Extraction Studies of Suberin from Bark and Synthesis of Lignin Isocyanate-Free Polyurethanes: Byproducts of the Beech Wood Industry

INAUGURALDISSERTATION

Zur Erlangung des Doktorgrades

der Fakultät für Chemie und Pharmazie

der Albert-Ludwigs-Universität Freiburg im Breisgau

vorgelegt von

Anna Fichtner

aus Nikopol (Ukraine)

Freiburg im Breisgau, November 2017

Vorsitzender des Promotionsausschusses: Prof. Dr. Stefan Weber

Dekan: Prof. Dr Manfred Jung

Referent: Prof. Dr. Dr. Christian Friedrich

Korreferent: Prof. PhD Marie-Pierre Laborie

Datum der mündlichen Prüfung: 17.01.2018

Albert-Ludwigs-Universität Freiburg I

Für die Familie und Freunde, die mich mit Ihrer Liebe begleiten, beschützen, auffangen und wenn ich doch stürze mir immer wieder aufhelfen.

Albert-Ludwigs-Universität Freiburg II

“The illiterate of the 21st Century are not those who cannot read and write but those who cannot learn, unlearn and relearn.”

“Die Analphabeten des 21. Jahrhunderts werden nicht diejenigen sein, die nicht Lesen und Schreiben können, sondern diejenigen, die nicht lernen, verlernen und wieder lernen“

Alvin Toffler (October 4, 1928 – June 27, 2016) American writer and futurist

Albert-Ludwigs-Universität Freiburg III

Die präparativen und analytischen Arbeiten im Rahmen dieser Dissertation wurden im Zeitraum vom Juni 2013 bis April 2017 am Institut für Makromolekulare Chemie sowie an der Professur für forstliche Biomaterialien der Albert-Ludwigs-Universität Freiburg in den Arbeitskreisen von Prof. Dr. Dr. Christian Friedrich und Prof. PhD Marie-Pierre Laborie durchgeführt. Nur durch die Unterstützung zahlreicher Personen war eine solche Arbeit möglich gewesen; deshalb gilt mein herzlicher Dank allen Beteiligten, die zum Gelingen dieser Arbeit beigetragen haben.

Prof. PhD Marie-Pierre Laborie danke ich für die Beauftragung mit dem interessantesten und innovativen Thema, für die fachliche Unterstützung, für das entgegengebrachte Vertrauen und die damit eingehenden Freiheiten.

Prof. Dr. Dr. Christian Friedrich danke ich für die Betreuung und konstruktiven Treffen, die diese Arbeit immer wieder in reale, zielgerichtete Bahnen lenkten.

Weiterhin gilt mein Dank meinen Kollegen und Freunden sowohl bei der Professur für Forstliche Biomaterialien (BIOMAT), Dr. Heiko Winter, Dr. Tiago von Lossau, Dr. Ricarda Böhm und Jamerson Oliveira, dem Freiburger Materialforschungszentrum (FMF), Levente Szantó, Dr. Gregory Stevens und Uli Matthes, wie auch dem Zentrum für Biosystemanalyse (ZBSA).

Dr. Heiko Winter gilt mein besonderer Dank, der mich mit viel Geduld, an seiner reichhaltigen Erfahrung hat teilhaben lassen.

Prof. Dr. Bernd Kammerer danke ich für das Ermöglichen selbständiger Messungen am GC-MS in seinem Arbeitskreis.

Prof. Dr. Rolf Mühlhaupt danke ich für das zur Verfügung stellen des Druckreaktors für meine Arbeit.

Elke Stibal und Beate Albrecht, den guten Seelen des Instituts danke ich für die Fürsorglichkeit und Hilfe in allen Belangen wärend der gesamten Zeit.

Meinen Kollegen Dr. Gopakumar Sivasankarapillai, der mir mit den 31P-NMR Messungen half, Jan Badorrek für die OL Daten zur Bestimmung des HSP danke ich für das Teilen an Wissen und gegenseitige Hilfe.

Ich danke den Studenten Sona Othman und Uasmin Zidanes, deren Master- bzw. Bachelorarbeiten ich betreuen durfte.

Ein herzlicher Dank gilt auch allen Korrekturlesern, die sich durch meine Arbeit gekämpft haben und darunter ganz besonders Dr. Gregory Stevens.

Albert-Ludwigs-Universität Freiburg IV

Ein besonderer Dank gebührt vor allem all den Menschen, die ich als ein Privileg sehe in meinem Leben zu haben:

Meinen Mädels: Veronika Kaufmann, Inna Weber, Nelli Nekrasov und Sabrina Zinn, die gerade durch ihre Unterschiede, mich in allen Lebenslagen sicher halten und nie fallen lassen.

Meinen „Kollegen“, die für mich Freunde wurden und ein Stück weit Ersatzfamilie.

Und zum Schluss gilt mein unendlicher Dank meinen Eltern, die viel in Ihrem Leben für meinen Bruder und mich geopfert haben.

Albert-Ludwigs-Universität Freiburg V

Table of Contents Table of Contents ...... VI

Zusammenfassung ...... XI

Abstract ...... XII

List of Abbreviations ...... XIII

List of Symbols ...... XVI

1. Introduction ...... 1

1.1 Problem Statement and Motivation ...... 1

1.2 Research Objectives ...... 3

1.3 Proposed Strategy ...... 4

2. Literature Review ...... 6

2.1 Beech (Fagus sylvatica)-Important Values and Characteristics ...... 6

2.1.1 Occurrence of Beech ...... 6

2.2 Bark ...... 7

Biological Function ...... 8

Utilization ...... 8

2.2.1 Chemical Composition of Beech Bark ...... 9

2.2.1 Cell Structure Beech Bark ...... 11

2.3 Suberin ...... 13

2.3.1 Assumed Suberin Models ...... 14

2.3.2 Extraction of Suberin ...... 18

2.3.3 Applied Analytical Methods ...... 21

Chromatography and Mass Spectroscopy for Suberin Analysis ...... 21

Nuclear Magnetic Resonance Spectroscopy (NMR) ...... 22

Other Optical Methods Used to Analyzed Suberin and Suberized Cells ...... 23

2.3.4 Application of Suberin as Bio-Based Polymer ...... 26

2.3.1 Treatment of Beech Bark Suberin ...... 26

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2.3.2 Suberin of Beech Bark ...... 27

2.3.3 Summary Suberin ...... 28

2.4 Lignin ...... 29

2.4.1 Technical Extraction of Lignin ...... 30

2.4.2 Characterization of Lignin and Lignin Polymers ...... 33

2.4.3 Application and Utilization (Possibilities) of Lignin ...... 35

2.4.4 Non-Isocyanate Polyurethanes (NIPU) ...... 38

2.4.5 Organosolv Lignin from Beech Wood ...... 39

2.5 Hansen Solubility Parameter ...... 40

3. Results and Discussion ...... 41

PART I ...... 42

3.1 Analysis of Beech Bark and Bark Extracts ...... 44

3.1.1 Process Efficiency Observation by Solid Bark (Residues) ...... 44

Natural Water Content of Different Bark Samples and Pre-Dried, Milled Bark Samples ...... 44

FTIR Measurements of Solid Bark (Residues) ...... 45

NMR Measurements of Solid Bark (Residues) ...... 46

Thermal Analysis of Solid Bark Residues ...... 47

3.1.2 Solvent Extracts of Beech Bark ...... 50

FTIR Measurements of the Solvent Extracts ...... 50

Ultraviolet-Visible Spectroscopy (UV-Vis) Measurements of Solvent Extracts ...... 52

NMR Measurements of Solvent Extracts ...... 54

Chromatographic (GC-MS) Measurements of Solvent Extracts ...... 55

3.1.3 Summary Measurements of the Solvent Extractives and the Treated Barks ...... 57

3.2 Suberin Monomer Analysis ...... 59

3.2.1 FTIR Measurement of Depolymerized Suberin ...... 59

3.2.2 UV-Vis Spectroscopy Measurement of Suberin ...... 61

Total Phenol Content and Indirect Determination of Tannins (Folin-Ciocalteu-Method) ...... 61

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Determination of Condensed Tannins ...... 61

3.2.3 Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) ...... 62

Proton NMR ...... 62

Total Correlated Spectroscopy (1H-1H TOCSY) ...... 65

13 1 13 C-, H- C HSQC and HMBC NMR of SM Spectra ...... 68

Calculation of Chain Length and Number of Functionality per Chain ...... 70

Diffusion-Ordered NMR Spectroscopy (DOSY) ...... 73

3.2.4 Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin ...... 74

3.2.5 Elemental Analysis Measurement of Suberin Monomers ...... 79

Conclusion of PART I ...... 82

PART II...... 83

3.3 Analysis of Organosolv Lignin and its Derivative ...... 85

3.3.1 FTIR Measurements of OL, EL and CL ...... 86

3.3.2 NMR Measurements of OL, EL and CL ...... 89

Proton NMR of OL, EL and CL ...... 89

Phosphorus NMR of OL and EL ...... 92

3.3.3 Thermal Analysis of Lignin (Derivative)...... 94

3.3.4 SEC Measurements of the OL and EL ...... 96

3.3.5 Hansen Solubility Parameter of the OL, EL and CL ...... 97

3.3.6 Summary Lignin Modification ...... 99

3.4 Lignin Based Non-Isocyanate Polyurethane ...... 100

3.4.1 Monitoring of Curing by FTIR ...... 100

3.4.2 Solubility of NIPU’s ...... 104

3.4.3 Thermal Analysis of the Lignin Based NIPUs ...... 104

Conclusion of PART II ...... 106

4. General Conclusions and Perspectives ...... 107

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4.1 Summary Measurement of Suberin ...... 107

4.2 Using Lignin as Precursor of NIPUs ...... 108

5. Experimental ...... 109

5.1 Materials and Chemicals ...... 109

5.2 Bark preparation and extraction...... 113

5.2.1 Solvent Extraction of Bark ...... 113

5.2.2 Depolymerization (Methanolysis) of Bark Suberin ...... 113

5.2.3 Purification of Suberin Monomers ...... 114

5.3 Lignin Preparation ...... 115

5.3.1 O-Glycidylation of Organosolv Lignin ...... 116

5.3.2 Carbon Dioxide Addition Reaction with Lignin Epoxides ...... 118

5.3.3 Hansen Solubility Parameter (HSP) Determination ...... 118

5.4 Lignin and Suberin Model Compounds Modification ...... 120

5.5 NIPU Synthesis ...... 122

5.6 Characterization Methods ...... 123

5.6.1 Fourier Transform Infrared Spectroscopy (FTIR) ...... 123

5.6.2 UV-Vis Spectroscopy ...... 123

Total Phenol Content (Folin-Ciocalteu-Method) ...... 123

Indirect Determination of Tannins ...... 124

Determination of Condensed Tannins ...... 124

5.6.3 Nuclear Magnetic Resonance (NMR) Study...... 124

Derivatization with TAI ...... 125

Derivatization with TMBP ...... 126

5.6.4 Differential Scanning Calorimetry (DSC) ...... 127

5.6.5 Thermogravimetric Analysis (TGA)...... 127

5.6.6 Gas Chromatography-Mass Spectrometry (GC-MS) ...... 127

5.6.7 Elemental Analysis ...... 128

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5.6.8 Gel permeation Chromatography (GPC) ...... 129

6. Supporting information ...... 130

6.1 Additional Figures ...... 130

6.2 Additional Tables ...... 143

7. References ...... 157

List of Tables ...... 170

List of Figures ...... 173

Albert-Ludwigs-Universität Freiburg X

Zusammenfassung

Das Ziel dieser Dissertation war es, die Extraktstoffe aus der Europäischen Rotbuche (Fagus sylvatica) zu gewinnen, charakterisieren und daraus ein Isocyanat-freies Polyurethan (NIPU) Copolymer herzustellen. Dazu wurden im ersten Teil der Arbeit diverse Extraktstoffe durch sukzessive Soxhlet- Extraktion mit Dichlormethan (DCM), Methanol und Wasser aus der Buchnenrinde gewonnen. Im Anschluss wurde das natürliche Polyester Suberin durch Methanolyse aus der verbliebenen extraktfreien Rinde gewonnen. Mit Hilfe von Gas-Chromatographie mit Massenspektrometer (GC-MS), Kernspinresonanzspektroskopie (NMR), Fourier-Transformations-Infrarotspektroskopie (FTIR), Elektronenabsorptionsspektroskopie, Elementaranalyse, Thermogravimetrischen Analyse und dynamischen Differenzkalorimetrie wurden sowohl die Extraktstoffe, wie auch die vorwiegend aliphatischen Suberinmonomere (3,7 ±0,9 Gew.%) untersucht. Der DCM-Extrakt besteht vor allem aus so genannten „Wachsen“, d. h. aus phenolischen Verbindungen sowie langkettigen Alkanen, welche keine oder nur eine einfache Funktionalisierung aufweisen und sich somit für eine Polymerisation nicht eignen. Mit Methanol und Wasser wurden vorwiegend lösliche Saccharide gewonnen, wobei auch 4.9 Gew.% des Atro-Rindengewichts als kondensierte Tannine identifiziert werden konnten, die sich bereits bei der Polyurethan (PU) Herstellung bewährt haben[1]. Die gewonnenen Suberinmonomere erwiesen sich im GC-MS als langkettige, lineare Säuren oder Alkohole, die drei- bzw. vierfach funktionalisiert vorliegen (mit Doppelbindungen, Epoxiden und Diolen in den Ketten). Das mit Abstand größte Signal im Total Ionen Chromatogramm wurde als 9, 10, 18-Trihydroxy-octansäuremethylester identifiziert. NMR-Untersuchungen, die unter Verwendung einer eigens hergeleiteten Formel die Berechnung der durchschnittlichen Funktionalität per Suberinmonomer ermöglicht, liefern eine Funktionalität von 1,5 pro Monomer, wohingegen GC-MS Messung Werte von 3 bis 4 liefern.

Im zweiten Teil dieser Arbeit wurde überprüft, ob das bereits bekannte Konzept der NIPU Polymerisation auch bei einem komplexeren System wie Lignin angewandt werden kann. Dafür wurden die Hydroxylgruppen eines Organosolv-Lignins (OL), dass bei der Zellstoff Herstellung aus Buchenholz gewonnen wird, via Phosphorus NMR untersucht. Die Hydroxylgruppen des OL wurden durch nukleophile Substitution mit Epichlorhydrin derivatisiert und unter Hochdruck (20 bar) und Normaldruck wurde Kohlendioxid an den Epoxidring addiert. Die Bildung der zyklischen Karbonate am Lignin (CL) wurde mit FTIR und NMR nachgewiesen. Nach Zugabe von unterschiedlich langen, linearen Diaminen zum CL konnte mit FTIR, DSC und Löslichkeitsexperimenten gezeigt werden, dass bei Verwendung von kurzkettigen Diaminen, trotz höherer Reaktivität der primären Amine, es vorwiegend zur Pfropfcopolymerisation kommt, während es mit dem längerkettigen Diamin, bei richtiger Konzentration ein vernetztes Lignin basiertes NIPU entsteht.

Albert-Ludwigs-Universität Freiburg XI

Abstract

The aim of this dissertation was to characterize extracts from European beech (Fagus sylvatica) and to obtain and to utilize the extracts to produce bio-based Non-Isocyanate Polyurethane (NIPU) copolymers. In the course of the first part of this study, extracts were collected from the bark by successive Soxhlet extraction with dichloro methane (DCM), methanol and water. Then the natural polyester suberin was extracted with a rather low yield (3.7 ±0.9 wt.%) from the remaining extract-free bark by methanolysis. The extracts and the predominantly aliphatic suberin monomers were investigated using the analytical methods of Gas Chromatography–Mass Spectrometry (GC-MS), Nuclear Magnetic Resonance (NMR) spectroscopy, Fourier-Transform InfraRed (FTIR) spectroscopy, UltraViolet–Visible spectroscopy, Elemental Analysis, Thermal Gravimetric Analysis and Differential Scanning Calorimetry. The DCM extract was found to consist of "waxes", which were primarily composed of phenolic compounds as well as long-chain alkanes with either no functionalization or simple mono functionalization, and therefore this extract was not suitable for polymerization. Predominantly soluble saccharides were obtained in the methanol and water extracts, whereby condensed tannins (4.9% by weight of the dry bark) was identified as well. These are already known in the polyurethane (PU) production[1]. The suberin monomers obtained were found by GC-MS to be long-chain, linear acids or alcohols, which were tri- or tetra-functionalized compounds with double bonds, epoxides and diols. The largest Total Ion Chromatogram (TIC) signal was identified as 9, 10, 18-hydroxy octadecanoic acid. NMR studies allowed the calculation of the average functionality per suberin monomer using a specially derived formula. However, only 1.5 functional groups per monomer were found by NMR method, which is an average of the whole mixture. Whereas 3- 4 functional groups per monomer were found by TIC of GC-MS.

In the second part of this study, the method of NIPU polymerization was applied to the more complex lignin system. For this purpose the hydroxyl groups of beech Organosolv Lignin (OL), obtained by Phosphorus NMR (31P-NMR) of wood pulp, were derivatized by nucleophilic substitution with epichlorohydrin followed by addition of carbon dioxide to the epoxide rings under normal and high (20 bar) pressure. The formation of cyclic carbonates on the resulting Carbonate Lignin (CL) was detected by FTIR and NMR. After further addition of linear diamines of different chain lengths to CL, analysis by FTIR, DSC and solubility experiments showed that the use of short diamines leads to graft copolymerization despite the higher reactivity of the primary amine groups, whereas NIPU lignin with crosslinking was formed when the CL was cured with diamines with longer chain lengths.

Albert-Ludwigs-Universität Freiburg XII

List of Abbreviations

ATR Attenuated Total Reflection

BE Extract-free Bark

BN Milled or Natural Bark bp Boiling point

BR Remaining bark II or Bark Residue BS Bark Sample BuOH Butanol CBP Chemical-Biochemical Processes cf. Compare with (lat. confer) CF Folin-Ciocalteu CG Chain Group CL Cyclic Carbonate Lignin

〈퐶퐿〉 Average chain Length CP/MAS Cross-Polarization Magic Angle Spinning CZ Cambium Zone DEPT Distortionless Enhancement by Polarization Transfer DMF DiMethylFormamide DOSY Diffusion-Ordered SpectroscopY DSC Differential Scanning Calorimetry ECH EpiChloroHydrin EG End or terminal Group e.g. For example (lat. exempli gratia) EI Electron Impact ionization EI Epoxy Index EL Epoxidized Lignin ELM Epoxidized Lignin Model EPh Early Phloem Eq. Equation ESI-MS/MS ElectroSpray Ionization tandem Mass Spectrometry ESM Epoxidized Suberin Modele etc. Et cetera FC Folin-Ciocalteu FID Flame Ionization Detector FTIR Fourier Transform InfraRed spectroscopy

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G(-unit) Guaiacyl unit GA Gallic Acid GC Gas Chromatography GPC Gel Permeation Chromatography H(-unit) p-Hydroxypehyl unit HMBC Heteronuclear Multiple Bond Correlation HPLC High-Performance Liquid Chromatography HPSEC High Performance Size Exclusion Chromatography HSP Hansen Solubility Parameter HSQC Heteronuclear Single Quantum Coherence IB Inner Bark IR InfraRed (spectroscopy) KL Kraft Lignin LM Lignin Model LPh Late phloem m Multiplet M Middle lamella MALDI Matrix Assisted Laser Desorption/Ionization MAS Magic Angle Spinning MeOH Methanol MS Mass Spectrometry MS/MS or MS² Tandem Mass spectrometry MSTFA N-Methyl-N-(triMethylSilyl) TriFluoroAcetamide

MSTFA-SM Suberin Monomer derivatized with N-Methyl-N-(triMethylSilyl) TriFluoroAcetamide NIPU Non-Isocyanate PolyUrethane NIR Near-InfraRed (spectroscopy) NMR Nuclear Magnetic Resonance (spectroscopy) OB Outer Bark OL Organosolv Lignin P Primary wall PAS PhotoAcoustic Spectroscopy PLM Polarization Light Microscopy PU PolyUrethane PVPP PolyVinylPolyPyrrolidone q Quintet

Albert-Ludwigs-Universität Freiburg XIV

Ref. References RI Retentions index RI-(detector) Refractive Index detector s Singlet S(-unit) Syringyl unit S Secondary wall SD Standard Deviation SEC Size-Exclusion Chromatography

SM Depolymerized suberin or Suberin Monomers SM Suberin Model

SN2 Nucleophilic Substitution (bimolecular reaction mechanism) SPAD Suberin Poly(Aliphatic) Domain SPE Solid Phase Extraction SPPD Suberin Poly(Phenolic) Domain SSC Spin-Spin Coupling SU Suberin t Triplet T Tertiary wall TAC TrichloroACetate TAI TrichloroAcetyl Isocyanate TEM Transmission Electron Microscopy TGA ThermoGravimetric Analysis TIC Total Ion Chromatogram TLC Thin-Layer Chromatography TMDP 2-Chloro-4, 4 ,5 ,5-TetraMethyl-1, 3 ,2-DioxaPhospholane TMS TriMethylSilyl TOCSY TOtal Correlated SpectroscopY UV-Vis UltraViolet–Visible (spectroscopy) vic Vicinal W “Wax” film

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List of Symbols

Symbol Name of Symbol unit 퐴 Absorbance A Area 푐 Concentration mol/l D Self-diffusion coefficient m²/s Path length of the beam of light through the material cm 푑 sample δ Chemical shift ppm 훿 Total cohesive Hansen solubility parameter √푀푃푎

훿퐷 Dispersion cohesive Hansen solubility parameter √푀푃푎

훿퐻 Hydrogen bonding cohesive Hansen solubility parameter √푀푃푎

훿푃 Polar cohesive Hansen solubility parameter √푀푃푎 E Total cohesive energy J

ED Dispersion cohesive energy J

EH Hydrogen bonding cohesive energy J

EP Polar cohesive Hansen solubility ernergy J 푙 ∙ 푐푚 휀 Molar attenuation coefficient 푚표푙 J Spin-spin coupling Hz 퐾 Mn Number average molecular weight g/mol Mp Peak molecular weight g/mol Mw Weight average molecular weight g/mol Mz Size average molecular weight g/mol 푚 Mass g

푚푥 Mass of bark g μ Dipole moment D N Relative number of function Relative number of: function group (x); a chain group; a 푁 , 푁 , 푁 or 푁 푥 퐶퐺 퐸퐺 퐶 terminal group; or chains p Pressure bar 푝 Number of protons Pd Polydispersity (Mw/Mn) R² Determination coefficient T Temperature °C

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Symbol Name of Symbol unit t Time s, min, h, d.

Td Degradation temperature °C

Tg Glass temperature °C

Tm Meeting temperature °C 푢 Bark humidity % V Molar volume mol/cm³ 푣̃ Wave number cm-1 푤 Water content % wt.900°C Residue weight after heating to 900°C wt.% 푥(훿) Relative peak high as function of the chemical shift

〈푋퐶〉 Frequency of functional groups per chain

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Introduction 1

1. Introduction

1.1 Problem Statement and Motivation

Owing to the increasing scarcity of fossil resources and climate change, the demand for alternative, bio- based energy sources and bio-based materials has increased considerably. An efficient evaluation of the existing biomass as a resourse is mandatory in solving coming challenges.[2] 53.21 million m³ timber was cut down in Germany in 2013[3], producing considerable byproducts. These byproducts of the timber industry are potential renewable resources. In this work we will use these resources, which up to now have been under-utilized (usually burned for energy production).

Four tree species dominate in German forests: beech, oak, spruce, and pine. They occupy approx. 74% of the forest area[4]. The „Dritte Bundeswaldinventur (2012)“ reported that for Germany, the proportion of beech and oak increased by 0.9% to 15.4% and by 0.6% to 10.4%, and that of spruce and pine decreased by 2.2% to 25.4% and by 0.8% to 22.3% compared to the proportions quoted ten years earlier in the “Zweite Bundeswaldagentur (2002)”.[5] This active intervention of the last years should keep the German forest resistant against extreme weather events and promote flora and fauna in the future. In the German state of Baden-Württemberg, the proportion of beech, oak and spruce is 21.5%, 7.5% and 33.5%. Due to climate change, the aim in Baden-Württemberg is to reach amounts of 32% beech, 7% oak and to reduce the share of spruce to 29%.[6] According to the estimates of “Holzmarktbericht 2013”, the proportion of logged beech increased from 15.9% (2004: 8.668 million m³) to 22.7% (2013: 12,071 million m³) in ten years.[3]

The processing of beech wood results in several million cubic meters of bark biomass worldwide every year. This bark is currently used as bark mulch, or utilized for energy generation. In 2014 approx. 0.8 million m³ beech bark 1 was produced as a byproduct of the wood processing industry in Germany. With the trend towards increasing hardwood and beech species in German forests, the availability of beech bark is further expected to rise over the next decades, making available a valuable new resource for material use.

Therefore, urgent steps to be taken to find new value and applications for hardwoods and beech wood, in particular. The Ganzheitliche Nutzung von Laubholz am Beispiel Buche (GANUBU) project particularly addresses this problem, by aiming for a full exploitation of beech wood, i.e. not only the

1 calculated with an estimated 40 [5]cm three diameter and the equilibration Rössler to determinate the proportion on beech bark [7] and the reported beech logging 2013 [3]. The results are consistent to 7% volume fraction published by Schneider and Baums [8]. Albert-Ludwigs-Universität Freiburg 1

Problem Statement and Motivation 1.1 beech xylem should be used, but also beech bark. Both should be considered for the development of novel bio-based thermosetting polymers with applications as wood adhesives.

However, for an effective evaluation of the resulting beech bark, the chemical components should be known first and should also be quantitatively estimated. Unfortunately, the published literature is limited and often contradictory which shows the importance of initial, analytical experiments to estimate the chemical components of beech bark. In this work, special attention is given to aliphatic monomers of suberin and the aromatic tannins with a view to using them as precursors in aliphatic-aromatic polymer systems. The extraction and analytical characterization of beech bark extracts and possible polymer building-blocks, is thus the focus of the “first part” of this dissertation. Here suberin is particularly targeted, as it provides a source of highly functionalized and flexible aliphatic building-blocks. Furthermore, the aromatic extracts (tannins etc.) are also considered as valuable sources of rigid and functionalized building blocks.

In addition, it is significant that during the pulping process of lignocellulose, about 50 million tons per year [9] of lignin is produced as a byproduct. This could be considered as an alternative to tannins for bio- based aromatic polymer precursors. Up to now almost all the lignin (98%) produced is burned for energy production. Alternative pulping procedures that produce, organosolv lignin would enable the production of sulfur-free lignin, which is more homogeneous and reproducible form of lignin that could increase material utilization possibilities. Because of these new properties, a more effective utilization of lignin is possible with this type of sulfur free lignin.

The second part of this dissertation focuses on the development of novel, bio-based, thermosetting polymers from beech wood extracts and by products to replace traditional synthetic wood adhesives.

One of the most important adhesives types are polyurethanes (PUs). Due to their varied properties from thermosetting to thermoplastic polymers, PUs are the one of the most important commercial products. Usually PUs are produced by polyaddition of poly-(or di-) hydroxyl precursors and highly reactive and toxic poly- (or di-) isocyanate. A feasible alternative is to produce PUs by non-isocyanate methods, utilizing the addition of cyclic carbonate with amine components. The benefits of isocyanate free PUs include benign starting materialswith low toxicity[10], and improved mechanical properties due to the additional hydroxyl group.[11]

According to the isocyanate-free concept, we will try to produce bio-based isocyanate free polyurethane (NIPU) using suberin aliphatic monomers as soft segments and tannins or lignins as hard segments for improved mechanical properties.

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Research Objectives 1.2

1.2 Research Objectives

The main goal of the research project for this thesis was the synthesis of bio-based, Non-Isocyanate Poly- Urethane (NIPU) copolymers with both aliphatic and aromatic building blocks obtained and derived from timber industry byproducts of European beech (Fagus sylvatica).

The specific objectives to achieve this goal are:

PART I: Assess the potential of beech bark as source of bio-based building blocks.

(1) Identify and quantify potential polymer precursors contained in beech bark, to match the results with the literature and to single out the most promising ones. (2) Analyze the aliphatic suberin monomers as polymer precursors qualitatively and quantitatively, with specific emphasis on the statistical distribution of the functional groups employable in polymerization.

PART II: Develop novel, bio-based thermosets from valuable building blocks and byproducts of beech wood processing.

(3) Derive novel aromatic NIPU precursors from organosolv beech wood lignin using the addition reaction of carbon dioxide and epoxides under various conditions. (4) Produce NIPUs from the novel lignin-derived precursors and commercially available diamines as a proof-of-concept. (5) Produce a bio-based suberin-lignin NIPU and determinate its properties.

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Proposed Strategy 1.3

1.3 Proposed Strategy

An overview of the structure of this project, including both the completed work and (an intended portion which, due to time constraints, was not realized) (grey), is presented in schematic form in Figure 1.3-1. The diagram is divided into defined regions corresponding to the respective objectives (see Section 1.2 Research Objectives). The Results and Discussion which follows in Chapter 3 are structured with respect to the drafted research objectives. The more detailed performance of the scheme is described in PART I and PART II.

(1)

(3)

(2)

(5)

(4)

Figure 1.3-1: Diagram of proposed strategy Section 3 of Results and Discussion shows the solvent extraction of beech bark - see blue marked region of the diagram in Figure 1.3-1. The solvent extracts and the diverse barks were planned to be analyzed by several spectroscopic and photometric methods so as to compare the results with the literature, and to determine if and which extract might be suitable for polymer synthesis.

