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Synthesis 2023; 55(15): 2273-2284
DOI: 10.1055/a-2004-1228
short review
Special Issue dedicated to Prof. David A. Evans

Chemistry and Biology of Acyloin Natural Products

Mark Rizzacasa  ∗
,
Michael Ricca 

Australian Research Council
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Abstract

This review details the isolation, biosynthesis, biological activity, and synthesis of α-hydroxy ketone (acyloin) natural products. The role of these compounds as biosynthetic precursors to complex natural products and synthetic strategies to access the sensitive acyloin moiety and stereochemistry are highlighted.

1 Introduction

1.1 Xenocyloins

1.2 Biological Activity

1.3 Biosynthesis

1.4 Acyloins as Discoipyrrole Biosynthetic Precursors

2 Total Synthesis of Acyloin Natural Products

2.1 Kurasoins A and B

2.2 Soraphinol A and Circumcin B

2.3 4-Hydroxysattabacin and Actinopolymorphol A

2.4 Actinopolymorphol B

2.5 Sattazolins and Sattabacins

2.6 Catalyst Development and Application in Acyloin Synthesis

2.7 Xenocyloins

3 Conclusion


#

Key words

acyloin - natural products - total synthesis - biosynthesis - biological activity
1

Introduction

The acyloin natural products are a family of compounds that feature an α-hydroxy ketone moiety with a benzylic aryl group on one side and a simple alkyl group on the other, with many of these compounds isolated as pairs of tautomers. Despite their apparent simplicity, the acyloins have been isolated from a wide range of fungi and bacteria. Furthermore, they display varied and often unusual biological activities and chemical properties. For the synthesis of these compounds, the sensitive α-hydroxy ketone moiety and reactive substituents such as indoles and phenols require a careful synthetic strategy. This review details the isolation, biological evaluation, and the biosynthetic and chemical synthesis of a series of acyloin compounds.

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Mark Rizzacasa(left) obtained a Ph.D. in 1990 under the guidance of Professor Melvyn Sargent at the University of Western Australia. He then moved on to the USA and spent 2 years as a postdoctoral fellow in Professor Robert E. Ireland’s research group at the University of Virginia. In early 1993, he then returned to Australia to begin an appointment in the School of Chemistry at the University of Melbourne and was promoted to full Professor in 2008. Mark was awarded the RACI Rennie Memorial Medal in 1997, the 2005 Organic Reactions Lectureship, and the prestigious RACI A. J. Birch medal in 2010. His interests include the total synthesis of bioactive natural products and the development of synthetic methodologies. Michael Ricca (right) completed a B.Sc. and M.Sc. at The University of Melbourne. His masters research project was conducted in Professor Mark Rizzacasa’s lab, focusing on the synthesis of bioactive natural products. He began a Ph.D. in 2019 in the same lab, with research interests including total synthesis and the development of metal complexes for hydrogen atom transfer reactions.

Soraphinols A (1), B (3), and C (4)[1] [2] [3] are a family of acyloins isolated from the myxobacterium Sorangium cellulosum by Ahn and co-workers (Figure [1]). Soraphinol A (1) contains an indole moiety at C4, while soraphinol B (3) features a phenyl ring at C4. The soraphinols are structurally related to kurasoin B (2)[4] which is a deshydroxy derivative of soraphinol A (1). Despite their structural similarities, the kurasoins were isolated from the non-myxobacterial fungus Paecilomyces sp strain FO-3684.[4] Soraphinol C (4)[3] was isolated from S. cellulosum along with its tautomer, 4-hydroxysattabacin (7), which has also been isolated from Bacillus sp.[5] 4-Hydroxysattabacin (7) is thus both a fungal and myxobacterial metabolite. Acetylation of the α-hydroxy moiety gives the natural product actinopolymorphol A (8)[6] [7] which was isolated from the bacterium Actinopolymorpha rutilus. Actinopolymorphol B (9) was also isolated with 8 and is an indole-containing acyloin with a terminal methyl group. The original isolation of (S)-4-hydroxysattabacin (7a) from Bacillus sp. yielded the related compounds (S)-sattabacin (5a), the indolic sattazolin A (14), and its methylated derivative methylsattazolin (15).[5]

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Figure 1 Acyloin natural products

Further acyloins of the sattazolin class have been isolated by Hertweck and co-workers[8] from Clostridium beijerinckii. Sattazolin B (16) is the tautomer of 14,[5] while dehydrosattazolin (13) features a further degree of unsaturation and exists as the enol tautomer. Computational studies suggested this is likely present as the more stable Z isomer.[8] Compound 19 is the anthranilic acid ester of sattazolin A (14). When the 7-position of the indole ring is oxidised to a phenol, hydroxysattazolin (17) is obtained.

