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Cyclohelminthols Y1-Y4 Metabolites Possessing Two Spirocyclopropanes in their Structure Shizuya Tanaka, Kazuaki Tanaka, Hayato Maeda, and Masaru Hashimoto J. Org. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.joc.8b00727 • Publication Date (Web): 03 May 2018 Downloaded from http://pubs.acs.org on May 3, 2018

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The Journal of Organic Chemistry

Cyclohelminthols Y1-Y4 Metabolites Possessing Two Spirocyclopropanes in their Structure Shizuya Tanaka, Kazuaki Tanaka, Hayato Maeda, and Masaru Hashimoto*

Faculty of Agriculture and Life Science, Hirosaki University, 3-Bunkyo-cho, Hirosaki, 036-8561, Japan

Corresponding Author: [email protected]

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TOC Graphic:


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Abstract Cyclohelminthols Y1-Y4 (1-4) were isolated from the culture broth of Helminthosporium velutinum yone96.

These

compounds

are

3-azabicyclo[3.1.0]hexane-6-spirocyclopentane

diastereomers linked

with

to a

each

other

featuring

cyclopentanespirocyclopropane

framework. Their planar structures were established via the comparison of their spectra with the simpler analogue cyclohelminthol X as well as the analysis of their HMBC spectra. Although the proton-deficient core frameworks of 1-4 prevented us from obtaining configurational information via conventional NMR analysis, their total structures involving the relative and the absolute configurations were established using density functional theory (DFT)-based molecular modeling calculations. The present study demonstrates the effectiveness of the comparison between the theoretical and experimental δ13C values for stereochemical analysis by focusing on the carbons that show relatively large δ13C deviations among the isomers. The G-ring of these molecules most likely originates from the cyclopropanation of the C6C7 double bond with the carbene equivalent 6 derived from cyclohelminthol IV (7), which was isolated from the same producer fungus. Preliminary biological experiments revealed the potent cytotoxicity of the (6S)-isomers against COLO201 cells, whereas the (6R)-isomers exhibited weak activity. The antifungal assay with Cochiobolus miyabeanus showed slightly different profile.



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Introduction Structurally unique natural products have attracted attention from many organic chemists.1 These molecules sometimes show unique and clinically useful activities, which also led them to perform the total syntheses2 with the aim to exploit the activities of these products.3 The challenging structural elucidation of these unique structures has led to the development of many useful analytical methodologies. Nowadays, the NMR technique is one of the most powerful tools for the elucidation of not only planar structures, but also relative configurations in solution through the analysis of features such as 3J-couplings and NOE (ROE) signals.4 Although the development of 2D NMR spectroscopy has undoubtedly contributed to the clarification of structures via spectral interpretation, these experiments still sometimes require specific empirical skills in order to obtain satisfactory results, especially in the case of proton-deficient molecules. Thanks to the significant progress in computer technology, quantum chemistry-supported molecular modeling calculations have become quite an effective tool to circumvent this lack of experience.5 In this context, we applied DFT calculations in the structural investigations of epoxyroussoeone,6 neomacrophorin X,7 and cyclohelminthol X (5).8 Although the proton-deficient core frameworks of these molecules afforded only limited structural information by NMR spectroscopy, the DFT-based 13C chemical shifts and ECD spectral analyses were quite useful to solve their planar structures and absolute configurations. In this work, our extensive exploration of secondary metabolites from Helminthosporium velutinum yone96 led to the isolation of cyclohelminthols Y1-Y4 (1-4). These compounds, which are diastereomers to each other, possess the characteristic cyclopentanespirocyclopropanes (FG rings) linked with 3-azabicyclo[3.1.0]hexane-6-spirocyclopentane frameworks (CDE rings). Since only three protons (H-6, H-7, and NH) are linked directly with the core of the CDE-FG-ring systems in 1-4, their NMR spectra provide insufficient structural information. The 13C NMR spectra of these molecules give a small average standard deviation (SD) value for the 13C chemical shifts among 1-4, which is below the accuracy level of ACS Paragon Plus Environment

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DFT chemical shift calculations. This article describes the structural elucidation of these molecules by taking advantage of the DFT-based molecular modeling calculations and the discussions of their NMR and ECD spectra. Plausible biosynthetic pathway and biological activities are also described.



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Results and Discussion Isolation and determination of the planar structure of cyclohelminthols Y1-Y4. Cyclohelminthols Y1-Y4 (1-4, Figure 1) were isolated from the less polar fraction of the ethyl acetate extracts from Helminthosporium velutinum yone96 culture broth than that gave cyclohelminthol X (5).8 The mycelia of the fungus contain them more than the culture broth. Compounds 1-4, which were named cyclohelminthols Y1-Y4, respectively, according to the order of their elution, were successfully separated with a cholesterol-supported reverse phase HPLC column9 after trials with octadecylsilyl (ODS) columns failed (Figure S1). These molecules provide almost the same LCESIMS profiles (Figure S2) as well as quite resembled 1H and

13

C NMR spectra, suggesting that 1-4 are

diastereomers to each other. Figure 2 shows the 1.5–3.0 and 6.5–8.0 ppm regions of the 1H NMR spectra of 1-4 as well as cyclohelminthol X (5). Although the structural analyses of 1-4 were performed in parallel, hereafter we will limit the discussion mainly to 1 since similar analyses resulted in identical planar structures for 2-4. The

13

C and 1H NMR data of 1-4 as well as their signal assignments are

summarized in Tables S1 S2, and S3, respectively.

Figure 1. Structures of cyclohelmintols Y1-Y4 (1-4), cyclohelmintol X (5), carbene equivalent 6,

IV (7)


and cyclohelminthol

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Thus, cyclohelminthol Y1 (1) afforded the nominal protonated ion at m/z = 810.2412 together with the ammonium and sodium adduct ions (m/z = 827.2683 and 832.2242, respectively) in its ESI-TOF mass spectrum. The isotopologue patterns of these ions (approximately 10:5:8:4:2) revealed the presence of two chlorine atoms in the molecule.10 These results suggest its molecular formula is C42H45Cl2NO11 ([M+H]+: 810.2442), which was confirmed by the 42 resonances present in its 13C NMR spectrum. The elemental difference between 1 and 5 is C8H5ClO2 which is in accord with the elemental composition of the carbene equivalent 6. This unit is theoretically derived from cyclohelminthol IV (7) through the

Figure 2. Aliphatic and olefinic regions of the 1H NMR spectra of 1-5 in CDCl3.



