Synthesis of acylglycerol derivatives by mechanochemistry

  1. Karen J. Ardila-Fierro1ORCID Logo,
  2. Andrij Pich2,3,4ORCID Logo,
  3. Marc Spehr5ORCID Logo,
  4. José G. Hernández1ORCID Logo and
  5. Carsten Bolm1ORCID Logo

1Institute of Organic Chemistry, RWTH Aachen University, Landoltweg 1, D-52074 Aachen, Germany
2Functional and Interactive Polymers, Institute of Technical and Macromolecular Chemistry, RWTH Aachen University, Worringerweg 2, 52074 Aachen, Germany
3DWI-Leibniz-Institute for Interactive Materials, Forckenbeckstrasse 50, D-52074 Aachen, Germany

4Aachen Maastricht Institute for Biobased Materials (AMIBM), Maastricht University, Brightlands Chemelot Campus, Urmonderbaan22, 6167 RD Geleen, The Netherlands
5Department of Chemosensation, Institute for Biology II, RWTH Aachen University, D-52074 Aachen, Germany

  1. Corresponding author email

This article is part of the thematic issue "Mechanochemistry II".

Associate Editor: L. Vaccaro
Beilstein J. Org. Chem. 2019, 15, 811–817. doi:10.3762/bjoc.15.78
Received 10 Jan 2019, Accepted 18 Mar 2019, Published 29 Mar 2019

Abstract

In recent times, many biologically relevant building blocks such as amino acids, peptides, saccharides, nucleotides and nucleosides, etc. have been prepared by mechanochemical synthesis. However, mechanosynthesis of lipids by ball milling techniques has remained essentially unexplored. In this work, a multistep synthetic route to access mono- and diacylglycerol derivatives by mechanochemistry has been realized, including the synthesis of diacylglycerol-coumarin conjugates.

Keywords: ball mill; coumarin; diacylglycerols; lipids; mechanochemistry

Introduction

In addition to being guided by chemical signals, cells respond to mechanical cues by sensing and transducing external mechanical inputs into biochemical and electrical signals [1]. Consequently, every time a cell is subjected to mechanical loads, the biomolecules that constitute the cell do also experience the effects of the mechanical forces. For example, from the moment a nascent peptide begins growing in the ribosome, such peptide experiences mechanical signals that regulate the speed of protein synthesis [2]. Not surprisingly, the natural ability of peptides to endure mechanical stress in nature has allured scientists to evaluate the mechanical stability of proteins by using single-molecule nanomechanical techniques (e.g., magnetic and optical tweezers or atomic force microscopy) [3,4]. Additionally, the resilience of the peptide bond to mechanical loads has led to mechanoenzymatic transformations [5-7], and to synthesize amino acid derivatives [8-10] and peptides [11-13] by ball milling and extrusion techniques. Similarly, mechanochemical derivatizations of sugars and sugar derivatives such as cyclodextrins (CDs) have proven compatible with the use of ball mills. These reports showed advantages such as higher selectivity by ball milling compared to classic solution methods and the possibility to effectively react CDs and reactants of different solubility profiles [14-17]. The compatibility of synthesizing biologically relevant building blocks with mechanochemistry has further been shown by the recent mechanochemical protocols to transform nucleoside and nucleotide substrates (Figure 1) [18,19].

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Figure 1: Biologically relevant molecules made, used or derivatized by mechanochemistry.

On the other hand, reports on mechanochemical protocols for the synthesis or derivatization of lipids are scarce [20,21]. Among the variety of amphipathic or hydrophobic small molecules that exhibit a lipid structure, diacylglycerols (DAGs) are important due to their signaling functions in cells (DAG signaling) [22-24]. Structurally, DAGs are glycerolipids containing two fatty acids esterified to the alcohol glycerol (Figure 2).

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Figure 2: Isomeric diacyl-sn-glycerols (DAGs).

Biological routes that lead to the formation of DAGs include enzymatic degradation of glycerophospholipids and lipolysis of triacylglycerols (TAGs) [22,25]. However, due to the structural diversity of fatty acids present in acylglycerols and to the small structural differences among these fatty acids (e.g., chain length, degree of unsaturation, double bond position or stereochemistry), the access to pure DAGs and TAGs from natural sources by extraction is cumbersome. Alternatively, protected DAGs 5 can be chemically synthesized starting from glycerol [26] or glycidol [27], but either synthetic alternative involves multiple preparative steps in organic solvents (e.g., CH2Cl2, THF, Et2O). These considerations led us to explore a mechanochemical multistep route for the synthesis of protected DAGs 5 starting from glycidol (1) through the installation of a hydroxy protecting group, followed by epoxide ring-opening and esterification reactions with fatty acids 3 (Scheme 1).