Circumcins B (11) and C (12) were isolated from a bacterium Gordonia sp. found in the venom duct of the cone snail Conus circumcisus.[9] These compounds are known and have been synthesised before being identified as natural products.[10] [11] Furthermore, kurasoins A (18) and B (2), soraphinols A (1) and C (4), and 4-hydroxysattabacin (7) and related compounds were isolated from the same bacterium.[9]

The stereochemistry of the acyloins has been assigned primarily through synthesis and comparison to other known compounds. Soraphinols A (1)[1] and B (3)[2] were tentatively assigned as having the same S configuration as kurasoin B (2) based on a comparison to the literature.[4] The configurations of soraphinols A (1) and B (3) were eventually confirmed by synthesis.[11] [12] Similarly, the absolute configuration of 2 has been confirmed by synthesis.[13] The stereochemistry of (S)-sattabacin (5a),[14] 4-hydroxysattabacin (7),[6] [14] actinopolymorphol A (8),[6] and sattazolin A (14)[15] have all been assigned by total synthesis. The stereochemistry of soraphinol C (4) and sattazolin B (16) is unknown, as is the configuration of methylsattazolin (15), sattazolin anthranilic acid ester (19), and hydroxysattazolin (17), although these are likely to be the same as sattazolin A (14).

Both the (R) and (S)-enantiomers of certain acyloins occur naturally. The initial isolate of sattabacin (5a) from Bacillus sp. was assigned as S.[5] The R-configured natural product 5b was isolated from the thermophilic bacterium Thermosporothrix hazakensis.[16] A further natural product, hazakacin (6),[16] featuring a terminal ethyl rather than isobutyl chain, was isolated alongside (R)-sattabacin (5b) and was also assigned as the R configuration based on the similarity in specific rotation to 5b. Igarashi[17] has also isolated (R)-sattabacin (5b), hazakacin (6), and an oxidised derivative 2′-oxosattabacin (10), containing a Z-configured enol (4:1 enol/keto, CDCl3) from Thermosporothrix hazakensis. 2′-Oxosattabacin (10) has been previously described as a synthetic intermediate.[18]

4-Hydroxysattabacin (7) may also be a similar case; the specific rotation for natural (Bacillus sp. extract)[5] and synthetic[6] [14] materials were both similar, however when 7 was isolated from S. cellulosum the specific rotation was negative ([α]D –3.5 (c 0.5, MeOH)).[3] The magnitude was also smaller, however the solvent used for the measurement was different to the original isolation, which indeed casts doubt on the validity of the value. Lim and co-workers[19] have also isolated (R)-sattabacin (5b) and (R)-4-hydroxysattabacin (7b) from Bacillus sp. (strain SCO-147). In this case, the specific rotations were closer in magnitude to the original isolation.[5] [19] 16S rRNA analysis indicated the isolated strain could be Bacillus gibsonii (99.8% rRNA concordance).

1.1

Xenocyloins

Xenorhabdus spp. are remarkable Gram-negative bacteria that reside as a symbiont in the gut of nematodes. When a nematode infects an insect, the bacteria kill the host and allow the nematode to complete its life cycle. Interestingly, in the absence of bacteria the nematode can kill the host but cannot complete the life cycle.[20] Furthermore, it is believed that antimicrobial compounds produced by the bacteria are crucial in those processes, as they stop the dead insect from being attacked by other bacteria. Examples of xenocyloins are shown in Figure [2]. Nealson,[21] Webster,[22] Bode,[23] and Yu,[24] [25] have isolated a series of indolic acyloins from Xenorhabdus bovienii; xenocyloins A–J (20–29) which contain an α methyl group substitution distinguishing them from the sattazolins, except for xenocyloin I (28) which is an N-methyl derivative of sattazolin A (14). Xenocyloin A (20) contains an α isopropyl group; that is the chain is one methylene group shorter than sattazolin A (14). Acetylation of the hydroxy group gives xenocyloin C (21). Xenocyloin B (22) features a sec-butyl group, and also has an acetylated congener xenocyloin D (23). Xenocyloin E (24) is an unusual case, in which the hydroxy group is propionylated, likewise, xenocyloin F (25) is a propionylated congener of xenocyloin A (20). Xenocyloin G (26), H (27), and J (29) are N-methyl derivatives of xenocyloins A (20), C (21), and B (22), respectively. The stereochemistry of the xenocyloins have been primarily assigned by inference from chiroptical data. Huang[26] and co-workers have isolated xenocyloins B (22), C (21), and D (23) from a different bacterium Streptomyces sp. CB09001.

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Figure 2 Xenocyloin natural products

# 1.2

Biological Activity

The biological activity of the acyloins is varied, and these compounds display an impressive array of activities. Kurasoins A (18) and B (2) showed activity against protein farnesyltransferase (PFTase) with IC50 59.0 μM for kurasoin A (18) and 58.7 μM for kurasoin B (2).[4] Inhibition of PFTase is implicated in cancer prevention, although this is likely quite limited.[27]

Schmidt and co-workers[9] have reported that kurasoin B (2) and soraphinol A (1) are active in a human norepinephrine transport assay, with K 1 values of 2575 nM and 275 nM, respectively. Estrogen receptors (ERs) exist in two forms ERα and ERβ with opposing effects and are implicated in breast cancer growth.[6] Whilst ERα promotes cell growth in response to 17β-estradiol, ERβ decreases growth in response to the same ligand. When ERα and ERβ are co-expressed, the proliferation of cancer cells is reduced. Furthermore, the receptors are known to heterodimerise and ERα/β heterodimers are implicated in cancer prevention. Shen and co-workers have shown that actinopolymorphol A (8) induces ERα/β heterodimerisation (EC50 = 19 μM). Shen postulates that the phenol moiety is likely involved in the mechanism of action, possibly by formation of H-bonded networks.[6] Actinopolymorphol B (9) was not active in this assay.