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oxidation of its alcohol groups and successive carbene formation (elimination of the α-proton and the chloride ion at the α-position), since 7 was also detected from the same producer fungus.11 Interestingly, the framework of 7 is incorporated in 5 as the E ring.8 Accordingly, it seems reasonable to assume that 1-4 are biosynthetic congeners of 5 carrying two units of 6. The NMR spectral analysis of 1 revealed the presence of E-1-oxo-2-butenyl (C1’-C4’), E-1-propenyl (C6”-C8”), and a n-hexyl (C17’-C22’) side chains in 1. The E-configurations in the former two substructures were determined based on the large 1H vicinal olefin spin coupling constants (15.3 and 15.6 Hz, respectively). The characteristic H-6 (2.83 ppm, d, J = 8.5 Hz), H-7 (2.96 ppm, dq, J = 8.5, 6.3 Hz), and H3-8 (1.48 ppm, d, J = 6.3 Hz) signals were correlated in the COSY spectrum, which allowed the C6-C7-C8 linkage to be established, as shown in Figure 3. Both H-6 and H-7 gave rise to HMBC signals with C1” (43.4 ppm), suggesting that the C6, C7, and C1” atoms comprise a cyclopropane ring (F ring), which is supported by the δ13C values of C6 (32.1 ppm) and C7 (32.2 ppm). The H-6 resonance also correlated in the HMBC spectrum with C3 and C5 (153.1 and 189.7 ppm, respectively), while H-7 correlated with C4 (152.2 ppm). These HMBC signals indicate that the F ring is connected to the cyclopentenedione E ring. Although the 3JHH values is not conclusive for stereochemical assignment

Figure 3. Characteristic HMBC (pink arrows) correlations and ROEs (blue arrows) in 1.


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of substituted cyclopropanes,12 an ROE signal was observed between H-6 and H3-8, revealing a trans-relationship between C4 and C8 on the cyclopropane F ring. Interestingly, irradiation at H-6 gave another ROE at H-2’, providing useful information regarding the stable conformation of this molecule which will be described later. Comparison of the 13C NMR spectra of 1 and 5 revealed the presence of a bicyclo[4.3.0]nonane unit (AB rings) as well as the central 3-aza-bicyclo[3.1.0]hexane-2,4-dione (CD rings) skeleton. One of the methylene protons at C17’ appeared as an isolated signal at 2.39 ppm and gave rise to HMBC correlations with C1, C16’, and C26’ (47.5, 50.8, and 169.3 ppm, respectively), disclosing the C16’-C17’ linkage. This also allowed us to distinguish the two carbonyls (C25’ and C26’) on the C ring. Although no informative HMBC signal was obtained for the connectivity between the AB and CDE ring segments, the C14’ (195.4 ppm) and C15’ (52.4 ppm) were connected by considering their spectral resemblance to that of 5. Cyclohelminthol X (5) did not provide the corresponding HMBC signals, neither.8 Finally, the chlorine atoms must be attached to the C3 and C3” appeared at 153.1 and 144.8 ppm, respectively, to complete the molecule. Overall, these results establish the planar structure of 1. Additionally, the planar structures of 2-4 were deduced similarly.

Stereochemistry. The planar structural discussions have established the cis relationship between H-6 and C8 and the E-configurations for the C2’/C3’ and the C6”/C7” double bonds as described.

We then investigated the

remaining stereochemistry of 1-4. From their spectral resemblance and shared origins, i.e., same producer fungi, identical relative and absolute configurations can be envisaged for the common AB- and CDE-ring systems in 1-5, which is supported by the following results. The splitting profiles for Hα-6’ (1.64 ppm, dd, J = 11.8, 14.0 Hz), H-9’ (2.46 ppm, tt, J = 3.6, 12.2 Hz), and H-13’ (4.16 ppm, d, J = 11.6 Hz) in the 1H NMR spectrum of 1 were in good accordance with the corresponding signals of 5, although the details of the other 1H signals were undecipherable due to serious signal overlapping. The maximum ACS Paragon Plus Environment

13

C chemical shift


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deviation for C1’-C18’ was found to be 0.6 ppm (C16’) between 1 and 5, as shown Figure 4(A). This figure displays a plot highlighting the localization of the δ13C differences between these two molecules. Figure 4(B) shows the localization of the standard deviations (SD) of the

13

C resonances among 1-4.

This plot clearly indicates that the AB and CDE rings exhibit the same configuration for 1-4 as well as 5, i.e., 1S,5’R,7’S,9’R,10’S,11’R,12’S,15’R,16’S. Considerable deviations were found around the F ring, which indicates that the diastereomeric relationship among 1-4 originates from the F rings. The C23’ carboxyl carbons showed a non-negligible large δ13C SD value (0.58 m) among 1-4. This is likely due to the slightly different contents of residual TFA from the final HPLC purification, which affects the equilibrium between the carboxylic acid and the corresponding carboxylate ion. Cyclohelminthols Y2-Y4 (2-4) also provided ROE signals between H-6 and H3-8 (see the spectra S-20, S-28, S-34, and S-40 in SI), disclosing the same trans-configuration for C4 and C8 on the G ring. This allowed us to assign that 1-4 are either one of (6R,7R,1”R)-, (6R,7R,1”S)-, (6S,7S,1”R)-, and (6S,7S,1”S)-configuration,

Figure 4. Distribution of the δ13C deviations. standard deviation (ppm) among 1-4.

(A) δ13C differences (ppm) between 1 and 5 and (B) δ13C


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respectively. The H-6, H-7, and the N-H are the only protons directly linked to the CDE and FG ring frameworks. Therefore, neither 3JHH- nor NOE (ROE)-based analyses were useful to elucidate the configurational relationship between F ring and the stereogenic centers on the other rings. Furthermore, the CDE ring contains a pseudo-symmetric plane, as shown in Figure 5, where the carbonyl group at C14’ and the methylene C17’ asymmetrize the spiro carbon at C1. Interestingly, the NMR spectroscopic features of H-6 and H-7 still vary among the isomers (see Figure 2), although these protons are located very far from C14’ and C17’ in their structures. Molecular modeling calculations were performed to gain more insight into the spatial relationships of H-6 and H-7 with other functionalities in the molecule. The structures depicted in Figure 6 were selected as models for this study. Although we assigned the (1S)-configuration, models of the (1R)-isomers were also calculated to verify our assignment. In these models, the C17’-C22’ n-hexyl groups were substituted for n-propyl groups to reduce the number of conformers to be examined, and the C23’ carboxylic acids were removed in the same purposes. In addition, a model carrying a n-hexyl side chain (Mo-S-RR-hex) was designed and calculated in the later discussion to investigate the effect of the C17’-C22’ hexyl group.

Figure 5. Pseudo-symmetric plane in cyclohelminthol Ys.



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Figure 6. Structures of the models used for the calculations.

