Discovery of a parallel family of euglenatide analogs in Euglena gracilis

1 Introduction

Polyketides and nonribosomal peptides are complex natural products that are produced on large enzymatic assembly lines known as polyketide synthases (PKSs) and nonribosomal peptide synthetases (NRPSs), respectively [1, 2]. Many of these natural products have been developed into important therapeutics, and thus discovery of structurally novel polyketides and nonribosomal peptides from underexplored sources is of interest. One of these potential sources is the unicellular flagellate algae Euglena gracilis, which can grow through photoautotrophic growth, using light as energy, heterotrophic feeding, using an external carbon source as energy, or a mixture of the two growth modes [3]. Although E. gracilis has been studied extensively as a potential source of food and biofuel, relatively little is known about its natural product biosynthetic potential. Even though the genome of E. gracilis has been only partially sequenced, these efforts indicate that the genome includes a large number of PKSs and NRPSs and other enzymes involved in secondary metabolism [4, 5].

Recently, E. gracilis was shown to produce a family of hybrid polyketide-nonribosomal peptides, euglenatides A–E, with euglenatide B being the most abundantly produced (Fig. 1) [6]. Interestingly, the euglenatides are structurally similar to a family of hybrid polyketide-nonribosomal peptides previously identified in the nematode Caenorhabditis elegans [7-9]. The euglenatides have antiproliferative activity against multiple cancer cell lines in the mid-nanomolar range and several fungal pathogens in the low micromolar range, but have no activity against bacteria [6]. A survey of additional Euglena species suggests that they also produce similar natural products. Interestingly, production of the eugenatides did not occur in E. gracilis grown in rich medium and only occurred in E. gracilis grown in minimal medium containing specific amino acids, either glutamate or asparagine, under photosynthetic conditions [6].

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Here, in analyzing extracts from E. gracilis, we discovered a new, parallel family of analogs that are produced in roughly equivalent amounts to euglenatides A–C. In comparison to the euglenatides, these analogs have the same molecular weights. However, the triene that is present in the euglenatides is disrupted in the analogs in that one of the double bonds is shifted by two carbons. The analogs are tenfold less bioactive against A549 lung adenocarcinoma cells than the euglenatides, indicating that the triene motif is important for activity.

2 Results and discussion

In order to determine whether E. gracilis produces additional euglenatides or euglenatide biosynthetic intermediates, we cultured E. gracilis using similar methods to those reported by Aldholmi et al. [6]; that is, we used minimal medium in which the carbon source was sodium acetate and the nitrogen source was glutamate, and we cultured the algae under photosynthetic conditions. In our hands, euglenatide B was more than tenfold more abundant than euglenatides A and C, euglenatide D was only minorly present, and euglenatide E was not detectable. Using molecular networking and LC–MS analysis of E. gracilis crude extracts, we noted that E. gracilis produces not only the euglenatides, but also a corresponding set of euglenatide analogs that we named euglenatides A₂-C₂ (Fig. 2) [10, 11]. These analogs have the same exact masses and MS fragmentation patterns as euglenatides A-C (Fig. 3, Fig. S1), but are slightly more polar based on their retention times (Fig. 2b). Unlike euglenatides A–C, which have a UV–Vis chromophore characteristic of a conjugated triene, euglenatides A₂–C₂ do not absorb at 280 nm and do not possess a conjugated triene (Fig. 2b, c). Each euglenatide is produced at roughly similar levels as its corresponding euglenatide analog. We have not been able to enhance the production of the euglenatide analogs relative to the euglenatides by harvesting the E. gracilis culture at a particular time point or utilizing a particular nitrogen source (glutamate versus asparagine).

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To further characterize the chemical structures of the euglenatide analogs, we purified the most abundant one, euglenatide B₂, as well as euglenatide B for comparison purposes. Chromatographic fractionation of defatted methanolic extracts of E. gracilis by reversed phase C18 flash chromatography, LH-20 Sephadex chromatography, and semipreparative HPLC allowed the purification of the two euglenatides (Fig. S2). Analysis of the 1D and 2D NMR (DMSO-d₆) data for euglenatide B (Table 1; Table S1 and Figs. S3, S4) in comparison to literature [6] confirmed its identification (Fig. 1).