Some activities are, however, puzzling. Despite the structural similarities between soraphinols A (1) and B (3)[1] [2] and the kurasoins,[4] the former did not show any activity against PFTase up to 100 μg/mL. Similarly, soraphinol C (4) was found to be a potent antioxidant with a similar value to Trolox (0.956 ORAC), a water-soluble vitamin E derivative, whereas the isomer 4-hydroxysattabacin (7) was significantly less potent (0.617 ORAC).[3]

Satta and co-workers[5] studied the activity of sattabacins and sattazolins for cytotoxicity and antiviral activity (Table [1]). The free acyloin hydroxy group appears to be important for activity, as its methylation (as in compound 15) leads to a significant loss of activity in all assays. 4-Hydroxysattabacin (7a) was the most cytotoxic compound and displayed the greatest antiviral activity, with the free phenol contributing significantly to the activity. When the phenol was absent, as in sattabacin (5a), the activity was reduced across all studies. The indole moiety was intermediate between the phenyl and phenol in both cytotoxicity and antiviral activity. In a separate study, sattabacin (5a) displayed antiviral activity against Varicella zoster (VZV) with an IC50 of 58 μM.[28] (R)-Sattabacin (5b) and hazakacin (6) did not display activity against Candida albicans or Micrococcus luteus, however they were slightly cytotoxic to human T lymphoma (Jurkat) cells.[16]

Hertweck and co-workers[8] studied the antimicrobial and antiproliferative activities of the acyloins isolated from Clostridium beijerinckii, and the results are listed in Table [2] and Table [3]. Moderate cytotoxicity in the ~50 μM range, in HUVEC, K-562, and HeLa cell lines was observed. Sattazolin anthranilic acid ester (19) showed the highest potency in the low μM range. This result contrasts with its antimicrobial activity, where 19 showed no inhibitory activity in any assay except against P. notatum, where it was the least active of the tested compounds. The sattazolins induced an inhibition zone of up to 22 mm against various microbes summarised in Table [3]. None of the derivatives were active against E. coli or P. aeruginosa, only sattazolin B (16) was active against C. albicans (14 mm inhibition zone).

Table 1 Biological Activity of Acyloins Reported by Satta[5]

Compound

[3H]-leucine uptake MTD50 (μg/mL)a

Viral plaque inhibition ID50 (μg/mL)

HSV1

HSV2

VV

1S

sattabacin (5)

20

3

3

7.5

7.5

4-hydroxysattabacin (7)

0.8

0.32

0.08

0.8

0.8

sattazolin A (14)

4.5

1.5

1.5

4.5

4.5

methylsattazolin (15)

70

35

>70

50

50

a MTD50 = median toxic dose 50%.

Table 2 Cytotoxic Activity of Acyloins Reported by Hertweck[8]

Compounda

HUVEC
GI50 (μg/mL)

K-562
GI50 (μg/mL)

HeLa
CC50 (μg/mL)

sattazolin A (14)

>50

>50

46.9 ± 1.9

sattazolin B (16)

35 ± 1.9

34.1 ± 1.3

40.3 ± 1.3

dehydrosattazolin (13)

29.9 ± 1.2

19.4 ± 0.3

31.3 ± 0.9

sattazolin ester 19

5.2 ± 0.5

4.4 ± 0.4

7.7 ± 1.8

4-hydroxysattabacin (7)

>50

>50

>50

a GI50 = concentration for 50% of maximal inhibition of cell proliferation; CC50 = concentration required to reduce cell viability by 50%.

Table 3 Antimicrobial Activity of Acyloins Reported by Hertweck[8]

Compound

Minimum inhibition zone (mm)

B. subtilis

S. aureus

M. vaccae

P. notatum

sattazolin A (14)

 0

13

18

17

sattazolin B (16)

20

18

21

22

dehydrosattazolin (13)

13

13

19

15

sattazolin ester 19

 0

 0

 0

14

4-hydroxysattabacin (7)

11

 0

19

15

Igarashi[17] has examined the bioactivity of 2′-oxosattabacin (10) in preadipocyte differentiation, which has implications for type 2 diabetes treatment, and autophagy induction, which is important in tumour growth and inflammation. Compound 10 induced 69% and 85% preadipocyte differentiation at 10 μM and 20 μM in murine ST-13 preadipocytes. Autophagy was assessed in PC12 cells, and fluorescence imaging indicated the induction of autophagy at 1–10 μL, by an increase in red puncta area. 2′-Oxosattabacin (10) showed no cytotoxicity nor antimicrobial activity.

The inhibitory activity of (R)-sattabacin (5b) and (R)-4-hydroxysattabacin (7b) in melanogenesis has been investigated, which has implications for hyperpigmentation disorders.[19] Murine melanoma cells (B16F10) were stimulated with α-melanocyte-stimulating hormone (α-MSH) and the effects of (R)-sattabacin (5b) and (R)-4-hydroxysattabacin (7b) were evaluated. (R)-Sattabacin (5b) was cytotoxic to these cells, and not probed further. However, (R)-4-hydroxysattabacin (7b) inhibited melanogenesis and was not cytotoxic, exhibiting increased potency relative to positive controls.