The n-propyl groups were fixed as the anti-conformation (∠C16’C17’C18’C19’ = 180°) during the conformational search also to reduce the number of initial conformers. The candidate stable conformers were narrowed down stepwise using geometrical optimization with HF/321G followed by that with ωB97X-D/6-31G*.13 The final candidates were optimized employing the BHLYP/def2-SVP level.14 The entropies, obtainable by the vibrational analysis, were considered by taking the large conformational freedom of these models into account. The Boltzmann distributions of the conformers were obtained based on the free energies. Since the 1H NMR profile of 1 remained virtually unaltered in different solvents (CDCl3 and CD3OD), the solvent effect was not considered in these calculations. The results obtained from the calculations of the Mo-S-RR, Mo-S-RS, Mo-S-SR, and Mo-S-SS models suggest that the C13’-C14’(=O)-C15’ moiety adopts the same conformation in most of the stable conformers regardless of the configurations at C6, C7, and C1”. In these conformers, the C1’-C5’ side chain exhibits an expanded conformation and blocks the left side of the E ring plane as shown in Figure 7, which sends the F ring to the right side. This geometry was confirmed by the appearance of an ROE


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Figure 7. Models applied for the calculations and stable conformations of the (6R)- and (6S)-isomers.

at H-6 upon irradiation of the H-2’ of 1 (see spectrum S-21 in SI). This conformational property differentiates the H-6 by its configuration. In the models with the (6R)-configuration (Mo-S-RR and Mo-S-RS), the bulky chlorine atom at C3 (Van der Waals radii: 1.8 Å)15 hinders the G ring to render the dihedral angle ∠C3C4C6C1” almost perpendicular in their stable conformers, which brings the H-7 atom close to the E-ring plane. Accordingly, H-7 is magnetically deshielded by the diatropic ring current of the C3C4 double bond. The carbonyl group at C5 may also contribute to the deshielding. This conformation also places the H-6 atom near the E ring plane, suggesting that H-6 might be weakly deshielded. In contrast, the G ring can approach the less bulky oxygen atom at C5 (Van der Waals radii: 1.5 Å)15 more closely in the (6S)-isomers (Mo-S-SR and Mo-S-SS), thereby placing the H-7 out of the E-ring plane. Therefore, the H-7 atom would not be deshielded in this conformation, i.e., H-7 of (6S)-isomers should resonate at a lower frequency than that of (6R)-isomers. These results allowed us to assign the (6R)-configuration for 1 and 3 and the (6S)-configuration for 2 and 4. The chemical and geometrical environment of the C6”-C8” side chains of 1-4 resemble that of C6-C8 in 5. This suggests that the spectroscopic features of H-6” and H-7” in the 1H NMR spectra of 1-4 should



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be similar to that of H-6 and H-7 in 5, respectively. However, in 2 and 3, H-6” and H-7” resonate at higher frequencies than those in 1 and 4, which evidences the similarities between 1 and 4 on one hand, and 2 and 3 on the other. This pairing pattern is different from that obtained based on the NMR spectroscopic features of H-6 and H-7. Since the configurations of C6 and C7 contribute to the latter pairing pattern, the configuration at C1” is expected to contribute to the present pairing pattern. The chemical shifts of H-6” and H-7” for 2 and 3 are almost identical to the corresponding olefinic protons in 5. Thereupon, we examined the spatial environments of C6”-C8” propenyl groups in these molecules. The most stable conformers of the models Mo-S-RS, Mo-S-SR, Mo-S-RR, and Mo-S-SS are summarized in Figure 8. The second most stable conformers are the rotamers at the C16’-C17’ single bond in these models. The C17’-C19’ propyl groups are highlighted with a red sphere. In the model of 5, the C6-C8 side chain protrudes from the molecular surface in the most stable conformer,8 which is the same in the C6”-C8” side chains in the stable conformers of Mo-S-RS and Mo-S-SR. Accordingly, H-6” and H-7” in these models are expected to behave similarly to the H-6 and H-7 atoms of 5 in the

Figure 8. The most stable conformers of model for 5, and models of cyclohelminthol Y (Mo-S-RS, Mo-S-SR, Mo-S-RR, and Mo-S-SS). The propyl groups corresponding to C17’-C19’ are highlighted with red surfaces.


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corresponding 1H NMR spectra. For cyclohelminthols Y2 (2) and Y3 (3), this feature seems to be valid. In contrast, the corresponding chains in Mo-S-RR and Mo-S-SS do not protrude from the molecular surfaces, which implies that the H-6” and H-7” atoms would be magnetically affected within the same molecule. Since the H-6” and H-7” atoms of 1 and 4 resonate at a slightly lower frequency than the corresponding protons of the others, 1 and 4 may correlate with either Mo-S-RR and Mo-S-SS. Comprehensively combining above results allowed us to assign the configurations of 1-4 as (6R,7R,1”R), (6S,7S,1”R), (6R,7R,1”S), and (6S,7S,1”S), respectively.

Confirmation of the configurational assignments based on DFT calculations Next, we considered whether the theoretical chemical shifts would confirm this assignment of configurations. Since cyclohelminthol Ys (1-4) afforded very similar

13

C NMR spectra, giving rise to

very small δ13C SD value (0.17 ppm) which is much smaller than the accuracy level in the DFT calculations (approximately 2 ppm),16 conventional statistical analyses with absolute chemical shifts proved to be useless in the present analysis. Subsequently, we decided to focus on the carbons that showed characteristic δ13C deviations in the experimental data, and their relative chemical shifts were compared. Table 1 shows the experimental 13

C resonances that afforded δ13C SD values larger than 0.20 ppm among 1-4 in descending order. The

carboxy carbon at C23’ (δ13C SD =0.58 ppm) was not discussed, because this carbon is omitted in the models. Interestingly, for most of the examined carbon nuclei, the NMR signals corresponding to the four compounds can be divided into two groups, those with the large chemical shifts (bold) and those with the small chemical shifts (plain). For example, the C1” resonance of 1 and 3 appears at higher frequencies (43.4 and 43.2 ppm, respectively) than that of 2 and 4 (41.3 and 41.2 ppm, respectively). The C6 follows the same tendency. In the case of the C3 and C5, that corresponding to compounds 1 and 3 resonates rather at lower frequencies than the other two. On the other hand, the C4 and C7” atoms



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Table 1. Comparison of the 13C chemical shift with the characteristic deviations among 1-4 and that of the models based on ωB97X-D/6-31G*. Compound or model

C1"

C6

C5

C3

C4

C2

C7"

C7

C5"

1 2 3 4 SD (ppm)