As suberin is one such valuable building block, analysis ofits composition after depolymerization was planned using GC-MS and NMR to find the specific and average functionalities of the monomers (Section 3.2 green in Figure 1.3-1). This corresponds to the second research objective. By having a statistical description of the functional groups of suberin, its possible utilization in polymer synthesis can best be determined.

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Proposed Strategy 1.3

An innovative2 attempt would be made to synthesize cyclic carbonate derivatized lignin (CL) using an epoxy lignin (EL) obtained from organosolv pulping of beech wood to produce organosolv lignin (OL). This section of the proposed strategy is marked red in the Figure 1.3-1, and deals with the third research objective. The subsections of Section 3.3 explain special NMR, FTIR and GPC techniques used to follow the reaction and to quantify the new functionalities of the resulting lignin. Thermal properties and the Hansen solubility parameter were to be determined in each step.

The fourth research objective (orange) was to prove the NIPU concept by curing the new CL with simple, linear, terminal diamines, and to analyze their thermal and mechanical properties. An analysis of this portion of the work is to be found in Section 3.4.

The last research objective (grey) was not implemented due to time limitations and is not part of this dissertation. However, it is a possible approach for future work. The intention was to develop a new synthetic approach in order to produce polymeric material with the flexible aliphatic modified suberin monomers as one part and the stiff aromatic modified lignin as the second part of the bio-based NIPU.

2 The synthesis of lignin carbonate using kraft lignin is published also by Salati, Zoia and Orlandi [12] in the meantime. Albert-Ludwigs-Universität Freiburg 5

Literature Review 2

2. Literature Review

Growing demand for renewable energy resources and substitute sources for fossil resources is leading the science communityto search for “green” chemicals.[13] Production of bio-based polymers is one field of research. A large number of natural substances are appropriate precursors of bio-based polymers[14– 17]. For example, oils or the fatty acids of oils[18–27], tannins[28], terpenes, suberin monomers[29–35] or monolignols of lignin, and also macro- and polymer like lignin[9, 24, 36–41], and cellulose, etc.[21, 42, 43]

The idea of this work is to use two natural polymers: Suberin of beech bark and lignin of beech wood.

2.1 Beech (Fagus sylvatica)-Important Values and Characteristics

This Section focuses in particular on beech. The values and characteristics known in the literature are summarized in subsections: beginning with the occurrence of beech; the structure and chemical composition of beech bark while the component suberin is discussed separately; closing with the lignin of beech wood. Consequently, this section is about existing resources obtainable from beech, and gives the literature on the first three research objectives.

2.1.1 Occurrence of Beech

European beech (Fagus sylvatica) is one of the three main species of trees in Germany. Only the coniferous trees spruce and pine are more frequently represented. Beech is the most common deciduous tree, occupying 15.4% (10,887,900 ha) forest area and this proportion is increasing.[4, 6] It is a widely distributed, deciduous tree in Europe, see Figure 2.1.1-1. The tree begins to reproduce only after 40-50 years, grows to 30-40 m height and lives for up to 250 years in age.[44] Typically beech is harvested after 80-120 years.[45–47] The volume of harvested wood in Germany is 53 Mio. m³ per annum, of which 22.7% is beech.[3]

Beech wood has a high degree of stiffness, wear-resistance, strength, excellent bending capabilities, low elasticity and thereby can be used for several applications, the most important being for furniture production, flooring, boat building, staircases and musical instruments. Additionally it is used in pulping and fabrication of various boards, veneers, panels and plywood. Due to its relatively high energy content, it is also used directly as fuel and for charcoal production.[45]

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Bark 2.2

Figure 2.1.1-1: Relative presence probability of European beech [44]

2.2 Bark

Bark (Cortex) is a highly specialized and complex tissue defined as all the tissues outside of the cambium, see Figure 2.1.1-1. Consequently it comprises all the dermal tissue of vascular plants and the cover layer of trunk, branches, steams and roots [48] of trees, woody vines and shrubs. Depending on the species, the trunk of the beech tree consists of approximately 10% - 20% bark, 20% - 35% branches, and a still higher percentage in the roots and the stump than in the trunk.[49] In the literature bark is often divided into inner bark or phloem and outer bark or Figure 2.1.1-1: Scheme of bark with the various tissues [70]. rhytidome.

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Bark 2.2

Biological Function

Tree bark is the primary, protective barrier against mechanical damage caused by animals, physical influences such as excessive heating or frost and biological attacks by pathogens[50–52], decay-producing organism and parasitic plants. Furthermore, bark is responsible for water balance[50, 53–55], water transport and for the accumulation and assimilation of products from the leaves to the rest of tree.[56] This tissue is also important as a repository for substances that are harmful to the metabolism of plants. The anatomical composition of bark can be divided into cambium, phloem and periderm:

Utilization

As a byproduct of the forest industry, bark is almost wholly used as a biofuel for thermal energy generation despite its low fuel value.[57] Compared to wood, bark differs by the presence of polyphenols and suberin, it is less anisotropic, possesses a lower heat transfer coefficient and, swells differently. Bark is also richer in extracts, it has weaker mechanical properties, fewer polysaccharides and less cellulose, and it has the same crystalline lattice but a lower degree if crystallinity than wood.

Until today only the bark of some specific species are used for non-energy industrial applications. Bark from Quercus suber (or variablis[58]) is the source of industrial cork which has numerous uses including as stoppers for bottles, flooring, insulating material, and as an anion exchanger for removing heavy metals from water. Applications of the extractable substances from bark include:

a) tannins from e.g. Acacia mearnsii, Tsuga heterophylla, Acacia mearsii, Picea abies, Picea brutia and Pinus halepensis bark used in leather manufacture [57], b) condensed tannins and phenols often used as adhesives in e.g. particleboard resins, cold setting laminating and corrugated paper bonding.[56, 59–62] They also have an important role as bioacids. c) bast fibers as a composite of polymers.[57] d) Pharmaceutical applications of barks (summarized by Laks [56]) are obvious because of their important biological roles and are used inter alia as folk medicines: Complex prenylated flavonoids are inhibitors for the cyclic adenosine monophosphate (AMP) phosphodiesterase; methylated epigallocatechin affects muscle relaxation; antifungal (ichthyoxoic and phytogrowth-inhibitory) bark extract quercetin is used for antibiotics; catechin ((+)- Cyanidanol-3 © from Zyma SA of Switzerland) is used against liver disease; betulin (acid esters) is used as a disinfectant and is a component of various cosmetics and shampoos, and is also supposed to have anticancer properties; cumarins are diuretic and anti-infammatory agents; hydrolysable tannins inhibit adrenaline-induced lipolysis and are active against reverse transcriptase from RNA tumor virus.

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Chemical Composition of Beech Bark 2.2.1

2.2.1 Chemical Composition of Beech Bark

The composition of bark is more complex than wood. It can firstly be divided into lignin, suberin, cellulose [63], inorganic material and extractable components. The first two components are described in detail in subsections 2.3.1 and 2.4.5. The cellulose of bark has been analyzed as to be very similar to xylem cellulose. A detailed composition of beech bark is shown in Table 2.2.1-1.

Table 2.2.1-1: The beech bark composition and ash content refer to dry bark (db). IB: inner bark; OB: outer bark; rA: number refers to relative peak area

Ash 4.0%-7.3%[63]; 9.90% [66] Proteins 2.4%-4.1%[63]; IB223 µmol/g db, OB159 µmol/g db[67] Starch 0.4%-0.8%[63] Holocellulose 20.9%[63]; α-Cellulose 11.8% Hemicellulose 9.1% Carbohydrates 43.1%-64.8% [63] Cellulose 10.2%-26.2[63] Glucomannan 0.3%-1.2%[63] O-Acetyl-4-O-methylglucuronoxylan 10.8-24.6%[63] Pectin 7.4%-10.1%[63] Saccharose 0.6%[63] Glucose 0.5%[63] Fructose 0.3%-2.9%[63] Fixed glucose 8.6%-28.8%[63] Soluble carbohydrates IB216 µmol/g db, OB33 µmol/g db[67] Total phenols 31-65 mg quercetin/g dry bark [68] Flavanols rA3.5%-16.5%[68] Procyanidines rA0.3%-12.5%[68] IB86 µmol/g db, OB105 µmol/g db[67] Taxifolin glycerols rA38.6%-57.3%[68] Extractable components in bark are present in large quantities and varieties compared to wood.[57, 64, 65] Typically, they are dissolved by three steps of successive (Soxhlet) extraction:

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Chemical Composition of Beech Bark 2.2.1

I. A non-polar solvent such as chloroform (CHCl3), dichlormethane (CH2Cl2) or diethyl ether is used to remove the nonpolar “waxes” constituted of alkanes, fatty acids, fats, resin acid, phytosterols, terpenes.[56] II. A semi-polar solvent such as methanol or ethanol dissolvs condensed tannins, flavonoids, and phenols.[56] III. A polar solvent, typically water or alkaline water, extracts condensed tannins and water-soluble carbohydrates.[56]

The quantity and composition of extracts of beech bark are summarized in Table 2.2.1-2.

Table 2.2.1-2: Yield of beech bark extractable materials refers to dry bark and their composition refers to the extract amount.

I. Non-polar extract Ether 2.1%-6.0%[63]; Chloroform 6.0% [69] II. Medium-polar extract Ethanol 6.4-20.4%[63]; 20.3% [66]; 13.6% [69] - Saccharose 0.5%[63] - Fructose 0.4%[63] - Galactose 0.1%[63] - Glucose 0.4%[63] III. Polar extract Water 2.9%-14.4%[63]: 14.4% [69] - Hollocellulose 45%-61.5% (28.0%-29.6% are α-Cellulose) - Lignin 44.3%[63]; klason lignin: 11.5%, kürschner lignin 43.0%[66]; 39.0% [65] - Proteins 0.2%[63] - Saccharose 0.7%[63] - Fructose 0.5%[63] - Galactose 0.2%[63] - Glucose 0.5%[63]

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Cell Structure Beech Bark 2.2.1

2.2.1 Cell Structure Beech Bark

The cambium is the living cell layer that separates bark and wood (Xylem) in the stem and roots [48] of a vascular plant (see Figure 2.2.1-1 CZ: cambium cells). It produces cells for the secondary xylem growth towards the inside of the stem and the secondary phloem growth outwards in the bark.[70]

The secondary phloem (“functional phloem”) includes living cells (sieve cells or tubes) similar to the xylem which are responsible for the transport of photosynthesis products from the leaves to the roots, and also transport in a radial direction (Xylem: transport of water and nutrients from the roots to the leaves) (see EPh: early phloem and LPH: late phloem in Figure 2.2.1-1 or collapsed and non-collapsed phloem (Figure 2.2.1-2) It occupies approx. 93% of the whole beech bark thickness.[59] In deciduous trees, longitudinally arranged sieve tubes consisting of narrow, single living cells connected end to end act as a conduit for photosynthesis products. The cell walls of the tubes are perforated by collections of pores known as “sieve plates”.[70, 71]

The polygonal stone cells (sclereids) belong to the sclerenchymatous cells, whose shape and content vary depending on the species, the external condition and Figure 2.2.1-2: Structure of the age of the tree. They beech bark observed via light microscopy [59]. have a transport function and are responsible for mechanical properties.[56, 70] The last important cell types in phloem are the longitudinal arranged parenchyma cells, which constitute the main part of the phloem and are dispersed among the sieve cells. Their function is the storage of nutrients.[70]

In contrast to the phloem, the outer bark (Periderm) is mainly composed of dead tissue.[70] Beech belongs to the Figure 2.2.1-1: Structure of the cambium “peridermal trees”, which means they permanently retain zone (CZ), early and late phloem (EPh and LPh) of beech bark observed via light microscopy [59].

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Cell Structure Beech Bark 2.2.1 their first periderm. The periderm (often called secondary covering) is formed by the phellogen, phellem and phelloderm (illustrated in Figure 2.2.1-3).

Kramar et al. and Prislan et al. examined cellular and topochemical features of beech bark with different microscopic techniques.[66] Prislan observed almost 90% collapsed phloem in bark that was more than 100 years old.[59]

The phellogen (cork or bark cambium) is a layer of meristem, producing exocrine phellem (cork) cells and endocrine phelloderm cells that normally form the outer cortex, but for “peridermal trees” (e.g. Fagus, Ribes, Carpinus, Corylus, Abies, Acer) the periderm replaces Figure 2.2.1-3: Structure of beech bark periderm [59]. the epidermis in roots and stems and remains for most of the life of the tree.[56]

In contrast to Quercus suber the periderm of beech bark survives the radial growth and the formation of the cork cambium.

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Suberin 2.3

2.3 Suberin

Suberin is a natural polymer almost ubiquitous in the vegetable realm. It is found in bark and subterranean (underground) plant periderm e.g. roots [48], tubers, stolon and, rhizomes. The bark of all plants undergo secondary thickening. Suberized cells are found in normal and especially in wounded tissues of plants [52, 72, 73], and mainly in the outer bark of higher plants. The thin natural polyester of suberin is responsible for cell wall strength (makes the tissue flexible and smooth [56]), is a moisture diffusion barrier [53, 55], a pathogen-induced defense [51] and influences germination.[54] Historically, the physical and biological function of suberin is confusing and varied, depending on the author.

The term “suberin” was coined by the French chemist M. Chevreul in 1815.[74] He extracted substances from cork several times at first with hot water followed by hot alcohol, and to the insoluble part of cork he gave the name suberin. Since then, the term has been repeatedly redefined and specified. In 1836 Boussingault [75, 76] found that “Chevreul’s suberin” is partly soluble in potassium hydroxide. He presumed that this was the important substance responsible for the properties of cork. After Doepping succeeded in extracting cellulose from “Chevreul’s suberin” in 1843 [77], the discovery that suberin contained cellulose at first met with resistance. Later, when cellulose was extracted in even larger quantities from various suberin sources, opinion changed rapidly to the other extreme: Suddenly suberin was regarded as a form of contaminated cellulose, or chemically modified form of cellulose.

As a result of microscopic observation of suberin in suberized cells, see Figure 2.3.3-1, von Höhnel assigned the term ‘suberin’ to one of the three observed cell wall layers, as suberin was the characteristic constituent of cork (Figure 2.3.2-1)[76] Additionally, using alkaine hydrolysis, von Höhnel found a chemical similarity between suberin and cutin whereby he mistakenly termed suberin as cutin and vice- versa.[54, 72] However, cutin is quite distinct from suberin in both cell location and chemical composition.[72, 78–87]

The lamella structure observed by electron microscopy (see Figure 2.3-1) generated theories that suberin enclosed “waxes” or that extractable “waxes” were part of suberin. For a long time the aromatic part of suberin was spuriously assumed to be lignin or lignin-like. [72]

Figure 2.3-1: Isolated periderm of potato tuber fixed with OsO4. [73]

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Assumed Suberin Models 2.3.1

Since the 1970s researchers have appropriated two suberin model types: aliphatic or aliphatic-aromatic crosslinked polymers. Depending on the publisher, terms like “desuberize”[88], “suberin phenolics” [89, 90] or “suberin poly(aliphatic) domain”[91] were used to, describ the preference at that time for the then current suberin model.

Comparable figures are hard to find in the literature due to the following: the number of definitions of suberin (just aliphatic or aliphatic aromatic polymer, with or without extracts); as different extraction methods [81, 92] (hydrolyzis or transesterification, with or without catalysts, different times for the procedure [92]); various material sources (whole bark or outer bark, extractive-free or just milled tissue, particle size [92], freeze dyed or fresh) of diverse plant species and their geographical locations [93], thus yielding unequal results. Nonetheless, the current state-of-the-art method to identify suberized cells is the observation of typical ultra-structure by electron microscopy and the analysis of suberin aliphatic or/and aromatic monomers by various methods.[56]

2.3.1 Assumed Suberin Models

Over time, different analytical methods generated information about the monomers. Several models for the description of the chemical structure of suberin were generated. However, all of them can be classified into one of two fundamental concepts, which exist in parallel to each other and yet still provide confusion. One approach considers only the aliphatic structure of the insoluble polymer as suberin [81, 88, 94–100] and the other one defines suberin as an aliphatic-aromatic polymer.[70, 72, 90, 101–105]

One of the first attempts to picture the chemical structure of suberin was by Esau 1977. His idea of suberin was as a “fatty substance in the cell wall of cork tissue and in the casparian band of the endodermis”. In 1973 Holloway and Deas described suberin as a long chain polyester on the basis of the discovery of saponification products (linear alkane chains with carboxyl or hydroxyl end groups) with probably cross-linkages to the vic-diols or epoxides.[81] Also Gil et al. used the aliphatic suberin model, in which aliphatic monomers connect by ester bonds to carbohydrates and lignin, to Figure 2.3.1-1: Proposed suberin organization in cork periderm. [95] describe their 13C solid state NMR Albert-Ludwigs-Universität Freiburg 14

Assumed Suberin Models 2.3.1 results of cork suberin.[88] Lopes at al. composed an improved model (see Figure 2.3.1-1) where the secondary wall was attached to, though spatially separated from, the carbohydrate-lignin middle lamella and the primary cell wall of cork.[95] Long chain ω-hydroxy acids were suggested to be ω-hydroxy acids which were more resistant to cleavage compared to aliphatic acids. α, ω-Dicarboxylic acids were found to be directly connected to middle lamella (low COOR/CH2 ratio), which in turn connect to the n- alcohols, fatty acids, α, ω-dicarboxylic acids (high COOR/CH2 ratio). The large amount of carbohydrates suggests that suberin is covalently attached to the cell wall carbohydrates, and that the linkages between suberin and carbohydrates may be similar to those proposed for lignin to carbohydrate [71].

Around the same time, Kolattukudy developed an aliphatic-aromatic model [94, 106] of the suberin polymer (Figure 2.3.1-2): An aromatic basic (core) structure such as an alkyl-aryl ether linked lignin with esterified hydroxy cinnamic acids covalently connected to the cell wall [72]. The α, ω-dicarboxylic acids and ω-hydroxy acids may serve to cross-link an aromatic matrix and form a linear polyester domain with terminal groups comprising mono functionalized fatty acids and alcohols. Kolattukudy’s model was confirmed by Scott and Peterson’s observation of phenolic components in the suberized cell walls using a positive staining test. The nitrobenzene oxidation concretizes the aromatics as benzaldehydes as in lignin but with a lower degree of substitution and less methoxy groups [107]. The extraction of suberin by Perra et al. as a polymer with dioxane (appropriate for lignin extraction in wood) confirms the results of IR and 13C NMR, agreeing with the aromatic- (lignin-like) aliphatic model of Kolattukudy [101, 108]. In addition, it corresponde with Garbow’s et al. 13C solid-state NMR study of suberized potato [109] or Quercus suber L. [88]. Also the complex polymer model was based on insights into the biosynthesis of the aromatic [110] and aliphatic acids of suberin [72]. At the same time, Kolattukudy agreed with Holloway [52, 81] that the aliphatic and the aromatic [84] part of suberin differ substantially from cutin which was confirmed by the lamella structure (alternating aliphatic and aromatic layers) observed by TEM. However Kolattukudy disagreed that the aliphatic monomers of suberin with the carboxyl-hydroxyl ratio can form an extensive, cross-linked polymer.

Doubts about the lignin-like polymer were Figure 2.3.1-2: 1980 proposed aliphatic-aromoatic raided for the aromatic structure (domain) of polymer structure of suberin [72, 94, 106].

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Assumed Suberin Models 2.3.1 the model with the development of non-destructive 13C solid-state NMR. Using proton and carbon NMR for Björkman protocol treated tissues and extraction by dioxan in dimethylformamide (DMF), Zimmermann et al. showed that the aromatic part of suberin is not made by syringyl and guaiacyl units, as proposed to lignin found in wood [111]. Also Bernards et al. [112] contradicted the previously accepted suberin model of Kolattukudy, since evidence was shown that the aromatic domain of suberin largely consist of hydroxycinnamate derived polymer and not of monolignols as in lignin[100].

The new findings about suberin were summarized by Bernards and Lewis 1998. They summarized that the polyaromatic and polyaliphatic layers have their own unique chemical composition (not lignin-like and not similar to cutin) in distinct entities within suberized cells [113]. The crosslinked aromatic domain consists primarily of hydroxycinnamic acids (see Figure 6.1-2) and their derivatives, the aliphatic domain contain mainly ω-hydroxy acids and α, ω-dicarboxylic acids (see Figure 6.1-1).

Simultaneously, Graça and Pereira revised the aliphatic composition of suberin.[90] They took notice that in most of the publications working with suberin monomers that glycerol was absent. The simple reason for the absence of glycerol was that it was washed out with water during the purification process [114]. Although glycerin had already been determined for the first time in 1887 to be part of suberin by Kügler [115], it was not considered in the model of Kolattukudy [72, 106] or Bernards [113]. In addition, glycerol proved to be a major component of the aliphatic suberin polymer (up to 48%) [90, 96, 96, 114, 116]. Several works used a mild methanolysis to produce monoacylglycerols which were separated and identified by GC-MS. With these, a renewed model was generated which consist of an aromatic polymer (which could be lignin-like or not) and an aliphatic fatty acid polyester cross-linked with glycerol. Both polymers are “chemically and structurally independent” [90] but peripherical links should be some linkages of glycerides and the aromatic part, possible through the ω-hydroxylated acids. Graça and Pereira summarized that the extracted suberin had enough glycerol to esterify all acid groups of the monomers [114]. Bento et al. confirmed the existence of mono-, and di-substituted glycols with MALDI-MS measurements [116].

The current valid version of a suberin model [117] was published by Bernards 2002. He used all preceding insights and versions, and defined new terms ‘Suberin Poly(Aliphatic) Domain (SPAD)’ and ‘Suberin Poly(Phenolic) Domains (SPPD)’. SPAD is represented by a linear, glycerol-branched polyester mainly made up of long chain ω-hydroxy acids and α, ω-dicarboxylic acids, with esterified hydroxycinnamates. In other words SPAD is all aliphatic substances extractable by ester linkage cleavage (see Extraction of Suberin pp. 18), including the organic solvent extractable “waxes”. SPPD’s are two different types of “suberin”. The first SPPD is mainly composed by hydroxycinnamic acids also extractible by ester cleavage, and is alternated with SPAD (summarized to be the observed electron-dense layers monitored by TEM). The second SPPD is a monolignol-hydroxycinnamoyl three-dimension linked co-polymer

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Assumed Suberin Models 2.3.1 with thr following properties: resides in the primary cell wall; is covalently linked to primary cell wall carbohydrates; contains mainly hydroxycinnamic acids, some monolignols, tyramines and phenol derivative, and is covalently linked by ester or ether bonds to the first SPAD lamella.

Suberin is reasoned to be organized by enzyme-mediated process, therefore it is appears uniform and regularly organized (according to the lamellae structure observed by TEM). The two domains are spatially distinct as previously summarized, but still covalently linked [107]. The suberin model presented by Bernards (see Figure 2.3.1-3) includes the important role of glycerol linking the linear arranged SPAD with the three-dimensional SPPD, and also justifies the constant 3-4 nm thickness of the lamellae observed by TEM.

Suberin and lignin, in contrast to e.g. natural rubber or cellulose, have a non-unique structure (i.e. non- repeating units). Synoptically, the combination of chromatography methods (TLC, GC, HPLC) with those of molecule-selective detection like MS gives the most information about the constitutive moieties of suberin [118].

Figure 2.3.1-3: Suggested suberin structure of potato periderm with C: carbohydrate, P: phenolic; S, suberin (phenolic or aliphatic)[91].

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Extraction of Suberin 2.3.2

2.3.2 Extraction of Suberin

The extraction of suberin in its native form is still impossible. An examination of this biopolymer is possible only in its (partial[119]) depolymerized condition. Different depolymerization methods were developed to extract suberin monomers which were generally based on hydrolysis or trans-esterification of the ester bonds. The most relevant procedures are: hydrolysis; transesterification and redox depolymerization (shown in Figure 2.3.2-1).

Hydrolysis

i. Alkaline hydrolysis (saponification) with NaOH or KOH is one of the oldest methods to release primarily the aliphatic monomers (see Figure 6.1-1), but also some aromatics like benzaldehyde (vanillin, p-hydroxybenzaldehyd), benzoic acids (vanillic acid), hydroxycinnamate etc., (see Figure 6.1-2) and triterpenes. Conditions: Water/ethanol solvents were used with a hydroxide concentration of 0.5-2 mol/l at 70-160°C for 1-3 h. Examples: Betula verrucosa outer bark [92, 120], Quercus suber L. [98, 102, 121–123], Betula pendula L. [105, 121, 122], Solanum tuberosum L. [124] and Fagus sylvatica L. [123].

The benefit of alkaline hydrolysis is the faster reaction time compared to methanolysis [92]. But hydrolysis and subsequent acidification during the depolymerization can open up the epoxide rings giving rise to vic-diols [81, 114]. Ekman et al. [92] preserved epoxies in aqueous ethanol with KOH and short reaction times (15 min).

Transesterification methods are the most common techniques [117]. They avoid inter-esterification of monomers during isolation and storage, while aldehydes and epoxide derivative are stable and still identifiable [114]. The amount of aromatic monomers is lower using transesterification in comparison with hydrolysis [93, 120, 125, 126]. A full depolymerization (just aliphatic domain) of suberin with mostly unmodified epoxy rings is possible using a small amount of sodium methoxide [114, 116]. If the epoxy rings were opened by the methanolate, they can be distinguished from the natural vic-diols.[81, 95]

ii. Transesterification without catalysts: Condition: 3-4 h reflux with methanol or ethanol Examples: betula verrucosa outer bark [81, 92] and Quercus variablis [58] iii. Alkaline transesterification [Bolmgren 1981] (methanolysis): Condition: 2-18 h at ~70°C with 0.01-3% NaOMe or Na in dry methanol Examples: Quercus suber [32, 81, 88, 90, 93, 93, 95, 97, 98, 102, 114, 116, 121, 122, 125–127], Betula pendula L. [121, 122].

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Extraction of Suberin 2.3.2

Figure 2.3.2-1: Famous suberin depolymerization methods and their products

iv. Partial depolymerization (mild methanolysis): Bark reflux in Ca(OH)2 or CaO -methanol solution for 15 min-6 h reflux at 50°C [90, 96, 97].

This method keeps the bonds between feruloyl esters, aliphatic chains and acylglycerols (oligomeric structure) e.g. α, ω-dicarboxylic acids, linked at both ends to glycerol units. Also, dimeric and trimeric structures, including ferulic acid linked to ω-hydroxy acids and the ω-hydroxy acids esterified to glycerol were found.[128]

Using this method a variation of monomer “families” was described depending on the catalyst concentration[95, 116]. In addition, Pinto et al. [122] showed that the yields of depolymerized suberin using methanolysis or hydrolysis do not differ very much from each other.

v. BF3-MeOH transesterification with a 14-16% BF3-methanol solution over 24-48 h under nitrogen at 70°C of Fagus sylvatica [87, 108], or of Solanum tuberosum [50, 83], or of different root tissues [83, 87].

A known side reaction of this method is the possible addition of methanol to the double bonds.

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Extraction of Suberin 2.3.2

vi. Acid transesterification (methanolysis) with approx. 2 mol/l sulfuric acid in methanol was used for depolymerization of betula verrucosa outer bark [92]

Redox depolymerization (reductive cleavage) reductive processes can also open ether bonds but this induces loss of structural information.

[50, 72, 82, 84] vii. LiAlH4 degradation in THF 24-48h in reflux of different tubers .

This method converts, carbonyls (esters, carboxylic acids, etc.) and epoxides into alcohols, thereby losing pristine information. The reaction with LiAlD4 was used in the structural studies of aliphatic chain functional/connecting groups [54, 72] and phenylpropanoids [84]. viii. Alkaline nitrobenzene oxidation with nitrobenzene and 2 mol/l sodium hydroxide at 160°C for

2 h of Solanum turberosum [85], Fagus sylvatica [108], Agave americana [86], Rubus idaeus, potato [111], Q. suber [111] was used to cleave phenylpropanoids side chains of suberin. The resulting benzaldehyde, benzoate and acetophenone derivatives are also typical for lignin.

A differentiation of polymer monomers is not possible as phenylpropanoide (monolignols), tyramine or hydroxycinnamic acid derivatives are oxidized to the same products and information about the prior molecule becomes lost.

Other methods found in literature:

ix. Dioxane and water (9:1, v/v) was used under inert atmosphere to extract the nondestructive suberin-lignin polymer of Fagus sylvatica. Lignin was separated by precipitation with ethyl ether. The extraction with dioxane is a well-known method to extract lignin from wood but Perra et al. also tried this method to extract lignin and suberin from bark [101, 108, 111].

x. Thioacidolysis with BF3/CH3CH2SH cleavage is selective to alkyl-aryl ether bonds β-4-O and produces 1, 2, 3-trithioethane derivatives. The method is more selective compared to nitrobenzene oxidation and shows that just 20-40% of the yielded monomers are typical monolignols for lignin, the rest are hydroxycinnamic acids, tyramine and p-hydroxy-substituted phenolics. xi. Ionic liquid Ferreira used cholinium hexanoate to extract suberin of extract-free outer bark of Betula pendula 9:1 wt./wt. for 4h at 100°C partial depolymerized polyester [105]. Garcia et al. studied the effect of three different cations by varying six anions using a volume ration of two parts of Ionic liquid and Part of extracted free cork was heated at 100°C for 4 h. The product was analyzed by FTIR [104]. xii. Enzymatic isolation of cork from Q. suber [129] xiii. Microwave hydrolysis [130]

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Applied Analytical Methods 2.3.3 xiv. Alkali fusion using 26 mol/l potassium hydroxide solution in an open nickel crucible [92]

2.3.3 Applied Analytical Methods

In this subsection current knowledge about the most important analytical methods used for analyzing suberin, and its characteristics and composition are presented.