A cell-free tyrosinase assay indicated that (R)-4-hydroxysattabacin (7b) did not modulate enzyme behaviour, which is often implicated in melanogenesis, while controls had inhibitory effects. This was further supported by the observation that tyrosinase activity was not affected by (R)-4-hydroxysattabacin (7b) in human melanoma cells (MNT-1), whilst a decrease in pigmentation was observed. Real time PCR analysis suggested that the activity of 7b was mediated by inhibition of enzyme expression, rather than modulation of the enzyme’s activity. Finally, in artificial human skin models, (R)-4-hydroxysattabacin (7b) caused colour lightening without appreciable toxicity to epidermal cells.

Some aspects of the activity of the xenocyloin class of acyloins are difficult to decipher. Akhurst[29] has shown that Xenorhabdus cultures act as antibiotics, and likewise Nealson[21] and Chang[30] report extracts containing xenocyloins inhibited a variety of bacteria. Notably, these reports make use of either live bacteria or complex indole containing mixtures. More recently Nguyen[31] and Bode[23] reported that testing of pure xenocyloin isolates showed no significant biological activity, including against strains such as E. coli which were inhibited by the semi-purified extracts. As such, it is likely that other compounds are responsible for the activity. It has been found that xenocyloins B (22) and D (23) were active against hemocytes from Galleria mellonella at 69 μg/mL for 22 and 33 μg/mL for 23.[23]

Huang and co-workers[26] have reported the anti-inflammatory activity of several xenocyloin natural products in a RAW264.7 cell model (lipopolysaccharide stimulated) by detection of inhibition of inducible nitric oxide synthase (iNOS) and cyclooxygenase-2 (COX-2) protein expression. Xenocyloin B (22) was active in the inhibition of iNOS protein expression (~70% inhibition at 20 μM) while xenocyloins C (21) and D (23) showed reduced inhibitory effects, implying that the free hydroxyl group and terminal ethyl chain are important in the bioactivity. COX-2 and iNOS interact, and the xenocyloins were inactive against COX-2 protein expression; and thus the xenocyloins could be used to study the interplay between these two enzymes.[26]

Yu[25] has investigated the anti-platelet aggregation activity of xenocyloins G (26) and J (29) and both compounds were active. Aspirin was used as a positive control, giving an IC50 value of 289.5 ± 15.7 μM. At the 50 μM level, inhibition levels between 52.9 ± 5.5% to 96.0 ± 0.1% were observed for all xenocyloins, with xenocyloins G (26) and H (27) being the most active compounds. Xenocyloin H (27) was the most active compound, and an IC50 value of 27.5 ± 3.5 μM was obtained, whilst xenocyloin G (26) was slightly less active, exhibiting an IC50 of 31.7 ± 4.4 μM, both compounds were significantly more active than the aspirin control.


# 1.3

Biosynthesis

The biosyntheses of several acyloins have been studied,[8] [11] [16] [17] and involves a thiamine pyrophosphate (TPP) dependent enzyme. For example, Thzk0150 (from Thermosporothrix hazakensis) showed similarity to other known acyloin synthase enzymes, which catalyse acyloin formation by a reaction between two α-keto acids.[16] When phenylpyruvate (acyl donor) and 4-methyl-2-oxovalerate (acyl acceptor) were treated with the enzyme Thzk0150, TPP and Mg2+, sattabacin (5) was obtained, however the product was racemic. The substrate specificity of the enzyme was probed further with several acyl donors and acceptors. When the donor was changed to indole-3-pyruvate a sattazolin precursor was obtained, however when 4-hydroxyphenylpyruvate was used no 4-hydroxysattabacin-type intermediate was obtained. 2-Oxobutanoate 31, 2-oxoisovalerate, and (±)-3-methyl-2-oxovalerate were all active as acceptors, while pyruvate was inactive. Igarashi[17] has conducted 13C-labelling experiments to probe the biosynthesis of hazakacin (6). This showed that phenylalanine (30) and 2-oxobutanoate 31 are involved in the biosynthesis of hazakacin (6). It was also postulated that 2-oxobutanoate 31 is derived from threonine (32), and this was demonstrated with a feeding study (Scheme [1]).[17]

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Scheme 1 13C-Labelling experiments, demonstrating the incorporation of phenylalanine, 2-oxobutanoate, and threonine in the biosynthesis of hazakacin

A similar spontaneous decarboxylation of a spent biosynthetic mixture was also reported by Walsh.[11] Soraphinol A (1) and its tautomer (1:1.4 ratio respectively) were isolated when a mixture of indole-3-pyruvic acid and 4-hydroxyphenylpyruvic acid was incubated with the TPP dependent enzyme NpR1276 and allowed to sit at room temperature after the enzymatic reaction was complete. The acyloins were not, however, obtained directly, suggesting that the decarboxylation was likely a spontaneous process.

Similarly, the Hertweck group[8] found that the enzyme Cbei2730 was responsible for the formation of sattazolin A (14) in C. beijerinckii. When Cbei2730 was incubated with indole-3-pyruvate, 4-methyl-2-oxovalerate, TPP, and Mg2+, sattazolins A (14) and B (16) were obtained, along with homocoupled products of indole-3-pyruvate. The enzymatic reactions were monitored by LC-MS, and the optical purity of the products was not determined. Bode and co-workers[23] have investigated xenocyloin biosynthesis and found that it proceeds via a TPP-dependent acyloin condensation type mechanism. A gene cluster xclABCDEF was identified as responsible for the xenocyloins A–E (20–24).