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Scheme 1: Synthetic route to access protected DAGs; PG = protecting group.

If successful, developing a multistep approach to prepare DAGs would contribute to the expansion of synthetic mechanochemical methodologies in ball mills [28-31], which are often limited to single-step transformations. Additionally, synthesizing lipid structures mechanochemically would complement the preparation of biologically relevant building blocks (amino acids, peptides, saccharides, nucleosides, etc.) by mechanochemistry, thereby highlighting the importance of mechanical forces in the chemistry of life.

Results and Discussion

To commence, we focused on the synthesis of tert-butyldimethylsilyl glycidyl ether (2) by reacting glycidol (1; 0.67 mmol) and tert-butyldimethylsilyl chloride (TBDMSCl) in the presence of imidazole in a mixer mill (MM, Scheme 2) [32].

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Scheme 2: Protection of glycidol (1) with TBDMSCl in the ball mill. MM = mixer mill, PBM = planetary ball mill.

After 2 h of milling at 25 Hz full consumption of the starting materials was observed by thin-layer chromatography (TLC), with exclusive formation of the corresponding silyl ether 2. However, the high volatility of 2 during posterior vacuum drying processes prevented the isolation of the TBDMS-protected glycidol 2 in higher yields after separation by column chromatography. A 5-fold scaled up version of the reaction using 3.37 mmol of 1 was carried out in a planetary ball mill (PBM) using 45 mL milling containers at 600 rpm, under otherwise identical conditions to afford product 2 in a similar yield.

With TBDMS-protected glycidol 2 in our hands, a selective epoxide ring-opening reaction with fatty acids 3 leading to the formation of the corresponding sn-1,3-protected monoacylglycerols (MAGs 4) was attempted (Scheme 3). Initially, commonly used solution-based protocols were tested in the ball mill [33]. For example, 2 was reacted with stearic acid (3a) in the presence of amines such as pyridine or tributylamine. However, the analysis of the reaction mixture only showed unreacted starting materials. In previous work, an acceleration of the oxirane ring-opening reaction with carboxylic acids [34] or alcohols [35] by using Lewis acid catalysts such as iron(III) chloride or bismuth(III) triflate was reported. However, the implementation of these protocols in the ball mill only led to trace amounts (less than 5% yield) of the protected monoacylglycerol 4a. Finally, one of the well-established Jacobsen catalysts for the epoxide ring-opening reaction of 2 with stearic acid (3a) was evaluated (Scheme 3a).

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Scheme 3: Cobalt-catalyzed epoxide ring-opening in the ball mill.

Specifically, we focused on the use of Jacobsen cobalt(II)-salen complex (S,S)-cat (Scheme 3b), since similar salen complexes had originally been reported to facilitate epoxide ring-opening reactions with carboxylic acids as nucleophiles [36]. Moreover, salen complexes endure mechanochemical conditions, as proven during their preparation in ball mills [37]. In addition, various related Jacobsen salen complexes have shown catalytic activity under solvent-free conditions [38]. Collectively, these precedents made this synthetic route a promising one to mechanochemically access MAGs by ball milling.

Experimentally, we attempted the cobalt-catalyzed epoxide ring-opening reaction by milling 2 with stearic acid (3a) in the presence of (S,S)-cat (2.5 mol %) and N,N-diisopropylethylamine (DIPEA; 1.0 equiv, Scheme 3a). Mechanistically, it is known that Co(II) complex (S,S)-cat is catalytically inactive and its oxidation is required to facilitate the reaction [39]. Aware of this, we began by relying on the atmospheric dioxygen inside the milling container to oxidize complex (S,S)-cat. This approach has been applied before in the mechanochemical synthesis of Cu–carbene complexes from N,N-diarylimidazolium salts, dioxygen and metallic copper [40], which involved a mechanochemical reaction with gaseous reagents [41]. Pleasingly, this time the reaction afforded product 4a, although the yield remained low (59%), even after three hours of milling. In order to improve the yield, we carried out the activation of the Co(II) complex (S,S)-cat by milling under a balloon pressure of dioxygen. Now, the yield of 4a was boosted up to 84% yield (for a representation of the set up used and experimental details, see Scheme 3c and Supporting Information File 1). Analysis of the reaction mixture by 1H NMR revealed that the epoxide ring-opening had occurred preferentially to give sn-1,3-protected monoacylglycerol 4a over its regioisomer counterpart sn-2,3-protected monoacylglycerol 4a’ (4a/4a’ 3.6:1; for details, see Supporting Information File 1). However, purification of 4a by column chromatography on SiO2 favored acyl migration in 4a, thereby dropping the yield of 4a by increasing the amount of the isomeric sn-2,3-protected monoacylglycerol 4a’ (Figure 2) [42]. Typically, cobalt complex (S,S)-cat has been used for kinetic resolution of racemic epoxides, for which the maximum theoretical yield of the reaction is 50%. Here, however, the yield of the sn-1,3-protected monoacylglycerol was high, and consequently we expected the enantiomeric excess of the product to be low. This, assumption was confirmed by analysis of the sample by high-performance liquid chromatography-chiral stationary phase (CSP-HPLC, for more details, see Supporting Information File 1). Access to enantiopure MAGs could be achieved under similar reaction conditions by starting from optically active commercially available silyl-protected glycidol derivatives [27,43].