Chromatography and Mass Spectroscopy for Suberin Analysis

The development of gas chromatography (GC) in 1951 opened new analytical possibilities. Eglinton and Hunneman, especially established GC for fatty acids analysis as a standard method using a selective mass spectrometer (MS) as the detector [32, 78–80, 90, 105, 114, 116, 122, 123, 130], or unselective flame ionization detector (FID) [114]. Kolattukudy used this new technique of GC-MS to qualify and quantify the suberization of different underground plant cell tissues [50] and the barks of different trees [125]. The results were that the monomer composition varied depending on the plant species but was consistent within a family [83, 131]. Despite the differences in the plant species or the depolymerization methods used [95], all researchers found the existence of α-ω-alkanes diol, ω-hydroxy acids, long chain n-alkanols, and fatty acids [125, 131] with and without mid-chain derivatives like epoxide, vic-diole or double bonds. Kolattukudy and Dean independently concluded that for all suberized plants, 18-hydroxy-octadec-9-enoic acid and octadec-9-ene-1,18-dioic acid are the major components [82, 132]. Holloway and Brown Deas focused on the epoxy octadecanoic acids that are also abundant suberin and cutin of different plants and their fragmentation in MS detection [81]. Graça and Pereira proved the existence of, and the important role of, glycerol in suberin which was an idea that had fallen out of favor for a long time [90, 95, 116].

To apply GC, the boiling points of the partly polar monomers were reduced to less than 300°C. For example the carboxyl groups of octadecanoic acid (steric acid) with a b.p. of 370°C were converted into methyl ester groups.[92, 120] This sample octadecanoic acid methyl ester reduced the b.p. to 181°C. The hydroxyl groups were converted to trimethylsilyl ethers with an excess of trifluoroacetamide or N,O- Bis(trimethylsilyl)trifluoroacetamide (BSTFA) [84, 90, 114] with trimethylchlorosilane [92, 95–97, 105, 116, 120, 122, 123, 130].

Using partial depolymerization methods, monoacylglycerols could be observed with GC-MS [90, 96, 97]. Studies using GC-MS by Electron Ionization (EI) [90] by Matrix Assisted Laser Desorption/Ionization (MALDI) [116] and Electro-Spray Ionization tandem Mass Spectrometry (ESI-MS/MS) [96, 97] revealed higher molecular weight dimers- and trimers e.g. linearly-linked ω-hydroxy acids and α, ω-dicarboxylic acids and glycerol-ω-hydroxy acid-ferulic acid [96]. This was an important step in order to understand the native suberin structure. Ferreira et al. [105] used ionic liquids to extract suberin in its polymer form and measured the polymer before and after hydrolysis by GC-MS. Using Albert-Ludwigs-Universität Freiburg 21

Applied Analytical Methods 2.3.3

Liquid Chromatography with a multi tandem Mass Spectrometer (LC-MSn), Wang et al. [124] observed triglycerides in potato suberin.

Several studies dealing with the aromatic compounds of suberin often produced completely different results. For example, Perra et al. [108] claimed that the aromatic domain of suberin in Fagus sylvatica L. is represented by 65% ferulic acid, while Riley and Kolattukudy [84] detected not even 0.3 % in suberin isolated from various bulbous vegetables. In Quercus suber suberin, ferulic acid was established as the main phenolic compound, where Holloway determined it to 1.5% [133] but García-Vallejo et al. 5.3-9.1% of the content [126]. Graça [100] summarized published yields of ferulic acid depending on species and depolymerization technique. It is assumed that the large quantity variations depend on the depolymerization method used. Alkaline hydrolysis is more aggressive and produces a higher amount of aromatics, thought to be from other cell wall polymers. Minor proportion aromatics are vanillin, syringic, vanillic, m-anisic, 4-hydroxy-3-methoxy benzene acetic, cinnamic acid, coniferyl alcohol, hydroxycinnamic amines, caffeic acid [95] and tyramine. Betulin and some betulin belonging to triterpenes derivative were also found by GC-MS in Q. suber.[126, 133]

Furthermore, GC has been used to investigate biological functionalities and processes. Using GC-MS Kolattukudy was able to show that the suberin in potatoes is important as a water barrier, especially for injured skins [50], and to understand the biosynthesis of alkanes [134].

It is important to note that the most GC-MS results published in papers are, at best, semi-quantitative. Most studies have been analyzed with Total Ion Chromatogram (TIC) areas as a reference [95] without the use of calibration standards, and have only looked at the relative abundance of TIC which is sometimes used a factor to calculate quantitative results [123]. An important, but usually unstated fact, is that just 27-74% of the monomers are detected by GC-MS [32, 105, 114]. The non-detected material, for example suberin oligomers would be non-volatile, high molecular weight components resistant to the depolymerization methods [95, 114]. However, components with very high volatility could also be missed in the spectra, since their retentention time could be so short that the signal would have been lost before the beginning of detection.

Nuclear Magnetic Resonance Spectroscopy (NMR)

NMR was used to obtain information about suberin such as the type of functional groups present in suberin [32] and structural information regarding the complex polymer: Garbow et al. [109] used solid state-state 13C-NMR to examine suberized Solanum tuberosum L. (potato) cell walls and concluded that suberin is a polyester with unsaturated carbons and aromatics such as polypropanoides. He suggested that suberin is chemically bound with aromatic groups linked to the cell wall, and that most of the aromatics should be hydroxycinnamates and covalently linked to each other. Additionally, ether bonds

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Applied Analytical Methods 2.3.3 between aromatics and saccharides of the primary cell wall and between aromatics and aliphatics [112] cross link the complex polymer.

Proton NMR was used to analyze the extracts and the depolymerization suberin products [95]. Sousa et al. [102] used the Proton NMR analysis of Trichloro-Acryl Isocyanate (TAI) marked hydroxyl groups to

[102] determine the carbonyl-/ hydroxyl- groups of depolymerized suberin (“dep-suberin” or SM) and for differentiation of primary and secondary hydroxyl groups.

A number of scientists [88, 95] used various harsh methanolysis methods to “desuberize” cork and analyze the molecular dynamics with 13C Cross-Polarization-Magnetic Angle Spinning (CP-MAS) and Solid Phase Extraction (SPE) NMR spectra. They concluded that suberin: is spatially separated from carbohydrate or lignin; experiences a higher motional freedom, contains motionally unhindered and hindered methyl groups depending on distance to the linkages; linked by esters; and they inferred that suberin is spatially separated from carbohydrate. The metabolic fate of specific carbons in NMR studies of Bernads et al. [112] shows mainly hydroxycinnamic acids and only a few monolignols found in lignin were the main components [105, 135–137]. This supported the conclusion that the components were covalently cross-linked via bonds other than ester bonds. Stark and Garbow concluded from their MAS 13C NMR [109] studies that aromatic and aliphatic carbons build in two distinct areas located far distant to the other cell wall structure components such as polysaccharides.

2D NMR-methods Heteronuclear Multiple Bond Correlation (HMBC), Heteronuclear Single Quantum Coherence (HSQC) and TOtal Correlation SpectroscopY (TOCSY) aided the identification of glycerol and its derivatives [124, 129].

However, no investigator has tried to use NMR data to give absolute average information about the functional groups and the chains sizes. Section 3.2 of our work is to give this kind of information to provide an idea of the monomer functionalities and the applicability for a new polymer system.

Other Optical Methods Used to Analyzed Suberin and Suberized Cells

Non-destructive optical methods have yielded histochemical, physical and biological findings for suberin in its native form. Some of the techniques have also used for treated suberin or just for the characterization of different suberin domains.

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Applied Analytical Methods 2.3.3

Microscopic observation of suberin in suberized cells

Franz von Höhnel used optical microscopy to treat the microscopic sections of different plants with potassium hydroxide, nitric acid or urea. With this he observed the unequal behavior of the cellulose membrane and the two visible lamellae between cork cells. One was a potassium hydroxide soluble lamella suberin and the other he called the other one “Mittellammele” (meddle lamella) which was a highly lignified cellulose layer, see Figure 2.3.3-1. Von Höhnel compared also the suberin and cellulose lamellae of cells of several plant species and discovered that plants such as Boswellia Papyrifera, Strychnos Innocua, Platanus Occidentalis, Pyrus Communis, Camellia Japonica and Viburnum Prunifolium revealed cellulose lamella as the main structure in the cell wall whereas in Salix, Fagus, Castanea, Virgilia Lutea Figure 2.3.3-1: Cell model with the tree lamellea proposed by von Höhnel 1877 with m: meddle lamella, and Pyrus malus suberin is the dominant material s: suberin lamella, c: cellulose lamella (tube) [115] of their wall lamella [76].

Polarization Light Microscopy (PLM)

The PLM enabled the observation of anticlinal lamella, showing a negative birefringence in the suberin layer, and x-ray diffraction allowed the determination of suberin as an amorphous substance.

Transmission Electron Microscopy (TEM)

The TEM technique developed in the 1930s was used to obtain high resolution images of suberized membranes. TEM using osmium tetroxide (OsO4) for fixation, generated images of the multi-lamella structures of suberized tissues such as the periderm of a potato tuber [53, 55, 72, 132, 138, 139], wounded xylem of Betula pendula [73], the inner seed coat of grapefruits[54] Quercus suber [55, 140]and parts of other plants [52, 141]. The observation of a few nanometer thick, alternating lamellae can be explained as being due to varied polar regions. The dark electron-lucent and bright electron-dense regions have different polarities, so that OsO4 is taken up in the polar lamella. Bright electron-dense layers are assumed to be aliphatic structures and the dark layers to be aromatic compounds, [73] as schematically illustrated in Figure 2.3.3-2. The electronic microscope observation of beech bark and wood was used by Alojz Kramar et al. to analyze the aromatic structure and chemical composition [66].

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Applied Analytical Methods 2.3.3

The treatment of the wounded periderm of Solanum tuberosum with trichloroacetate (TAC), which is an inhibitor for “wax” synthesis, resulted in differences in the monomer composition of the hydrolysis product and modified the suberin lamella observed by TEM [53]. Soliday et al. inhibited the biosynthesis of long chain (C> 20) fatty acids and alcohol by TAC whereby the concentration of ω-hydroxy acids and dicarboxylic acids was marginally affected, and thereby also the structure of the layers observed by TEM. They monitored a concomitant reduction in the diffusion resistance, and the disappearance of light bands. This seemed to confirm the theory that suberin associated “waxes” are located in the light bands [54, 55] of the periderm layer, and that they are necessary for the development of diffusion resistance. The dark bands are a phenolic matrix covalently attached to the cell wall and cross-linked by the bifunctional ω- hydroxy acids and dicarboxylic acids.

These findings from the optical images confirm a schematic representation of a suberized cell wall which was generated by Sitte [140] (Figure 2.3.3-2). The medial lamella (M) mainly consisted of lignin and the primary wall (P) consisted of lignin and the Figure 2.3.3-2: Schematic illustration of suberized cell carbohydrates. The primary wall was covered by a wall: medial lamella (M); primary wall (P); secondary wall (S); suberin (SU); wax films (W); tertiary wall secondary wall (S) containing suberin (SU) and (T); pores (PO) [140] anticlinal arranged waxes in approx. 3 nm thick films (W) permeated the suberin. The distance between the wax films increased from the primary wall to the tertiary wall (t) which is a plasma membrane containing carbohydrates and lignin [138].

Soliday et al. [53] suggested that suberin based on polar and nonpolar structures auto-organize in polar lamella polymer and nonpolar bands of “waxes”. However, based on Schmidt and Schönherr’s TEM observation of non-effected potato tuber periderm lamellae extracted by chloroform and methanol. According to Kolattukudy’s model [72, 106] the extractable non-polar waxes somehow interact with the aliphatic polymer portion but don’t generate the characteristic films.

Furthermore, Kolattukudy used this technique to demonstrate that suberin is tightly bound to cell walls.[72]

Fourier Transform Infrared Spectroscopy (FTIR):

FTIR was used to show the typical ester bonds in suberin. Rocha et al. used PhotoACoustic Fourier Transform Infrared Spectroscopy (FTIR-PAS) to observe differences between natural cork and extract- free cork [32, 129]. Graça and Pereira used FTIR to analyze the effect of “desuberization” of Q. suber and Pseudotsuga menziesii bark. [105, 114]

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Application of Suberin as Bio-Based Polymer 2.3.4

Ultraviolet–Visible (UV-Vis) Spectroscopy

UV-Vis spectroscopy was used to determine the cell wall bond aromatic (polyphenolics with e.g. the fluorescent probe berberine/aniline blue) and aliphatic (with e.g. neutral red/toluidine blue O) compounds of cells. With this technique, Lulai showed that the aromatic and the aliphatic domains are in distinct regions of the cell wall membrane.

2.3.4 Application of Suberin as Bio-Based Polymer

Suberin is the main component of cork which is part of the other bark of Quercus suber L. (cork oak). Approximatly 280,000 tons of cork is produced for stoppers, isolating corkboards, composites, and other useful and decorative items produced by the worldwide cork industry [32, 33]. Garcia et al.[30] synthetized suberin polyester (PE) films by homopolymerization. Sousa et al.[99] polymerized suberin monomers in a microwave bulk reaction. Heredia-Gguerrero et al.[29] apply a self-assembly and self-esterification based on the suberin monomer 9(10), 16-dihydroypolmic acid. Olsson, Lindström and Tommy [142] used the epoxidized suberin monomer cis-9, 10 epoxy-18-hydroxyoctadeacoic acid and cured it with Novozyme 435®. Li, Iversen and Ek[21] used this monomer and copolymerizd it with cellulose. Corserio et al.[31–35] was more focused on kinetic and characterizations studies to create polyurethanes (PU) from suberin monomers. Corderio et al.[35] used suberin as an additive in offset lithographic printing inks to improve the viscosity.

Because suberin monomers resemble fatty acids is it possible to use them like fatty acid-based polymers.

2.3.1 Treatment of Beech Bark Suberin

To use bark as a source of natural raw material has diverse challenges. Starting with varying concentration of extractives based on external influences like regional climate and soil. But also harvesting season, tree age and the processing of the tree like debaking, storage milling and drying affect the yield of useable substances.

Form an economic viewpoint the usage of just 5% of the dry bark is not economical. So if bark should be used as renewable material it may be more interesting to use the co-substances like lignin, tannin or cellulose, too. One approach to reduce process costs was taken in the Bachelor thesis of Sona Othman[212], she examined in cooperating with this work the influence of the yield and functionality of suberin from non-previously extracted bark. We found out that the yield is 2% increasing but the chain length and the functionality per chain decrease. Via GC-MS we did not find a significant extension on aromatics or saccharides but more “Waxes”. So the assumption is if the high functionality is necessary for the

Albert-Ludwigs-Universität Freiburg 26

Suberin of Beech Bark 2.3.2 following steps an extraction of “waxes” with non-polar solvents is sufficiently, if not the time consuming extractions are pointless. In this bachelor thesis Othman could also show that using just the periderm of the bark would also increase significant the suberin yield.

The method using for the extraction could be more adjusted to the intended application. Methanolysis was fine for the analysis, since it was possible to quantify the functional groups but for usage may be as a copolymer like proposed (gray) in the Introduction, see Figure 1.3-1 would be epoxides or hydroxyl groups preferred in stand of methyl esters. Alternative methods to break the ester bonds are listed in

Subsection 2.3.3. Very interesting is the Redox depolymerization with LiALH4 (vii p. 20) because it reduces all esters to hydroxyls and does not react with the double bonds, see Figure 2.3.2-1.

Since the quantitative determination of the monomer functional groups is still not consistent using EA, GC-MS and NMR other methods like HPLC or MALDI could give further insight in the monomer structure.

2.3.2 Suberin of Beech Bark

Almost all suberin is located in the periderm of the bark [70].Perra, Haluk and Metche [101, 108] extracted 3.2 wt.% suberin and lignin in an approx. ratio of 3:7 form beech bark aqueous dioxane and precipitated the lignin with ethyl ether. Residual suberin is 0.95 wt.% of the dry weight of bark (Kramár and Ebringerová present a divergent amount of suberin in Fagus sylvatica L. 4.3% [143] quoted from [70]) was analyzed by IR, 1H and 13C-NMR depolymerized and purified using liquid-liquid extraction. They found a 26 wt.% phenolic fraction and a 69 wt.% aliphatic fraction. The fatty acids and phenolic were determined by GC-FID. They found saturated and unsaturated fatty acids with a chain length of C12-C24.

The most common represented fatty acids were: myristic acid (C14); behenic acid (C22); palmitic acid

(C16); and oleic acid (C18:1). This is in agreement with the results of Ghezzi-Perra, Haluk and Metche [128]. The phenolic fraction consisted of vanillic acid, syringic acid, vanillin and ferulic acid. Parahydroxyphenyl units (sinapic acid and syringaldehyde) were not detected.

After alkali hydrolysis of Fagus sylvatica L. bark Strebl et al. [144] found β-sitosterol, aliphatic alcohols with chain lengths of between C12 and C28, C8-C24 long fatty acids, unsaturated fatty acids (oleic acid and linoleic acid) and Oxoacids. In a later work Strebel and Mahdalik [69, 145] quantified the fractions of molecule classes via column chromatography and used GC to analyze the sizes of the molecules. They obtained fractions of: 73% oxygen containing substances (aliphatic n-alcohols C12-C31, steroidal alcohols like β-sitosterol, fatty acids C12-C24, saturated hydroxy acids C16-C24, unsaturated fatty acids and hydroxyl acids); 7.3% hydrocarbons and oxygen containing substances; 2.8% alkanes; 0.6% alkanes and

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Summary Suberin 2.3.3

alkenes; 2.1% n-alkanes (C15-C35) and some 2(3)-methyl alkanes; and 5.8% branched, cyclic hydrocarbons and triterpenic diene (determinate by MS and NMR).

Table 2.3.2-1: Suberin monomer types fond in beech (Fagus sylvatica) via GC-MS, cf. Figure 6.1-1. The values are given in areas found in total ion chromatogram (TIC).

Matzke and Ribechini Spielvogel et al. [123] Source Holloway [133] Riederer [87] et al. [130] Extraction method ii. v. xiii. i. % of TIC % of TIC TIC based calculation in Unit % of TIC area area area µg/g of dry material of roots

n-alkyl alcohol 6.8 9.9 5.3 3,034 (C16-C26)

Carboxylic acids 4.4 10.7 9.2 3,339 (C14-C28)

α, ω-dihydroxy alkane 0.8

α, ω-dicarboxylic acids 11.9 11.8 35.1 4,634 (C14-C24)

ω-hydroxy acids 73.8 62.5 50.4 17,605(C16-C24)

Aliphatic fraction 69% of dep suberin

Phenols 26% of dep suberin

Spielvogel et al. [123] tried to quantify the amounts of the monomers “according to Mendez-Millan et al. (2011)“ via GC-MS using some internal and external standards. Their absolute quantity of the found aliphatic monomers in µg/g of dry root material is summarized in Table 2.3.2-1.

2.3.3 Summary Suberin

This Subsection shows that the term “suberin” was redefined several times, and highlights the importance of knowing the definition or model used by the author of a publication. The current model of Bernards with the aromatic and aliphatic domain conforms to the results of modern analytical techniques. The qualitative and quantitative recovery of suberin monomers is directly dependant on the depolymerization method. The determination of monomers can be conducted by GC-MS. This method provides information on qualitative composition. Due to the time consuming calibration and lack of available standards, GC-MS analysis often enables only semi-quantitative information of depolymerized suberin

(SM). To make suberin useful for normal application, other analytical methods are more practical: NMR gives only an average for chain length and functional groups and the aliphatic monomer mixture, but this could be sufficient to produce polymer composites. The time factor especially makes NMR attractive.

Albert-Ludwigs-Universität Freiburg 28

Lignin 2.4

2.4 Lignin

Lignin is one of the most prominent plant polymers beside cellulose and hemicellulose. These three polymers build up the cell walls of all vascular plants[146, 147]. The mass of lignin of land-growing plants is 20-30 wt.%.[148] Currently, lignin is chiefly observed a co-product of the cellulose pulp manufacture for paper.[149]

Lignin is formed by three typical monolignols (cinnamyl alcohols) which are connected by various inter- unit linkage types, see Figure 2.4-1. Depending on the species, the content and constituent of monolignol ratio varies. For example sinapyl alcohol (S-units) and coniferyl alcohol (G-units) are predominant in hardwoods, whereas in softwoods G-units are predominant.[150] In coniferous woods coniferyl alcohol and coumaryl alcohol (H-unit) are the major constituents of the lignin. The molecular mass ranges between 100 and 300,000 g/mol due to the high the degree of polydispersity[146]. Furthermore, lignin is chemically linked to the (poly) saccharides of the cell wall. It accumulates especially in the secondary wall and the middle lamella of plant cells, see Figure 5.2.3-1. Because of lignin’s complex structural changes during isolation, identification or structural determination remains a challenge.

Figure 2.4-1: Determination of most common inter-unit linkages in lignin macromolecules: R1 and R2 are protons or methoxy groups depending on the type of monolignol hydrogen or methoxy group

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Technical Extraction of Lignin 2.4.1

2.4.1 Technical Extraction of Lignin

Lignin is generated when plant biomass is separated in molecular fragments, typically – cellulose and hemicelluloses – lignin. During pulping the structure of lignin is modified depending on the pulping process[149, 151–155].

a) The delignification process of Kraft lignins is an effective method with a broad range of applicability for delignification of various sources of biomaterials.[156] It is performed under harsh conditions, but it is the most prevalent method. Worldwide about 85% of produced lignin are Kraft lignins.[157] Howeverup to mow, Kraft lignins are not used for industrial applications. Biomass like wood is dissolved in hot sodium hydroxide and sodium sulphide solution. This treatment cleaves lignin-carbohydrate linkages, hydrolyzes most of the β-O-4’ bonds, and other aryl ether bonds such as 4-O-5’. Soluble fragments (phenols) can be recovered from the alkaline environment by acidification, this initiates self-aggregation of the fragments. The molecular mass ranges between 150 and 200,000 g/mol[146]. The recovered Kraft lignin differs from native lignin: the polydispersity is greater, the phenylpropane side chains are eliminated or modified (hydroxyl groups), oxygen-liked carbons are reduced, and sulphur residual as sulphide is in the macromolecule. Since the Kraft lignins are not very soluble in water, the purification for

removing salts and sugars is simple. The Tg of softwood is of hardwood 110°C. b) Sulphite lignin is the most widely used lignin product[149] obtained by pulping various types of biomass with sulphite. It is a highly water soluble – due to the incorporated sulphonate groups – polydisperse macromolecule with a molecular weight range of 1,000 to 150,000 g/mol [149]. This pulping method cleaves almost all lignin carbohydrate linkages, but also some β-O-4’ bonds. In comparison to Kraft lignins, the side-chain groups are mostly not reacting but the condensation between C and C6 is a predominant side-reaction. The sulphite lignin extraction results in weaker pulp products, and a less robust chemical recovery. This method is not selective by extracting lignin. Carbohydrates and hemicelluloses are also soluble, with negative implications for applications. Thus, various purification methods are necessary to obtained sulphite lignin[149]. Sulphite lignin has applications as dispersants and binders. c) The organosolv process[158] is a promising approach to fractionate cellulose with a high carbohydrate content and high purity lignin. Under high pressure and temperature biomass is treated with organic solvents (such as alcohols, organic acids, etc.). Low molecular weight lignin with a narrow dispersity can be produced.[156] The resulting Organosolv Lignin (OL) has a low molecular weight distribution low polydispersity, is highly soluble in organic solvents but nearly insoluble in water.[153]

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Technical Extraction of Lignin 2.4.1

Other popular approaches of pulping are Chemi-thermo-mechanical pulping methods by soda pulping and the thermos-mechanical pulping by steam explosion.

Johansson, Aaltonen and Ylinen[156] graphically compared the two common pulping processes with different variants of organosolv pulping, using published data of the techniques for spruce and pine, see Figure 2.4-1. They demonstrated that organosolv pulping for the production of pulps is equal to that of the preferred conventional industrial processes.

Perez-Cantu et al.[154] investigated Organosolv, Steam exposin and Liquid-Hot-Water treatments on lignocellulose to observe the effect on cellulose, hemicellulose and lignin. They show that organosolv pulping gives the largest extraction of hemicellulose and lignin. The product has shown the highest glucose recovery in a second enzymatic hydrolysis step.

Spruce Pine

Figure 2.4.1-1: Purity as a function of kappa (top) spruce (left) and pine (right) pulps and resulting paper strength properties of this two species (below) [156]. The numerated lines represent the various solvents used during the organosolv pulping: 1. peroxoic acid, 2. methanol, 3. cresol (spruce)/phenol (pine), 4 propanol (spruce)/ amine (pine), 5. and 6 ethanol, 7 acetic acid, 8. phenol. Albert-Ludwigs-Universität Freiburg 31

Technical Extraction of Lignin 2.4.1

Table 2.4.1-1: Comparison between commercial lignin types and organosolv lignin, their economic interest and properties[159]

Organosolv Kraft process Sulfite process Soda process process (ethanol/water)

Proportion of industrial 85% 6% 9% 0% pulp production

organosolv Lignin type kraft lignin lignosulfonate soda lignin lignin

Actual/potential ~0.25%/<10% 25%/ <90% low/<90% 0%/60%-90% utilization degree

Molecular weight (MW) 2,000-3,000 2,000-5,000 20,000-50,000 5,000-6,000 [g/mol] 8,300[161] 2,100-8,000[153]

Polydispersity 2-3; 8.2[161] 6-8 9-10 2-6

Sulfur content 1%-1.25% 1.25%-2.5% 0% 0%

Ash content 1%-6% <25% 2%-4% <0.1%

water (insoluble alkaline solution alkali, organic Solubility in organic alkaline solution and organic solvents solvents) solvents

high; Phenolic OH/C9 high low low HW 0.3-0.6[153]

Color dark brown light brown light brown light brown

Purification costs high very high high none

[160] [160] Glass temperature (Tg) 124-174 91-97

[°C] [162] SW 142; no Tg 109 HW 110[162] HW 97[153]

Maximum thermal 319.7[163] 363[162] degradation (Td) 330-365[162]

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Characterization of Lignin and Lignin Polymers 2.4.2

On the other hand Puls[159] compared the factual and potential utilization degree, availability, and property of the various processes byproduct lignin, see Table 2.4.1-1. He clarified the benefits of the organosolv lignins compared to common procedures. Organosolv lignin combines positive properties such as: sulfur free like soda lignin (low odor nuisance), solubility and aromatic hydroxyl content similar to Kraft lignin, low ash content, and no need for purification. Furthermore, organosolv lignin has the lowest Tg, butthis is still higher that the natural value in wood. It is also observed that Tg is affected by

[160] the source: hardwood lignin usually has a lower Tg than softwood lignins.

2.4.2 Characterization of Lignin and Lignin Polymers

The complexity of lignin’s chemical structure was shown in beginning of this Section 2.4. followed, by some technical extraction methods. This Subsection deals with the typical characterizations techniques used for lignins, lignin derivatives and lignin based polymers. Distinct analytical methods are used to determine: a) the kind and portion of the monolignols; b) their inter-unit linkage types; c) molecular size respectively weight clarification, their inter- or intramolecular interactions[160] which are particularly relevant for possible application mechanical tests d)were used to obtained especially the influence of concentrations, copolymers, reactions conditions, etc. on the mechanical and thermal properties.

a) The monolignol quantification is usually a statistical determination of elemental analysis (carbon, hydrogen, nitrogen and optional sulfur) and investigation of methoxy groups. Based on the C9-monolignol unit (phenyl propane), a useful information of overall structure is possible to obtain. Various techniques are used rely on the analytic of the hydroxyl groups such as NMR of lignin (derivative): Proton NMR [157, 164, 165] of acetylated lignins; Carbon NMR [166]; and Phosphorus NMR[164] can be used to determinate quantitatively the monolignols after a derivatization of hydroxyl groups, but also FTIR [157, 167] could be used. Faix classified lignins in G type, GS type and HGS type lignins via FTIR.[168, 169] UV-spectroscopy with special dyes. b) The determination of intermolecular linkages is usually performed by various types of chemical degradation like thioalysis, hydrogenysis, acidolysis, thioacidolysis and acetyl bromide- zinc/acetic anhydride, etc..[160] The fragments could be chronographically separated and analyzed with molecule specific detectors. A chemical definition of “hydrolysis ratio” allows estimations if the polymer is primary bonded by alkyl-aryl ether bonds (hydrolysis ratio < 0.4) or carbon-carbon bonds (hydrolysis ratio > 0.6). According reverse to the hydrolysis ration is the determination of benzoic to (iso-)phthalic acid units. Diverse non-destructive NMR methods are also used for the determination of linkages especially two-dimension NMR for soluble lignin (fragments).