A proposed mechanism for TPP 34 dependent acyloin biosynthesis is shown in Scheme [2]. Attack of the TPP ylide 34 on the α-keto acid 33 followed by decarboxylation yields an intermediate enamine 37. Addition of 37 to the acceptor 36 forges the C–C bond giving 38. Deprotonation and elimination allow regeneration of the TPP ylide 34, and affords α-hydroxy acid 39, which undergoes decarboxylation to ene-diol 40. Tautomerisation could give either acyloin 41 or 42 which are interchangeable via 40. The origin of the enantioselectivity is unknown, however an enzyme-controlled decarboxylation may be involved.

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Scheme 2 Proposed TPP-dependent biosynthesis of acyloin natural products

# 1.4

Acyloins as Discoipyrrole Biosynthetic Precursors

The acyloin 4-hydroxysattabacin (7) has been shown to be an intermediate in the biosynthesis of discoipyrrole natural products[32] (see Figure [3]). 4-Hydroxysattabacin (7) and enol 47 were isolated alongside discoipyrroles A–D (43–46) from Bacillus hunanensis.[32] The discoipyrroles are discodin domain receptor (DDR2) signalling pathway inhibitors. Mutant DDR2 has been found in around 4% of human squamous cell lung carcinomas and is implicated in tumour growth. Inhibition of this pathway has been shown to kill these cells. Discoipyrroles A–D (43–46) showed cytotoxicity against mutant lung cancer cells (non-small-cell lung cancer line HCC366) in the μM range.

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Figure 3 Discoipyrroles, with the incorporated acyloin highlighted in red

13C-Labelling studies have shown that 4-hydroxysattabacin (7) is incorporated into discoipyrrole A (43), and by analogy sattabacin (5) is incorporated into discoipyrrole B (44). This led MacMillan and co-workers to perform both a one-pot total synthesis of 43 in organic solvent, and a synthesis by treating the starting materials 7, 48, and 55 with spent bacterial media, both strongly suggesting a non-enzymatic process.[32]

Scheme [3] shows the biosynthesis of discoipyrrole A (43) from 4-hydroxysattabacin (7). Oxidation of 4-hydroxysattabacin (7) to enol 47, followed by aldol condensation with p-hydroxybenzaldehyde (48) and further oxidation yielded triketo species 49. Condensation with anthranilic acid (50) gives enamine 51, which undergoes hemiaminal formation to give seco acid 52. Adduct 52 was detected but rapidly lactonises to form discoipyrrole A (43). Several other total syntheses of the discoipyrroles have been reported.[33] [34] [35] [36]

When 4-hydroxysattabacin (7), p-hydroxybenzaldehyde (48), and anthranilic acid (50) were treated with either spent bacterial media (enzymes denatured by heating), or catalytic TFA in DMSO, discoipyrrole A (43) was obtained (Scheme [3]). It is believed that the α-oxidations are catalysed by various metals present in the spent media, or in the case of the ‘traditional’ synthesis by atmospheric O2, as when degassed DMSO was used as solvent and the reaction was conducted under a N2 atmosphere, no discoipyrrole A (43) was produced. Soraphinol A (1) has been proposed as a biosynthetic intermediate in the synthesis of the cyanobacterial sunscreen scytonemin (77).[11]

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Scheme 3 Proposed biosynthesis of discoipyrrole A (43)

#
# 2

Total Synthesis of Acyloin Natural Products

2.1

Kurasoins A and B

The Omura group reported the synthesis[4] [13] of kurasoins A (18) and B (2). Addition of BnMgCl to racemic Weinreb amide 53 yielded kurasoin A (18) in 65% yield, while addition of BnMgCl to 54 furnished kurasoin B (2) in 49% yield (Scheme [4]). These short syntheses served to confirm the structure of the natural products.

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Scheme 4 Racemic synthesis of kurasoins A and B

The enantioselective synthesis of kurasoin A (18) began with chiral resolution by Sharpless asymmetric epoxidation of 55 to yield epoxide 56 in 35% yield and >90% ee (Scheme [5]). Protection of the hydroxy groups as the TBS ethers, and epoxide ring opening with PhMgBr and CuI yielded diol 57. Moffat oxidation and deprotection then afforded kurasoin A (18). The magnitude and sign of the specific rotation confirmed the absolute configuration as S.

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Scheme 5 Omura’s enantioselective synthesis of kurasoin A

Scheme [6] details the asymmetric synthesis of kurasoin B (2). Sharpless asymmetric epoxidation of alkene 58 furnished epoxide 59 in 38% yield and >90% ee, and the alcohol was further oxidised to the ketone. Ring opening of the epoxide with indole in the presence of SnCl4 produced kurasoin B (2) in a low 22% yield over the two steps. The absolute configuration of kurasoin B (2) was also confirmed as S.