43.4 41.3 43.2 41.2 1.03

32.1 31.1 32.1 31.1 0.50

189. 7 190. 7 189.9 190.7 0.46

153.1 153.9 153.2 153.9 0.38

152.2 151.9 151.6 152.5 0.34

185.6 185.8 185.2 186.0 0.30

146.3 146.0 146.1 146.7 0.27

32.2 32.2 32.3 31.7 0.23

194.7 194.6 194.4 195.0 0.22

Mo-S-RR Mo-S-SR Mo-S-RS Mo-S-SS Judgement

45.2 41.0 44.6 40.8 agree

32.2 30.6 32.5 29.8 agree

192.9 195.8 194.0 196.0 agree

147.4 150.5 148.7 150.3 agree

157.2 154.6 155.0 157.0 agree

193.7 193.5 193.9 194.7 agree

150.7 147.3 147.8 152.1 agree

31.3 32.2 32.0 30.5 agree

200.4 200.4 199.9 201.2 agree

Mo-R-SS Mo-R-RS Mo-R-SR Mo-R-RR Judgement

44.5 40.2 44.3 40.1 agree

32.2 31.6 32.1 31.9 agree

198.6 200.8 199.1 195.8 disagree

153.1 154.4 153.4 153.3 disagree

151.7 152.7 151.1 153.1 disagree

189.2 190.1 189.3 191.3 agree

148.9 147.5 147.8 148.2 agree

31.5 31.7 31.2 32.2 disagree

200.5 201.1 200.1 201.8 agree

Pairs or groups of larger values are highlighted with bold letters.

exhibit a different pairing pattern, where the 13C signals of 1 and 4 appear at higher frequencies than that of 2 and 3. For the C2 and C5” signals, that corresponding to compound 4 stands out with higher frequencies than that of 1, 2, and 3. In contrast, the C7 atom of compound 4 gives rise to remarkably small chemical shift among the four compounds. These 13C signals were compared with the theoretical chemical shifts obtained with ωB97X-D/6-31G*, which revealed that the pairing patterns were in complete agreement with that of the assigned configurations. No other combinations satisfy these pairing patterns. Furthermore, they were also computed using the B3LYP/6-31G*, EDF2/6-31G*, and ωB97X-D/6-311G* levels, and no inconsistency was found (Table S3), which supports the configurational assignment as described in the preceding section. Although we assigned the (1S)-configuration for 1-4, the theoretical chemical shifts for the (1R)-isomers were also investigated. However, Mo-R-RS, Mo-R-SR, Mo-R-RR, and Mo-R-SS did not satisfy the abovementioned pairing patterns with any combinations. Although the combination shown in Table 1 is


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the best match to the experimental data, there are inconsistencies at the C3 and C4 atoms, despite their considerably large δ13C SD values (0.38 and 0.34 ppm, respectively). These calculations simultaneously provided the 1H chemical shifts despite of low accuracy because these sensitively dependent on solvents. It was found that the calculations based on ωB97X-D/6-31G* satisfactorily reproduced the NMR behaviors of H-6 and H-7, as shown in Table 2. However, the theoretical result of H-7” is not in accord with the experimental data. The theoretical resonance of H-7” in Mo-S-RR (assigned as 1) appears at the highest frequency, whereas it resonates at a lower frequency than that of 2 and 3 in the experimental spectra. Calculations performed with other DFT functionals also led to the same results. Interestingly, the models with the (1R)-configuration exhibit an accordance in the case of the chemical shifts of H-7”. We did not examine any combinations comprised of the isomers with (1R)- and (1S)-configurations, because these are impractical from the viewpoint of their biogenesis, as will be described later.

Table 2. Comparison of the 1H chemical shift with the characteristic deviations among 1-4, and that of the models

based on ωB97X-D/6-31G*. Compounds or model 1 2 3 4 SD (ppm)

H-6

H-7

H-7”

H-6”

2.83 2.72 2.85 2.67 0.07

2.96 2.59 2.98 2.50 0.21

7.35 7.50 7.50 7.35 0.08

6.40 6.50 6.47 6.41 0.04

2.57

3.15

8.08

6.35

Mo-S-SR

2.37

2.54

7.95

6.51

Mo-S-RS

2.58

3.05

7.97

6.46

2.14 agree

7.95 disagree

6.29 agree

Mo-S-RR

Mo-S-SS Judgement

2.34 agree

Mo-R-SS

2.34

2.93

7.79

6.40

Mo-R-RS

2.07

2.06

7.96

6.50

Mo-R-SR

2.33

2.94

7.91

6.48

2.05 agree

7.59 agree

6.42 agree

3.44 2.97

8.25 7.85

6.38 6.39

Mo-R-RR Judgement Mo-S-RR-hex conf-A conf-B

2.20 agree 2.67 2.51

Pairs or groups of lager values are highlighted with bold letters.



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As previously mentioned, the C17’-C22’ hexyl groups in natural 1-4 were replaced with n-propyl groups in these models for computational time saving. The results suggest that the n-propyl chain locates spatially close enough to the C6”-C8” propenyl side chain in Mo-S-RR for some interaction between both groups to occur¸ whereas the long distance between those of the other models hinders such interaction (see Figure 8). In fact, irradiation at H-7” of 1 afforded weak ROEs at H-17’, H-18’, and H-19’ (2.16, 1.78, and 1.27, respectively, see the spectrum S-22 in SI). To further investigate this interaction, a model carrying the C17’-C22’ n-hexyl group, Mo-S-RR-hex, was also subjected to the calculations. This model was constructed by elongating the n-propyl group of the most stable conformer of Mo-S-RR to the n-hexyl chain. Although comprehensive conformational search was impractical because of the large conformational freedom of the hexyl group, a restricted conformational search performed with relaxation of only the n-hexyl group led to conf-A and conf-B as stable conformers (Figure 9). Despite being quite similar, these conformers show a clear 1H chemical shift difference at H-7” (conf-A: 8.25 ppm and conf-B: 7.85 ppm). This is likely caused by the difference of the deshielding effect due to the carbonyl group at C26’. Accordingly, we concluded that the simplified model Mo-S-RR could not satisfyingly reflect the spectral properties of 1. Cyclohelminthol Ys 1-4 were also subjected to ECD spectroscopic analysis to further ascertain our configurational assignment. Although the experimental ECD spectra show some differences (colored

Figure 9. Stable conformers of Mo-S-RR-hex and theoretical chemical shifts at H-7” (ωB97X-D/6-31G*).


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solid lines in Figure 10), they are not diagnostic enough to directly discriminate between the four candidate models. Thus, we attempted to compare the experimental ECD spectra with that obtained theoretically for the models with the assigned configurations. The stable conformers theoretically dominating more than 90% abundance were subjected to ECD calculations at the BHLYP/def2-SVP level.8 The theoretical ECD spectra were obtained after corrections based on the Boltzmann distributions. The ECD intensity of Mo-S-RS was normalized to that of the experimental spectrum of 2 at around 236 nm, and the same scale was used for the other theoretical

Figure 10. Experimental ECD spectra of 1-4 and the theoretical spectra of the models based on BHLYP/def2-SVP.