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Characterization of Lignin and Lignin Polymers 2.4.2

c) Chromatographic methods like the high performance size exclusion chromatography (HPSEC) or gel permeation chromatography (GPC) using multiple detections devices monitor the size polydispersity of the macromere.[157, 164]

Figure 2.4.2-1: First derivative of the TGA thermogram: L1 is Kraft lignin; L2 soda/anthraquinine lignin; and L3 organosolv lignin. [157] d) Inter- and intra-molecular interactions are often determined by thermal analysis methods. Free volume between the lignin chains, attractive intra- and inter-molecular forces, the freedom of chains, segments, side groups or branches, stiffness of chains and molecular size influences physical measurable parameter: such as the heat capacity or thermo dynamical behavior such as changing in Young’s (E)- and shear (G)- modulus; or dielectric property’s[160]. According to this essential parameter various techniques could be used to determinate them. The most common are differential scanning calorimetry (DSC)[157], dynamic mechanical thermal analysis (DMTA), or Dielectric thermal analysis (DTEA). All these techniques enable observation of the

transmission of lignin between glassy and rubbery state defined by the glass temperature (Tg). For example Glasser and Jain [162] showed that KL lignins have higher Tg compare to OL of softwoods. Thermal gravimetric analysis (TGA) gives information for the thermal stability of samples. It is important information for limiting the treatment conditions used. In the literature usually only the temperature is given, which shows the maximal weight loss. In Figure 2.4.2-1, the first derivative of the TGA curve of three different extracted lignins is shown. It shows clearly that the natural material lignin could show diverse thermal stability, depending on the pulping method, but also that the species, source and, pretreatment etc. can influence the thermal behavior.[40, 170]

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Application and Utilization (Possibilities) of Lignin 2.4.3

2.4.3 Application and Utilization (Possibilities) of Lignin

Despite that a large number of studies have pointed repeatedly to lignin’s potential contribution, most of obtained lignin from biomass is directly used as fuel. Less than, 2% (1 Mio. t/a) [148, 149, 153, 171] world- wide is used as a raw material for valorized gasification products, as a source of aromatic and for macromolecular development [160]. Bio-refineries, for example working on lignocellulose, could produce cellulose, hemicellululoses, waxes, proteins and, terpenes and in for example organosolv process. Furthermore, secondary product like benzene, toluene, xylene, ethanol, vanillin derivatives, or carbon fiber can be derived from lignin.[148, 150, 153, 171–174] Technical lignin-macromeres are polyols having both aliphatic and aromatic hydroxyl groups, carbonyl, and carboxyl moieties, methoxy substituents and aromatic rings as functionalities. This natural material could be utilized directly or modified by various methods [162, 172, 175] into bio-based copolymer, blend,[40] or polymer composite. Usually, lignin based materials have thermoplastic [176] or thermosetting behavior. Typically derivatization types for modification of lignin or lignin derivative are [150, 176]:

a. Epoxidation: - Glycidylation with epichlorohydrin (ECH) of aromatic phenols producing ethyl ether derivative.[161, 163, 176–184] - By nucleophile addition of 1, 3-gycerol diglycidyl ether[185] to hydroxyl groups - Of unsaturated aliphatic bonds[39] b. Oxyalkylation: - propylene oxide (alkoxylation) [164, 186] forms lignin grafts with poly(propylene oxide) by homopolymerization - by decarboxylation of carbonate derivative[41] - by oxirane rong-opening [39] - OS lignin hydroxyl groups of F. sylvatica L. was successful reacted first direct and second after esterification with maleic anhydride with propylene carbonate by Kühnel et al.[37]

c. Alkylation destroyed intra-molecolar hydrogen bonds, reduce the Tg of derivatized lignin - methylation of lignin hydroxy groups increase the thermal stability and solubility of Kraft lignin[163, 164, 186] - monolignols hydroxyls with dimethyl carbonate producing methoxy and methyl carbonate derivative[187] or Kraft lignin methylated and /or methoxycarbonylated by dimethoxy carbonate[173, 188, 189] observing an increasing thermal stability.

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Application and Utilization (Possibilities) of Lignin 2.4.3

- Acetylation is one of the famous modifications methods, it improve the solubility of lignin so that lignin acetate are better usable for analytical techniques like NMR or GPC.[151] d. Allylation OL by esterification of aromatic hydroxyl groups with dialylcarbonate, and coincident carboxyallylation of the aliphatic hydroxyls [41]

Lignin (modified) was used as filler[190], copolymers, blends and composite[150] in the production of thermoset[42] and thermoplastic[191] polymers the most important are:

i. Phenol formaldehyde (PF) resins by alkoxylation modified lignins used to produce formaldehyde resins[191] which are primarily used in in the manufacture of wood panels such as playwood, medium-density fibreboard (MDF), particle-bord and oriented strand board (OSB).[192] Tejado et al. [157] tested Kraft pine lignin, soda–anthraquinone flax lignin, and ethanol–water wild tamarind lignin as PF recourse and decide form structural and thermal characteristics that KL is most suitable precursor. Podschun et al. increased the phenol hydroxides by phenolation of OL and increase the activity especially of hardwood OL for PF formation.[36] ii. Epoxy resins [193] - EL [161] or lignin derivatives [176] with diamines, imidazole, [185] tri- and tetraamines [180], 1-(2-cyanoethyl)-2-ethyl-4-methylimidazole.[182] For example Sun et al.[184] synthetize an epoxidized KL with bisphenol A diglycidyl ether and cured with diethyltoluene diamine and observed the mechanical, morphological, and thermal properties of the composite depending on the bisphenol A content. - catalytically with aromatic hydroxyl groups of lignin with EL[179] iii. Polyester (PE) was formed by lignin[194] or lignin derivative with diacides like dicarboxyl- terminated polybutadiene.[191] Demaret and Glasser synthetized a cellulose triacetate –

hydroxypropylated lignin block copolymer, the soft segment influences with a Tg lower room temperature elastomeric behavior, melt rheology and toughness in rigid materials. Furthermore, thermoset lignin ester.[195] iv. Radical polymerization of alkene derivative - Lignin graft copolymer synthesis with for example dimer acid[196] v. Polyurethane (PU) reaction of lignin hydroxyl groups by isocyanates forming urethane linkages.[38, 172, 175, 197] Aliphatic hydroxyl groups with isocyanates are significantly higher compared than aromatic OH. This is one of the important reasons why lignin precursors are hydroxyalkyladed.[197]

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Application and Utilization (Possibilities) of Lignin 2.4.3

- Poly(ethyleneadipate), ethylene glycol and 4, 4’diphenylmethanediisocyanate was blended by flax soda lignin.[40] - OSL polyol precursor produced by esterification of oleic acid and modification of the unsaturated bonds was reacted linear poly(propylene)glycol building blocks having terminal isocyanate groups, the resulting copolymer show a higher thermal stability[39] - KL, poly(ethylene)glycol (soft segment) are cured with diisocyanete to produce copolymer with better mechanical properties.[38] The thermal decomposition change to the typical lignin degradation. - KL reacted with polybutadiene diisocyanate ito produce a thermoplastic lignin-based PU[198] vi. Lignin polymer blends: - Hydroxypropyled lignin was used to produce blends with Polyethylene (PE), Polyvinyl alcohol (PVA), poly(methyl methacrylate) (PMMA), and poly(vinyl alcohol) (PVA) by Ciemniecki and Glasser.[199] - Alkylated OL by esterification with various linear acid chlorides were used to prepare thermoplastic blands with cellulose acetate butyrated (CAB), poly(hydroxybutyrate) (PHB), and starch-caprolactone copolymer (SCC) blends.[200] vii. Non-isocyanate PU (NIPU)[201] lignin is produced - by cyclocarbonated lignin for the production of lignin based NIPUS were developed in parallel to this work by Salanti, Zoia and Orlandi. Using with ECH deoxidized soda pulped lignin, they studied the influence of diverse solvents (DMF, DMSO, Meacac

[12] and Dioxane) and various catalysts for the CO2 addition , and for the determination of the highs cyclic carbonate index via FTIR. Chen et al.[202] used linin based bisphenols. They epoxidized the bisphenols and convert the epoxides to cyclic carbonates. NIPU was polymerized by adding of diamins. - by aminated lignin[190] with dicyclocarbonate

The Lignin modification is one method to improve lignin properties: increase solubility, or reactivity, reduce the viscosity, be a precourser for polymerization, or function as copartibilizer, plastsizer and lubricants.

The thermal behavior of lignins implies that lignin can act as a thermoplastic with intra- and inter – molecular hydrogen bonds as well as a thermoset material forming cross-linked structures[150].

The research objective is to integrate lignin into existing polymer formulations to improve their mechanical and thermal properties, and to substitute petrochemical educts.

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Non-Isocyanate Polyurethanes (NIPU) 2.4.4

2.4.4 Non-Isocyanate Polyurethanes (NIPU)

PU can vary from thermoplastic to thermosetting materials with broad applications such as flexible or rigid foams, thermoplastic elastomers, adhesives and coatings[201]. Currently, 2 million tons of PU was produced in Europe in a conventional way.[203] This process is based on an extremely dangerous process (see phosgene release 15. Jun last year in BASF Ludwigshafen[204]) with high health hazards and toxic substances. A lot of reviews constituted problems of the controversial process and pointed alternative manner[10, 16, 201, 205, 206]. One of them is phosgene, it is known as chemical weapon during the first World War and became important in the chemical industry for synthesis of carbonates and isocyanates. A good graphical display of the prosess and the possible alternatives is made by Blattmann et al.[10], see Figure 2.4.4-1.

The strategy to produce NIPUs is a simple polycondensation of linear carbonates or polyaddition (ring opening reaction of five or six membered cyclic carbonate with amine. Nohra et al.[16] gave a good overview about materials and strategies used to produce cyclic carbonates. One of them is the addition of carbon dioxide to epoxides. Foltron, Meareau and Tassaing[207] studied theoretical the chemical fixation of carbon dioxide with propylene oxide using ammonium and guanidinium catalysts. Kirsch, Millini and Wang[208] tested the effectiveness of diverse catalysts with model systems. Last year Salanti, Zoia and Orlandi[12] used classical catalysts such as: potassium iodid; 1, 8-diazabicycloundec-7-ene; N, N-dimethyl-aminopyridine, tetrabutylammonium bromide (TBAB) or ionic liquids (1-allyl-3-methyl imidazolium chloride or bromide) for fixation carbon dioxide on a deoxidized soda pulping hardwood lignin. They determined that the best results were obtained with the ionic liquids, followed by TBAB. This year Salanti at al.[209] first demonstrated a successful synthesis of Lignin based NIPU.

Figure 2.4.4-1: Conventional PU synthesis (left) and the isocyanate-free way producing NIPUS (right)[10] Albert-Ludwigs-Universität Freiburg 38

Organosolv Lignin from Beech Wood 2.4.5

2.4.5 Organosolv Lignin from Beech Wood

The European beech (Fagus sylvatica) wood contains 43% cellulose and hemicellulose; Xylan 28% and Mannan 4%; Lignin 23% and Extractives 1.5%. After the organosolv pulping process is it possible to produce a sulfur-free lignin with a low saccharide content.

Over and Maier used a typical organosolv pulping of beech wood and analyzed it by various methods.[41] The resulting OL characteristics found by GPC, 31P-NMR and EA are listed in

Table 2.4.5-1: OL of beech bark produced and analyzed by Over and Maier[41].

GPC Mn [g/mol] 1200 D 3.1

31P-NMR aliphatic OH [mmol/g] 3.83 aromatic OH [mmol/g] 2.23

S-units[mmol/g] 1.39 G-units[mmol/g] 0.84

EA C=61.53% H=6.15% N=0.26% O=32%

Elemental analysis (EA) of beech organosolv (OL) showed a quite good fitting amounts of hydrogen, and oxygen per C9 (monolignol): H=7.49-8-50; O=2.53-2.95.[160] Glasser calculated with this EA values a methoxy (OCH3)=1.39-1.46.

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Hansen Solubility Parameter 2.5

2.5 Hansen Solubility Parameter

A prediction of miscibility, permeability or solubility of polymers, solvents, pigments, fibers, fillers etc. is important for diverse industrial production. The calculation of the Hansen solubility parameter (HSP)[210] is a common approach to predict interactions between two or more materials by giving a figure of merit to quantify the statement “like dissolves like”. Just as different forces play a role in interactions of molecules, the total energy of attraction is divided into three individual forms of energy. The sum of the dispersions cohesive energy 퐸퐷, the polar cohesive energy 퐸푃 and hydrogen bonding cohesive energy

퐸퐻 result in in the total cohesive energy 퐸.

퐸 = 퐸퐷 + 퐸푃 + 퐸퐻 Eq. 1

Inspired by the Hildebrand[211] solubility parameter 훿 calculated with the molar volume of the pure solvent:

훿 = √퐸/푉 Eq. 2 the Eq. 3 definite the relation between the total HSP 훿2 and individual HSP.

2 2 2 2 훿 = 훿퐷 + 훿푃 + 훿퐻 Eq. 3

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Results and Discussion 3

3. Results and Discussion

This chapter contains the experimental results, which are evaluated and discussed. The organization is adapted to both parts of the research questions as outlined in Section 1.2 of the Introduction.

PART I: (1) Analysis of beech bark and their solvent extracts - Section 3.1 (2) Analysis of beach suberin monomers – Section 3.2

PART II: (3) Synthesis and analysis of lignin derivative - Section 3.3 (4) Synthesis and analysis of lignin based NIPU - Section 3.4

Each section is composed of subsections containing the results of the specific analytical method used. All results are valuated, discussed and compared with the literature, where possible. Each section concludes with a summary subsection, which brings together the findings from all previous sections.

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PART I

PART I

The goal of the first part of this work is to ascertain if the byproduct beech bark is a potential source of polymer building blocks. Therefore, the extracts were analyzed qualitatively and quantitatively for the components. Tannins are already of great interest as aromatic ingredients for polymer products. The Soxhlet extraction method is displayed schematically in Figure PART I-1.

A poorly understood constituent of bark is the solvent-insoluble polymer suberin, on which the focus

DCM-extract

MeOH-extract

water-extract

Figure PART I-1: Schematic representation of the processes done in Part I. The blue-labeled steps are analyzed in Section 0 and the green-marked product is examined in Section 3.2. (s: solid, l: liquid) and a photo of the three solvent extracts. will lie. After depolarization by methanolysis the resulting monomers of the complex aliphatic-aromatic polymers were extracted and analyzed by diverse methods. Conventional techniques were used including: Fourier Transform InfraRed spectroscopy (FTIR) to give an overview of the type of functional groups; Gas Chromatography–Mass Spectrometry (GC-MS) a time-consuming method to quantify the

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PART I individual detected molecules; quantitative aromatic analysis via UltraViolet–Visible (UV-Vis) spectroscopy of detectable complexes; and nonspecific Elemental Analysis (EA). The Nuclear Magnetic Resonance spectroscopy (NMR), so far usually used for qualitatively or semi-quantitatively functional group analysis, was also used to determine statistical key-values such as average chain length and the number of functional groups per chain.

The efficiency of the depolymerization method was monitored in a number of ways: at first we observed the chemical modifications in the insoluble products (native bark BN, solvent extracted bark BE and

13 residual bark BR) via FTIR and C-NMR, and changes in their thermal behavior were also observed.

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Analysis of Beech Bark and Bark Extracts 3.1

3.1 Analysis of Beech Bark and Bark Extracts

This Section is divided into two main topics: the solvent extractable components of the bark in Subchapter 3.1.2 and the solid beech bark (before and after treatments) in Subchapter 3.1.1. The monitoring of the bark BN and bark residues BE and BR gave information about the efficiency of different process steps, see Figure PART I-1. The analysis of the extracts with regard to the first research objective: to identify and quantify potential polymer precursors, see Section 1.2. Following the procedure described in Section 4, the solvent extractable material will be analyzed in Subsection 3.1.2, and treated barks in Subsection 3.1.1 of these ingredients and the resulting modification of the bark and its solid residuals with different spectrometric and thermal methods. Both Subsections are divided into paragraphs dealing with the results and discussion of an analytical method. Conclusive, in Subsection 3.1.3 the results and reflections are brought together. The determination of the suberin monomers (SM) is analyzed in detail in Section 3.2.

3.1.1 Process Efficiency Observation by Solid Bark (Residues)

We were interested to see the effect of solvent extraction and transesterification of the beech bark on the extraction of beech suberin via transesterification. For comparison beech bark was extracted successively with solvents of increasing polarity to remove all extractable components, which could influence the results, before the extraction of suberin. The milled and differently treated bark samples were analyzed by spectroscopy via FTIR and NMR, and thermally via DSC and TGA.

Natural Water Content of Different Bark Samples and Pre-Dried, Milled Bark Samples

The water content 풘 for the fresh Bark Sample (BS) was measured to determine the dry matter of the bark. For BS-1 w was 41.7 ±3.5 wt.%, for BS-2 w was 43.0 ±2.5 wt.% and for BS-1+BS-2 pre-dried at 75°C w was 3.9 ±0.3 wt.%.

The average water content 푤 of the pre-dried freshly fine-milled bark samples (BN) is 3.42 ±0.12 wt.%., the water content of each sample is listed in Table 6.2-1 more detailed.

The lower humidity/water content of BS-3 compared with fresh BS-1 and BS-2 can be explained by its storage in the freezer for one year before measurement: The cold environment had removed water from the bark.

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Process Efficiency Observation by Solid Bark (Residues) 3.1.1

BS-3 was separated into two parts of approx. 10% by mass Inner Bark (IB) and approx. 90% by mass Outer Bark (OB), according to the results of Prislan et al.[59], and the beech periderm was determined to be 7% of the bark thickness.

The water content 푤 of the defrosted non-milled OB was 27.2 ±1.3 wt.%, and for the IB it was 27.2 ±3.4 wt.%. The bark humidity 푢 for the defrosted non-milled OB was 37.4 ±2.5% wt.% and IB 37.5% ± 6.4%.

FTIR Measurements of Solid Bark (Residues)

The solvent-soluble components and suberin from bark were measured in the FTIR spectrometer in ATR mode, for Non-extracted (BN), Extracted bark (BE) and Residual bark after depolymerization (BR) (see Figure 3.1.1-1). It was necessary to normalize the three spectra to get comparable, quantitative bands after baseline and ATR correction. Three bands seem to be suitable for normalization due to their high intensity: the absorption band at 푣̃ =1615 cm-1 pertaining to C=C stretch vibrations of aromatic rings (typical for lignin); a typical band for (poly)saccharides at 푣̃ =1023 cm-1, and the C-H vibration of aliphatic chains methylene 푣̃ =2916 cm-1.

Figure 3.1.1-1: ATR-FTIR spectrum of native milled bark (BN) is black, of Soxhlet extracted bark (BE) is red and of bark residue after methanolysis (BR) is blue. Observed transmittance band are marked with dotted lines and labeled with corresponding wavenumber and listed in Table 6.2-2.

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Process Efficiency Observation by Solid Bark (Residues) 3.1.1

However; none of these bands alone is well suited for normalization because each treatment step (extraction or methanolysis) affected the aromatic, saccharide and especially the aliphatic chain concentration. Therefore nit was decided to use the band of aromatic vibration for the normalization

[63, 65, 114] between BN and BE since it is known that during extraction saccharide is usually removed. Normalization to the aliphatic band was also rejected, as the transmittance functions for each sample were significantly different in the region around this wavenumber. Thus the saccharide band was used to normalize BE and BR, since this band was least influenced by methanolysis.

However, it must be said that this normalization is not ideal, since aromatic compounds were found in the extracts, and saccharides were found in the extracted suberin monomer (SM) fraction, cf. Subsection 3.1.2. A quantitative evaluation was feasible only to a limited extent, since it was not possible to find another explicit assignable absorbance band with significant intensity for which it could be certain, would not change in intensity during the process steps.

The observed wavenumber of vibration absorbance bands are marked with dotted lines in Figure 3.1.1-1, and listed with the assigned molecular vibration in Table 6.2-2. The biggest difference between BE (red)

-1 -1 and BN (black) was found at 푣̃ =2917 cm and 푣̃ = 2850cm . These bands pertain to asymmetric and symmetric C-H vibrations of alkanes, and in this case are inducted by “waxes” in the milled bark. The intensity degradation of a typical ester or carboxylic acid carbonyl group at 푣̃ =1736 cm-1 supports this suggestion. Furthermore, saccharide bands around 푣̃ =1023 cm-1 decrease, which was expected, since hemicellulose and other soluble saccharides should be removed by solvents. If we assume that aromatic compounds were removed during the solvent extraction, the extent of saccharide and “wax” degradation should increase.

-1 -1 The transmittance of BR was less than that of BE especially at 푣̃ =2917 cm , 푣̃ =1736 cm and

-1 -1 푣̃ =1615 cm . These reductions were in proportion to 푣̃ =1023 cm . This indicates loss of aromatic and aliphatic compounds, and indicates that suberin was removed by methanolysis.

NMR Measurements of Solid Bark (Residues)

Another method to monitor the course of suberin extraction is to observe the aliphatic chain carbons via

13 a solid-state C-NMR of Non-extracted (BN), Extracted (BE) and Residual bark (BR) after depolymerization, see Figure 3.1.1-2. The resulting spectra were normalized to the highest shift signal (72 ppm) corresponding to polysaccharide carbons.

No significant differences were observed between BE (red) and BN, but compared to BR (blue), BR loses nearly all aliphatic carbons, see peak at δ=30 ppm in Figure 3.1.1-2. Also the peak of carboxylic acids carbonyl carbons (C=O) at δ=172.1 ppm decreases. At the same time a sharp signal at δ=168.4 ppm

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Process Efficiency Observation by Solid Bark (Residues) 3.1.1 seems to increase. It is suggested that this peak contributes to carbonyl carbons (C=O) of methyl esters. It is expected that after methanolysis all carboxylic acids, and hopefully all ester bond linkages, would be transesterified to methyl esters. The methylene carbons of aliphatic chains of suberin at δ=30 and

δ=33 ppm disappeared. This is also a sign of successful suberin monomer (SM) extraction from BE by methanolysis. All other signals were identified from the literature [88, 95, 105, 109, 129], see Table 6.2-2 in Chapter 6 Supporting information.

Figure 3.1.1-2: Solid-state 13C-NMR of solvent extracted bark BE (red) and rest bark after methanolysis BR (blue). The red arrows mark the increasing signals, the blue arrows the decreasing signals after the methanolysis of the BE. The signals are normalized at δ=105.2 ppm, which supposed to correspond to polysaccharides (cellulose) carbons.

Thermal Analysis of Solid Bark Residues

It was also of interest to investigate the thermal degradation of the residual bark, or see if the observed glass temperature Tg is influenced by suberin. Therefore, analysis via TGA and DSC was performed for the treated bark samples. One example thermogram for each sample is shown in Figure 3.1.1-3. The corresponding first derivative of the thermogram is shown on the right side. All results from TGA for different Td, onsets of the maximal Td and ash content after heating to 900°C, and the Tg from the DSC are summarized in Table 3.1.1-1.

Raw bark (BN) was measured five times, since it showed large temperatures variations, especially for the first two degradations. The average results and their standard deviation are shown in Table 3.1.1-1. Albert-Ludwigs-Universität Freiburg 47

Process Efficiency Observation by Solid Bark (Residues) 3.1.1

It was possible to observe up to six different degradation temperatures (maxima of the first derivative curves labeled with the related temperature). The broadest degradation (max) was monitored at approximately Td3: 360°C for BN and BE. This temperature is characteristic for cellulose. The shoulder

[170, 213] at approximately Td2: 294°C pertain to lignin . Furthermore, it was observed that the degradation maximum of cellulose Td3 had fallen by approx. 40°C after depolymerization of the suberin. Possible reasons for the observed decrease in the degradation temperature include manifold: changes in the crystallinity of cellulose; side reactions in the harsh methanolysis conditions, for example carboxylic acids of cellulose may be esterified; suberin polymer may be protecting cellulose; hydrolysis of cellulose etc.

Unexpectedly, it was observed that with each processing step Td3 decreased after each treatment. The mass portions, as inferred from integration of Td3 decreased with each processing step: 59.0% for BN,

56.2% for BE and up to 51.6% after depolymerization. However, these results could be massively influenced by the weight loss caused by Td2. Lignin showed typically a second degradation temperature

[170] [213] approx. at 750°C cf. Figure 2.4.2-1 or Zhou et al. , which fits well to the observed Td5. In the case

[107] of bark, Td2 and Td5 may belong (partially) to the aromatic domain of suberin (“lignin-like” structure).

Td1 Td2 Td3 Td4 Td5 Td6

Figure 3.1.1-3: TGA thermograms of different treated bark sampels are on the left side, and the first derivatives of the thermograms are on the right side. On the top (black) is untreated bark, in the middle (red) is extracted bark and below (blue) remaining bark after the methanolysis.

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Process Efficiency Observation by Solid Bark (Residues) 3.1.1

This assumption refers to the thermogram of BR (blue), where the degradation peaks at Td2 and Td5 almost disappear. The typical ether bonds between monolignols should be not influenced by a transesterification reaction, but some bonds are esters and they could have been opened, and the lignin macromere could have been dissolved in methanol. Transesterification may also have influenced the Tg3 of BR.

During the five repetitions of TGA measurements of BN, a variation of Td1 higher than 10%, and of Td2 higher than 2% of the heating temperatures was observed. Whereby a deviation of Td2: 300 ±6°C is negligible because it belongs to a shoulder of the maximum peak Td3,MAX , so it was difficult to analyze the absolute temperature of Td2. This declares also the 3% standard deviation of the onset temperature

Td3, onset. Solvent extracted bark BE and depolymerized bark BR did not show degradation temperature variations of greater than 2%.

Table 3.1.1-1: Mean degradation temperatures, onset and end temperature of the maximal degradation step and the mass loss after heating to 900°C of four milled bark samples detected by TGA. SD is the standard deviation. The glass temperature and glass temperature onset was detected by DSC.

Raw bark (BN) Soxhlet extracted Suberin extracted bark Mean SD bark (BE) bark (BR)

Td1 [°C] 64 6.9 55

Td2 [°C] 300 6.1 296

Td3, max [°C] 360 1.3 361 324

Td3, onset [°C] 294 7.8 250

Td4 [°C] 523 1.5 505 497

Td5 [°C] 702 2.4 687 640

Td6 [°C] 783

wt.900°C [%] 21.8 1.1 7 25

Tg [°C] 45 41

Tg, onset [°C] 17 21

The large shifts in the Td1 values of BN (64.3±6.9°C) suggests that the degradation temperature was influenced by substantial deviations in the water content and amount of the other volatile matter in a sample, which depends on pretreatment, storage and milling[214].

The evaporation of residual methanol was responsible for the degradation peak Td1 of BR at 55°C of BR. This was used for the methanolysis reaction, and is therefore irrelevant.

After heating to 900°C in the TGA, the volatile components 76.9 wt.% BN and 75.5 wt.% BR. This is, comparable with that of our project partner EnBW, who determined values of 74.3 wt.% and 71.5 wt.%

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Solvent Extracts of Beech Bark 3.1.2

(using DIN EN 15148) for BN and BR respectively. Other important values if the rest bark is to be burned for energy production are ash content and calorific value. EnBW investigated values for fresh (wet) bark, for BN and BR. Their results are summarized in Table 6.2-3. Furthermore, they measured the phenolic index in the eluent and the trace elements potassium, nitrogen, calcium, and sodium.

The caloric value of BR (16,592 kJ/kg) decreased in comparison to BN (14,548 kJ/kg). However, more problematic was the higher ash content, inter alia, produced due to the higher sodium content (see Table

6.2-3). This is an important issue when using BN for energy production by burning. An extraction and/ or neutralization of the BN bark could wash out the main part of sodium.

The phenolic index in eluent of BN (89.6 mg/l) and BR (28. mg/l) samples indicates the expected decrease after depolymerization, see Table 6.2-3.

The DSC measurements of dry BN result in a glass temperature for bark Tg(1)=54.2°C during the first heating cycle and Tg(2)=44.7°C during the second, for BE is the Tg(1)=40.7°C and Tg(2)=46.42°C. The BR thermogram does not show any Tg.

The loss of Tg could also indicate that the previously observed Tg in BE and BN was generated by suberin.

3.1.2 Solvent Extracts of Beech Bark

The fine milled beech bark was successively extracted with solvents of increasing polarity to remove all soluble material so that they do not influence the monomers of the methanolysis, see Section 3.2. The extraction solvents were evaporated and the extractable mass determined by gravimetric analysis. It was possible to extract 1.3% of DCM-, 5.1% of MeOH- and 3.2% of water-soluble material with reference to dry bark mass (determinated in Subsection 3.1.2) with successive Soxhlet extractions.

Compared to the extractable material found in the literature, see Table 2.2.1-2, the extracted absolute amounts were lower than expected. However, the proportions corresponded to those ones found in the literature.

FTIR Measurements of the Solvent Extracts

The solvent free extracts were measured via attenuated total reflectance (ATR)-FTIR, and the spectra obtained are shown in Figure 3.1.2-1. The FTIR spectra of solvent extracts are normalized at wave number 푣̃ =2916 cm-1, wich pertains to an asymmetric carbon-hydrogen bond vibration ν (C-H) for alkanes. This is not optimal to show absolute quantities, and is used merely to get an idea of the extract composition. The observed transmissions bands are marked with vertical dotted lines, whereby gray

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Solvent Extracts of Beech Bark 3.1.2 dotted lines are previously observed bands, cf. FTIR spectra of barks Figure 3.1.1-1, newly found bands are labeled and indicated by dotted black lines. The wavenumber bands and their related kind of vibrations are listed in Table 6.2-2.