Comparison of the 1D NMR (DMSO-d₆) data for euglenatide B₂ with that of euglenatide B disclosed common peptide and polyketide substructures, with the significant differences being the presence of two extra allylic methylene resonances in euglenatide B₂ at C-29 (δ_2H 2.10, m; δ_C 32.2) and C-30 (δ_2H 2.04, m; δ_C 31.7), accompanied by a change in the pattern of olefinic methine resonances (Fig. 4, Table 1; Table S2, and Figs. S5–S13). These NMR changes in euglenatide B₂ suggested that one double bond (Δ³¹) is separated from the diene (Δ²⁵ and Δ²⁷) by two methylenes at C-29 and C-30, which explains the absence of the conjugated triene chromophore in the UV–Vis spectrum. Diagnostic 2D NMR correlations (shown in Fig. 4) confirmed the double bond shift and the molecular connectivity of euglenatide B₂, while NMR similarities with euglenatide B, including comparable J values, and biosynthetic considerations suggested a common absolute configuration. Finally, Double Quantum Filtered (DQF)-COSY NMR analysis of euglenatide B₂ allowed the assignment of the Δ³¹ double bond geometry as E (trans) based on a coupling constant (J_{31, 32}) of 16.2 Hz (Figs. S10, S11). This assignment is further supported with the upfield chemical shifts of the allylic methylenes at C-30 (δ_C 31.7) and C-33 (δ_C 31.9) [12-15]. Thus, we conclude that the double bond at C-29 in euglenatide B has been shifted by two carbons to C-31 in euglenatide B₂ (Fig. 5a).

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Similarly, our data suggest that euglenatides A₂ and C₂ are exactly the same as euglenatides A and C, but one of the double bonds in the triene has been shifted by two carbons (Fig. 5a). Specifically, our data show that (1) euglenatides A-C and A₂-C₂ have the same exact masses and MS/MS fragmentation patterns (Fig. 3, Fig. S1), (2) the conjugated triene chromophore in euglenatides A-C is absent in A₂-C₂ (Fig. 2b, c), and (3) the euglenatides and their analogs have regularly spaced elution times by LC–MS (Fig. 2b). Several trials were made to purify euglenatides A, A₂, C and C₂ for NMR analysis, but unfortunately their low abundance and co-elution with other metabolites hindered their purification.

Previously, it was shown that euglenatide B had antiproliferative activity in the mid-nanomolar range against mammalian cancer cell lines, including A549 lung adenocarcinoma cells. To test the antiproliferative activity of the euglenatide analogs and thus the importance of the conjugated triene motif, we tested the activity of both euglenatides B and B₂ against A549 cells (Fig. 5b). Our data confirm the potency of euglenatide B in the mid-nanomolar range, with an IC₅₀ of 358 nM, in comparison to the previously reported IC₅₀ of 773 nM. The data also show that euglenatide B₂ is about tenfold less potent than euglenatide B, with an IC₅₀ of 2987 nM. Thus, the conjugated triene is quite important for the activity of the euglenatides, and this information should inform future efforts to characterize the structure–activity relationships of the euglenatides.

3 Conclusions

Here, we show that E. gracilis produces not only the previously reported euglenatides A–C, but also a related family of euglenatide analogs, euglenatides A₂–C₂, in roughly equal amounts. In the euglenatide analogs, one of the double bonds in the conjugated triene has been shifted by two carbons, thereby disrupting the chromophore. The euglenatide B₂ analog was fully characterized by NMR spectroscopy and mass spectrometry and shown to have antiproliferative activity that is roughly tenfold less than euglenatide B against A549 lung adenocarcinoma cells, indicating that the conjugated triene is important to the activity of the euglenatides. This information helps to characterize the structure–activity relationships of the euglenatides.

4 General experimental procedures

4.1 Cultivation of E. gracilis

E. gracilis var. saccharophila (CCAP 1224/5T) was obtained from the Culture Collection of Algae and Protozoa (United Kingdom). An axenic stock was generated by plating the algae on plates containing a 1:1 mix of E. gracilis medium (EG) and Jaworski's medium (JM) and 15 g/L of agar and picking a single colony. EG and JM were made using a previously described method [6]. E. gracilis was seeded typically in a 1:10 ratio into EG: JM medium in an Erlenmeyer flask and incubated at room temperature under a daylight lamp for 7d on a 150 rpm shaker. The lamp was set at a 12 h light/12 h dark cycle with an intensity of 2000 lumens.