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Scheme 6 Omura’s asymmetric synthesis of kurasoin B

Ring opening of epoxide 59 proceeded in a low 27% yield in Omura’s synthesis. This ring opening was revisited by Fernandes,[37] who found that the yield could be increased to 49% when the solvent was changed to DCM/nitromethane, or to 62% when the Lewis acid was changed to Yb(OTf)3 in DCE as solvent. Yuste[38] has reported a formal total synthesis of kurasoin B (2) via a chiral sulfoxide, intersecting an intermediate reported by Omura.[13] When ester 60 was treated with the anion of chiral sulfoxide 61, β-keto sulfoxide 62 was obtained in 82% yield. Further elaboration afforded epoxide 63, which was an intermediate reported by Omura,[13] thus constituting a formal total synthesis of kurasoin B (2) (Scheme [7]).

The Omura group[39] have revisited the synthesis of kurasoin B (2) taking a slightly different approach, that nonetheless pivots on the ring opening of an epoxide (Scheme [8]). Opening of epoxide 65 with indole with Yb(OTf)3 in DCE furnished ester 66 in 77% yield. The optimisation of the yield to 77% was conducted on the epoxide of bearing the opposite stereochemistry. Hydrolysis of the ester, formation of Weinreb amide 54, and displacement with BnMgCl furnished kurasoin B (2) in 77% yield over the three steps.

Zoom Image
Scheme 7 Formal total synthesis of kurasoin B
Zoom Image
Scheme 8 Omura’s 2nd generation synthesis of kurasoin B

Andrus and co-workers have reported syntheses of kurasoins A (18)[40] and B (2)[41] via enantioselective alkylation using chiral phase transfer catalysis. These routes introduce the acyloin and requisite stereochemistry as part of a C–C bond forming step (Scheme [9]). Alkylation of benzyl bromide derivative 67 with diphenylmethyl-protected α-hydroxy ketone 68 in the presence of the chiral cinchonidine-derived catalyst 69 afforded the adduct 70 in 95% yield and 83% ee. Elaboration to the Weinreb amide 71 proceeded smoothly and in highest yield when a TES protecting group was used. Treatment with BnMgCl, cleavage of the TES ether with TBAF, followed by pivaloyl ester hydrolysis delivered kurasoin A (18) in 53% yield over the three steps. When the final deprotection was mediated by NaOH then HCl, or TBAF then NaOH isomerisation (up to 25%) to soraphinol B (3) was observed, which predates the isolation of this compound.

Zoom Image
Scheme 9 Synthesis of kurasoin A via phase transfer catalysis

Kurasoin B (2) was synthesised by a similar approach (Scheme [10]). Alkylation of protected acyloin 74 with indole bromide 73 using dimeric catalyst 72 furnished adduct 75 in near quantitative yield and 99% ee. Functional group manipulations yielded Weinreb amide 54, which was the same compound used by Omura for the synthesis of 2. Kurasoin B (2) was prepared by first masking the free hydroxyl as the TES ether, then conducting the Grignard displacement with BnMgCl. Desilylation afforded kurasoin B (2) in 53% yield over the three steps.

Zoom Image
Scheme 10 Synthesis of kurasoin B via phase transfer catalysis

# 2.2

Soraphinol A and Circumcin B

Zoom Image
Scheme 11 Walsh’s synthesis of soraphinol A and circumcin B

The biosynthesis of cyanobacterial sunshield natural product scytonemin (77) has been elucidated by Walsh,[11] and it has been proposed that soraphinol A (1) is a biosynthetic intermediate from keto acid 76 to scytonemin (77). The TPP-dependent enzyme NpR1276 is responsible for the formation of keto acid 76. As part of the biosynthetic elucidation of 77 a concise synthesis of soraphinol A (1) was reported[11] following an approach to other acyloins as described previously (Scheme [11]). Displacement of TBS-protected Weinreb amide (S)-78 with the Grignard reagent 79 afforded globally protected soraphinol A in 58% yield. Cleavage of the TBS ethers with TBAF furnished soraphinol A (1) in 60% yield, however no specific rotation was reported. Walsh has also reported a one-step racemic synthesis of circumcin B (11)[9] via a Stetter reaction using aldehyde 80 and catalyst 81 in 39% yield.


# 2.3

4-Hydroxysattabacin and Actinopolymorphol A

The Shen[6] and Xu[7] groups (Scheme [12]) have reported the discovery and total synthesis of actinopolymorphol A (8), as well as (S)-4-hydroxysattabacin (7a) which was an intermediate towards 8. Weinreb amide 83 was prepared from known α-hydroxy acid 82 in several steps and displacement with i BuMgCl gave the protected 4-hydroxysattabacin derivative 84 in 67% yield. Cleavage of the silyl ethers furnished (S)-4-hydroxysattabacin (7a), thus confirming the absolute configuration as S. Diacetylation of 7a followed by selective removal of the phenolic acetate using pyrrolidine afforded actinopolymorphol A (8) in 48% yield over the two steps. The magnitude and sign of the specific rotation for both synthetic compounds matched that for the natural material.

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Scheme 12 Synthesis of (S)-4-hydroxysattabacin and actinopolymorphol A

# 2.4

Actinopolymorphol B

A synthesis of actinopolymorphol B (9) has been reported by Tamariz[42] by a Michael addition of indole (64) to acceptor 85 to afford masked acyloin 86 in 98% yield. Basic hydrolysis afforded actinopolymorphol B (9) in 89% yield (Scheme [13]).