Page 20 of 33

spectra. The obtained theoretical ECD spectra (colored dotted lines in Figure 10) show acceptable accordance with their experimental counterparts. The theoretical ECD spectra of the models with the (1R)-configuration (Mo-R-RR, Mo-R-RS, Mo-R-SR, and Mo-R-SS) were also compared (black dotted lines in Figure 10) using the combination that exhibited a best match with the theoretical NMR chemical shifts. Although the model Mo-R-RS afforded an ECD spectrum that was in agreement with the experimental data, other models exhibited substantial differences. Especially, theoretical ECD spectrum of Mo-R-SR did not show resemblance with any experimental spectra. The mirror image (ent-Mo-R-SR) may accord with 4. But, chiral reversing for only one model would be an arbitrary interpretation, when their biogenesis is considered. These results further substantiate that the (1R)-configuration can be ruled out. Overall, the DFT-based NMR and ECD analyses allowed us not only to eliminate the possibility of the (1R)-isomers but also to verify our assignment based on the experimental NMR discussions.

Plausible biogenesis. The biogenetic pathways of cyclohelminthols Y1-Y4 (1-4) are depicted in Scheme 1. We propose a mechanism in which cyclohelminthol X (5) is derived biosynthetically from maleimide 8 and chloroenolate 9 through Michael addition/alkylation sequences that involves intermediate 1017. Chloroenolate 9 acts as the carbene equivalent 6. Since cyclohelminthol IV (7) was isolated from the same producer fungus, H. velutinum yone96,11 the oxidation of 7 in vivo would likely generate 9. We consider that this cyclopropanation takes place through an enzymatic process, because 5 was isolated as a single diastereomer despite four diastereomers possible in this transformation. The electron-withdrawing carbonyl groups at the C2 and C5 in 5 renders the C6C7 double bond more electrophilic, which enables the second Michael addition of another 9 to C7, giving intermediate 11. The subsequent cyclopropanation eventually provides cyclohelminthol Ys 1-4. Although the stereochemical information at C6 is lost in 10, the steric repulsion between C4 and C8 may force their trans


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Scheme 1. Proposed biogenesis of 1-4.

configuration on the newly furnished cyclopropane F ring. The second cyclopropanation is assumed to take place nonenzymatically, since diastereomers 1-4 were detected in the similar level in the HPLC chromatogram (Figure S1). The configurations at C6, C7, and C1” are defined by the following two factors: 1) the Re-face/Si-face selection of the C6C7 double bond upon the approach of 9, and 2) the exo/endo-selection of 9 as shown in Figure 11. For example, the (6R,7R,1”R)-isomer 1 results from the process in which 9 approaches 5 from the Re-face of its C6C7 double bond in an endo manner. Although the C1’-C4’ side-chain blocks the front side of the C7C8 double bond in the figure, rotation of the C4C6 single bond may cause the Re-side/Si-side of the C6C7 double bond to switch, which enables the formation of all possible diastereomers. However, the s-trans conformations at C4C6 predominate in 5 (>95%) at room temperature according to our calculations, which contrasts with the similar yields of 1-4 obtained from the producer fungus.



Page 22 of 33

Figure 11. Mechanism for determining the configurations of C6 and C1”.

There is an alternative pathway that rationally explains the yields of 1-4. Homo-cyclopropanation of 9 (or its equivalent) may occur to afford the dimeric cyclopropane 12 (or its equivalent). Assuming a nonenzymatic reaction for this process, the reaction takes place in non-stereo selective manner to give 12 as a mixture of diastereomers. Since 9 is achiral, a nonenzymatic process should give them in racemic forms. In addition, the exo/endo selection would not be affected due to the low bulkiness of 9. Accordingly, four possible isomers of 12 might be produced in nearly equal amounts. The enzyme responsible for the formation of 5 would likely accept 12 as the substrate. This enzymatic cyclopropanation takes place in stereoselective manner to afford only 1S,15’R,16’S -configuration, but does not concern the configurations at C6, C7, and C1”, which allow to produce 1-4 in nearly equal amount. This lies on our assumption that the enzyme hardly discriminates the bulkiness and configuration of the cyclopentanespirocyclopropane moiety, because this part is far from the reaction site C1.

Biological properties. Preliminary biological experiments revealed that the (6S)-isomers 2 and 4 show potent cytotoxicity (IC50 11 and 10 µM, respectively) against COLO201,18 in contrast with the (6R)-isomers 1 and 3 (IC50


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>180 and 34 µM, respectively). Cyclohelminthol X (5) inhibited COLO201 in lower concentration (IC50 1.6 µM). Antifungal assay using Cochiobolus miyabeanus showed slightly different profile; 2 afforded the largest inhibition zone (approximately 30 mm) in the paper disk assay (50 µg on the 8.0 mm paper disk) on potato-dextrose-agar gel, while 1 and 4 resulted slightly smaller inhibition zones (both approximately 20 mm) under the same conditions (Figure S5). Cyclohelminthol Y3 (3) afforded only the trace of inhibition zone even in higher concentration (100 µg/disk). Since biological activities of 1-4 depend on their configurations, the specific molecule may respond to each activity.

The details are

under investigation in our laboratories.

Summary We

discovered

cyclohelminthols

Y1-Y4

(1-4),

unique

natural

products

containing

two

spirocyclopropanes in their structure. Small number of protons around the core-framework of these molecules renders NMR analyses impractical to obtain stereochemical information. Furthermore, their similar NMR profiles hinder the conventional statistical comparison employing absolute chemical shifts. However, DFT-based molecular modeling calculations revealed the stable conformations of 1-4, which helped us to interpret the 1H NMR signal behaviors of H6, H-7’, H-6”, and H-7” to the configurations. The assigned configuration was verified with theoretical chemical shift by focusing on the carbons with characteristic chemical shift dependence on the configurations. That was further verified by theoretical ECD calculations. The present study proves that DFT calculations are a powerful strategy in structural analysis.5,19 Furthermore, preliminary biological experiments revealed that their cytotoxicity against COLO201 cells and their antifungal activity against Cochiobolus miyabeanus depend on their configurations, which suggests the presence of a target molecule responsible for their activity.

Experimental Section



Page 24 of 33

General Experimental Procedures. The 1H (500 MHz) and 13C (125 MHz) NMR spectra were recorded on a JEOL JNM-ECX500 spectrometer. Tetramethylsilane (0 ppm) was used as the internal standard for both 1H and

13

C NMR spectra. UV spectra were obtained on a HITACHI U-2010 spectrometer.