Next, we targeted the mechanosynthesis of the DAGs by reacting MAG 4a and fatty acids 3 in the ball mill. Such esterifications required the activation of 3 with N,N’-dicyclohexylcarbodiimide (DCC), thereby complementing other recently developed solvent-free carboxylic acid activations towards amidation or esterification reactions by ball milling [44]. In practice, we milled a mixture of MAG 4a, stearic acid (3a), DCC and 4-dimethylaminopyridine (DMAP) at 25 Hz for 2 h in a mixer mill. Separation of the product by column chromatography gave DAG 5a in 97% yield (Scheme 4).

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Scheme 4: Mechanosynthesis of DAGs 5.

Alternatively, 5a could be prepared following a one-pot two-step approach in the ball mill by beginning with the cobalt-catalyzed epoxide-ring opening of 2 with 3a, followed immediately by the esterification of the corresponding MAG 4a with stearic acid (3a), DCC and DMAP. Although successful, this strategy led to DAG 5a in lower yield (50%). Then, in order to expand the library of DAGs, other fatty acids containing various degrees of unsaturation and chain length were tested. For instance, esterification of MAG 4a with oleic acid (3b), linoleic acid (3c), and arachidonic acid (3d) underwent smoothly in the ball mill affording DAGs 5bd in yields up to 91% (Scheme 4). Particularly interesting was the formation of DAG (18:0/20:4) 5d, an important lipidic backbone present in the biologically relevant phosphatidylinositol 4,5-bisphosphate (PIP2) [22]. In fact, diacylglycerols have proven to play vital roles in regulation of lipid bilayer and in the catalytic action of various membrane-related enzymes, such as protein kinase C (PKC) isoforms [45]. Therefore, the development of strategies for visualization of acylglycerols in cellular environments by their fusion with fluorescent molecular labels is in high demand [46]. As a result, once the mechanosynthesis of DAGs 5 was established, we turned our efforts towards the conjugation of DAG 5a with 7-hydroxycoumarin (9) (Scheme 5).

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Scheme 5: Conjugation of DAG 5a with 7-hydroxycoumarin (9).

Initially, removal of the TBDMS protecting group of 5a was attempted by milling. However, reacting DAG 5a with a mixture of BF3·CH3CN and silica gel followed by an aqueous work-up gave DAG 6a in only 31% yield, together with concomitant acyl migration of the corresponding sn-1,2-diacylglycerol 6a into sn-1,3-diacylglycerol 6a’. Therefore, the desilylation reaction was carried out by stirring 5a and BF3·CH3CN (Scheme 5a). Next, DAG 6a was reacted with 4-nitrophenyl chloroformate (7) and triethylamine in the ball mill to form the activated DAG anhydride derivative 8a (Scheme 5b). Analysis by 1H NMR spectroscopy of the reaction mixture revealed the presence of DAG 8a along with its isomeric DAG 8a’. Formation of the latter compound could have been facilitated through acyl migration of 6a under the basic milling conditions. Subsequently, the reaction mixture containing 8a and 8a’ was milled with 7-hydroxycoumarin (9) and triethylamine to achieve the conjugation of the DAGs 8 in 53% yield after two steps (Scheme 5c). A mixture of 10a and 10a’ (10a/10a’ 72:28) was separated from the unreacted starting materials, and analyzed by UV–vis spectroscopy (Figure 3). Comparison of the UV–vis spectra of 6a and 10a/10a’ showed the successful conjugation of the DAG with the coumarin moiety [47].

