ~ found that selective reduction of the 3-formyl group of Bmph-e was possible using NaBH, in wet THF. Thus, treatment of 18 with NaBH, in 10% H20/90% THF at 0 “C for 15 min gave the 3-hydroxymethyl compound 23 (Scheme VI) in an isolated yield of 57 5%. However, when this dihydroxy compound was treated with NaBH4/TFA, it behaved in the same unsatisfactory way as the 3-formyl compound itself. The failure of this reaction should not be a total surprise, because Gribble et al.& found little or no reduction when benzyl alcohol was submitted to these reaction conditions. Therefore, we investigated other methods. Reduction of the aldehyde and ketone functions with NaBH, proceeded smoothly to give the Bmph-e triol 24. Me
Me02C’
@ / ’\ \
Rd
24
logues the P-meso proton is downfield relative to the CYmeso proton. (ii) The relative positions of the methyl peaks (from left to right in the spectra) is 6, 7-OMe, (5) 3, 8, 1. Preparation of “Bacteriopetroporphyrinse “. The bacteriochlorophyll c (15) and e (17) series have the same carbon skeleton except that bacteriochlorophyll e has no 5-methyl homologues; thus, the petroporphyrins that would be expected to be produced from both under geologic conditions should be the same. We therefore undertook to prepare the 22 homologous series from bacteriochlorophyll e. It was thought best to take a synthetic route that would intersect the path taken for the bacteriochlorophyll c as early as possible, making it unnecessary to carry out steps all the way to the porphyrin. It was therefore hoped that using the NaBH,/TFA methodology developed above would provide the deoxo compounds 19 in one step, and the rest of the procedure would be the same and therefore not necessary to carry out. As will be described, the NaBHJTFA methodology did not work satisfactorily so another route had to be devised. The bacteriochlorophylls e (17) were initially characterized by B r o ~ k m a n n . ~ Simpson ~’ and Smith determined the correct stereochemistries of the 2-(l-hydroxyethyl) group for the homologues.20 Growth of the bacteriochlorophyll e producing bacteria (Chlorobium pheouibroides) was performed according to the literature procedure.20 About 30 days was typically allowed for maxi(58) Brockmann, H.; Gloe, A,; Risch, N.; Trowitzsch, W. Liebigs Ann. Chem. 1976,566. (59) Brockmann, H. Philos. Trans. R. SOC.London, B 1976,273, 277. (60)Risch, N.; Brockmann, H. Liebigs Ann. Chem. 1976, 578. (61)Risch, N.; Kemmer, T.; Brockmann, H. Liebigs Ann. Chem. 1978, 585.
Me H
Me”’
/
‘M’”
-N ,~
,‘
\N
‘ \,
Et
OH Me02C
24 M = 2 H
25 M = Z n
Attempts to remove the three benzylic hydroxy groups simultaneously by catalytic hydrogenolysis in THF failed. Catalytic hydrogenolysis in the presence of acetic acid or formic acid32provided minor amounts of a material that coeluted on TLC with an authentic sample of 19 prepared earlier, but the main result was decomposition to polar material. Isolation of the TLC-mobile material was possible, even though it was in very small amount, and the NMR spectra confirmed the identity as 19. However, due to very low yields, a better method was sought. Recently, Lau et al.62reported reductive deoxygenations of aryl aldehydes and ketones and allylic, benzylic and tertiary alcohols using Zn12/NaCNBH3 in 1,2-dichloroethane, In comparison with the NaBH,/TFA method, Lau et a1.62 found benzophenone was reduced to diphenylmethane in 95% yield. This method seemed well suited for our purposes since 18 contains aryl aldehyde, aryl ketone, and benzylic alcohol functions. Teatment of 18 under the Lau et a1.62conditions (1.5 equiv of Zn12,7.5 equiv of NaCNBH3, in refluxing 1,2-dichloroethane) provided only a mixture of several minor unidentified compounds and, again, only a small amount of the desired material. It was then thought possible that reduction of both the 3-formyl (62) Lau, C. K.; Dufresne, C.; Belanger, P. C.; Pietre, S.; Scheigetz, J.
J. Org. Chem. 1986, 51, 3038.
Energy & Fuels, Vol. 4, No. 6, 1990 685
Preparation of Bacteriopetroporphyrins Scheme VI1
>