At first glance huge differences are recognised between the non-polar DCM (green) extract and the polar MeOH (blue) and water extracts (red). As one might expect a broad O-H vibration band (푣̃ =3800- 2300 cm-1) wass obseved with the polar solvents, since in DCM it has lower untensity. The predominant bands in DCM extract are aliphatic C-H vibrations of asymetric methyl (CH3) and methylene (CH2) group (푣̃ =2958; 2916 cm-1), symmetric methylene group (푣̃ =2850 cm-1) and asymmetric deformation vibration (푣̃ =1461 cm-1) of alkane methylene or methyl groups. Additionally the symmetric deformation band 푣̃ =1379 cm-1 and rocking (푣̃ =725 cm-1) of methylend confirms the high aliphatic chain content in the DCM extract.

-1 The band 푣̃ =1379 cm could also be caused by a phenolic –OH strech vibration. This proposition is supported by the presence of typical linin units absorptions bands at 푣̃ =1260 and 1167 cm-1. The

Figure 3.1.2-1: ATR-FTIR measurements of the DCM-extract (green), methanol-extract (blue) and water (red). Observed transmittance bands are marked with black dotted lines and labeled with corresponding wavenumber, the grey dotted lines are already found in Figure 3.1.1-1 and listed in Table 6.2-2.

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Solvent Extracts of Beech Bark 3.1.2

carbonyl (C=O) vibration bands (푣̃ =1763; 1714 cm-1) indicate the presence of ester, acid or ketone groups, also found in MeOH extract.

In contrast to the aliphatic extract, methanol and water extracts show predominant typical absorption bands for saccharides (푣̃ =3800-2300; 1235; 1113; 1075; 1032; 1023; 883; 820 cm-1). The finding of saccharides was expected in polar and medium-polar solvent, cf. Table 2.2.1-2.

Ultraviolet-Visible Spectroscopy (UV-Vis) Measurements of Solvent Extracts

Some amounts of aromatic compounds were found in measurements by FTIR, NMR and GC-MS. The MeOH-extract especially showed proton shifts, which indicates that condensed tannins (proanthocyanidins) could be one of the main components.

In this paragraph we discuss the results of a photometric method (UV-Vis) using two different complexing agents: Folin-Ciocalteu (FC) and Vanillin-HCl to quantify the aromatic content in the solvent extracts. The FC agent stains phenols and tannins, allowing determination of the total phenol content. Vanillin-HCl agent dyes condensed tannins pink, thus an observation of the substances can be made by UV-Vis spectrometry.

Total Phenol Content and Indirect Determination of Tannins (Folin-Ciocalteu Method)

A calibration of the absorbed visible wavelength related to concentration is necessary for the determination of the total phenol content. Gallic acid was chosen as the reference substance. Before preparing the linear calibration function, the absorbance spectrogram of gallic acid was measured in the range 400-1000 nm at defined time intervals after preparation of the sample.

In Figure 3.1.2-2a. the time-dependent spectra are shown, and Figure 3.1.2-2b. reveals an application of the absorbance as a function of time for the wavelength at 725 nm, as recommended in the literature[215, 216], and at 745 nm since the absorbance maximum was at this wavelenght. After spectra measuring other samples containing phenolic groups, see Figure 3.1.1-3c., we decided to measure the samples at 745 nm in order to minimize errors.

According to the literature [215], it is necessary to measure the absorption after a 40 min incubation period (Singelton et al. [217] recommend 2 h). To check this, the influence of incubation time on the absorption was measured at various times. Figure 3.1.1-3b clearly shows that after 10 min the absorbance at both wavelengths stabilizes and do not change the next 40 min. This shows that a measurement after 40 min is not influenced anymore that much by time and that it is not necessary to measure in minute steps.

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Solvent Extracts of Beech Bark 3.1.2

Figure 3.1.2-2: Measurements for application of the Folin-Ciocalteu-Method: a-b. Kinetically observation of gallic acid spectra, c. spectra of different substances containing phenol structures, d. linear calibration curve with confidence band and forecast ribbon. The red vertical lines in a. and c. mark the wavelength recommended in the literature [108, 109] and the dashed red line was the one used in this work. The blue vertical line in b. marks the time that elapsed before the samples were measured.

For the calibration the absorbance of eight different concentrations of gallic acid solutions was measured at 745 nm. Figure 3.1.1-3d shows the absorbance as a function of the GA concentration. The slope of the regressions line is 29.68 ml/µg ±0.25 ml/µg with a determination coefficient of 푅2 = 0.99945.

The amounts of phenolic content were 1.52 wt.%, 17.96 wt.% and 20.35 wt.% of the DCM, MeOH and water extract. The amount of aromatics that were not removable with PVPP was 0.88 wt.%, 0.76 wt.% and 4.81 wt.%. Consequently 0.64 wt.%, 17.20 wt.% and 15.54 wt.% of proanthocyanidins [218] were found in the extracts (see Table 3.1.2-1).

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Solvent Extracts of Beech Bark 3.1.2

Table 3.1.2-1: Summary of the photometric analysis. Total phenols, PVPP non-precipitated phenol content and condensed tannins content in percent by weight of the extract mass for the different solvents.

PVPP non precipitate total phenols condensed tannins phenols

DCM extract 1.52% 0.9% 0.7%

MeOH extract 17.96% 0.8% 62.9%

Water extract 20.35% 4.8% 52.6%

It is important to interpret these yields with caution, since the calibration is based on the gallic acid model. The phenols in our sample could have a shifted absorption maxima, different mass and the absorption intensity could also be influenced by substituents and matrix. Additionally, the FC reagent could also stain other antioxidants in the extracts.

Determination of Condensed Tannins

The percentage of condensed tannins was 0.7 wt.% for the DCM-extract, 62.9 wt.% for the MeOH- extract and 52.6% for the water extract, as determined by the Vanillin-HCl method. These weight percentages correspond to 4.9 wt.% of dry bark. This means that over 50 wt.% of the whole extracted mass were phenolics.

As for the Folin-Ciocalteu-Method, all values (see Table 3.1.2-1) should be considered with caution, since they were calculated with a molar attenuation coefficient for (2R, 3S, 4R)-leucocyanidin, which does not necessarily correspond to the real system.

NMR Measurements of Solvent Extracts

As with FTIR measurements, NMR is a quick method to obtain an overview of the solvent extracted beech bark material. The NMR-spectra of the DCM extract were measured in CDCl3, the MeOH-extract in d6-DMSO and the water extract in deuterium oxide. The proton spectrum of all three fractions is shown in Figure 3.1.2-3.

It is clearly seen that the ingredients vary widely, and an identification of the individual substances is challenging. However, it is possible to give an overview of the contained substances because of the shifts of the typical proton signals.

It was expected that non-polar substances or “waxes” accumulate in the non-polar solvent DCM, as indicated in the literature[56]. The green spectrum of DCM extract verifies this assumption. Almost none of the signals were shifted more downfield than the frequency region marked by green. This region shows

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Solvent Extracts of Beech Bark 3.1.2

CHCl3 DMSO

H2O

Figure 3.1.2-3: 300 MHz proton NMR of all solvent extract fractions: green is the DCM-extract in CDCl3; blue the MeOH-extract in d6-DMSO; and red the water-extract in deuterium oxide. the typical proton shifts of aliphatic chains without groups such as oxygen containing groups that would increase the frequency sweep 푣0.

The more-polar methanol (blue) and water (red) extracts do not show many aliphatic protons. The main part of the peaks are shifted downfield to δ=3-5 ppm (red highlighted), this indicates the presence of heterogeneous atoms like oxygen and is a typical for saccharides and tannins.[56] Additionally some aromatic protons which shifted to δ=6-7 ppm (blue highlighted), were extracted with methanol.

Chromatographic (GC-MS) Measurements of Solvent Extracts

The GC-MS analysis of the three solvent extracts is summarized for the DCM extract in Table 6.2-6, for the MeOH in Table 6.2-7 and the water extract in Table 6.2-8. For simplicity the analyzed components of the dissolved material are reported in their acid or alcohol form.

To obtain a better overview, the data of Table 6.2-6, Table 6.2-7 and Table 6.2-8 are aggregated to the substance classes in Table 3.1.2-2 and visually supported by Figure 3.1.2-4.

The fraction extracted by DCM contained the greatest variety of substance classes, thereby just 56% on average of the TIC peak areas could be identified from the NIST library. They consist of aliphatic acids (25%) with chain length between 6 to 26 carbons (C6-C26), aromatics (17%) belonging to methoxy

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Solvent Extracts of Beech Bark 3.1.2

Table 3.1.2-2: Heavily summarized presentation of identified substances groups found as areas in TIC chromatogram of the solvent extracts GC-MS measurements.

DCM MeOH water dihydroxy alkanes 0.14 0.00 0.10 aliphatic alcohols 7.91 0.00 0.00 aliphatic acids 24.51 0.00 0.57 dicarboxylic acids 0.06 0.00 0.00 hydroxy acids 0.77 0.00 0.21 aromatics 17.26 0.92 1.30 saccharides 0.64 59.31 59.18 other 5.04 5.06 7.31

Figure 3.1.2-4: Graphical representation of Table 3.1.2-2. benzyl aldehydes, alcohols or acids, methoxy hydroxy benzyl alcohols or aldehydes, etc. and aliphatic alcohols (8%) with chain length of between 18 and 24 carbons for the most part, see Table 3.1.2-2. In addition, ω-hydroxy acids, mono- and di-saccharides, dicarboxylic acids, dihydroxy alkanes, aliphatic aldehydes, anthracenediones, phytosterols, amyrins, glycerol and saccharide- or fatty acid esterified glycerol were found in the DCM fraction.

Comparable data were found by Streibl, Konečný and Mabdalic [69, 144, 145] in petroleum extract of Fagus sylvatica bark. They found aliphatic alcohols with chain length of between C12 and C31, saturated fatty acids of C8-C24, and unsaturated C12-C24 fatty acids, hydroxy carboxylic acids in the range f C12-C24) and β-sitosterol (phytosterols) by GC.

The water and methanol extracts were found to have NIST identifiable substances for 69% and 65% of the total TIC peak areas. These two fractions mainly consist of mono and disaccharides starting with triose up to hexoses and their alcohol or acid derivatives (e.g. thretiol; arabitol; or threonic acid). Also glycerol esterified saccharides were observed in the extracts. In addition small quantities of some simple phenols were found, but also catechine (flavonoid). In the water fraction the polar propane-1, 3-diole short aliphatic acids or hydroxy groups containing aliphatic acids are found in small amounts.

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Summary Measurements of the Solvent Extractives and the Treated Barks 3.1.3

3.1.3 Summary Measurements of the Solvent Extractives and the Treated Barks

 The extracts from beech bark were on average 10 wt.%. Compared to extracts from Quercus suber L. 16 wt.%[125], various Pinuses (28-68 wt.%)[70] or Betula pendula R. (29 wt.%)[125] the yield from beech was low. Dietrichs et al. [63] found 34 wt.% solvent extractives in Fagus sylvatica using diethyl ether (6.0 wt.%), ethanol (13.6 wt.%) and hot water (14.4 wt.%). This is three times more than we found, but the yields in each solvent type (maximal yield rate with medium-polar solvent (MeOH) and minimal with the non-polar DCM) are proportional to the yields rations of Dietrichs et al. [63] cf. Table 2.2.1-2.  The extraction result of Streibl, Konečný and Mabdalic [69] (they extract with petroleum 0.77%) are somewhat comparable with our DCM-extract, whereas Dietrichs et al. [63] (extracted beech bark with ether – ethanol – water ) obtained significantly higher amounts of extractable material 6.0% – 13.6% – 14.4%.

 The spectra of FTIR spectra of BE indicate the extraction of saccharides and “waxes” with solvents. This confirm with the according bands of saccharides and “waxes” were found by FTIR and the typical shifts observed by NMR in the spectra of the solvent extractives.  Despite the low amount of DCM extractable material, the proportion of “waxes” is minor. It was

confirmed with the solid state NMR of BN and BE. Just in the analyzed DCM extract is via FTIR, NMR and GC-MS the extraction of waxes observable.  The aliphatic part (“waxes”) of the DCM-extract had chiefly just one functional group, and the chains were usually saturated.  Saccharides were the main components in the water and methanol extract, as observed by NMR, FTIR and GC-MS. This corresponds well with the high sugar amounts found by Dietrichs et al. [63] for beech bark. The NMR and FTIR data made it clear that a high amount of sugar was expected in the methanol and water fraction.  The aromatic content of DCM-extract is not very consistent, and was clearly detected by the various methods. While the FTIR, NMR, UV-Vis data showed only a small quantity of aromatic substances, the GC-MS data contradicted this finding. It can be suggested that the largest error lies in the GC-MS method itself: It is known that not all substances in the sample were volatile for example ternary and dual esterified glycerols (typical fats) and other macromolecules. This substances cannot be seen in the TIC that the proportion of observable substances apperes to be greater. A transesterification or hydrolysis should be done before taking measurements so as to minimize losses by evaporation.

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Summary Measurements of the Solvent Extractives and the Treated Barks 3.1.3

 The low concentration of aromatics in both the methanol and water extracts, found by GC-MS is explainable with the same arguments, but instead of non-evaporating “waxes”, complex aromatics e.g. tannins or lignin were not detected, just simple aromatic molecules or monomers such as catechine could be observed. This theory confirms the data received from the photometric measurements, see the amount after the precipitation of complex molecules with PVPP.

 From the FTIR data of BE it is probable that an aromatic content could be extracted. This would

mean that the aromatic band of BE should decrease, and a normalization to BN would be an

overestimation of the bands. This automatically gives an overestimation of BR. However, with

the overestimated bands of BR, there should also be a decrease in the aromatic band monitored, and the TGA data showed a decrease in aromatics. This would confirm the thermal behavior of

lignin in BR as well.

 In contrast to BR, the thermal analysis of BE showed no significant differences. The loss of waxes,

saccharides and aromatics only had a minor influence on the thermal behavior of BE.  TGA and the phenolic index of EnBW showed that phenolic compounds are extracted during depolymerization. We expect that these are lignin or/and an aromatic domain of suberin.

 The solid-state NMR spectra of BR point out that the amounts of the carbon signals for aliphatic chains and for carboxylic decreased after the transesterification. This is a confirmation of the transesterification method for suberin extraction.

 The BR is suitable for energy production. Only the high sodium content and thus the increasing ash content could be problematic.

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Suberin Monomer Analysis 3.2

3.2 Suberin Monomer Analysis

The analytical results of the extracted suberin monomers (SM) by transesterification from solvent extracted bark (BE), see Figure PART I-1. Techniques were used to provide information on the monomeric mixture of SM.In the context of using the monomers as a polymer precursor, a statistical determination of the aliphatic chains via NMR was used and compared with the analytical results of other techniques, such as highly specific GC-MS, or nonspecific elemental analysis, thus this Section corresponds to second research question (refer to Section 1.2).

Depolymerization and purification (see Figure 5.2.3-1) resulted in 3.7 wt.% ±0.9 wt.% suberin monomers (SM) referred to solvent extracted free bark (BE) using 13 repetitions. Thereby, a 9 wt.% mass loss of BR was obtained gravimetrically compering with BE.

Based on the assumption that transesterification results in addition of methoxy groups, and thus a total mass increase, three conclusions can be made: the observed mass loss of BE should be higher than 9 wt.%, the real yield of SM 5.2 wt.% ±0.4 wt.% was lower than previously obtained. This indicates that during the purification the main part of SM was lost. Along with the salts other water soluble components like glycerol are removed during purification [90, 105, 114, 116]. For comparison, Perra et al. [108] extracted suberin and lignin from Fagus sylvatica L. with a dioxane/water solution following the Björkman method [219], and precipitated the lignin (yield 0.16%). After precipitation of lignin 29% of the extract was defined as suberin.

This assumption is also confirmed with the yields of non-extracted bark depolymerization yield of 5.21 wt.% ± 0.3 wt.% [212]. The yield was less than expecteded (minimum SM + MeOH extract should be 3.7 +5.1 wt.%)

Beech bark contains significantly less amount of suberin comparing with suberin yields of Quercus suber L. (to 60%) [72, 90, 94, 105, 114, 116, 122, 125, 126].

3.2.1 FTIR Measurement of Depolymerized Suberin

Suberin monomers were extracted from beech bark and analyzed via FTIR in ATR mode. The resulting spectrum is shown in Figure 3.2.1-1. The gray lines correspond with the previously observed and discussed bands of different treated beech barks (BN, BE, BR), see Figure 3.1.1-1 and of the solvent (DCM, MeOH, water) extracts of the beech bark, see Figure 3.1.2-1. Here, the newly observed bands are shown as dotted black lines, labeled with the corresponding wave number, and also summarized in Table 6.2-2.

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FTIR Measurement of Depolymerized Suberin 3.2.1

-1 The afore monitored bands fitted well with the new spectrum: 푣̃ =2916 and 2850 cm pertains to asymmetric and symmetric stretch vibration; 푣̃ =1461 and 1379 cm-1 deformation vibration; 푣̃ =725 cm- 1 rocking of methylene groups; 푣̃ =1736; 1714 cm-1 is typical for carbonyl vibration; and 푣̃ =1513, 1423, 1369, 1260, and 1167 cm-1 is a characteristic band for substituted phenols likely to occur in lignin.

All things considered, the spectrum is unambiguous. The main components of the SM are aliphatic chains and some substituted phenols. The aliphatic chains seem to be very long and terminal functionalized, since the characteristic bands for methyl (푣̃ =2958 cm-1) are missing.

Figure 3.2.1-1: ATR-FTIR spectrum of suberin monomers

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UV-Vis Spectroscopy Measurement of Suberin 3.2.2

3.2.2 UV-Vis Spectroscopy Measurement of Suberin

As in Subsection 3.1.2, this method was used to determinate the total phenol, tannin and condensed tannin content in the SM, and in the initial solvent water after purification in DCM (cf. Figure 5.2.3-1).

Total Phenol Content and Indirect Determination of Tannins (Folin-Ciocalteu-Method)

The determination of total phenol content via the Folin-Ciocalteu-Method was performed and calculated as before for the solvent extracts in Subsection 3.1.2, see page 52. SM was suspended overnight in water, centrifuged and decanted. The measurable phenol content before and after the precipitation with PVPP was 1.49 wt.% and 1.35 wt.% respectively, see Table 3.2.2-1.

During the purification a substantial mass loss was observed. The liquid-liquid extraction using water was extracted again three times with diethyl ether. Both solvents were dried and treated as SM before.

The solid matter from the purification water contained only small almonds of phenols. Approximately half of this was precipitated with PVPP (cf. Table 3.2.2-1). This was expected as the main mass fraction should be salts, which were produced by neutralization of the methoxides.

The phenol content of the solid matter from the purification ethyl ether was higher than 150 wt.%. This is possible, hence some phenols have a higher molecular absorbance coefficient [217] than galic acid (GA) which is used for the calibration. A content of phenols with smaller molar mass than GA would lead the calculated percent by weight are higher than 100 wt.%. This theory is backed by the observation that the simple phenols are not precipitable with PVPP.

Determination of Condensed Tannins

The amount of condensed tannins with the Vanillin-HCl method was not detectable (NQ), as one might expect (cf. Table 3.2.2-1). Marginal quantities found in diethyl ether could be extracted from the remaining bark during the alkali methanolysis process. In any case the value was so low that it can be ignored.

Aromatic compounds were also found by B. Perra, J.-P. Haluk and M. Metche [108] in Fagus sylvatica L. after extracting with dioxane/water, precipitating of lignin and depolymerizing with boron trifluoride. They found 30% of the chloroform extractable suberin belonged to a phenolic fraction and 70% to an aliphatic fraction. The much higher ratio of aromatic compounds was affected mainly by the extraction method developed by Andreas Björkman for the extraction of lignin and cellulose [219]. In contrast to this method usually used for dissolving the aromatic lignin the approach of methanolysis is depolymerizing mainly the aliphatic domain. This results in the weigh lower mass of aromatics.

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

Table 3.2.2-1: Photometric analyzed total, PVPP non precipitate phenol content and condensed tannins content in percent by weight referred to the extract mass of the samples. The calculated weight percent of total and PVPP non precipitate phenols rest on the gallic acid calibration, condensed tannins based on the molar attenuation coefficient for (2R, 3S, 4R)-leucocyanidin [220].

total phenols PVPP non precipitate condensed tannins

[wt.%] phenols [wt.%] [wt.%]

SM 1.49 1.35 NQ Purification water 0.52 0.25 NQ Purification diethyl ether 156.05 166,85 0.05%

3.2.3 Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI)

For the analysis of suberin monomers (SM) and their functionality various Nuclear Magnetic Resonance

(NMR) methods were used. Proton NMR was used before and after derivatization of SM with trichloroacetyl isocyanate (TAI) samples. Carbon NMR gives more information about the aliphatic/aromatic character of SM. Phosphorus NMR after derivatization of SM with 2-chloro-4, 4, 5, 5- tetramethyl-1, 3, 2–dioxaphospholane (TMDP) permit insights about the hydroxyl nature in the monomers. Diverse two-dimensional techniques like Diffusion-Order SpectroscopY (DOSY), Two- dimensional nuclear magnetic resonance spectroscopy (TOCSY), Heteronuclear Single Quantum coherence Spectroscopy (HSQC) and Heteronuclear Multiple Bond Correlation (HMBC) shows more about the SM character and can be used to verify the conclusions from the one-dimensional measurements.

Proton NMR

Depolymerized suberin (SM) was measured via proton NMR in deuteron chloroform, see Figure 3.2.3-1.

This shows two proton NMR spectra: the red spectrum is a typical SM proton spectrum; the blue is the

SM spectrum after the derivatization of the hydroxyl groups with TAI (TAI-SM). The peak shifts assigned to protons depended on the functional group, the latter labeled peaks are listed in Table 3.2.3-1 with their assigned shifts or shift ranges.

Signals at δ=0.56-0.93 ppm and δ=0.93-2.38 ppm are due to methyl and methylene groups of aliphatic chains. A small singlet found at δ=2.88 ppm could be due to epoxide α protons [21, 39], this signal disappears in the TAI-SM spectrum. Various peaks shifted to δ=3.40-3.66 ppm pertain to α protons of remaining methanol, aromatic secondary or primary hydroxyl groups disappear after forming an urethane linkage with TAI, see the zoom in spectrum in Figure 3.2.3-2. Albert-Ludwigs-Universität Freiburg 62

Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

Otherwise new peaks are monitored in the TAI-SM spectrum (blue). The triplet signal at δ=2.81 ppm (2H, t, J=7.27 Hz) is suggested to be α-protons of a TAI derivative carboxylic acid forming a O-acylurea bond. During the neutralization of the methoxide solution there might be some methyl esters that were hydrolyzed to carboxylic acids and methanol.

The new triplet and quintet at δ=4.2 ppm (2H, t, J=6.72 Hz) and δ=5.0 ppm (H, quint, J=4.90 Hz) [102, 121] pertain to α protons of primary and secondary hydroxyls forming urea bonds with TAI.

The advantage of using TAI is especially obvious in consideration the square marked shift region in Figure 3.2.3-1. This paragraph is magnified illustrated in Figure 3.2.3-2. Three different proton signals overlap in the region δ=3.4-3.7 ppm, the peaks belong to α protons of: primer hydroxy groups (predicted

solvent

b i solvent j h f c a d m l k g e

Figure 3.2.3-1: Proton NMR (300 MHz) spectra of SM (red) and derivatized TAI-SM (blue) dissolved in CDCl3. triplet); seconder hydroxyl groups (predicted quintet); and methyl protons of methyl ester groups (singlet). This dilemma was solved using the derivatization agent TAI. The appropriate proton NMR spectrum (blue) is illustrated in Figure 3.2.3-1, the framed area is enlarged in Figure 3.2.3-2. SM and

TAI-SM spectrum are still comparable, only the overlapping peaks α-protons of prim and sec. hydroxyl

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3 groups (δ=3.4-3.7 ppm) are shifted downfield. However, the singlet of the methyl ester methyl group is still there. A typical triplet and quintet is observed for the α-protons of the new urethane bonds. This enables separate integration of the peak areas. The integrals of these regions were used for all calculations at later paragraph: “Calculation of Chain Length and Number of Functionality per Chain” p. 70.

The disappearing epoxide protons at 2,88 ppm can be explained by the reaction of epoxides with isocyanates to 2-Oxazolidones [221, 222]. The expected proton peak should shift around 6 ppm [222] downfield. However, it was not possible to be certain about the identity of this peak.

Another observable phenomenon is marked with a circle in Figure 3.2.3-1. The hydroxyl derivative deshielded not only the α-protons but also β-protons of the hydroxyl and carboxylic acids groups. TAI also influenced the effective field strength of the β-protons, which are shifted to higher frequency δ=1.6 ppm (2H, quint, J=7.38 Hz). A separation of the methylene groups was not possible to observe, since the chemical shift is too small [102].

j i

k h

l

Figure 3.2.3-2: Enlarged section of the 300 MHz proton NMR spectra presented in Figure 3.2.3-1. The red spectrum belongs to the SM sample and the blue spectrum belongs to TAI-SM.

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

A lake of aromatic protons was observed in the SM spectrum after the transesterification, in agreement with Cordeiro et al. [32]. An increase in baseline of only around δ=6.8 ppm could indicate the existence of different substituted aromatic components. One of the common substituents are methoxy groups. Typical chemical shift due to aromatic methoxy methyl protons is around δ=3.8 ppm, depending of the substituents.

Additional peaks δ=6.1 ppm, δ=6.7 ppm and between δ=8.2-10.6 ppm in the TAI-SM spectrum is assigned to byproducts of TAI reacting with the air humidity and also due to aging processes.

To ensure that the allocation of signal shifts are correctly assigned to the kind of protons so far indicated in the literature, two-dimensional NMR’s were taken and analyzed before a statistical calculation of the chain parameters was undertaken, as discussed in the following paragraphs.

Total Correlated Spectroscopy (1H-1H TOCSY)

A Tocsy spectrum is a two dimensional spectrum showing bond correlations by spin-spin coupling (SSP). This measurement should help to divide the superimposed signals. A TOCSY-Spectrum exemplifies the typical TAI-SM-spectrum, as illustrated in Figure 3.2.3-3. The correlation signals are stressed with doted lines, which are labeled alphabetically. The same labeling was used also in Table 2.2.1-1. The red dotted lines mark cross-peaks which are not listed Table 3.2.3-1 because this signal peaks were not distinguishable from the baseline noise. The volume of these protons was assumed to be so small, that they were ignored.

Our observation and conclusion for each peak shift of Figure 3.2.3-3 are as follows:

Protons of methyl group (a) interact only with methylene group (b-d). The methylene protons (b) couple foreseeable with nearly all signals (a-h) and j-m. Since a cross-sections to (i) was not observed (i) was expected to belong to methyl group protons of methyl esters. Methylene protons (b) also do not interact with (n-t) (TAI by-products), this is positive assessed since there are no superimposed proton shifts belonging to SM.

Methylene protons (c) exhibit sections with the methylene groups (b) and (f) refer to α methylene of carbonyl function. So it was suggested that (c) belongs to β methylene protons of the methyl ester carbonyl groups.

Methylene protons (d) generate cross-section with (k) and (l), therefore it was assumed that this signal belong to β methylene protons of urethane.

Methylene protons (e) of protons in β position of allylic bonds couple with the connected methylene protons (b-d) and methine protons (m) in α position of double bond. Albert-Ludwigs-Universität Freiburg 65

Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

Methylne protons (f) show a cross sections to (c and b) methylene in β, γ position of methyl ester and its methane group (i). The interaction with (i) methyl protons of methyl ester has to be a five-bond spin- spin correlation, therefore (f) protons α methylene of carbonyl groups and (g) are α protons of TAI derivatized carboxylic acids couple with d and b.

a

b c, d e f

g

h, i j k

l m

n-t

m l k j i h g f e d c b a

Figure 3.2.3-3: TOCSY spectrum of TAI-SM sample in CDCl3, measured in deuterated chloroform. The by Proton NMR applied letters for labeling the signals see Figure 3.2.3-1 or Table 3.2.3-1 are used to describe the two dimension spectrum.

α Methylene protons of primary alcohol derivative of TAI (k) and α methine protons (l) of secondary alcohol display cross sections to (d) and (b) methylene.

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

Table 3.2.3-1: Observed peak shifts using proton NMR and TCSY with the observed kind of splitting and correlations signals by spin-spin coupling (SSC). The monitored shifts are given in the range which was used for the integration and the TAI-SM spectrum signals are labelled alphabetically. Consequentially the most probable kind of protons are described and marked in red in the formula.

Peak shift δ [ppm] SSC Kind of protons Formula

1 SM TAI-SM H TOCSY

a 0.56-0.93 0.56-0.93 b-d Aliphatic methyl group protons -CH2CH3

b- 0.93-1.80 0.93-1.80 a-h, j-m Aliphatic methylene group -CH2- d protons e 1.80-2.11 1.80-2.11 m β allylic methylene group - protons CH2CH=CHCH2-

f 2.11-2.38 2.11-2.38 t b, c, i α methylene protons of carbonyl -CH2CO- groups 2.86-2.90 s Epoxide α protons [21, 39] -HC(O)CH-

g 2.76-2.90 t b, d α protons of carboxylic acid -CH2CO2- groups as (Trichloroacetyl) CONHCOCCl3 carbamate

h 3.20-3.50 3.20-3.50 b Methoxy group protons >CH-OCH3

3.40-3.44 s Rest methanol methyl groups CH3OH protons Or aromatic methoxy group 3.41-3.66 Alpha protons of sec. hydroxyl >CHOH groups

i 3.56-3.66 3.54-3.63 s f methyl group protons of methyl -COOCH3 esters

j 3.82 s b, d, l methyl group protons of CH3O-Ph aromatic methoxy derivative

3.63-4.00 3.66-4.00 m methyl group protons of CH3O-Ph aromatic methoxy derivative

k 4.12-4.32 t b, d α protons of prim. hydrogen -CH2O- groups as (Trichloroacetyl) CONHCOCCl3 carbamate l 4.97-5.13 q b-d α protons of sec. hydrogen >CHO- groups as (Trichloroacetyl) CONHCOCCl3 carbamate m 5.20-5.37 5.20-5.37 t b, e α allylic methine group protons -CH=CH- n 5.50-11.0 Protons of TAI byproducts

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

13 1 13 C-, H- C HSQC and HMBC NMR of SM Spectra

Interpretation of carbon (13C) NMR and Heteronuclear Single Quantum and Multiple Bond Correlation (HSQC and HMBC) is necessary to confirm the findings and to classify functional groups of the proton NMR.