4.2 Large scale cultivation of E. gracilis for euglenatide production

The EG: JM culture of E. gracilis was adjusted to 4 g/L (Abs 0.1 at 740 nm). 75 mL of this suspension was harvested by centrifugation at 3000 g at 4 ℃ for 20 min, washed three times with sterile Milli-Q water to remove any trace components of EG: JM medium, and seeded into 1.5L of synthetic medium (JM + 1 g/L sodium acetate trihydrate + 0.1 g/L calcium chloride) with 30 mM Glu (pH adjusted to 4.8 prior to autoclaving) in each bioreactor. The culture was incubated at room temperature under a daylight lamp at 12 h light/12 h dark cycle for 10d before collecting the cells for further extraction. Approximately 12 L of culture was generated in all. The cultures were centrifuged at 3000 g for 20 min, and the culture medium was discarded.

4.3 Purification of euglenatides

12L worth of cells were extracted with 2L of 90% MeOH. The 90% MeOH was extracted with 2L of hexanes. The MeOH layer was evaporated to yield the methanolic extractives (1.6 g) that were further fractionated on a C18 reversed phase flash column (300 mm × 46 mm, 80 g silica) with a 10% (200 mL) step-wise gradient elution from 20 to 100% MeCN. HPLC–DAD-MS analysis revealed the fraction eluting with 60% MeCN to be rich in euglenatides (Fig. S2). This fraction (80 mg) was chromatographed on an LH-20 Sephadex gel column, eluting with MeOH. Based on HPLC–DAD-MS analysis, subfractions containing mainly metabolites of m/z(–) 808 (euglenatide B and its analog) were combined (25 mg) and subjected to repeated semi-preparative reversed phase HPLC (HS C18-10, 250 mm × 10 mm, 10 μm, 3 mL/min) with an isocratic elution over 35 min with 34% MeCN/H₂O to yield euglenatide B (3 mg) and its new isomer, euglenatide B₂ (3 mg) (Fig. S2).

4.4 Euglenatide B

White powder; NMR (600 MHz, DMSO-d₆) see Table 1 and Table S1 and Figs. S3, S4; ESI(−)MS m/z 808 [M − H]⁻ and m/z 844 [M − H + Cl]⁻; HRESI(+)MS m/z 810.4618 [M + H]⁺ (calcd for C₃₈H₆₄N₇O₁₂, 810.4613).

4.5 Euglenatide B₂

White powder; NMR (600 MHz, DMSO-d₆) see Table 1 and Table S2 and Figures S5-S13; ESI(−)MS m/z 808 [M − H]⁻ and m/z 844 [M − H + Cl]⁻; HRESI(+)MS m/z 810.4617 [M + H]⁺ (calcd for C₃₈H₆₄N₇O₁₂, 810.4613).

4.6 Antiproliferation assay

A549 cells were cultured in DMEM medium supplemented with 10% FBS, 100 U/mL penicillin and 100 μg/μL streptomycin. 6 × 10³ A549 cells per 100 μL were seeded in a 96-well plate and incubated at 37℃ with 5% CO₂. After 22 h of incubation, each well was treated with 1 μL of dimethyl sulfoxide (DMSO) or 1 μL of euglenatide B or euglenatide B₂ in DMSO at final concentration of 10⁵, 10⁴, 10³, 10², 10, or 1 nM. Each concentration was tested in triplicate. The plate was incubated for 72 h, and cell viability was tested with the MTS assay (Promega). IC₅₀ values were calculated using GraphPad. Wells with only culture media were used as a blank, and wells seeded with A549 cells and treated with 1% DMSO were used as a negative control. The viability of treated A549 cells relative to the negative control was calculated after a blank correction.

Notes

Acknowledgements

The authors would like to thank J. Rocca for his assistance in acquiring NMR spectra, I. Ghiviriga for additional NMR advice, and R. Yu for help preparing samples. NMR spectra were acquired in the McKnight Brain Institute at the National High Magnetic Field Laboratory's Advanced Magnetic Resonance Imaging and Spectroscopy (AMRIS) Facility, which is supported by National Science Foundation Cooperative Agreement DMR-2128556 and the State of Florida. Mass spectrometry analysis was performed at the University of Florida's Mass Spectrometry Research and Education Center, which is supported in part by the National Institutes of Health (S10 OD030250-01A1). This work was supported by a grant from the National Institutes of Health (R35 GM144076 to R.A.B.).

Author contributions

A.H.E. purified compounds and elucidated their structures. X.K. cultured algae, purified compounds, and performed mammalian cell culture assays. S.R. tested growth conditions for enhancing analog production. D.V.P. performed mass spectrometry and molecular networking analysis. R.A.B. conceived the study, and R.A.B. and A.H.E. wrote the manuscript, which was approved by all authors.

Availability of data and materials

The authors will provide access to all datasets upon request.

Declarations

Competing interests

The authors declare no conflict of interest.