Measurements of ECD spectra were performed on a JASCO J-1100 spectropolarimeter with a 10 mm length cell. The ∆ε values were obtained by dividing the output values (mdeg) by the concentration (mol/L) and 33000. Electrospray ionization time of flight (ESI-TOF) MS spectra were obtained from a HITACHI NanoFrontier LD spectrometer equipped with a HITACHI 2100 HPLC pump, a HITACHI L-2420 UV detector, a HITACHI L-2300 column oven, and a HITACHI L-2200 autosampler. Calibration was performed with tetrabutylammonium ion (m/z 242.2848) and reserpine (m/z 609.2807). IR spectra were obtained with a HORIBA FT-720 Fourier transform infrared spectrometer on a KBr cell. Chemicals and solvents were purchased from Wako Pure Chemical Industries and Sigma-Aldrich Co. LLC. Those were used without further purification. TLC analyses were carried out using Merck silica gel TLC silica gel 60 F254 plates (No. 5715). The column chromatography was carried out using Merck 707734. Analytical HPLC was performed using a Waters 1525 binary pump equipped with a Waters 996 photodiode array detector and a Waters 2707 auto sampler. Preparative HPLC was performed with a Waters 1525 binary Pump equipped with a Waters 2489 UV/visible detector. COSMOSIL Cholester columns (nacalai tesque, 4.6mml.D.×250mm, and 10mml.D.×250mm) were used for both analysis and separations in the final step. Conformation searches and chemical shift calculations were performed with Spartan’16 (Wavefunction, Irvine, CA, USA) using a PC (operating system: Windows7 Professional; CPU: Intel Xeon E5-1660 v2 processor, 3.70 GHz, 6 cores; RAM: 64 GB). Free energies of conformers and ECD spectra were calculated using Turbomole 7.0.1 with TmolX 7.3.2 (COSMOlogic GmbH & Co., Leverkusen Germany) on a PC workstation (operating system: CentOS 7.1.1; CPU: Intel Xeon E5-2687W V4, 3.0 GHz, 12 cores ×2; RAM: 256 GB). Reproduction of the ECD spectra were performed using Microsoft Excel2016 on a commercial PC (Windows 7).


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Fungus. Helminthosporium velutinum yone96 was isolated from the dead twigs of a woody plant on Yakushima island, Kagoshima prefecture, Japan in 2007. The fungus was deposited at the Genbank Project, NARO, Japan (ID: MAFF 243859).

Isolation and physical properties of 1-4. Helminthosporium veltinum yone96 was cultured (10 L) in the similar manner we obtained cyclohelminthol X (5).8 The culture broth was filtered, and the residual fungus body was suspended in MeOH (1.0 L). After 24 hours, the suspension was filtered, and the filtrate was concentrated in vacuo to give aqueous suspension (200 ml). The suspension was extracted with EtOAc (300 mL ×3) and organic layers were combined, washed with brine, dried over MgSO4, and concentrated in vacuo to give the crude extract (1.2 g). After re-dilution with EtOAc (20 ml), silica gel (20 g) was added and the solvent was removed with a rotary evaporator. The residual silica gel was placed on a silica gel column (80 g) and eluted with 0, 20, 40, 60, 80 and 100% EtOAc in hexane (300 mL each). The fraction eluted by 60% EtOAc/hexane was recovered to give a residue (350 mg) after concentration in vacuo. The residue was loaded on Sep-Pak ODS (10g) and eluted with 50, 60, 70, 80, 90, 100 % MeOH in H2O (200 mL each). The fraction eluted by 90 % MeOH/H2O was collected and the following concentration gave residue mainly containing 1-5 (58 mg) after concentration in vacuo. The residue was subjected to preparative HPLC (COSMOSIL, Cholester, 10 ×250 mm, CH3CN:H2O = 65:35, 5.0 ml/min, detected by UV at 302 nm) to give cyclohelminthol Y1 (1, 7.0 mg), cyclohelminthol Y2 (2, 11.0 mg), cyclohelminthol Y3 (3, 3.0 mg), cyclohelminthol Y4 (4, 6.0 mg) and cyclohelminthol X (5, 5.0 mg) as each an amorphous solid; tR 26.9, 28.0, 29.5, 31.3 and 14.3 min, respectively under the above conditions. The 1H NMR spectrum of 1 was identical to that we recently reported.8 Physical properties of 1. ESIMS (re lint. %, assignment): m/z 792.2304 (55, [M-OH]+, calcd. for C42H44Cl2NO10+: 792.2337), 810.2412 (100, [M+H]+, calcd. for C42H46Cl2NO11+: 810.2442), 832.2242 (50, [M+Na]+, calcd. for C42H45Cl2NNaO11+: 832.2262), IR (film) 3260, 2930, 2860, 1715, 1630 cm-1, 1

H NMR (500 MHz, CDCl3) 0.79 (3H, d, J = 6.2 Hz, H3-24’), 0.85 (3H, t, J = 6.7 Hz, H3-22’), 1.11 (m,



Page 26 of 33

H-18’), 1.25-1.28 (6H, H2-19’, H2-20’, H2-21’), 1.30-1.32 (2H, H-8’, H-10’), 1.45 (1H, m, H-7’), 1.48 (3H, d, J = 6.3, H3-8), 1.56 (1H, m, H-11’), 1.64 (1H, dd, J = 11.8, 14.0 Hz, H-6’), 1.74 (1H, m, H-18’), 1.92 (3H, dd, J = 1.2, 6.9, H3-1’), 1.92 (1H, m, H-12’), 1.96 (3H, dd, J = 1.5, 6.9 Hz, H3-8”), 2.00 (1H, m, H-10’), 2.14-2.16 (3H, H-6’, H-8’, H-17’), 2.39 (ddd, J = 4.6, 12.0, 14.5 Hz, H-17’), 2.46 (tt, J = 3.6, 12.2 Hz, H-9’), 2.83 (d, J = 8.5 Hz, H-6), 2.96 (1H, dq, J = 8.5, 6.3 Hz, H-7), 4.16 (1H, d, J = 11.6 Hz, H-13’), 6.40 (1H, dq, J = 15.6, 1.6 Hz, H-6”), 6.51 (1H, dq, J = 15.3, 1.2 Hz, H-3’), 6.92 (1H, dq, J = 15.3, 6.9 Hz, H-2’), 7.35 (1H, dq, J = 15.6, 6.9 Hz, H-7”), 7.76 (1H, br, NH),