The allocation of the functional groups shown in Table 3.2.3-1 can be confirmed by the two dimensional

NMR HSQC and HMBC. Therefore a typical carbon NMR of TAI-SM, shown in Figure 6.1-4, was measured to observe the carbon shift signals. The monitored signals are numbered consecutively and shown in Table 3.2.3-1.

In the heteronuclear, two dimensional NMR some of the carbon signals show chemical correlations – 1J in HSQC and 3J, 4J in HMBC – with protons which are labeled alphabetically, see Table 3.2.3-1. The detected correlation peaks for each attached proton to a carbon (HSQC) being considered are shown in Figure 6.1-5 with positive (blue) and negative (green) magnetization. The magnetization is dependent on the number of attached aliphatic carbons. If those carbons have an odd number (one or three) of aliphatic carbons, the magnetization sign is negative, whereas an even number of aliphatic carbons show a positive magnetization. In Table 3.2.3-2 the negative magnetization direction is marked with a minus symbol in front of the letter labeling the corresponding protons.

The 1H-13C HMBC spectrum is shown in Figure 6.1-6 and Figure 6.1-7. The monitored cross sections are listed in Table 3.2.3-2 while to each carbon peak set the 3J and 4J correlating protons (letter marked) out.

For some carbons a cross section was not observed in HSQC or in HMBC. For the identification of these carbons only the chemical shifts can be used and therefore peak no. 3 and 10 are still unknown. For the other signals the identification was possible due to collective information of all the different NMR methods taken together. The assumed allocations are also listed in Table 3.2.3-2 and the relevant carbon is marked red.

All types of NMR indicate: allylic methylene group (δ=129 ppm); TAI derivatized hydrogen groups (δ=76 ppm and δ=67 ppm); β methylene of carbonyl groups (δ=33 ppm); aliphatic methylene (δ=22- 33 ppm) and aliphatic methyl groups (δ=13 ppm). This is comparable with the carbon NMR published by Bose and Srinivasan [223].

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

The colour of the cross peaks in the HSQC indicates the phase and thus the multiplicity. The blue colored cross peaks show CH2 correlations and the green CH and CH3 correlations in aliphatic chains. This also gives an indication of the accuracy of the allocations of the peaks.

In the HMQC it is also possible to see carbons without any protons, such as the carbonyl carbon of the methylester (δ=173 ppm), which has cross peaks with the protons of methylester and the carbonyl carbons of TAI derivative (δ=162-150 ppm).

Carbons belonging to tertiary aliphatic alcohols δ=84,59 ppm [223] were not observed.

Table 3.2.3-2: Generated NMR data observed peaks from 13C-NMR spectra and correlation peaks detected by HSQC and HMBC. Proton signals are labeled as in Paragraph “Proton NMR” or in Table 3.2.3-1. Carbon signals observed in 13C-NMR are labelled numerically. The non-numbered carbon or not labeled proton signals were not visible in HSQC or HMBC, but were observed in the carbon or proton NMR (Figure 6.1-4; Figure 3.2.3-1; Figure 3.2.3-2).

Peak Interaction Interaction in δ [ppm] Assumed functional group no. in HSQC HMBC Carbonyl carbon of methyl ester group or 1 173.13 TAI derivative carboxylic acid c, f, g, i

CH2COOCH3 and CH2COO-CONHCOCCl3 Carbonyl carbon of the TAI urethane linkage 2 162.39 (j) -CONHCOCCl3 3 159.29 4 157.75 Carbonyl carbons of TAI by-products 8.57 5 156.81 Carbonyl carbons of TAI by-products 8.57; 8.44 6 156.62 Carbonyl carbons of TAI by-products 8.57 Carbonyl carbon of the TAI urethane linkage 7 148.61 with primer or seconder hydroxyl groups j, k

-CONHCOCCl3

8 144.99 -CONHCOCCl3 9 129.27 α allylic methylene group carbon -CH=CH- -m 10 123.86

11 91.35 -CONHCOCCl3

12 90.16 -CONHCOCCl3 TAI-derivatized secondary alcohol [223] 13 76.26 -l 7.57; 6.87; >CHO-CONHCOCCl3 TAI-derivatized primer alcohol [223] 14 66.60 k d, (e) -CH2O-CONHCOCCl3 Methyl of aromatic methoxy groups or 55.21 methylene group of different substituted -(3.63-4.00) methyl phenylacetate

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

Peak Interaction Interaction in δ [ppm] Assumed functional group no. in HSQC HMBC Methyl of aromatic methoxy groups or TAI- 15 52.96 -j derivatized methanol Methyl of aromatic methoxy groups or -(3.63- 51.82 methylene group of different substituted 4.00) methyl phenylacetate Methyl carbons of a methyl ester group 16 50.43 -i h, j -CH2CO2CH3 α-Metylene carbons of carboxylic acid groups 46.09 derivatized with TAI -CH2CO2- g CONHCOCCl3 α-Metylene carbons of carbonyl groups i, 2.44, 20.01, 17 33.08 f -CH2CH2CO- c,

18 32.95 -CH2CH2CO- f Methylene carbon of aliphatic chain 19 30.94 b a -CH2CH2CH2-

20 29.54 -CH2CH2CH2- 1.582

21 28.71 -CH2CH2CH2- b a, b, e, f

22 28.47 -CH2CH2CH2- b a, b, f

23 28.37 -CH2CH2CH2- b a, b, f

24 28.27 -CH2CH2CH2- b a, b, f

25 28.17 -CH2CH2CH2- b a, b, f

26 28.03 -CH2CH2CH2- b k, b, f

27 27.90 -CH2CH2CH2- b k, b, f

28 27.41 -CH2CH2CH2- d k, f

29 26.17 -CH2CH2CH2- e,-b k, m, f

30 23.97 -CH2CH2CH2- -b, c k, d, f

31 23.80 -CH2CH2CH2- b, c k, d, f, g Methylene carbon of aliphatic chain - 32 21.70 b a CH2CH2CH3

33 13.10 Methyl carbon of aliphatic chain -CH2CH3 -a

Calculation of Chain Length and Number of Functionality per Chain

Form the literature it is known that the aliphatic chains of SM are highly functionalized by carboxylic acids, hydroxyl and epoxy groups. A simple quantification of these groups is still challenging. NMR is one of the simplest methods in modern labs. However, to use this analytical technique quantitatively, two points must be observed: the sample used must be completely soluble in deuterated solvent and the

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3 decisive peaks must not overlap. The solubility issue was satisfied by using deuterated chloroform.

However, use of proton NMR for the received SM directly did not result in clearly quantifiable or sharply separated peaks. Overlapping of the alpha protons of primary and secondary hydrogen groups was observed, as was overlapping with the methyl groups of the methyl ester around δ=3.6 ppm. A derivatization of the hydroxyl groups with TAI overcame this problem, while the peaks due to the α- protons of the new urethane bond (before α-protons of the hydroxyl bond) shifted downfield, see section: Proton NMR pp.62.

For the computation of the chemical functionality of SM, only the proton spectra with derivatized.

Depolymerized Suberin (TAI-SM) was used, see the blue coloured spectrum in Figure 3.2.3-1, since this will ensure that there are no overlapping peak areas of methyl esters, primary and secondary alcohols. The integration area limits used to calculate the chain building blocks are listed in Table 3.2.3-1. The following assumptions (from GC-MS and other publications) were made:

 All aliphatic chains are linear and without any sidechains, as shownin Figure 6.1-1 [50, 69, 83, 90, 92, 96, 105, 108, 114, 144]. Therefore each chain had two terminal or end groups (EG).

 Possible chain end (terminal) groups (EG) are: aliphatic methyls (-CH2CH3), methyl esters

[50, 69, 83, 90, 92, 96, 105, 108, 114, 144] (-COOCH3) and primary alcohols as shown in Figure 6.1-1 (-OH) . Therefore, each chain had two terminal groups (EG).  Within a chain (chain groups CG) simple methylene groups (-CH2-) are expected. Also, monounsaturated compounds (-HC=CH-) and their oxidation products such as epoxides and diols (secondary alcohol) might be present, as shown in Figure 6.1-1.  The methyl ester carbons do not have a proton signal, so the number of the methyl ester was used twice as an EG and as a CG as well.  The relative number of the function 푁 for example unsaturated function was calculated with α allylic methylene group. The number of protons in the methane group is 푝 = 1. But two methylene groups require a double bond, that is why 2푝 is used in this case. The accounting

frequency region of this functional groups (〈푋퐶〉) is δ=5.20-5.37 ppm.  To find out the relative number of the functional groups (푁), all relative areas were divided by the number of protons (푝) in this group.

∫ 푥(훿) 푁 = 푋 푝 Eq. 4

For the calculation of the average chain length (〈퐶퐿〉) the effective number of the end groups (푁퐸퐺) and the chain groups (푁퐶퐺) are summarized and used as follows.

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3

2 ∙ ∑ 푁퐶퐺 ∑ 푁퐶퐺 〈퐶퐿〉 = + 2 = + 2 Eq. 5 ∑ 푁퐸퐺 푁퐶

Using the number of chains (푁퐶) to account frequency of functional groups (〈푋퐶〉) in a chain.

푁푋 2 ∙ 푁푋 〈푋퐶〉 = = Eq. 6 푁퐶 ∑ 푁퐸퐺

The chain lengths and probability of a functional group in a chain were calculated for nine different batches of bark samples, different extractions runs and different methanolysis, as indicated in Table 3.2.3-3. The last row shows the results from the combined samples.

Table 3.2.3-3: Calculated average chain length and percentage portion of functional groups in chains for the TAI derivatized Suberin monomers.

prim. Sample 〈푪 〉 average sec. alcohol unsaturated methyl methyl 푳 alcohol name chain length [%] bonds [%] ester [%] group[%] [%]

141216I 15.8 26.6 28.8 7.3 84.1 89.4 141216I-Cr 15.1 31.2 33.3 9.5 83.0 85.8 20140218I 16.7 30.9 35.3 10.5 85.4 83.6 141216II 15.2 31.6 30.7 6.6 83.6 84.7 140218II 16.3 20.0 27.2 8.7 88.2 91.8 150413II 14.6 8.4 20.1 10.9 91.7 99.9 150413I 16.2 13.7 43.1 15.0 88.8 97.5 150413II 14.8 11.5 26.3 14.1 80.5 108.0 150413III 16.2 15.4 28.2 15.4 89.8 94.8

Average 15.7 21.0 30.3 10.9 86.1 92.8 SD 0.8 9.2 6.5 3.3 3.7 8.1

Mix 15.3 22.0 32.3 11.9 85.0 93.0 This table shows that the chains consist of 16 carbons on average. Almost all (92.8%) of the chains had one methyl group and one methyl ester group (86.1%), and approximately every fifth had a hydroxyl group. Every third contained diols in the chain and just 11% an unsaturated group. Furthermore a high standard deviation was determined for unsaturated bonds, and primary and secondary alcohols. The observation of epoxides such as 2-Oxazolidones [221, 222] was not observed. This could also influence the calculated number of the chain conformation. Considering that bark is a natural product, the level of the

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Nuclear Magnetic Resonance (NMR) Study of Suberin Monomers: With and Without Using Trichloroacetyl Isocyanate (TAI) 3.2.3 standard deviation is acceptable. Oxidation processes initiated by physical, chemical or biological treatment during storage or sample preparation are also possible.

An important source of errors is the adaption of the spectra. How to set the frequency correction or base line and where the integrations limits are. This is often depends on the individual and leaves space for interpretation. In this work spectra were processed as uniformly as possible.

Diffusion-Ordered NMR Spectroscopy (DOSY)

The diffusion NMR plots of SM (red) and TAI-SM (blue) are illustrated in Figure 3.2.3-4. A DOSY spectrum is typically presented as a two dimensional spectrum. The x-axis represents the chemical shift δ in ppm, and the the self-diffusion constant (D) in m²/s is plotted on the y-axis. According to the Stokes- Einstein equation the molecular diffusion coefficient is inversely proportional to the molecular size. Consequently, molecules with a high diffusion coefficient are smaller than those with a lower diffusion coefficient.

At first, the self-diffusion coefficient (D) of the suberin monomer mixture is quite constant for SM at

-9.15 -9.5 around 10 m²/s and for the TAI-SM around 10 m²/s. Some signals are out of the characteristic D- areas marked in Figure 3.2.3-4 with the red and blue bar: The SM spectrum (red) displays two signals with higher D belonging to chemical shifts δ=2.05 ppm, δ=3.42 ppm, and one with lower D pertains to chemical shifts between δ=6.4-7.1 ppm; Almost all signals of the TAI-SM spectrum (blue) are shifted to lower D with the exception of δ=2.81 ppm (2H, t, J = 7.27 Hz), 3.91 ppm, 4.00 ppm and 5.36 ppm.

It was possible to mark the typical D-area with bars, this means that the monomer chain length did not vary widely. The decrease in D after derivatization with TAI was expected, since the molecular mass increase, caused by addition of derivative. The general decreasing of D demonstrate the existence of hydroxyl groups in the molecules. The exceptions are tied to explain:

The general decrease in D demonstrates the existence of hydroxyl groups in the molecules. The exceptions are explained as follows:

The chemical shift in SM spectra δ=3.42 ppm (3H, s) is similar to the D of solvent, this indicates that the singlet refers to the methanol used for sample preparation; signals pertaining to δ=2.05 ppm could be due to ethyl acetate. The TAI-SM spectrum shows three signals with higher D, all three are non-significant peaks in the 1H-NMR-spectra; Lower diffusion coefficients δ=3.97 ppm (assuming triplet, J=6.39 Hz) and 3.64 ppm (3H, s), 3.61 ppm could be side reactions or addition of more than one TAI (diole).

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Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin 3.2.4

Thus it was possible to show that the SM monomers had a narrow distribution in size, with hydroxy

groups which could be derivatized by TAI, since the D decrease for almost all TAI-SM.

diffusion constant

-

self

lower molecular size molecular lower higher

solvent

Figure 3.2.3-4: DOSY spectra of SM (red) and TAI-SM (blue) in CDCl3 stacked atop each other. The bars mark the main diffusion area (D-area).

3.2.4 Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin

The suberin monomers (SM) were analyzed via GC-MS after derivatization of the hydroxyl groups with N-Methyl-N-(trimethylsilyl) trifluoroacetamide (MSTFA). A typical total ion count (TIC) spectrum of

MSTFA-SM is shown in Figure 3.2.4-1. The peaks assigned to substances with a high match (>750) by NIST library using fragmentation spectra and retentions index (RI) were numbered consecutively. The UPAC names of the substances found are listed in Table 6.2-9. For simplicity the substrates found are presented in their pre-derivatization form. This means for example that No. 29 was detected as methyl 16-((trimethylsilyl)oxy) hexadecanoate and in this list is labeled as 16-hydroxyhexadecanoic acid. All trimethylsilyl (TMS) ether and methyl or TMS esters are given in their hydroxyl or carboxylic acid

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Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin 3.2.4 forms. This is also the reason why some of them are listed twice, for example No. 20 and 25 are 9,12- octadecadienoic acid. The first peak was found as a methyl ester the second one as a TMS ester.

Figure 3.2.4-1-: Example of total ion chromatogram (TIC) of the GC-MS measurement of MSTFA-SM

Over a hundred small peaks were found in the TIC spectrum with only 42 being identified unambiguously according to the NIST library. A typical SM TIC displayed in Figure 3.2.4-1, dotted lines mark the 42 identified signals. Six different samples were analyzed and their spectra evaluated. The results are given in condensed form in Table 6.2-9. In the chemical components fond are summarized in substance classes as percentage of TIC. The six substance types are presented in Figure 6.1-1.

Before discussing the values it is to be noted that peak areas are only a first approximation, Thus only an indication of relative quantities is made, and not the absolute quantity of the material is made. Furthermore, only a fraction (27-74%) of the whole sample is displayed in the GC-MS spectrum [32, 105, 114].

In Figure 3.2.4-1 peak 37 is by far the biggest signal in the whole observed spectrum, with an area often of greater than 50% of the total TIC area. This is assigned to 9, 10, 18-hydroxy octadecanoic acid methyl ester (see Figure 6.1-1 molecule 5.3). This is a highly interesting substance, as it potentially has four functional groups for producing crosslink polymers. The other peaks are discussed in terms of utilization of substance classes, see Table 3.2.4-1. All discussion in terms of quantity is based on the TIC integrations area.

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Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin 3.2.4

The main components found were ω-hydroxy acids, with their proportion being over half of the substances determined (this is due to inclusion of 9, 10, 18-hydroxy octadecanoic acid methyl ester). The chain length was observed to be between 16 and 22 carbons, with unsaturated bonds, epoxy and vicinal (vic)-dihydroxy groups. Aliphatic acids were about 20% of the identified substances with chain length C16-C26 in partial polyunsaturated form. Dicarboxylic acids with around 10% were found in varying sizes ranging from C8 to C18, several with vic-diols. Up to 5% of aromatics and of long chain C16-C26 aliphatic alcohols with partly vic-diols in the chain were analyzed in the SM samples. Dihydroxy alkane C22 was found in trace amounts.

Table 3.2.4-1: Area proportion in percentage of the different substance classes for the six MSTFA-SM measured samples, illustrated in Figure 3.2.4-3

ω- α, ω- α, ω- Sample aliphatic aliphatic hydroxy dicarboxylic dihydroxy aromatics other Name acids alcohols acids acids alkanes I140218 3.33 62.42 2.65 3.37 0.12 0.84 0.55 I141216 5.65 58.40 2.30 3.26 0.20 1.07 0.72 II140218 6.59 44.82 1.90 3.96 0.05 3.13 0.89 II141216 4.47 61.99 2.11 1.67 0.08 1.07 0.33 III141216 6.87 45.54 3.26 5.21 0.05 2.69 1.35 IV141216 9.97 32.67 3.82 4.36 0.08 1.65 0.91

Between 54% and 74% of the TIC peak areas could be referred to substances in the NIST library, see Figure 3.2.4-3. These values are dependent on systematic and random errors. Especially using different blanks stored in plastic vails, shows partially catastrophic influences of contaminating substances, that some samples could not be used for this work. Signals found in the blanks were not used in the analysis. However, in the best cases contamination signals were modified the TIC total areas so that also the getting results are influenced

A TIC spectrum is not only influenced by the GC response factor, it is often more influenced by the fragmentation of a molecule. A substance which fragments into more ions shows a higher TIC signal than one that fragments into fewer ions. Another important point is the overlapping of signals. Sometimes signals are not visible in TIC spectrum, but are observable in specific fragment spectra. This can be seen for example in the fragment spectra m/z=317 (red), m/z=427 (yellow) and the TIC spectrum in Figure 3.2.4-2. Here both fragment peaks at 40.528 min clearly show a shift in relation to each other. Although the TIC peak does not show any evidence, an overlapping of two components is indicated by the fragment peaks.

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Figure 3.2.4-3: Bar charts of analyzed six MSTFA-SM samples bear on the various substance classes refer to the monitored TIC peak area (left side) and refer to identify components (right). Additional a bar of Fagus sylvatica L. roots SM composition is showed (right) found by Spielvogel et al. [77].

Absolute quantification of SM components is only possible when standards for each substance are used. For some of these a calibration curve was produced, while some were calculated using the assumption that if the same substance class is used plus-minus a few carbons that it does not significantly change the slope of the calibration, especially if characteristic mass-to-charge peaks are used. This idea was tested with the FAME standard mix using m/z ratio 74 which is characteristic for the carboxylic acid methyl

+• ester fragment (C3H6O2 ). The result is shown in Figure 3.2.4-4. Using this suggestion we tried calculated the absolute volumes, using a few standards, see Table 3.2.4-2. However, for most of the standards, especially No. 37 (9, 10, 18-hydroxy octadecanoic acid) it was impossible to obtain sufficiently pure substances. This prevented further investigation in this direction.

Figure 3.2.4-2: TIC spectrum versus fragments spectrum, a screen shot of AMDIS Spielvogel et al. [123] analyzed suberin monomers of Fagus sylvatica L. roots by GC-MS after extraction and methanolysis. The absolute quantitative results in µg/g was based on comparison between the TIC peak area of an external standard and the component, and adjusted by a GC response factor. For better comparability the absolute amounts were used to calculate the percentage of the components. The outcome of this work is shown in Figure 3.2.4-3 as the bar on the far right labeled “Ref”. Aromatic or other components were not considered. While these volumes are not comparable, they do show that

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Gas Chromatography-Mass Spectrometry (GC-MS) Measurement of Suberin 3.2.4 despite different sources (root or trunk) and depolymerization methods (hydrolysis or methanolysis) similar proportions of substance groups were found.

Table 3.2.4-2: Determinate calibration curve slope of specific m/z standards

component m/z Slope [ml/mg] aliphatic alcohols TMS icosan-1-ol 75 45245 hexadecane-1-ol 75 46944 α, ω-dihydroxy alkanes TMS dodecane-1, 12-diol 103 53692 aliphatic acids methyl ester hexadecanoic acid 74

9-octadecenoic acid (Z) 74 aliphatic acids TMS hexadecanoic acid 9, 12-octadecanoic acid (Z, Z) Octadecanoic acid Other ß-sitosterol TMS 129 4500 357 3242

A high proportion (30%) of aromatics were found in suberin by B. Perra, J.-P. Haluk and M. Metche [108, 128]. They determined 70% as aliphatics and analyzed the fatty acids with GC-ionizing flame detector, finding carboxylic acids between C12-C24 with partial (poly)unsaturated chains. This fits well to the results in this work.

The finding of mid-chain mono hydroxyl groups, as discussed in the publication of Spielvogel et al. [123] is conspicuous. Except for hydroxyl groups as vic-diols such functional groups were not detected in this work. This can be explained by formation of hydroxyl groups by the addition reaction of double bonds and water during alkaline hydrolysis. The method Spielvogel et al. [123] used to calculated the absolute values of the components may also have led to errors.

Spielvogel et al. [123] made a quantitative analysis based on TIC using standards. They maintain to quantify like Mercedes-Millan et al. [224]. But Mercedes-Millan et al. used an FID detector for the quantification of the monomers without any explanation to the calculation method. It is possible that they use response factors for the flame ionization detection published from different authors [225].

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Elemental Analysis Measurement of Suberin Monomers 3.2.5

However, this kind of calculation is usable with a thermal ionization system. That is why the absolute results of this paper are questionable, see Table 2.3.2-1.

Figure 3.2.4-4: Calibration diagram for different carboxylic acid methyl esters using characteristic m/z ratio 74.

3.2.5 Elemental Analysis Measurement of Suberin Monomers

Elemental analysis (EA) provides weight percent of carbon, hydrogen, nitrogen and sulfur in a sample, as shown for example in Table 3.2.5-1. The measurements were repeated with two different samples. Since no conspicuous variation was observed, a third measurement was not required. The value of oxygen was calculated from the remaining mass, which was assumed to be due to oxygen. However, this is known to overestimate the amount of oxygen since other elements could be in the sample. The average was charged with the individual molecular mass of the elements.

Table 3.2.5-1: Elemental analysis of SM. Carbon, hydrogen, nitrogen and sulfur were determined experimentally, with oxygen being estimated. Values are in percent by weight [wt.%].

C H N S O [wt.%] 62.81 8.62 0.13 1.21 27.23 [wt.%] 62.61 8.69 0.10 0.98 27.62 Average 5.23 8.66 0.01 0.03 1.71 [mol/100g]

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Elemental Analysis Measurement of Suberin Monomers 3.2.5

It can be seen from Table 3.2.5-1 that traces of nitrogen and sulfur were also found. While both may be naturally found in the samples tested, it is possible that the sulfur fraction could be increased due to the use of H2SO4 for neutralization of SM during purification, see Figure 5.2.3-1 step 3.1. If sulfate did contaminate the SM the amount of oxygen would also be increased. For each 0.03 mol sulfate in the SM sample, 0.12 mol oxygen would also be found, thereby increasing the already overestimated oxygen proportion. This would decrease the amount of oxygen up to 1.59 mol/100g in SM.

The resulting proportion per one carbon: 1C – 1.7H – 0.3O

Since 9, 10, 18-hydroxy octadecanoic acid methyl ester was the highest peak observed by GC-MS and also the other chain classes shows chain length around 18, the C18 chain model was used for comparing the results in Table 3.2.5-2.

It is possible to find the 18 carbon chain molecular formula for some chain classes partial with the mid chain functionalities: double bonds, epoxides and vic diols. Because of the esterification of carboxylic acids the sum of carbons increases to 19 or 20.

Using the calculated proportion from the elemental analysis following molecular formulae were received:

C16H27O5

C17H29O5

C18H31O5

C19H32O6

C20H34O6

Comparing the formulae in Table 3.2.5-2 the hydrogen ratio was lower, and the oxygen ratio was higher than expected from previous results of GC-MS. One possible explanation for the low amount of hydrogen is the presence of aromatic components, which decrease the average proportion of hydrogen.

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Elemental Analysis Measurement of Suberin Monomers 3.2.5

Table 3.2.5-2: Theoretical molecular formula for 18 carbon long chain molecules, which are found in a high amount in GC-MS measurements.

average area [%] molecular formulae for C18- chain class (GC-MS) chain

ω-hydroxyalkanoic acids 7.60 C19H38O3

α.ω-Dicarboxylic acids 1.97 C20H38O4

Alcohols 2.08 C18H38O

α.ω-Alkandiole 1.27 C18H38O2

9.10-Dihydroxy-α.ω-dicarboxylic acids 1.45 C20H38O6

9.10.18-Trihydroxycarboxylic acids 52.44 C19H38O5

9.10-Epoxy-ω-Hydroxycarbonsäre 4.14 C19H36O4

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Conclusion of PART I

Conclusion of PART I

In this first part, it was possible to show by solid-state 13C-NMR, that the depolymerization via methanolysis works well. That a statistical evaluation of suberin monomers (SM) by NMR is possible it the sampel SM is derivatized with TAI and if using assumptions based on literature and own GC-MS results. The calculaded 16 carbon long chains with 1.4 functional groups per chain (without counting the double bonds) of suberin are not usible for polymerization. Additionally, the small yield (lower than 4 wt%) and the contrast NMR to GC-MS data prospects that SM is only usably as filler/ copolymer in another system.

The yield of phenolic compounds, especially tannins, in the solvent extracts was low and to be useful needs further investigation into preparation and purification. Thus it was decided to use another aromatic building block as polymer precursor. It was decided to use the beech wood organosolv lignin, see second part of this thesis.

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PART II

PART II

Diverse methods are already known for the preparation of NIPUs from the more simple petrochemical systems such as ethylene carbonate[226], bisperoxide[227] glycerol[228, 229] or oil based[25, 27, 230] than foe a natural product such as lignin. In this study, the preferred polymerization concept is the ring opening reaction of cyclic carbonates with amines. Two possible strategies could be used to produce lignin precursors: first the conversion of hydroxyl groups to amines and the second approach is to cyclic carbonates of lignin. We chose the second route in which the intermediate of epoxide lignin (EL) was prepared at first for the synthesis of carbonate lignin (CL The reactions products were monitored analytically and tested to find out if the known process is applicable for lignin, see Section 3.3. Various problems on the solubility of OL, EL and especially CL, the purification of intermediate and product, reaction conditions and also analytical determination were solved during the process.

For the NIPU’s synthesis, three commercial terminal diamines were used and the reaction conditions and mixing ratios were tried (Section 3.4) for producing new bio-based NIPUs. FTIR was used to demonstrate the success of the reaction, and the thermal properties of the polymers were analyzed by TGA and DSC. Depending on the amount and the length of the diamines, an endcapping or cross linking of the CL was observed.

Both processes are schematiclly presented in Figure PART II-1.

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PART II

Figure PART II-1: Schematic representation of the processes done in Part II. The derivatization steps (marked red) are investigated in Section 3.3 and the polymer synthesis is examined in Section 3.4 (marked yellow).

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Analysis of Organosolv Lignin and its Derivative 3.3

3.3 Analysis of Organosolv Lignin and its Derivative

This Section deals with the synthetic process analysis of organosolv lignin derivatives (see Experimental Section 5.3), their analysis, and yield quantification. The results of the educts: organosolv lignin (OL); the intermediate lignin epoxide (EL); and the product, lignin carbonate (CL) are included in this section. FTIR spectra showed that the desired reactions successfully take place. NMR gave additional information about the amount of hydroxyl in OL (1.45 mmol/g aliphatic and 2.61 mmol/g aromatic hydroxyl groups) and the epoxy index (EI) of EL samples (0.32-1.48). According to the NMR analysis 8-36% of the hydroxyl groups were converted to epoxides.

