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Comunicación oral presentada en la XXIX Conferencia Internacional sobre Polifenoles y la 9ª Conferencia de Taninos, del 16 al 20 de julio de 2018 Madison, WI, USA, MAIN CONCLUSION Lignin biosynthesis is uniquely malleable, allowing a variety of phenolics to be utilized as lignin monomers and therefore allowing some tailoring of its structure, reactivity, and value. We are now at a juncture where actual ‘design’ of the polymer can be contemplated and envisioning an ideal polymer for lignin valorization can be entertained. INTRODUCTION Evidence continues to mount regarding lignins’ inherent structural malleability from studies on lignin pathway mutants and transgenics as well as on various ‘natural’ plants discovered to possess unusual lignins. [1-4] Most of the monomers previously considered were from the monolignol biosynthetic pathway itself. More recently, phenolics from beyond the monolignol pathway have been shown to be authentic monomers in some plants, including the flavone tricin in all grasses (and beyond),[5,6] various hydroxystilbenes in some palm endocarp tissues,[7,8] and now possible nitrogenous compounds such a putrescine in maize kernel lignin. [9] As we have frequently noted from some time back, but not in print until 2008: “any phenolic transported to the lignifying zone of the cell wall can, subject to simple chemical compatibility, be incorporated into the polymer.”[10] That does not automatically mean that the r sulting polymer will be well tolerated by the plant, but researchers are now able to contemplate some degree of actually designing lignins for improved utilization and value, and muse over what might constitute a lignin that is ideally suited for conversion to phenolic monomers, adding value to the biorefinery. RESULTS AND DISCUSSION As José Carlos del Río will cover the other pathways that are now known to contribute to lignification, we shall provide an update to how lignins can be designed to fall apart more readily during processing (the so-called ‘zip-lignin’ approach) and describe a lignin that is natural in some tissues (but not yet in plant stems) that is one example of an ideal lignin. Zip-lignins. We have now shown that it is possible to engineer weak bonds (esters) into the lignin backbone,[11] thus facilitating lignin depolymerization during pretreatment or processing (such as in pulping).[12] It turns out that Nature is already making lignins this way, at low levels, in a variety of plants;[13] we may have even inadvertently selected for this trait in targeting woody species that pulp most easily, for example. An “ideal lignin.” It is now a realistic juncture to posit the characteristics for an “ideal lignin” archetype for biomass processing. For the depolymerization of the polymer to monomers, one ideotype is a lignin that has at least the following three characteristics. First, it should be stable under acidic conditions to prevent condensation and the generation of undesired new C–C bonds, during pretreatment. Second, it should contain only ether (C–O) inter-unit linkages in its backbone so that it can be fully depolymerized. Last, it should be generated in planta from a single phenylpropanoid monomer to allow the production of the simplest array of compounds. C-lignin, such as that found in vanilla and various cacti seed coats, [14,15] is essentially a homopolymer synthesized almost purely by β–O–4-coupling of caffeyl alcohol with the growing polymer chain, producing benzodioxanes as the dominant unit in the polymer, is an example of such an “ideal lignin” that can, in principle, be depolymerized to a single monomeric product in high yield. Here we will describe the ideal nature of this lignin via a revised compositional characterization of the vanilla seedcoat fiber, new features of the C-lignin’s reactivity and stability, and our successful attempts at converting it to monomers in near-quantitative yields., REFERENCES [1] Y. Mottiar, R. Vanholme, W. Boerjan, J. Ralph, S. D. Mansfield, Current Opinion in Biotechnology 2016, 37, 190. [2] R. Vanholme, K. Morreel, C. Darrah, P. Oyarce, J. H. Grabber, J. Ralph, W. Boerjan, New Phytologist 2012, 196, 978. [3] J. Ralph, Phytochemistry Reviews 2010, 9, 65. [4] W. Boerjan, J. Ralph, M. Baucher, Annual Review of Plant Biology 2003, 54, 519. [5] W. Lan, J. Rencoret, F. Lu, S. D. Karlen, B. G. Smith, P. J. Harris, J. C. del Rio, J. Ralph, The Plant Journal 2016, 88, 1046. [6] J. C. del Río, J. Rencoret, P. Prinsen, Á. T. Martínez, J. Ralph, A. Gutiérrez, Journal of Agricultural and Food Chemistry 2012, 60, 5922. [7] J. C. del Río, J. Rencoret, A. Gutiérrez, H. Kim, J. Ralph, Plant Physiology 2017, 174, 2072. [8] J. Rencoret, H. Kim, A. B. Evaristo, A. Gutiérrez, J. Ralph, J. C. del Río, Journal of Agricultural and Food Chemistry 2018, 66, 138. [9] J. C. del Río, J. Rencoret, A. Gutierrez, H. Kim, J. Ralph, Journal of Agricultural & Food Chemistry 2018, in press (accepted 4/20/2018). [10] J. Ralph, G. Brunow, P. J. Harris, R. A. Dixon, P. F. Schatz, W. Boerjan, in Recent Advances in Polyphenol Research, Vol. 1 (Eds.: F. Daayf, A. El Hadrami, L. Adam, G. M. Ballance), Wiley-Blackwell Publishing, Oxford, UK, 2008, pp. 36-66. [11] C. G. Wilkerson, S. D. Mansfield, F. Lu, S. Withers, J.-Y. Park, S. D. Karlen, E. Gonzales-Vigil, D. Padmakshan, F. Unda, J. Rencoret, J. Ralph, Science 2014, 344, 90. [12] S. Zhou, T. Runge, S. D. Karlen, J. Ralph, E. Gonzales-Vigil, S. D. Mansfield, ChemSusChem 2017, 10, 3565. [13] S. D. Karlen, C. Zhang, M. L. Peck, R. A. Smith, D. Padmakshan, K. E. Helmich, H. C. A. Free, S. Lee, B. G. Smith, F. Lu, J. C. Sedbrook, R. Sibout, J. H. Grabber, T. M. Runge, K. S. Mysore, P. J. Harris, L. E. Bartley, J. Ralph, Science Advances 2016, 2, e1600393. [14] F. Chen, Y. Tobimatsu, L. Jackson, J. Nakashima, J. Ralph, R. A. Dixon, The Plant Journal 2013, 73, 201. [15] F. Chen, Y. Tobimatsu, D. Havkin-Frenkel, R. A. Dixon, J. Ralph, P Natl Acad Sci USA 2012, 109, 1772., This work was funded in part by the DOE Great Lakes Bioenergy Research Center (DOE BER Office of Science DE-FC02-07ER64494 and DE-SC0018409).