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Figure 3: UV−vis spectra of DAG 6a (dotted line) and conjugated DAGs 10a and 10a’ as a mixture (10a/10a’ 72:28) in toluene at a concentration of 0.5 μM.

Conclusion

The implementation of ball milling techniques has provided the opportunity to extend the applicability of mechanochemistry to the synthesis of architecturally complex targets, such as mono- and diacylglycerols. Altogether, the mechanosynthesis of lipids and lipid derivatives complements the current systematic work towards the synthesis of other biologically relevant molecules under environments of high mechanical stress. Specifically, the synthesis of mono- and diacylglycerols required first, the application of solventless functional group protection chemistry in ball mills, second, the implementation of metal-catalyzed epoxide-ring opening, and third, the development of solvent-free ester formation between monoacylglycerols and fatty acids to afford DAGs. Moreover, the synthesis of conjugated DAGs 10 represents a step forward towards the establishment of mechanochemical conjugation reactions for linking fluorescent materials to lipids at the proof-of-concept level.

Supporting Information

Supporting Information File 1: Experimental procedures, set-ups and characterization data.
Format: PDF Size: 2.4 MB Download

Acknowledgements

This research was possible thanks to the financial support from the RWTH Aachen University through the Seed Fund Projects 2016. We kindly acknowledge Marvin Mendel for his experimental work in this project (RWTH Aachen University). We are grateful to Prof. Dr M. Rueping (RWTH Aachen University) for kindly allowing us to use a UV–vis spectrometer and to Cornelia Vermeeren (RWTH Aachen University) for technical assistance.