Reactions with model compounds showed that under anhydrous conditions (OL dissolved in ECH) used for lignin:

- The lignin model (LM) compound shows an epoxidation yield of between 40-45% (ELM-1, ELM- 3, ELM-4). This was observed by proton NMR. - The epoxidation of Suberin Model (SM) compound representing aliphatic hydroxyl groups was not observed by NMR or FTIR in ECH (reaction EMS-1a in Section 5.4). However, epoxidation was observed in around 35% of the hydroxyl groups after the addition of NaOH solution (EMS- 1b) and longer reaction times did not change the yield significantly ESM-1.

An aqueous reaction system used by Malutan, Nicu and Popa[181] converted nearly 100% of the LM to epoxide (ELM-2), as observed by proton NMR. But the OL in this medium led to an organic solvents insoluble product (EL-5). Therefore, an anhydrous reaction system was used for further synthesis.

The performance by Karuana et al.[231] was quite successful. ESM-2 proton NMR shows a 100% conversation of aliphatic hydroxyl groups, without losing much material.

The addition of carbon dioxide with and without using high pressure conditions was applied to synthesize CL. The absolute quantification of cyclic carbonates per gram lignin (CI) was not possible. However, the curing with diamines (Section 3.4) gave an idea of the cyclic carbonate concentration (≤0.5 mmol/g) in CL. In addition, Hansen solubility parameters were determined to observe change in intramolecular interactions.

This section is structured in subsections based on the specific analytical methods used. A short summary at the end of this section bring the results together and demonstrates a possible approach to produce a novel lignin based precursor to produce NIPUs, see Section 3.4.

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FTIR Measurements of OL, EL and CL 3.3.1

3.3.1 FTIR Measurements of OL, EL and CL

Organosolv lignin (OL) was derivatized with epichlorohydrin (ECH). The resulting lignin epoxide (EL) was used to synthetize lignin carbonate (CL) adding carbon dioxide, see Section 5.3. FTIR was used to monitor the reactions. The ATR-FTIR spectra are shown in Figure 3.3.1-1. The displayed transmission band are summarized in Table 6.2-4 and assigned the assumed bond vibration.

The OL spectrum (black) in Figure 3.3.1-1 shows the typical bands for lignin. If in particular, the typical syringyl (S) bands 푣̃=1326 cm-1(C-O stretch); 1113 cm-1(aromatic C-H deformation); 830 cm-1(C-H deformation) and guaiacyl (G) band 푣̃=1266 cm-1(C-O stretch) are considered, a S-G unit relationship could be calculated as S/G= 1.06.[169] However, beech wood OL has a slightly higher amount of S-units. However, compared to other hardwood lignins (S/G=2.60[169]) OL has relatively low S-units proportion. This fits well with the EA results of Glasser[160], where he found a methoxy content of 1.39-1.46 per monolignol in beech OL. Further lignin specific bands observed in FTIR spectra are listed in Table 6.2-4.

The red EL spectrum in Figure 3.3.1-1 displays new bands after the glycidylation with ECH in the fingerprint area: 푣̃=1254 cm-1; 960 cm-1; 925 cm-1; 880 cm-1; 851 cm-1; 757 cm-1; 732 cm-1; and 695 cm-1. The bands 푣̃=1254 cm-1, 851 cm-1, and 757 cm-1 were identified from the literature [12, 163, 181, 231] as typical oxirane ring (epoxide) vibration. The band at 푣̃=910 cm-1 is also often assigned to an epoxy ring vibration in the literature[163, 180–184], but this measurement is not consistend to be an evidence for epoxidation of EL. The other bands were already observed in OL and did not show any significant changes in the EL spectrum. The band at 푣̃=851 cm-1 should be an alkyl ether band vibration according to Kühnel et al.[37]. Two unknown bands 푣̃=732 cm-1; and 695 cm-1 appeared could not assigned from the literature of epoxied Lignins. However, the bands at 푣̃=1254° cm-1; 732 cm-1 and 695 cm-1 could also be assigned to aliphatic chlorides vibration[232]. This could indicate that ECH is still part of the sample, and that the reaction follows the SN2 (II) mechanism (see Figure 5.3.1-1) resulting in an open ring product (3). So the EL spectrum confirmed the successful O-glycidylation of OL with ECH.

The Figure 3.3.1-1 shows two CL spectra and a pure DMSO spectrum. The cyan dashed spectrum is the CL product solution in DMSO and the blue curve is the spectrum of CL which was collected after drying for one week at 80 C in vacuum (< 50mbar). Due to the difficulty in complete removal of DMSO from the product both spectra are given to recognize possible influences of DMSO in the lignin carbonate reaction. Although precipitation of the CL in various solvents (acetone, methanol, xylene, toluene and THF) was possible, the precipitated CL was no longer completely soluble in these solvents, (see Hansen solubility parameter in Subsection 0 FTIR spectrum of precipitated CL with acetone as solvent in Figure 6.1-8.)

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FTIR Measurements of OL, EL and CL 3.3.1

Figure 3.3.1-1: FTIR spectra of OL (black), EL (red), ECH (pink), CL– vacuum dried (blue), CL - dissolved in DMSO (Cyan deshed), and pure DMSO (grey). For OL, EL and CL (vac.) the reference band at 1597 cm-1 was used. All important bands are marked with dotted lines and labeld on the top. New obands are labeld with the coresonding colour of the spectrum observed. Observed transmission bands are marked with dotted lines and labeled Newly identified bands are indicated by the color of the label at the top: black in the OL spectrum; red in the EL spectrum; and blue in CL (vac.) spectrum.

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FTIR Measurements of OL, EL and CL 3.3.1

Significantly different bands were found in the spectrum in acetone: 푣̃=3267 cm-1; 2961 cm-1; 2937 cm-1; 2840 cm-1; 1707 cm-1; 1327 cm-1; and 925 cm-1. However, those bands did not give enough information about the insolubility of the precipitated CL products. It was presumed that they could be due to the formation of crosslinked lignin during the precipitation process. If the polymers are to be dissolved again, the small molecules which may have allowed solubility are missing. Another reason could be that the hydroscopic CL becomes deposited in water which may be preventing the disintegration in solvents. However, the typical EL bands (indicated in red) disappeared from the spectrum of CL-vacuum dried (blue) after the addition of carbon dioxide. This suggests that the band is more likely to be assigned to the epoxide ring in EL. On the other hand the bands at 푣̃=757 cm-1 and 960 cm-1 are the only two peaks which did not disappear completely, so it is suggested that these bands pertain to the alkyl ether bonds.

The spectrum of the CL-vacuum dried sample (blue) shows some new bands compared to the EL spectrum (red): 3267 cm-1; and 1793 cm-1. The band at 푣̃=1793 cm-1 are identified in the literature at 푣̃=1780-

Figure 3.3.1-2: FTIR spectra of CL-1, CL-2 and a mix of CL-2 and CL-3 (high pressure) and CL-4 (normal pressure).

1805 cm-1 as C=O vibration of cyclic carbonate.[12, 24, 25, 27, 209, 228–230, 233] The other new bands at 푣̃=3267 cm- 1, 2255 cm-1, 2127 cm-1 and 1000 cm-1 could also belong to vibrations of the catalyst TBAB. Nevertheless, the FTIR results show that both reactions were successful.

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NMR Measurements of OL, EL and CL 3.3.2

The addition of carbon dioxide for the preparation of CL 1-3 was done in a high pressure reactor, but also done under normal pressure conditions for CL 4. FTIR spectra of both products, which are normalized to 푣̃=1592 cm-1 are shown in Figure 3.3.1-2. The biggest differences between these spectra are the intensity difference of cyclic carbonate peak at 1792 cm-1. It can be concluded that the reaction under 20 bar pressure works better than that under normal pressure with a higher yield.

3.3.2 NMR Measurements of OL, EL and CL

Organosolv lignin (OL), the lignin after the O-glycidylation (EL) and the carbon dioxide addition (CL) were analyzed by different NMR techniques. 31P-NMR[234] was used to determine the ratio of S/G units and for quantifying hydroxyl and carboxylic acid groups in OL and epoxide groups in EL. A quantification of cyclic carbonates was not possible due to overlapping signals. 1H-NMR was used to study the reaction process of O-glycidylation and addition of carbon dioxide.

Proton NMR of OL, EL and CL

Proton NMR spectra were performed for organosolv lignin (OL), lignin R O O after O-glycidylation (EL) and the lignin derivatized with cyclic H carbonates (CL). Examples of spectra are given in Figure 3.3.2-2. 4 1 Typical chemical shifts for epoxide rings are marked in red, and for H H 3 2 cyclic carbonates in blue. Observed proton shifts for OL (black), EL (red), and CL (blue), Figure 3.3.2-2 are summarized in Table 3.3.2-1. Figure 3.3.2-1: Ethylene oxide derivative of EL (R=lignin unit)

The proton NMR of OL shows the typical shifts of lignin, except for some signals due to acetone and acetic acid. Traces of these chemicals are suspected to be part of the production solvents of OL, and were removed in the next process step of O-glycidylation.

The EL proton NMR spectrum was monitored after the nucleophilic substitution of OL with ECH (Figure 5.3.1-1). This shows that the expected proton shifts for the new derivative, shown in Figure 3.3.2-1: δ=3.89 ppm (4, dd, 1H, J=11.73 Hz, J=3.73 Hz); 3.54 ppm (4, dd, 1H, J=11.76 Hz, J=6.92 Hz); 3.23 ppm (3, m, 1H); 2.84 ppm (1, dd, 1H, J=5.06 Hz, J=4.07 Hz); 2.69 ppm (2, dd, 1H, J=5.07 Hz, J=2.49 Hz).

The internal standard, 4-Nitrobenzaldehyde was used for the determination of the EI, see Eq. 10. Causes of the overlapping of the typical derivative signals are the shift signals of the methylene protons of the epoxide ring at δ=2.84 ppm and 2.69 ppm were used for EI calculation. The area of aldehyde proton of the internal standard at δ=10.21 ppm was used as reference. The different batches of EL show varying EI, see Table 5.3.2-1.

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NMR Measurements of OL, EL and CL 3.3.2

A high decrease of EI values is observed after the extra purification step. This indicates that the whole unreacted ECH had not been removed. However, it is not clear that whether the unreacted ECH is removed, or the EL molecules of high epoxide amounts are eliminated during the purification process.

Figure 3.3.2-2: 300 MHz proton NMR spectra of OL (black), EL (red) and CL (blue) in DMSO. The typical region for epoxide ring is shown in red and for cyclic carbonate in blue.

Moreover, a decrease of EI values was observed after storage of the sample at room temperature. After two and a half months, the EI of EL-3 decrease from the average value of 3.5 mmol/g to 2.3 mmol/g and for, EL-2, the value decreased to 2.2 mmol/g after three months. Therefore the products should be used immediately after the preparation.

The EL-4 showed higher EI. It is not known, whether the higher EI of EL-4 is based on the longer reaction time (24h), the addition of sodium hydroxide, or the direct measurement without an aging process.

The NMR spectra of CL-1 and CL-2 were not used for the analysis since the reactions were done in non- deuterated DMSO. The trials to remove DMSO via solvent exchange with different solvents (acetone, methanol, xylene and toluene), or via vacuum distillation resulted insoluble products.

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NMR Measurements of OL, EL and CL 3.3.2

Table 3.3.2-1: Proton NMR (300 MHz, DMSO) chemical shifts observed for OL, EL and CL.

Peak shift δ [ppm] Nature of protons Formula OL EL CL 7.7-5.7 7.7-5.7 7.7-5.7 Aromatic protons, α- and β-protons [165] m Ar-H 7.3-7.1 7.3-7.1 Toluene m Ar-H

[165] 5.0-3.4 5.0-3.4 5.0-3.4 Methoxy protons, γ-protons ; Secondary -OCH3; >CH-

hydroxyl α-protons; primary hydroxyl α-protons OH; -CH2-OH

[24, 227, 228] 4.9-4.7 Methine of ethylene carbonate CH2-CH-

(CO3)CH2

[24, 227, 228] 4.5 Methylene of ethylene carbonate CH2-CH-

(CO3)CHH

[24, 227, 228] 4.3-4.2 Methylene of ethylene carbonate CH2-CH-

(CO3)CHH 4.0-3.9 trans position methylene proton[178, 235] dd CH-CHH-O 3.7-3.5 cis position methylene proton [177, 178, 231] dd CH-CHH-O

3.4-2.9 3.4-2.9 3.4-2.9 water s H2O

[178, 180, 235] 3.3 Methine of the ethylene oxide m -CH-OCH2 2.9 trans proton of the methylene ethylene oxide [177, dd -CHO-CHH 178, 231]

3.2-3.1 TBAB methylene protons t N-CH2-CH2 2.8 cis proton of the methylene ethylene oxide [177, 178, dd -CHO-CHH 180, 231, 235]

2.5-2.4 2.5-2.4 2.5-2.4 DMSO q O=S-(CH3)2

[165] 2.1 Acetone or methyl ester s O=C-(CH3)2

1.9 Acetic acid s CH3-COOH

1.5-1.6 TBAB methylene protons m CH2-CH2-CH2

1.4-1.3 TBAB methylene protons m CH2-CH2-CH3

[228] 1.4-0.9 1.4-0.9 1.4-0.9 Aliphatic methylene protons m -CH2- 1.0-0.9 TBAB methylene protons

[228] 0.9-0.7 0.9-0.7 0.9-0.7 Methyl protons m -CH3 The batches, CL-3 and CL-4 were done in deuterated DMSO whereby CL-3 was synthesized in a pressure reactor and CL-4 at ambient pressure. However, identification of the typical ethylene carbonate peaks is challenging because the signal to noise is insufficient in most of the measurements. The identified chemical shifts for the ethylene carbonate were: δ=4.84-4.75 ppm (2, m, 1H); 4.53-4.51 ppm (1, dd, 1H);

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NMR Measurements of OL, EL and CL 3.3.2

4.32-4.24 ppm (2, dd, 1H), see Figure 3.3.2-3). The methylene protons 3 (CH2) seems to be shifted upfield, but specific identification was not possible. The coupling parameter of the ethylene carbonate ring protons was also not able to be determined.

Table 3.3.2-2: Calculated EI for all EL batches using an IS. EL-1, EL-2 and EL-3 were measured without an additional purification step. EL-2 and EL-3 were purified and measured afterwards, EL-4 directly after the reaction.

Sample EI (2.7 ppm) [mmol/g] EI (2.8 ppm) [mmol/g] Average [mmol/g]

EL-1 0.84 0.93 0.89

EL-2 1.00 1.02 1.01

EL-2-purified 0.32 0.38 0.35

EL-3 1.26 1.48 1.37

EL-3-purified 0.33 0.47 0.35

EL-4-purified 0.38 0.43 0.41

All NMR spectra of lignin show broad peaks, whereby the interpretation of the NMR spectra of CL is complicated because of its hygroscopic behavior. Due to the low signal noise relation the use of other NMR methods like carbon NMR or two dimensional NMR (2D-NMR) are not applicable. Very intense peaks of water and DMSO protons are appeared which are superimposed to proton signals of typical lignin signals derivative. Figure 3.3.2-3: Ethylene carbonate derivative of Phosphorus NMR of OL and EL CL (R=Lignin unit)

Organosolv lignin (OL) and ethylene oxide derivatized lignin (EL) were analyzed by Phosphorus NMR (31P-NMR). The 31P-NMR was performed for the determination of the monolignols ratio and for the quantification of aliphatic and aromatic hydroxyl groups and carboxylic acids in OL and EL[236], see Figure 3.3.2-4. According to the reported 31P-NMR procedure, the samples were derivatized with 2-Chloro- 4, 4, 5, 5-tetramethyl-1, 3, 2-dioxaphospholane (TMDP), see Subsection 5.6.3 p. 124.

The 31P-NMR measurement of the EL sample was analyzed twice using the same method. The representative spectrum of 31P-NMR is shown (red) in Figure 3.3.2-4. The evaluation of the spectra and integration of functional group peak areas was done using the normalized value of an internal standard (cyclohexanol). Observed areas A and the used mass of OL and EL (m) were utilized to calculate the proportion of lignols and functional groups (K) in mmol per gram of the OL and EL, see Eq. 7. The results are summarized in Table 3.3.2-3

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NMR Measurements of OL, EL and CL 3.3.2

After the glycidylation, the amounts of aromatic hydroxyl groups were decreased from 2.61 mmol/g to 0.68 mmol/g, this is approx. three-quarter of the aromatic hydroxyl groups reacted with ECH. From this measurement, it is not obvious that one of the aromatic phenols is somehow preferred for the glycidylation. The small change of p-hydroxyphenyl OH concentration could be justified by the low signal noise ratio.

0.006 ∙ 퐴 ∙ 푚 = 퐾 Eq. 7

The amount of the aliphatic hydroxyl groups seems to increase by 15% after epoxidation. This can be explained by some epoxide remaining in an open conformation, see Figure 5.3.1-1. The Epoxide may have reacted with water, or opened up by addition reaction of TMBP during derivatization, see Figure 5.6.3-2. But in any case it is not possible to determine how much of the aliphatic hydroxyl groups had reacted with ECH.

aliphatic OH phenols OH COOH

Catechol aliphatic aromatic S-OH G-OH -OH H-OH COOH COOH

Figure 3.3.2-4: 121 MHz 31P-NMR of OL-1 (black) and EL-2 (red) The values of aliphatic and aromatic hydroxyl of carboxyl groups are not completely reliable because of the low signal to noise ratio. However, it was clearly observed that the acids reacted during glycidylation. Esterification of the carboxyl groups is one expected reaction.[237]

The difference between the consumed hydroxyl (aromatic + carboxyl=1.97 mml/g) and additional aliphatic hydroxyl groups (0.22 mmol/g) was 1.75 mmol/g lignin. This could be due to the indirect calculation of epoxy index (EI) if elimination and substitution side reactions, or water addition are ignored.

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Thermal Analysis of Lignin (Derivative) 3.3.3

The precision of the method is quite good with a slight deviation even though the values are calculated from only two repeated reactions, and are therefore not statistically significant.

Table 3.3.2-3: 31P-NMR samples spectra of OL and two EL batches. The area was calculated relative to the internal standard, cyclohexanol. The EL sample was measured twice, from which the sample mean and covariance was calculated. Additionally, the increase or decrease in signals of EL and OL were calculated and compared.

Integration K(OL) Sample K(EL) [mmol/g] Change [%] region [mmol/g]

Aliphatic OH 150.0-145.3 1.45 1.67 ± 0.03 +14.9

Syringyl aromatic OH 144.7-140.5 1.89 0.50 ± 0.01 -73.7 Guaiacyl aromatic OH 140.5-139.0 0.64 0.16 ± 0.00 -75.4 Catechol OH 139.0-138.2 0.06 0.02 ± 0.00 -73.1 p-hydroxyphenyl OH 138.2-137.3 0.02 0.012 ± 0.00 -47.3 Σ of aromatic OH 144.7-137.3 2.61 0.68 ± 0.01 -73.9 aliphatic carboxyl OH 136.5-135.4 0.02 0.01 ± 0.00 -45.9 aromatic carboxyl OH 135.4-133.0 0.05 0.01 ± 0.00 -69.4 Σ of carboxyl OH 136.5-133.0 0.06 0.02 ± 0.00 -63.3

3.3.3 Thermal Analysis of Lignin (Derivative)

The organosolv lignin (OL) and the derivatized lignins (EL) and (CL) were analyzed by TGA and DSC. The TGA thermograms are shown in Figure 3.3.3-1. The most prominent minima of the first derivative graph are marked and labeled with the associated temperature. In Table 3.3.3-1, the observed degradation temperatures and the corresponding mass loss are listed. The mass loss at 88°C of EL is not included in the table since the difficulty to assign whether it is due to water elimination, or to degradation products.

TGA thermogram of the OL display expected degradation[236], see also Figure 2.4.2-1. LeVan,1989 cit. by Ciobanu et al.[40] describe the degradation in three stages: starting with breaking of α- and β-aryl-alkyl- ether linkages by dehydration and decarboxylation process. This corresponds well with the recognizable shoulder of the curve at 150-260°C. The second and third stage are the cleavage of aliphatic side chains from the aromatic backbone at 260-290°C followed by splitting of carbon-carbon bonds between monolignols at 290-400°C. However, the degradation process is sometimes varied depending on lignin sources (from different species) and the pulping process[170].

In this study, the thermal stability decreased with each derivatization step. EL shows the lowest degradation temperature at 183°C, with that for CL at 216°C. The onset temperatures of both EL and CL

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Thermal Analysis of Lignin (Derivative) 3.3.3 appeared at the same temperature of around 120°C, whereby CL show the highest mass loss in the first degradation step at 216°C, followed by a second degradation at 291°C. EL also shows a second degradation at higher temperatures (334°C). In contrast to CL, the third degradation temperature of EL was lower than that of OL.

Figure 3.3.3-1: TGA termograms of OL (black), EL (red) and CL (blue) left side; first derivatives (right side).

Table 3.3.3-1: Degradation temperature with the mass loss of OL, EL and CL by TGA and glass transition temperatures of OL and EL samples (EL-1, EL-2, EL-3) by DSC.

OL EL CL

Td1 183°C 7.5 wt.% 216°C 16.3 wt.%

Td2 334°C 19.2 wt.% 291°C 10.3 wt.%

Td3 373°C 58.1 wt.% 385°C 29.3 wt.% 380°C 26.8 wt.% not T 116.5°C 72.4 ±4.0°C g quantifiable

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SEC Measurements of the OL and EL 3.3.4

3.3.4 SEC Measurements of the OL and EL

Size exclusion chromatography (SEC) was used to determine the average molecular weight number (Mn), weight average molecular weight (Mw), size (Z-)average molecular weight (Mz), the peak molecular weight (Mp) and the polydispersity (Pd). The measurement of CL molecular weight was not possible since CL was insoluble in any solvents used in SEC, see Subsection 0. The evaluated chromatograms are shown in Figure 2.1.1-1. The most important data are summarized in Table 3.3.4-1.

The SEC data of OL form Fraunhofer CBP and the University of Hamburg, are given in Table 5.1-1. The Mn values are comparable with our data, but Mw values show difference between the two measurements. The explanation for the mismatch of Pd which is directly proportional to Mw could be the use of different calibration standards. However, this observation could be also be affected by ageing, and reactions such as hydrolysis could also have decreased the Mw from 2800 g/mol (university Hamburg) to 1600 g/mol.

Figure 3.3.4-1: SEC of OL (back) and EL (red) dissolved in THF detected by RI-detector (line) and UV- detector (dotted line) When comparing the Mn data of EL with that of OL, it was observed that the molecular weights were almost doubled, with a slight change in Pd. This might be the reason for the loss of solubility, see Subsection 3.3.5.

The big differences between the given data (see Table 5.1-1) and our own observed values demonstrate the imperative to analyze the OL and EL under the same condition, as was done by our partners.

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Hansen Solubility Parameter of the OL, EL and CL 3.3.5

Table 3.3.4-1: SEC data of OL and EL in THF detected by UV- and RI- detector.

OL-UV OL-RI EL-UV EL-RI Mn [103 g/mol] 1.1 1.1 1.8 1.6 Mw [103 g/mol] 1.6 1.6 3.2 2.9 Mz [103 g/mol] 2.4 2.4 5.3 5.2 Mp [103 g/mol] 1.4 1.3 2.4 2.5 Pd 1.5 1.5 1.7 1.8 As expected, an increase in Mw was observed after the derivatization of OL with ECH. The intermolecular side reaction such as condensation would lead to multiplication of molecular weight. However, the resulting data shows that Mw increased to double that of OL. The slight changes in Mw and Pd may be due to marginal intermolecular side reactions.

3.3.5 Hansen Solubility Parameter of the OL, EL and CL

OL and EL were dissolved in 32 different solvents to observe the variations in cohesive energy density, see Table 6.2-5. Based on the observed solubility behavior in the solvents used, and with the special software HSPiP the Hansen solubility parameters were calculated, see Table 3.3.5-1. A graphical representation, shown in Figure 3.3.5-1, shows a three dimensional coordinate system with HSP dispersion (D), polarity (P), and hydrogen bonding (H) on the three axes. The used solvents are shown in the graph depending on whether EL was soluble in the solvent (blue spheres), or not (red cubes). The large sphere indicates the predicted range of HSP for solvents which should dissolve EL. The values of focal point of this area are the calculated HSP, listed in Table 3.3.5-1. Propylene carbonate and 1,4-dioxane (blue dotted spheres) are outside the large sphere although EL was soluble in this solvents. The total HSP (휹) was calculated using Eq. 3.

The HSP calculation for CL was not possible because the vacuum dried CL was no longer soluble in any of the solvents listed in Table 6.2-5.

Comparing the HSP’s of OL and EL is the dispersions solubility parameter quite equal, but the hydrogen bonding parameter and in particular the polar cohesion solubility parameter decreased. Expected changes are: the decreasing hydrogen bonding due to reacted hydroxy groups; and increasing polarity since the dipole moment of the previously phenols (1.224 D[238]) or aliphatic hydroxides (1.69 D3) was lower than for the epoxide group (1.89 D4). However, the data shows a decrease in polar HSP.

3 Dipole moment of ethanol[238] 4 Dipole moment of ethylene oxide (oxirane)[238] Albert-Ludwigs-Universität Freiburg 97

Hansen Solubility Parameter of the OL, EL and CL 3.3.5

Due to misleading naming, the measured and expected values for 휹푷 and 휹푯 could not be reconciled. However, the polar HSP is not only influenced by the dipole moment, it is also due to “the combined polar and hydrogen parameter”[210] and the hydrogen bounding HSP is a experimentally measured parameter (“subtracting the polar and dispersion energy of the vaporization from the total energy of vaporization”[210]). However, it was still possible to observe a decrease in the δ . This was expected since Table 3.3.5-1: Generic Algorithm used to calculate H dispersion (D), polar (P), hydrogen bonding (H) and hydroxyl groups are converted in epoxide and total cohesion Hansen solubility parameter δ. The calculated radius of the green sphere (R) and fit consequently the hydrogen bonds were lost. quality (Fit) of OL and EL.

Eventually, the dispersion HSP predominant Parameter OL EL influenced by of van der Waals forces is consistent, as 휹푫 [√푴푷풂] 18.11 18.35 expected. Despite the larger dipole moment, it is 휹푷 [√푴푷풂] 14.89 10.16 possible for 휹푷 and 휹푯 to decrease, as hydroxyl groups 휹푯 [√푴푷풂] 11.63 10.15 consumption leads to lower hydrogen bonds. The 26.17 23.31 smaller radius R of the calculated HSP of EL implies 휹 [√푴푷풂] that the solvent parameters to dissolve EL are more R 10.6 6.4 restricted. Fit 0.926 0.938

Figure 3.3.5-1: 3D-application of OL (left) EL (right) using the dispersion (D), polar (P), and hydrogen bond (H) cohesive Hansen-solubility Paramers. Blue spheres are selvents desolving EL, red cubes mark the solvents non- disolving the sample. The green speres show mark the parameter with are necessery to desolve EL.

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Summary Lignin Modification 3.3.6

3.3.6 Summary Lignin Modification

In the Subsection of PART II, efficacy of epoxidation and carbonization derivatizations is clearly shown by the FTIR, NMR, TGA and SEC techniques. It was established that: sinapyl alcohol (S) is the main monolignol in beech OL; and O-glycidylation affects all phenols independent of the substituents (syringyl, guaiacyl, or catechol).

31P-NMR analysis demonstrated that approximately three quarters of the aromatic hydroxyl groups react with ECH, but only 15% (0.216 mmol/g) additional aliphatic hydroxides were found The decrease in the aliphatic hydroxyl groups from lignin is not an indication of reactivity itself. To prove the reactivity of aliphatic hydroxyl groups with ECH, a reaction was performed with a lignin model compound under the same anhydrous conditions. This confirmed that the aliphatic groups did not take part in the epoxidation reaction.

Epoxide Indices (EI) in the range 0.35-1.48 mmol/g were observed by 1H-NMR, which is higher than the amount of additional hydroxyl groups. This indicates that the aliphatic hydroxyls reacted with ECH. But some side experiments with model compounds of lignin and suberin (vanillin alcohol and 1, 12- dodecandiol), which are not described in detail here, showed that less than half of the hydroxyl groups of vanillin alcohol, and none of the hydroxyl groups in diol, reacted under the applied conditions (as shown by FTIR and NMR). This indicates that the phenol groups in lignin reacted, and this implies that optimization of the reaction conditions is required for higher EI.

The DSC measurement shows that EL had a low glass transition temperature (Tg) despite that its Mw (SEC) is higher than that of OL. During the epoxidation, even though a marginal level of condensation led to a doubling in the Mw.

A verification of cyclic carbonates in CL was done with FTIR and NMR. The insolubility behavior of dried and, precipitated CL could not be explained with the performed experiments. One reason might be the side reaction with the remaining phenols.[176] Both carbonates and epoxides could polymerized to produce condensed system under the harsh conditions in the pressure reactor or during the precipitation of LC in different solvents.

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Lignin Based Non-Isocyanate Polyurethane 3.4

3.4 Lignin Based Non-Isocyanate Polyurethane

The cyclic carbonates derivatized lignin CL was synthetized (see Section 5.3) and analyzed, and the results are given in Section 3.3. This CL is used to produce lignin based non-isocyanate polyurethane (NIPU) by ring opening polymerization with different size diamines as a proof–of-concept, see Section 5.5. Therefore, this chapter deals with the fourth research question, drafted in Section 1.2 and marked orange in Figure 1.3-1 or Figure PART II-2.

FTIR analysis helped to determine, reaction between cyclic carbonates with diamines and allow some estimation about the “cyclic carbonate index” (CI) – as used for by EI. Thermal properties of the polymers were also determined by TGA and DSC.