13

C NMR (125 MHz,

CDCl3) 11.7 (C8), 14.2 (C22’), 18.9 (C1’), 20.9 (C8”), 21.9 (C17’), 22.1 (C24’), 22.6 (C21’), 27.5 (C18’), 29.5 (C19’), 31.5 (C20’), 32.1 (C6), 32.2 (C7), 33.3 (C8’), 36.4 (C11’), 37.6 (C10’), 42.7 (C7’), 42.7 (9’), 43.4 (1”), 44.8 (C6’), 47.5 (C1), 50.8 (C16’), 52.4 (C15’), 52.6 (C12’), 60.5 (C13’), 86.9 (C5’), 119.6 (C6”), 125.3 (C3’), 144.8 (C3”), 146.3 (C7”), 146.7 (C2’), 147.8 (C4”), 152.2 (C4), 153.1 (C3), 167.9 (C25’), 169.3 (C26’), 179.0 (C23’), 185.6 (C2), 189.7 (C5), 190.4 (C2”), 194.7 (C5”), 195.4 (C14’), 199.6 (C4’), UV (2.0×10-5 mol/mL, CH3CN, ε), λ 226 nm (44000), 316 nm (12000). Physical properties of 2. ESIMS (re lint. %, assignment): m/z 792.2301 (55, [M-OH]+, calcd. for C42H44Cl2NO10+: 792.2337), 810.2414 (70, [M+H]+, calcd. for C42H46Cl2NO11+: 810.2442), 832.2234 (100, [M+Na]+, calcd. for C42H45Cl2NNaO11+: 832.2262), IR (film) 3235, 2925, 2955, 1715, 1625 cm-1, 1


H-18’), 1.26-1.35 (8H, H-8’, H-10’, H2-19’, H2-20’, H2-21’), 1.45 (1H, m, H-7’), 1.51 (3H, d, J = 6.3, H3-8), 1.55 (1H, m, H-11’), 1.65 (1H, dd, J = 11.4, 13.3 Hz, H-6’), 1.67 (1H, m, H-18’), 1.92 (3H, dd, J = 1.5, 7.0, H3-1’), 1.93 (1H, m, H-12’), 2.01 (1H, m, H-10’), 2.04 (3H, dd, J = 1.5, 6.9 Hz, H3-8”), 2.14-2.18 (3H, H-6’, H-8’, H-17’), 2.39 (ddd, J = 4.6, 12.0, 14.5 Hz, H-17’), 2.46 (tt, J = 3.4, 12.1 Hz, H-9’), 2.59 (1H, dq, J = 8.5, 6.3 Hz, H-7), 2.72 (d, J = 8.5 Hz, H-6), 4.16 (1H, d, J = 11.6 Hz, H-13’), 6.49 (1H, dq, J = 15.9, 1.5 Hz, H-6”), 6.52 (1H, dq, J = 15.3, 1.5 Hz, H-3’), 6.97 (1H, dq, J = 15.3, 7.0 Hz, H-2’), 7.50 (1H, dq, J = 15.9, 6.9 Hz, H-7”), 7.55 (1H, br, NH), 13C NMR (125 MHz, CDCl3) 11.6 (C8), 14.2 (C22’), 18.9 (C1’), 20.9 (C8”), 21.6 (C17’), 22.0 (C24’), 22.7 (C21’), 27.2 (C18’), 29.5


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(C19’), 31.1 (C6), 31.5 (C20’), 32.2 (C7), 33.3 (C8’), 36.4 (C11’), 37.6 (C10’), 41.3 (1”), 42.6 (C7’), 42.7 (C9’), 44.8 (C6’), 47.1 (C1), 50.9 (C16’), 52.5 (C15’), 52.7 (12’), 60.3 (C13’), 87.0 (C5’), 119.8 (C6”), 125.3 (C3’), 145.0 (C3”), 146.0 (C7”), 146.6 (C2’), 148.2 (C4”), 151.9 (C4), 153.9 (C3), 167.7 (C25’), 168.9 (C26’), 178.8 (C23’), 185.8 (C2), 190.7 (C5), 190.8 (C2”), 194.6 (C5”), 195.3 (C14’), 199.7 (C4’), UV (1.6×10-5 mol/mL, CH3CN, ε), λ 226 nm (41000), 316 nm (13000). Physical properties of 3. ESIMS (re lint. %, assignment): m/z 792.2310 (55, [M-OH]+, calcd. for C42H44Cl2NO10+: 792.2337), 810.2417 (100, [M+H]+, calcd. for C42H46Cl2NO11+: 810.2442), 832.2246 (55, [M+Na]+, calcd. for C42H45Cl2NNaO11+: 832.2262), IR (film) 3300, 2930, 2855, 1715, 1630 cm-1, 1


H-18’), 1.25-1.33 (8H, H-8’, H-10’,H2-19’, H2-20’, H2-21’), 1.45 (1H, m, H-7’), 1.48 (3H, d, J = 6.3, H3-8), 1.53 (1H, m, H-11’), 1.65 (1H, dd, J = 11.6, 13.4 Hz, H-6’), 1.67 (1H, m, H-18’), 1.91 (3H, dd, J = 1.5, 7.0, H3-1’), 1.94 (1H, m, H-12’), 2.00 (1H, m, H-10’), 2.04 (3H, dd, J = 1.6, 7.0 Hz, H3-8”), 2.12-2.17 (3H, H-6’, H-8’, H-17’), 2.43 (1H, m, H-17’), 2.46 (tt, J = 3.7, 12.4 Hz, H-9’), 2.85 (d, J = 8.5 Hz, H-6), 2.98 (1H, dq, J = 8.6, 6.3 Hz, H-7), 4.14 (1H, d, J = 11.6 Hz, H-13’), 6.48 (1H, dq, J = 15.9, 1.5 Hz, H-6”), 6.49 (1H, dq, J = 15.3, 1.5 Hz, H-3’), 6.91 (1H, dq, J = 15.3, 7.0 Hz, H-2’), 7.50 (1H, dq, J = 15.9, 7.0 Hz, H-7”), 7.65 (1H, br, NH),

13

C NMR (125 MHz, CDCl3) 11.7 (C8), 14.2 (C22’), 18.9

(C1’), 20.9 (C8”), 21.7 (C17’), 22.0 (C24’), 22.6 (C21’), 27.3 (C18’), 29.5 (C19’), 31.5 (C20’), 32.1 (C6), 32.3 (C7), 33.4 (C8’), 36.6 (C11’), 37.6 (C10’), 42.7 (C9’), 42.8 (C7’), 43.2 (C1”), 44.9 (C6’), 47.3 (C1), 50.9 (C16’), 52.2 (C15’), 52.3 (C12’), 60.5 (C13’), 86.9 (C5’), 119.7 (C6”), 125.3 (C3’), 145.0 (C3”), 146.1 (C7”), 146.4 (C2’), 148.1 (C4”), 151.6 (C4), 153.2 (C3), 167.9 (C25’), 169.2 (C26’), 178.6 (C23’), 185.2 (C2), 189.9 (C5), 190.8 (C2”), 194.4 (C5”), 195.2 (C14’), 199.6 (C4’), UV (8.0×10-6 mol/mL, CH3CN, ε), λ 226 nm (43000), 316 nm (11000). Physical properties of 4. ESIMS (re lint. %, assignment): m/z 792.2307 (55, [M-OH]+, calcd. for C42H44Cl2NO10+: 792.2337), 810.2416 (100, [M+H]+, calcd. for C42H46Cl2NO11+: 810.2442), 832.2241 (95, [M+Na]+, calcd. for C42H45Cl2NNaO11+: 832.2262), IR (film) 3260, 2930, 2860, 1715, 1625 cm-1,