References

  1. Petridou, N. I.; Spiró, Z.; Heisenberg, C.-P. Nat. Cell Biol. 2017, 19, 581–588. doi:10.1038/ncb3524
    Return to citation in text: [1]
  2. Fritch, B.; Kosolapov, A.; Hudson, P.; Nissley, D. A.; Woodcock, H. L.; Deutsch, C.; O’Brien, E. P. J. Am. Chem. Soc. 2018, 140, 5077–5087. doi:10.1021/jacs.7b11044
    Return to citation in text: [1]
  3. Garcia-Manyes, S.; Beedle, A. E. M. Nat. Rev. Chem. 2017, 1, 0083. doi:10.1038/s41570-017-0083
    Return to citation in text: [1]
  4. Schönfelder, J.; Alonso-Caballero, A.; De Sancho, D.; Perez-Jimenez, R. Chem. Soc. Rev. 2018, 47, 3558–3573. doi:10.1039/c7cs00820a
    Return to citation in text: [1]
  5. Bolm, C.; Hernández, J. G. ChemSusChem 2018, 11, 1410–1420. doi:10.1002/cssc.201800113
    Return to citation in text: [1]
  6. Ardila-Fierro, K. J.; Crawford, D. E.; Körner, A.; James, S. L.; Bolm, C.; Hernández, J. G. Green Chem. 2018, 20, 1262–1269. doi:10.1039/c7gc03205f
    Return to citation in text: [1]
  7. Hernández, J. G.; Ardila-Fierro, K. J.; Crawford, D.; James, S. L.; Bolm, C. Green Chem. 2017, 19, 2620–2625. doi:10.1039/c7gc00615b
    Return to citation in text: [1]
  8. Bolm, C.; Mocci, R.; Schumacher, C.; Turberg, M.; Puccetti, F.; Hernández, J. G. Angew. Chem., Int. Ed. 2018, 57, 2423–2426. doi:10.1002/anie.201713109
    Return to citation in text: [1]
  9. Jörres, M.; Aceña, J. L.; Soloshonok, V. A.; Bolm, C. ChemCatChem 2015, 7, 1265–1269. doi:10.1002/cctc.201500102
    Return to citation in text: [1]
  10. Baron, A.; Martinez, J.; Lamaty, F. Tetrahedron Lett. 2010, 51, 6246–6249. doi:10.1016/j.tetlet.2010.09.069
    Return to citation in text: [1]
  11. Métro, T.-X.; Colacino, E.; Martinez, J.; Lamaty, F. Amino Acids and Peptides in Ball Milling. Ball Milling Towards Green Synthesis: Applications, Projects, Challenges; Green Chemistry Series; Royal Society of Chemistry: Cambridge, United Kingdom; pp 114–150. doi:10.1039/9781782621980-00114
    Return to citation in text: [1]
  12. Maurin, O.; Verdié, P.; Subra, G.; Lamaty, F.; Martinez, J.; Métro, T.-X. Beilstein J. Org. Chem. 2017, 13, 2087–2093. doi:10.3762/bjoc.13.206
    Return to citation in text: [1]
  13. Yeboue, Y.; Gallard, B.; Le Moigne, N.; Jean, M.; Lamaty, F.; Martinez, J.; Métro, T.-X. ACS Sustainable Chem. Eng. 2018, 6, 16001–16004. doi:10.1021/acssuschemeng.8b04509
    Return to citation in text: [1]
  14. Kumar, V.; Taxak, N.; Jangir, R.; Bharatam, P. V.; Kartha, K. P. R. J. Org. Chem. 2014, 79, 3427–3439. doi:10.1021/jo5001753
    Return to citation in text: [1]
  15. Menuel, S.; Doumert, B.; Saitzek, S.; Ponchel, A.; Delevoye, L.; Monflier, E.; Hapiot, F. J. Org. Chem. 2015, 80, 6259–6266. doi:10.1021/acs.joc.5b00697
    Return to citation in text: [1]
  16. Jicsinszky, L.; Caporaso, M.; Martina, K.; Calcio Gaudino, E.; Cravotto, G. Beilstein J. Org. Chem. 2016, 12, 2364–2371. doi:10.3762/bjoc.12.230
    Return to citation in text: [1]
  17. Jicsinszky, L.; Caporaso, M.; Tuza, K.; Martina, K.; Calcio Gaudino, E.; Cravotto, G. ACS Sustainable Chem. Eng. 2016, 4, 919–929. doi:10.1021/acssuschemeng.5b01006
    Return to citation in text: [1]
  18. Eguaogie, O.; Vyle, J. S.; Conlon, P. F.; Gîlea, M. A.; Liang, Y. Beilstein J. Org. Chem. 2018, 14, 955–970. doi:10.3762/bjoc.14.81
    Return to citation in text: [1]
  19. Appy, L.; Depaix, A.; Bantreil, X.; Lamaty, F.; Peyrottes, S.; Roy, B. Chem. – Eur. J. 2019, 25, 2477–2481. doi:10.1002/chem.201805924
    Return to citation in text: [1]
  20. Tyagi, M.; Kartha, K. P. R. Carbohydr. Res. 2015, 413, 85–92. doi:10.1016/j.carres.2015.04.008
    Return to citation in text: [1]
  21. Hu, H.; Li, H.; Zhang, Y.; Chen, Y.; Huang, Z.; Huang, A.; Zhu, Y.; Qin, X.; Lin, B. RSC Adv. 2015, 5, 20656–20662. doi:10.1039/c5ra02393a
    Return to citation in text: [1]
  22. Eichmann, T. O.; Lass, A. Cell. Mol. Life Sci. 2015, 72, 3931–3952. doi:10.1007/s00018-015-1982-3
    Return to citation in text: [1] [2] [3]
  23. Yang, C.; Kazanietz, M. G. Trends Pharmacol. Sci. 2003, 24, 602–608. doi:10.1016/j.tips.2003.09.003