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Scheme 13 Synthesis of actinopolymorphol B by Michael addition

# 2.5

Sattazolins and Sattabacins

Miller and co-workers reported the synthesis of (S)-sattabacin (5a) and (S)-4-hydroxysattabacin (7a)[14] and later sattazolins A (14) and B (16) (Scheme [14]).[15] Phenylalanine (30) was converted into the Weinreb amide 87, and displacement with i BuLi furnished (S)-sattabacin (5a) in 80% yield. The data including the magnitude and sign of the specific rotation were identical to the natural product and the stereochemistry was assigned as S.[14]

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Scheme 14 Synthesis of (S)-sattabacin

A similar approach was used to access (S)-4-hydroxysattabacin (7a) as detailed in Scheme [15]. Reduction of keto acid 88 with (+)-DIPCl yielded α-hydroxy acid 82 in good yield and ee. Acid 82 was converted into the Weinreb amide 53 then displaced with i BuLi to furnish (S)-4-hydroxysattabacin (7a) in low yield.[14]

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Scheme 15 Synthesis of (S)-4-hydroxysattabacin

Scheme [16] details the synthesis of sattazolins A (14) and B (16).[15] Ring opening of commercially available epoxide 65 with indole (64) in the presence of Yb(OTf)3 gave methyl ester 66 in near quantitative yield, which was further elaborated into protected Weinreb amide 89. Treatment of 89 with i BuLi furnished globally protected sattazolin A derivative 90 in 82% yield. The deprotection step proved challenging, when the alcohol was unveiled with TBAF followed by thermolysis to remove the Boc group, an inseparable mixture (~93:7) of sattazolins A (14) and B (16) was obtained in 74% yield. Interestingly, sattazolin B (16) had not been reported at the time of synthesis; when the carbamate was thermolysed first, followed by treatment with TBAF, sattazolin A (14) was the only product obtained in 71% yield.[15]

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Scheme 16 Synthesis of sattazolins A and B

Several intermediates reported by Shen,[6] [7] Miller,[14] and Omura[13] have been synthesised by Tilve and Patil,[43] and constitute formal syntheses of kurasoin A (18), 4-hydroxysattabacin (7a), and actinopolymorphol A (8) (Scheme [17]). Sharpless dihydroxylation, followed by reduction of the benzylic alcohol and benzyl ether cleavage afforded known Weinreb amide 53 which was further protected as the bis TBS ether, yielding 83, thus completing the formal syntheses.

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Scheme 17 Formal synthesis of various acyloins

The Narsaiah group[44] reported concise asymmetric syntheses of sattazolin A (14) and (S)-sattabacin (5a) hinging on a ring opening of epoxide 95 (Scheme [18]). Sharpless asymmetric epoxidation and concomitant kinetic resolution of racemic alcohol 93 afforded epoxide 94 in 44% yield, which was oxidised and converted into acetal 95. Epoxide ring opening with PhMgBr followed by acetal cleavage with HCl afforded (S)-sattabacin (5a), whilst treatment with indole and Eu(OTf)3 opened the epoxide and concomitantly cleaved the acetal, affording sattazolin A (14).

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Scheme 18 Narsaiah’s synthesis of sattazolin A and (S)-sattabacin

The Shimada group[12] has recently developed a catalytic Weinreb amide formation from α-hydroxy acids and N,O-dimethylhydroxylamine using a diboronic acid anhydride (DBAA) type catalyst 96. This method is the first example of a catalytic dehydrative amidation for the formation of Weinreb amides, culminating in the synthesis of eight acyloins, including the first total synthesis of soraphinol B (3) in a one-pot fashion, which served to confirm the absolute configuration. Shimada achieved a one-pot, two-step, gram-scale synthesis of (S)-sattabacin (5a) in 75% yield without racemisation of the acyloin moiety.

Scheme [19] details the synthesis of various acyloins. When an aryl α-hydroxy acid such as 97 is treated with DBAA catalyst 96 and HNMe(OMe) at reflux in DCE Weinreb amides such as 98 are obtained. Upon displacement of the Weinreb amide with a Grignard reagent, a general acyloin such as 99 is obtained. This concise method allowed access to eight acyloin compounds, in good to excellent yields in one pot and two steps, or three steps where a deprotection was required.

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Scheme 19 Shimada’s development of a catalytic Weinreb amide formation and synthesis of acyloin natural products

# 2.6

Catalyst Development and Application in Acyloin Synthesis

The Rizzacasa group has achieved the racemic synthesis of seven members of the acyloin family by Mukaiyama enone hydration and acyloin rearrangement using newly developed Co(III) 100 and Co(II) 101 SALPN-type metal complexes (Figure [4]).[45] This approach allowed isomeric acyloins, such as sattazolin A (14) and sattazolin B (16), to be accessed from a common intermediate, in this case 107. The enone precursors 102–108 were prepared by Horner–Wadsworth–Emmons reaction (HWE) and subjected to Mukaiyama hydration.