The Journal of Organic Chemistry 1

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Page 28 of 33


H-18’), 1.26-1.30 (8H, H-8’, H-10’,H2-19’, H2-20’, H2-21’), 1.43 (1H, m, H-7’), 1.53 (3H, d, J = 6.3, H3-8), 1.53 (1H, m, H-11’), 1.65 (1H, dd, J = 11.6, 13.4 Hz, H-6’), 1.70 (1H, m, H-18’), 1.89 (1H, m, H-12’), 1.92 (3H, dd, J = 1.6, 6.9, H3-1’), 1.99 (1H, m, H-10’), 2.00 (3H, dd, J = 1.6, 7.3 Hz, H3-8”), 2.08-2.16 (3H, H-6’, H-8’, H-17’), 2.34 (1H, ddd, J = 4.4, 12.1, 14.3 Hz, H-17’), 2.45 (tt, J = 3.5, 12.0 Hz, H-9’), 2.50 (1H, dq, J = 8.5, 6.3 Hz, H-7), 2.67 (d, J = 8.5 Hz, H-6), 4.14 (1H, d, J = 11.5 Hz, H-13’), 6.41 (1H, dq, J = 15.9, 1.6 Hz, H-6”), 6.52 (1H, dq, J = 15.4, 1.6 Hz, H-3’), 6.96 (1H, dq, J = 15.4, 6.9 Hz, H-2’), 7.35 (1H, dq, J = 15.9, 7.3 Hz, H-7”), 7.77 (1H, br, NH),

13

C NMR (125 MHz,

CDCl3) 11.5 (C8), 14.2 (C22’), 18.9 (C1’), 20.9 (C8”), 21.8 (C17’), 22.0 (C24’), 22.6 (C21’), 27.4 (C18’), 29.5 (C19’), 31.1 (C6), 31.5 (C20’), 31.7 (C7), 33.3 (C8’), 36.4 (C11’), 37.6 (C10’), 42.6 (C7’), 42.8 (C9’), 41.2 (C1”), 44.9 (C6’), 47.0 (C1), 50.7 (C16’), 52.7 (C15’), 52.6 (C12’), 60.5 (C13’), 86.9 (C5’), 119.4 (C6”), 125.2 (C3’), 144.9 (C3”), 146.7 (C7”), 146.7 (C2’), 148.0 (C4”), 152.5 (C4), 153.9 (C3), 167.8 (C25’), 169.2 (C26’), 180.1 (C23’), 186.0 (C2), 190.7 (C5), 190.7 (C2”), 195.0 (C5”), 195.3 (C14’), 199.7 (C4’), UV (1.8×10-5 mol/mL, CH3CN, ε), λ 226 nm (42000), 316 nm (11000).

Calculations.

Models Mo-S-RR, Mo-S-RS, Mo-S-SR, Mo-S-SS, Mo-R-RR, Mo-R-RS, Mo-R-SR,

and Mo-R-SS were built in the program and conformational searches were performed with MMF. In the conformational search, the propyl group corresponding to CC7C19 was fixed in an expanded conformation (∠C16’C17’C18’C19’ = 180°) to reduce the number of initial conformations. Suggested stable conformers were optimized successively with HF/321G and ωB97X-D/6-31G*. Duplicate conformers were manually removed and missing conformers were manually added in each step. The relative free energy of each conformer was obtained by BHLYP/def2-SVP structural optimization and following vibrational analysis. The chemical shifts of each conformers were calculated with ωB97X-D/6-31G*, wB97-X-D/6-311G*, B3LYP/6-31G, and EDF2/6-31G*. The obtained chemical shifts were corrected using the Boltzmann distribution based on free energies to give series of


13

C and

Page 29 of 33 1

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H theoretical chemical shifts. The results obtained by the latter three are shown in the Supplemental

data (S-42). ECD calculations were carried out for stable conformers occupying more than 90% population based on BHLYP/def2-SVP. Fifty excitations were examined in these calculations. The UV and ECD spectra of each conformer were constructed based on frequencies and rotary strengths using the NORMDIST function in Microsoft Excel 2016. The wavelengths of the spectra were corrected (+22 nm). Theoretical ECD spectra were obtained after correction of the conformational distribution based on the free energy. The intensity of the theoretical ECD spectrum Mo-S-SR was normalized with that of the experimental spectrum of 2 at 236 nm, and the same scale was used for the other ECD spectra of the models with (1S)-configuration. The same scale was used also for models with (1R)-configuration. Mo-S-RR-hex was built by elongating the propyl group of Mo-S-RR to hexyl group. Conformational search by perturbing only the hexyl group using MMFF afforded 193 tentative stable conformers. The characteristic

conformers

were

chosen

and

structurally

optimized

and

calculated

with

wB-97X-D/6-31G* to find conf-A and conf-B shown in Figure 8.

Cytotoxicity assay.

The effect of compound 1-4 on human colon adenocarcinoma (COLO 201) cell

proliferation was measured by WST-1 assay.18 COLO 201 cells were cultured in RPMI medium (5×103 cells/well) containing compounds 1-4 in 96-well tissue culture plates with 0.01, 0.1, 10, 100, 1000 µg/mL concentrations. After 24 h, 10 µl of WST-1 reagent was added to each well and the absorbance measured at 450 nm using a titer-plate reader.

Antifungal assay.

Samples (10, 50, and 100 µg) in 50 µL ethanol were loaded on 8.0 mm dimeter

paper disks and these were dried in the vacuum conditions for 30 min. These were placed on the potato-dextrose-agar culture medium containing Cochiobolus miyabeanus spores on the Petri dish. After standing at 25 ºC for a week, the inhibition zones were measured with a ruler.



Page 30 of 33

ASSOCIATED CONTENT Supporting Information. Experimental and computational details, NMR spectra, NMR assignments, calculated chemical shifts, and stable conformers of the models. This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author * [email protected]. ORCID Masaru Hashimoto: 0000-0002-4508-2105 Hayato Maeda: 0000-0002-8546-6250 Kazuaki Tanaka: 0000-0002-7037-0774 Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENT We are grateful to Dr. Warren J. Hehre of Wavefunction Inc. for his kind advices. Part of this work was supported by a Grant-in-Aid for Scientific Research (B) (15H04491), a Grant-in-Aid for Scientific Research (C) (16K07474), and a Grant-in-Aid for Challenging Exploratory Research (16K14910) from the Japan Society for the Promotion of Science (JSPS). Biological experiments were supported by Hirosaki University Grant for Exploratory Research by Young Scientists and Newly-appointed Scientists.

The authors would like to thank go (www.enago.jp) for the English language review.


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