    Return to citation in text: [1]
  24. Brose, N.; Betz, A.; Wegmeyer, H. Curr. Opin. Neurobiol. 2004, 14, 328–340. doi:10.1016/j.conb.2004.05.006
    Return to citation in text: [1]
  25. Storck, E. M.; Özbalci, C.; Eggert, U. S. Annu. Rev. Biochem. 2018, 87, 839–869. doi:10.1146/annurev-biochem-062917-012448
    Return to citation in text: [1]
  26. Martin, S. F.; Josey, J. A.; Wong, Y.-L.; Dean, D. W. J. Org. Chem. 1994, 59, 4805–4820. doi:10.1021/jo00096a023
    Return to citation in text: [1]
  27. Fodran, P.; Minnaard, A. J. Org. Biomol. Chem. 2013, 11, 6919–6928. doi:10.1039/c3ob41483c
    Return to citation in text: [1] [2]
  28. Bonnamour, J.; Métro, T.-X.; Martinez, J.; Lamaty, F. Green Chem. 2013, 15, 1116–1120. doi:10.1039/c3gc40302e
    Return to citation in text: [1]
  29. Hernández, J. G.; Butler, I. S.; Friščić, T. Chem. Sci. 2014, 5, 3576–3582. doi:10.1039/c4sc01252f
    Return to citation in text: [1]
  30. Beillard, A.; Golliard, E.; Gillet, V.; Bantreil, X.; Métro, T.-X.; Martinez, J.; Lamaty, F. Chem. – Eur. J. 2015, 21, 17614–17617. doi:10.1002/chem.201503472
    Return to citation in text: [1]
  31. Crawford, D. E.; Miskimmin, C. K.; Cahir, J.; James, S. L. Chem. Commun. 2017, 53, 13067–13070. doi:10.1039/c7cc06010f
    Return to citation in text: [1]
  32. Giri, N.; Bowen, C.; Vyle, J. S.; James, S. L. Green Chem. 2008, 10, 627–628. doi:10.1039/b801455h
    Return to citation in text: [1]
  33. Rytczak, P.; Drzazga, A.; Gendaszewska-Darmach, E.; Okruszek, A. Bioorg. Med. Chem. Lett. 2013, 23, 6794–6798. doi:10.1016/j.bmcl.2013.10.020
    Return to citation in text: [1]
  34. Zhao, Y.; Wang, W.; Li, J.; Wang, F.; Zheng, X.; Yun, H.; Zhao, W.; Dong, X. Tetrahedron Lett. 2013, 54, 5849–5852. doi:10.1016/j.tetlet.2013.08.074
    Return to citation in text: [1]
  35. Cucciniello, R.; Ricciardi, M.; Vitiello, R.; Di Serio, M.; Proto, A.; Capacchione, C. ChemSusChem 2016, 9, 3272–3275. doi:10.1002/cssc.201600989
    Return to citation in text: [1]
  36. Tokunaga, M.; Larrow, J. F.; Kakiuchi, F.; Jacobsen, E. N. Science 1997, 277, 936–938. doi:10.1126/science.277.5328.936
    Return to citation in text: [1]
  37. Ferguson, M.; Giri, N.; Huang, X.; Apperley, D.; James, S. L. Green Chem. 2014, 16, 1374–1382. doi:10.1039/c3gc42141d
    Return to citation in text: [1]
  38. Jacobsen, E. N. Acc. Chem. Res. 2000, 33, 421–431. doi:10.1021/ar960061v
    Return to citation in text: [1]
  39. Schaus, S. E.; Brandes, B. D.; Larrow, J. F.; Tokunaga, M.; Hansen, K. B.; Gould, A. E.; Furrow, M. E.; Jacobsen, E. N. J. Am. Chem. Soc. 2002, 124, 1307–1315. doi:10.1021/ja016737l
    Return to citation in text: [1]
  40. Beillard, A.; Métro, T.-X.; Bantreil, X.; Martinez, J.; Lamaty, F. Chem. Sci. 2017, 8, 1086–1089. doi:10.1039/c6sc03182j
    Return to citation in text: [1]
  41. Bolm, C.; Hernández, J. G. Angew. Chem., Int. Ed. 2019, 58, 3285–3299. doi:10.1002/anie.201810902
    Return to citation in text: [1]
  42. Laszlo, J. A.; Compton, D. L.; Vermillion, K. E. J. Am. Oil Chem. Soc. 2008, 85, 307–312. doi:10.1007/s11746-008-1202-5
    Return to citation in text: [1]
  43. Fodran, P.; Das, N. J. L. C.; Eisink, N. N. H. M.; Welleman, I. M.; Kloek, W.; Minnaard, A. J. Eur. J. Lipid Sci. Technol. 2016, 118, 1768–1774. doi:10.1002/ejlt.201500547
    Return to citation in text: [1]
  44. Métro, T.-X.; Bonnamour, J.; Reidon, T.; Duprez, A.; Sarpoulet, J.; Martinez, J.; Lamaty, F. Chem. – Eur. J. 2015, 21, 12787–12796. doi:10.1002/chem.201501325
    Return to citation in text: [1]
  45. Sprong, H.; van der Sluijs, P.; van Meer, G. Nat. Rev. Mol. Cell Biol. 2001, 2, 504–513. doi:10.1038/35080071
    Return to citation in text: [1]
  46. Cai, X.; Mao, D.; Wang, C.; Kong, D.; Cheng, X.; Liu, B. Angew. Chem., Int. Ed. 2018, 57, 16396–16400. doi:10.1002/anie.201809641
    Return to citation in text: [1]
  47. Taniguchi, M.; Lindsey, J. S. Photochem. Photobiol. 2018, 94, 290–327. doi:10.1111/php.12860
    Return to citation in text: [1]

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