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Figure 4 Co(III) and Co(II) complexes developed Rizzacasa for Mukaiyama enone hydration

The synthesis of the acyloins is summarised in Scheme [20]. Hydration with catalysts 100 and 101 was compared directly against Mn(dpm)3. This catalyst gave a single isomer of the hydration product, giving access to soraphinol C (4) after debenzylation and sattazolin B (16) in a single step from 107. When the novel Co(III) and Co(II) catalysts 100 and 101 were used to hydrate enone 102, an acyloin rearrangement was observed, which allowed access to 4-hydroxysattabacin (7) after debenzylation, this product was then converted into actinopolymorphol A (8).[6] [7] The acyloin mixtures could be further equilibrated by resubjecting the reaction mixture to hydration conditions with Co(III) catalyst 100. Sattabacin (5), sattazolin A (14), and hazakacin (6) were accessed by the same approach in a single step from the enone precursors. The indole mixture was however not resubjected to hydration due to stability issues.

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Scheme 20 Rizzacasa’s synthesis of acyloin natural products by Mukaiyama hydration

# 2.7

Xenocyloins

Nguyen and co-workers[31] have reported a concise racemic synthesis of xenocyloin A (20) followed by resolution via chiral supercritical fluid chromatography (SFC) (Scheme [21]). Racemic indole methyl ester 66 was treated with i PrMgCl, to yield xenocyloin A (20) as well as undesired tautomer 110 in a 1.2:1 ratio and 67% overall yield. The isomers were inseparable by chromatography and attempts to ameliorate the issue by protecting the hydroxy group failed, as did the use of a Weinreb amide. However, the isomers were separable by chiral SFC, which allowed production of both sets of enantiomers (>96% ee). The acetates of a mixture of xenocyloin A (20) and isomer 110 were also synthesised, constituting a synthesis of xenocyloin C (21) however the isomers were not separated. The antibiotic activity of the xenocyloin compounds and the derived acetates was only very low (>80 μg/mL).

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Scheme 21 Nguyen’s racemic synthesis of xenocyloin A

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# 3

Conclusion

This review has detailed the rich chemistry and biology of acyloin-containing natural products. Despite apparent structural simplicity, the acyloins described herein have varied biological properties, a fascinating biosynthetic pathway, serve as biosynthetic precursors to complex natural products, and have inspired concise syntheses as well as method development.


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#

Conflict of Interest

The authors declare no conflict of interest.


Corresponding Author

Mark Rizzacasa
University of Melbourne, School of Chemistry, Bio21 Institute
Melbourne, 3010
Australia   

Publication History

Received: 16 November 2022

Accepted after revision: 27 December 2022

Accepted Manuscript online:
27 December 2022

Article published online:
06 February 2023

© 2022. Thieme. All rights reserved

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Mark Rizzacasa(left) obtained a Ph.D. in 1990 under the guidance of Professor Melvyn Sargent at the University of Western Australia. He then moved on to the USA and spent 2 years as a postdoctoral fellow in Professor Robert E. Ireland’s research group at the University of Virginia. In early 1993, he then returned to Australia to begin an appointment in the School of Chemistry at the University of Melbourne and was promoted to full Professor in 2008. Mark was awarded the RACI Rennie Memorial Medal in 1997, the 2005 Organic Reactions Lectureship, and the prestigious RACI A. J. Birch medal in 2010. His interests include the total synthesis of bioactive natural products and the development of synthetic methodologies. Michael Ricca (right) completed a B.Sc. and M.Sc. at The University of Melbourne. His masters research project was conducted in Professor Mark Rizzacasa’s lab, focusing on the synthesis of bioactive natural products. He began a Ph.D. in 2019 in the same lab, with research interests including total synthesis and the development of metal complexes for hydrogen atom transfer reactions.
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Figure 1 Acyloin natural products
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Figure 2 Xenocyloin natural products
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Scheme 1 13C-Labelling experiments, demonstrating the incorporation of phenylalanine, 2-oxobutanoate, and threonine in the biosynthesis of hazakacin
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Scheme 2 Proposed TPP-dependent biosynthesis of acyloin natural products
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Figure 3 Discoipyrroles, with the incorporated acyloin highlighted in red
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Scheme 3 Proposed biosynthesis of discoipyrrole A (43)
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Scheme 4 Racemic synthesis of kurasoins A and B
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Scheme 5 Omura’s enantioselective synthesis of kurasoin A
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Scheme 6 Omura’s asymmetric synthesis of kurasoin B
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Scheme 7 Formal total synthesis of kurasoin B
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Scheme 8 Omura’s 2nd generation synthesis of kurasoin B
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Scheme 9 Synthesis of kurasoin A via phase transfer catalysis
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Scheme 10 Synthesis of kurasoin B via phase transfer catalysis
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Scheme 11 Walsh’s synthesis of soraphinol A and circumcin B
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Scheme 12 Synthesis of (S)-4-hydroxysattabacin and actinopolymorphol A
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Scheme 13 Synthesis of actinopolymorphol B by Michael addition
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Scheme 14 Synthesis of (S)-sattabacin
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Scheme 15 Synthesis of (S)-4-hydroxysattabacin
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Scheme 16 Synthesis of sattazolins A and B
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Scheme 17 Formal synthesis of various acyloins
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Scheme 18 Narsaiah’s synthesis of sattazolin A and (S)-sattabacin
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Scheme 19 Shimada’s development of a catalytic Weinreb amide formation and synthesis of acyloin natural products
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Figure 4 Co(III) and Co(II) complexes developed Rizzacasa for Mukaiyama enone hydration
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Scheme 20 Rizzacasa’s synthesis of acyloin natural products by Mukaiyama hydration
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Scheme 21 Nguyen’s racemic synthesis of xenocyloin A