3.4.1 Monitoring of Curing by FTIR

For monitoring the curing process, firstly the amount of diamine equal to the correct ratio of carbonates for CL should be calculated. The difficulty to determinate the CI is described in Section 3.3. So the proper ratio of amine/CL hat to be found experimentally. For these experiments, batches 2-4 of the CL samples were mixed together. Three commercially available, terminal diamines: 1, 12-dodecan diamine; 1, 4- butanediamine; and 1, 6-hexanediamine were used to synthetize (NIPU-1); (NIPU-2); and (NIPU3) respectively.

Various proportions of diamines to CL were tried for the NIPU reaction using the carbonate band at 푣̃=1793 cm-1 observed by FTIR as reaction prove. For better comparison, all spectra were normalized to 푣̃=1597 cm-1 (T=60%) except the dotted diamine reference spectra, see Figure 3.4.1-1. Each NIPU was given an individual colored FTIR spectrum: NIPU-2 (red) using the shortest chain diamine (1, 4- butanediamine); NIPU-3 cured by 1, 6-hexanediamine (green) and the NIPU-1 (blue) by longest chain diamine, 1, 12-dodecan diamine (blue). The spectra are overlapped with the specific diamine spectrum (dotted). Additionally, the spectrum of CL (black) and the spectra of NIPUs cured with an excess of diamines is displayed in Figure 6.1-13 for compassion with the adjusted NIPUs.

So far, only the carbonate band at 푣̃=1793 cm-1 is the appropriate peak to monitor the reaction by FTIR. A typical, not superimposed urethane band or bands of non-reacting amines were not observed. All the expected bands of urethanes and amines are overlapped with bands of CL, see Figure 3.4.1-1 and Table 6.2-4.

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Monitoring of Curing by FTIR 3.4.1

Figure 3.4.1-1: FTIR spectra of the educts: CL (black) and the relevant diamine 1,12-dodecan diamine (blue spotted); 1,4-butanediamine (red spotted); and 1,6-hexanediamine (green spotted) and spectra of the resulting PU colored with the right ratio for reacting all groups.

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Monitoring of Curing by FTIR 3.4.1

Consequently, only the carbonate band was used for the experimental observation. An application of the height of the resulting band at 푣̃=1793 cm-1 is shown in Figure 3.4.1-2 which depending on molar ratio of amines per gram CL. The grey horizontal line illustrate the 100% transmittance at 푣̃=1793 cm-1. This line markes the moment when no carbonate band was observable in the FTIR after curring with the corresponding amount. As a result, the lower thr transmittance, the higher the amount of non-reacting carbonates in NIPU sample.

Figure 3.4.1-2: FTIR cyclic carbonate band high monitoring of δ=1793 cm-1 normed by band of δ=1597 cm-1 (60%) after reaction with the three diamines: 1, 12-dodecan diamine (blue); 1,4-butanediamine (red); and 1,6-hexanediamine (green) in different proportional with CL. It was possible to observe a high diversification in the monitored data points. This caused first of all on the challenge to get a homogenous mixture of CL solution and the solid diamines. A curing of these two reactants was not observed at room temperature, so the salts were melted; see reaction conditions in Table 5.5-1. Nevertheless, some of the samples show a visibly recognizable inhomogeneity. Of course, only the apparently homogeneous samples were used for the measurements.

However, it was observed that reaction trend lines shows significant deviations on the necessary diamine concentration per gram CL. Figure 3.4.1-2 shows FTIR transmission data points of the cyclic carbonate band as a function of the amount of diamines used for curing. A transmission of zero means that all carbonates were consumed in the reaction. It is clear that different amounts of diamines are necessary to

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Monitoring of Curing by FTIR 3.4.1 react with all carbonate groups of CL. The amount seems to strongly depend of the length of the amine chain. Two things could affect discrepancies: reactivity and stereochemistry.

The three amines used for the production of NIPU’s have different reactivity based on number of CH2 groups in the aliphatic chain. Shorter 1, 4-butanediamine has a higher reactivity (producing NIPU-2) and the long chain 1, 12-dodecan diamine needs more activation energy (for synthesis of NIPU-1). As shown in Figure 3.4.1-2, NIPU-1 batches (blue squares) reach the full consuming of the cyclic carbonate peak (T=0 highlighted by the black line) with already low amount of diamine (0.46 mmol of amines per gram CL). For the short diamin, 1, 4-butanediamine, nearly 35% more excess diamine was needed to consume all carbonates for the production of NIPU-2 (red) than for the NIPU-1 (blue).

Figure 3.4.1-3: Schematically illustration of endcapped lignin NIPU (left) and cross-linked lignin NIPU (right). Substituents of epoxidation are marked red; of carbon ´dioxide are blue; and the diamines green.

The possible explanation for this behavior is that just one side of the shorter diamine reacts with the sterically hindered three dimensional lignin molecules. This resulted in endcapped lignin, with extension of the sidechains with terminal amine groups, see Figure 3.4.1-3 (left). With increasing chain length of diamine the possibility of cross-linking increases, see Figure 3.4.1-3 (right), consequently less amines are necessary to consume all cyclic carbonates.

Table 3.4.1-1: Determination of CI by using 1, 4-butanediamine (NIPU-2); and 1, 6-hexanediamine(NIPU-3); and 1, 12- dodecan diamine (NIPU-1) as curing agent of CL.

NIPU-2 NIPU-3 NIPU-1 0.70 mmol/g 0.67 mmol/g 0.46 mmol/g

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Solubility of NIPU’s 3.4.2

3.4.2 Solubility of NIPU’s

NIPU’s were synthetized with empirically determined amount of diamines, see Table NIPU-1 3.4.1-1 and their solubility in ethanolamine was NIPU-2 NIPU-3 excess NIPU-1 tested. After two weeks NIPU-2 and NIPU-3 were partially soluble, while NIPU-1 was insoluble, see Figure 3.4.2-1. The NIPU-1 excess sample was also prepared with an excess of diamines, this sample was partially soluble as Figure 3.4.2-1: Photo of the solubility test of NIPU's in with the samples NIPU-2 and NIPU-3. This ethanolamine after two weeks at room temperature. supports the theory that the long chain diamine react as a cross-linker with CL.

3.4.3 Thermal Analysis of the Lignin Based NIPUs

The thermal behavior of the NIPU’s with balanced amine/cyclic carbonate concentration was determined by TGA and DSC. The resulting degradation and glass temperatures and residual mass after heating to 900°C is presented in Table 3.4.3-1. The thermograms and their first derivatives are shown in Figure 3.4.3-1. The weight loss of all three samples after heating to 900°C is around 60 wt.%. The glass temperature (Tg) increased with increasing diamine chain.

Table 3.4.3-1: Degradations temperatures Td and the residual mass after heating to 900°C of NIPU 1-3 observed by TGA measurements. The glass temperature Tg was determined by the first heating cycle of the DSC measurements.

CL NIPU-2 NIPU-3 NIPU-1

Td1 216°C 16.3 wt.% 250°C 15.5 wt.% 250°C 10.6 wt.% 208°C 7.29 wt.%

Td2 291°C 19.2 wt.% 332°C 34.1 wt.% 326°C 18.9 wt.% 315°C 23.0 wt.%

Td3 380°C 26.8 wt.% 390°C 22.0 wt.% 471°C 24.7 wt.%

wt.900°C 40.3 wt.% 37.3 wt.% 37.6 wt.%

Tg 52.9°C 56.1°C 64.6°C

The thermograms of the three NIPUs show surprisingly large differences between them. The thermogram of NIPU-2 and NIPU-3 with the short chain diamines 1, 4-butanediamine, and 1, 6-hexanediamine respectively have similar degradation products even at low temperatures starting around 150°C with the maximum at 250°C, and a second degradation at about 330°C.

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Thermal Analysis of the Lignin Based NIPUs 3.4.3

However, NIPU-3 was found to have lower mass loss, in contrast to NIPU-2 at the first td. Consequently, a third degradation step appeared at the higher temperature.

NIPU-1 degradation started at lower temperatures (125°C), but the mass loss up to 260°C was equivalent to the NIPU-3, but lower than of NIPU-2. However, the maximum degradation occurred at significantly higher temperatures of around 471oC

The combination of the FTIR amine-hydroxyl ratio was used to get the right proportion of the curing agent for polymers, NIPU-1-3. From the thermograms it was suspected that the Td of around 250°C was connected with unreacted terminal amines. Most of them are expected to be in NIPU-2, with a small amount expected in NIPU-1.

Figure 3.4.3-1: TGA thermogram of NIPU-1 (blue), NIPU-2 (red) and NIPU-3 (green)

Furthermore, an increase of Tg from NIPU-1 to NIPU-3 was observed by DSC, see Table 3.4.3-1.

Generally, increasing aliphatic sidechains that are attached to the lignin will decrease the Tg. The data obtained from DSC shows that the Tg increase by increasing the diamine length. Based on these results, explanation could be that the long chain amine more favorable for cross-linking which increases the Tg.

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Conclusion of PART II

Conclusion of PART II

In the PART II of the thesis, organosolv lignin (OL) was the bio-based source for the epoxidation. Using ECH in anhydrous condition gave an organic solvent soluble product (EL) - in contrast to aqueous conditions. This derivatization decreased Tg in spite of an increasing MW, see Figure PART II-2. In the second step, the derivatization of lignin with cyclic carbonates, lignin carbonate (CL) was done under normal and high pressure conditions. Whereby, the carbon dioxide addition seems to be faster and more effective at higher pressure.

Figure Part II-2: Change of Tg with the made processing steps and variations in diamines

The insolubility of CL in all tested solvents, even with the solvent used for preparation, DMSO was noticed after removig the CL from DMSO by vacuum distillation or precipitation with solvents. Hence, CL was used without further purification for NIPU synthesis with terminal diamines. FTIR observation of the cyclic carbonate band showed that the required amount of amines is depending on the type of diamine (DA). Due to the DA chain size and amine/CL ratio a mainly endcapped or crosslinked NIPU was prepared (see Figure 3.4.1-3). This was proved by the solubility test in ethanolamine in Subsection 3.4.2, by thermal behavior especially increasing the Tg of DSC measurements in Subsection 3.4.3 (see Figure PART II-2), and empirical amine/CL ratio determination in Subsection 3.4.1. Consequently, the degree of crosslinks increase with increasing chain length of DA.

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General Conclusions and Perspectives 4

4. General Conclusions and Perspectives

This chapter aims to draw attention to the difficulties and possible alternative solutions of this work. The Section 4.1 deals with the work relating to Suberin (PART I), the second Section 4.2 with the works relating to the lignin (PART II).

4.1 Summary Measurement of Suberin

Each of the analytical methods used to quantify and qualify the SM has their advantages and disadvantages. This gave results which validate, but also partly contradict each other:

 The proportion of Suberin in the beech bark was substantially lower compared to Quercus suber.  The observed difference between suberin mass and the lost mass is explained in the literature as the loss of glycerin, but we could also show that some phenolics also stay in the aqueous phase during the purification via liquid-liquid extraction.

 The phenolic content in SM was clearly observed in FTIR, UV-Vis and GC-MS. NMR did not show aromatic protons, only the methyl protons of methoxy groups give an indication that

aromatic groups were still present in the SM sample. On the other hand the EA show a proton/carbon ratio of 1.7. This indicates that we do not have just a trace of aromatics.  The observed phenols are simple and not precipitable with PVPP.  DOSY measurements showed a uniform diffusion coefficient, indicating a uniform molecular size. This contradicts the suggestion that a part of the sample was not detected in the GC-MS due to differences in the molecular size (smaller molecules RT< 5 min, or large molecules which are not gasifiable).  With the two dimensional NMR methods almost all 1H-NMR signals were identifiable. And with the TAI derivatization a separation and integration of the peaks was possible. This allowed calculation of the average CL, and location of functional groups in terminal or mid-chain positions.  The calculated CL of 16 carbons was less than 18 carbon chains expected from the GC-MS.  In the FTIR the typical methyl group vibration was not observed, and also from the GC-MS a higher amount of hydroxy and carboxylic acid terminal groups was expected, since α-hydroxy acids were the main part of the observed peaks. The large amount of terminal methyl groups observed in TAI-1H NMR was not expectable from the other methods and could be the reason for the shorter average CL.

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Using Lignin as Precursor of NIPUs 4.2

 Compared to “waxes” found in the DCM extract (see Subsection 3.1.2) the SM showed a greater amount of functionalities per chain.

4.2 Using Lignin as Precursor of NIPUs

In the PART II of this thesis the most important question was if lignin could be carbonated to be usable as a precursor in the NIPU synthesis. For the first time, parallel with Salanti et al.[12, 209] it was possible to show that carbonization of lignin works. But now and maybe for future work it is important to clarify the optimal conditions for each reaction step.

As we know now that EPH just reacts with the aromatic hydroxyls under anhydrous conditions. May be it is possible to increase the EI of EL using other conditions. Preliminary hydroxypropylation could solve the solubility problem.

Furthermore, it is still important to know if the distillation and toluene extraction effectively removed all unreacted ECH from EL, and what influence the ageing process of EI in EL has.

Also the condition for the addition of carbon dioxide could be enhanced. Salanti et al.[12] already made first tests with diverse catalysts. We could show that carbonization is also possible without using pressure, and maybe it is possible to find milder conditions with a good CI in CL.

Salanti et al.[209] solved the problem of the solubility of CL by acetylation of CL. This is interesting because it creates more analytical possibilities, and an amine functionalized macromere is obtained, since an excess of diamine has to be used for the reaction with carbonate.

It is interesting to find out if aliphatic carbonates from for example suberin could dissolve CL. If so, it would be possible to produce a non-isocyanate polyurethane copolymer, as planned for this work. It would be interesting to test this copolymer NIPUs for their thermal and mechanical properties.

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Experimental 5

5. Experimental

Materials, chemicals, reaction conditions, and analytical equipment used for the implementation of practical experiments are listed and described in this Chapter. Section 5.2, Bark preparation and extraction, describes the extraction analysis of suberin, see PART I of this thesis. Sections 5.3; 5.4; and 5.5 describe the processing of Organosolv Lignin (OL), of model substances (SM and LM), and the NIPU synthesis of PART II. The analytical characterization via diverse methods and appropriate sample preparation is described in Section 5.6.

5.1 Materials and Chemicals

The beech (Fagus sylvatica) bark used was provided by Holzwerk Keck at three different times. The dry mass of each bark samples (BS) was approx. 60-70 kg.

 BS 1: April 2013  BS 2: June 2013  BS 3: March 2014

The organosolv lignin “KS018” used in the experiments was provided from the Fraunhofer Center for Chemical-Biotechnical Processes (CBP) in Leuna (Germany). The beech wood lignin was analyzed by the project partners. Results of the elemental analysis, high-performance liquid chromatograph (HPLC) after hydrolysis, Size-Exclusion Chromatography (SEC) and Phosphorus-31 Nuclear Magnetic Resonance (31P-NMR) are listed in Table 5.1-1:

Table 5.1-1: Lignin data determined by Fraunhofer CBP and SEC by University of Hamburg.

Elemental analysis [%] C H O N S H/C O/C 64.71 5.99 28.99 0.31 0 1.10 0.34 Results from hydrolysis and subsequent HPLC ∑ Sugars Acid soluble ∑ Total Klason lignin ∑ Total lignin Glucose hemicellulose- lignin sugars based sugars 91.4 2.19 93.59 0.15 1.37 1.52 SEC (Uni Hamburg) SEC (Fraunhofer) Polydispersity Polydispersity M M M M n [g/mol] w [g/mol] Pd ( Mw/Mn) n [g/mol] w [g/mol] Pd (Mw/Mn) 1000 2800 2.8 900 2400 2.6 31P-NMR [mmol/g Lignin] Other methods aliphatic OH aromatic OH COOH Tg[°C] Ash content [%] 1.62 2.93 0.09 120.1 0.13

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Materials and Chemicals 5.1

Commercially available chemicals with their source of supply used in the experiments are alphabetically listed in Table 5.1-2.

Table 5.1-2: Alphabetical list of chemical used with source specification

Chemical Supplier 1, 4-Dioxane Merck, Germany 1-Butanol (BuOH) Grüssing, Germany 2-Chloro-4, 4, 5, 5-tetramethyl-1, 3, 2-dioxaphospholane Sigma-Aldrich, Germany (TMDP) 2-Ethyl-1-Butanol Aldrich, Germany 2-Propanol Sigma-Aldrich, Germany 4-nitrobenzaldehyde (IS2) Sigma-Aldrich, Germany Abietic acid 85% Acros, Germany Acetic acid Roth, Germany Acetic Anhydride Sigma-Aldrich, Germany Acetone Sigma-Aldrich, Germany Acetonitrile Sigma-Aldrich, Germany Acetophenone Acros Organics, US Aleuric acid Sigma-Aldrich, Germany Ammonium iron(III) sulfate dodecahydrat Applichem Argon gas Messer group, Germany Benzaldehyde Roth, Germany Butyl lactate Aldrich, Germany Catechine Sigma-Aldrich, Germany Chlorobenzene Bernd Kraft, Germany Chloroform Fisher, US Chromium(III) acetylacetonate (CrAcAc) Sigma-Aldrich, Germany Cyclohexanol (IS3) Sigma-Aldrich, Germany Cyclohexanone VWR, Germany

Deuterated chloroform (CDCl3) Euriso-top, France

Deuterated dimethyl sulfoxide (d6-DMSO) Euriso-top, France

Deuterium (D2O) Euriso-top, France Diacetone alcohol Sigma-Aldrich, Germany Dichloromethane (DCM) 99.9% Sigma-Aldrich, Germany Diethylene glycol monobutyl ether Aldrich, Germany Diethylether Sigma-Aldrich, Germany

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Materials and Chemicals 5.1

Chemical Supplier Dimethyl sulfoxide (DMSO) Sigma-Aldrich, Germany Dipropyl amine Aldrich, Germany 1, 12-Dodecandiol Sigma-Aldrich, Germany Dry methanol 99.8% (MeOH) BenrdKraft, Germany Epichlorohydrin (ECH) Sigma-Aldrich, Germany Ethanol (EtOH) VWR, Germany Ethanolamine Sigma-Aldrich, Germany Ethylacetate VWR, Germany Ethylbenzene Funca, Germany Folin-Ciocalteu reagent (FC) 2 mol/l Sigma-Aldrich, Germany Furfuryl alcohol Sigma-Aldrich, Germany Gallic acid Sigma-Aldrich, Germany Hexane VWR, Germany Hydrochloric acid (HCl) Sigma-Aldrich, Germany 4-Hydroxyl-3-methoxybenzyl alcohol Sigma-Aldrich, Germany Methanol (MeOH) VWR, Germany N, N-Dimethylacetamide Riedel De Haën, Germany NEOCHEMA GmbH & Co. KG, n-Alkane Mix 16 (C10-C40 even) Germany Nitrogen gas Messer group, Germany Nitrogen liquid N-Methyl-N-(trimethylsilyl) trifluoroacetamide (MSTFA) Sigma-Aldrich, Germany o-Dichlorobenzene Sigma-Aldrich, Germany Polyvinyl polypyrrolidone (PVPP) Applichem, Germany Potassium bromide (KBr) Sigma-Aldrich, Germany Propylene carbonate Sigma-Aldrich, Germany Propylene Glycol Sigma-Aldrich, Germany p-Xylene VWR, Germany Pyridine Sigma-Aldrich, Germany Rutine Abcr, Germany Sodium carbonate anhydrous Sigma-Aldrich, Germany

Sodium carbonate decahydrate (Na2CO3 x 10H2O) Sigma-Aldrich, Germany Sodium methoxide (MeONa) solution 25 wt.% in methanol Sigma-Aldrich, Germany

Sulfuric acid (H2SO4) 95% Fluka analytical, Germany Supelco® 37 Component FAME Mix (FEME) Sigma-Aldrich, Germany

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Materials and Chemicals 5.1

Chemical Supplier Tannin Tetrahydrofuran (THF) Sigma-Aldrich, Germany Tetra-n-butylammonium bromide (TBAB) Sigma-Aldrich, Germany Toluene Sigma-Aldrich, Germany Trichloroacetyl isocyanate (TAI) Fluka analytical, Germany Vanillin Acros, Germany

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Bark preparation and extraction 5.2

5.2 Bark preparation and extraction

BS 1 and BS 2 were dried at 75°C and stored in plastic barrels in cool condition. Similar parts of these samples were mixed with each other and roughly chopped with the cutting mill SM 300 using 2 mm square hole bottom sieves from Retsch (Germany). The fine milling was made using the vibrator disc mill RS 200 from Retsch (Germany) at 1200 rpm for 40-60 s. The mill shaker 60°mesh from Retsch (Germany) was used to separate bark particles >250 µm.

A part of BS 3 was separated in inner bark (IB) and outer bark (OB) manually. Separated and non- separated BS 3 was stored frozen.

The water content (푤), Eq. 8, and bark humidity (푢), Eq. 9, of the fresh, pre-dried and fine milled BS was determined three-times gravimetrically after drying at 103°C for a minimum of 48 h.

푚푤푒푡 − 푚푑푟푦 푚푤푎푡푒푟 Eq. 8 푤 = ∗ 100% = ∗ 100% 푚푤푒푡 푚푤푒푡

푚 Eq. 9 푢 = 푤푎푡푒푟 ∗ 100% 푚푑푟푦

푚푤푒푡: mass of wet bark; 푚푑푟푦: mass of dry bark and 푚푤푎푡푒푟: mass difference after drying correspond to the mass of water

5.2.1 Solvent Extraction of Bark

Approximately 40 g milled bark was extracted for 8 h by successive Soxhlet extraction in a cellulose thimble with Dichloromethane (DCM) at 60°C, methanol (MeOH) at 85°C, and water at 125°C, respectively (see the steps 1.1-1.3 in Figure 5.2.3-1). The extract free bark was dried in air, frozen and freeze-dried in an Alpha 1-2LDplus freeze-dryer, Christ (UK), for minimum of two days. Subsequently, the sample was crushed, mixed, frozen and freeze-dried for at least another two days (see the steps 1.1- 1.3 in Figure 5.2.3-1).

5.2.2 Depolymerization (Methanolysis) of Bark Suberin

The milled, extract-free bark was used for the methanolysis in dry methanol with Sodium methoxide (0.025 mol/l). The mechanism of the transesterification reaction is shown in Figure 5.3.1-1. The nucleophilic methoxide ion (2) attacks the carbonyl carbon of the ester (1) to give a tetrahedral intermediate (3), which can produce the methyl ester (4) and a new alkoxide (5) which is soluble in

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Purification of Suberin Monomers 5.2.3 methanol. The remaining bark I suspension was refluxed at 75°C. After 2 h the contents of the flask were filtered through the glass-filter. The solid bark residue was refluxed once more with methanol at 75°C for 1 h, filtered and dried at room temperature and finally in an oven at 75°C (see the steps 2.1-2.4 in Figure 5.2.3-1).

Figure 5.2.2-1: Reaction mechanism of methanolysis.

5.2.3 Purification of Suberin Monomers

The combined filtrates were acidified to pH 6-7 with dilute sulfuric acid (H2SO4). The suberin monomer solution was evaporated to almost dryness with a Heidolph G1 rotary evaporator. The monomers were suspended in water and DCM. The resulting phases were separated and the water phase was additionally extracted twice with approximately 40 ml DCM. The combined organic phase was concentrated in the rotary evaporator (see the steps 3.1-4.4 in Figure 5.2.3-1).

milled bark 2.4 filtration (solid) remaining bark II 1.1 DCM DCM extract 2.3 NaOMe in dried MeOH 8h at 60 °C reflux 1h at 70°C (liquid) 1.2 MeOH MeOH extract remaining bark I 8h at 75 °C (solid) suberin monomers in extraction 1.3 water water extract alkali MeOH solution 8h at 125 °C (liquid)

1. Successive soxhlet soxhlet 1. Successive 2.2 filtration of the 3.1 neutralization 2. Methanolysis suspension with diluted H SO extracted free 2 4 bark 2.1 NaOMe in dried MeOH of pH ~6 reflux 2h at 70°C 3.2 evoporated almost to dryness

3. Purification 3.3 suspended in water suberin monomers 3.4 liquid-liquid extraction with DCM suberin monomers in DCM water suspension >3 times + DCM

water phase

Figure 5.2.3-1: Process scheme of the suberin monomer (SM) extraction from bark.

The variously treated barks (BN, BE, and BR), were analyzed by solid state NMR, FTIR, TGA and DSC. Albert-Ludwigs-Universität Freiburg 114

Lignin Preparation 5.3

5.3 Lignin Preparation

The organosolv lignin (OL), “KS018”, provided from Frauenhofer Center was dried for 20 h at 70°C in a vacuum oven at a pressure of 200 mbar. The water content (푤) (see Eq. 8) was determined based on gravimetric measurements before and after drying. The OL was characterized by FTIR, NMR, TGA, DSC, GPC, and the Hansen Solubility Parameter (HSP) was determined. Detailed processing is described in Section 5.6. and the results are presented and discussed in Section 3.3.

Figure 5.3-1: Process scheme of lignin O-glycidylation and CO2 addition to produce EL and LC.

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O-Glycidylation of Organosolv Lignin 5.3.1

5.3.1 O-Glycidylation of Organosolv Lignin

The organosolv lignin was derivatized with epichlorohydrin (ECH). Two possible nucleophilic substitution reactions (SN2) are illustrated in Figure 5.3.1-1. The nucleophilic (electron donor) oxygen hydroxyl groups of lignin (shown schematically) (1) can attack ECH (2) on two possible electron deficient carbons: In one case, scenario (I), the nucleophile attac the methylengroup of the chain and chloride is the leaving group. An ether-bonded lignin epoxide (EL) (4) is formed. In the second case, scenario (II), the carbon of the epoxide ring methylene is attacked by the nucleophile so that a ring opening reaction is formed the product (3). In an alkali environment (3) is converted to propylene oxide (4) by an elimination reaction.

(I) S 2 O HO Cl N + O O - R O - Cl (4) H3C O -NaCl, - - 1/2 H O 2- O 2 +1/2 CO3 Cl - 1/2 CO + Na+ HO + 2 OH (II) O S 2 + N H O Cl Ring Opening R (1) (2) (3)

Figure 5.3.1-1: Reactions mechanism of epichlorohydrin reaction

The scheme of O-glycidylation is illustrated in Figure 5.3-1. The O-glycidylation was done once using aqueous and four times using anhydrous conditions. The procedures of the anhydrous condition reactions are described in Table 5.3.1-1. The aqueous reaction was done as described by Malutan, Nicu and Popa[181]. 10.56 g Lignin (undried) was suspended in 150 ml 20% NaOH solution by stirring for 30 mins at room temperature. Then 90 ml ECH was added to the suspension and stirred again for 3 h at 70°C. The suspension was neutralized, centrifuged and the settled brown material was washed twice with acid solution, and three times with water. The product, denoted as EL-5, was freeze-dried before characterization.

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O-Glycidylation of Organosolv Lignin 5.3.1

Table 5.3.1-1: Procedure and water- free using conditions for O-glycidylation of lignin to produce lignin epoxide (EL).

Step performance EL-1 EL-2 EL-3 EL-4

1.1 Lignin was dissolved in ECH in aninert Lignin: 11 g(dry) 21 g 30 g 31 g atmosphere (argon). The solution was ECH: 100 ml 200 ml 300 ml 260 ml heated to 70°C in reflux. Anhydrous

Na2CO3 was added to the solution. The Na2CO3: 20 g 30 g 40 g 5 g suspension was stirred for 7 or 24 h at NaOH: ------2 g 70°C and then cooled down to room Time: 7 h 7 h 7 h 24 h temperature.

1.2 The suspension was filtered with a glass

cellulose filter. filter jar

1.3 The lignin epoxide solution was vacuum distilled (60 mbar at 45°C) to almost dryness. Some toluene was added and distilled again, with more than three repetitions. Finally the material was distilled to dryness.

1.4 Additional purification: EL powder was suspended several times in toluene, centrifuged and the solvent decanted.

1.5 The solid lignin epoxide was vacuum dried at 45°C.

The Epoxy index of the samples was determined by proton NMR using Eq. 10. The amount of epoxides

푛푒푝표푥 per mass of epoxy-enhanced material 푚푟푒푠푖푛 used was calculated by using an internal standard.

푛푒푝표푥[푚표푙] 퐸퐼 = Eq. 10 푚푟푒푠푖푛[푘푔]

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Carbon Dioxide Addition Reaction with Lignin Epoxides 5.3.2

5.3.2 Carbon Dioxide Addition Reaction with Lignin Epoxides

Carbon dioxide was added to the epoxy-enhanced lignin (EL). The reaction mechanism is illustrated in Figure 5.3.3-1, the procedure is shown schematically in Figure 5.3-1 step 2, and the different conditions used are shown in Table 5.3.2-1.

Lignin carbonates (CL) 1-3 were prepared in a Limbo high pressure reactor from Büchiglasuster

(Switzerland), lignin carbonate CL-4 was prepared in reflux equipment with high-purity CO2 (N45). d6- DMSO was used for samples CL-3 and CL-4.

Table 5.3.2-1: Using conditions for addition of carbon dioxide.

Substance / Condition CL-1 CL -2 CL -3 CL -4

EL [g] 3 3 6 46

TBAB [g] 0.2 0.2 0.4 0.4

DMSO [ml] 20 100 40 100

Pressure p [bar] 20 25 25 1

Temperature T [°C] 130 120 120 120

Time t [d] 3 2 2 7

In the catalyzed reaction mechanism [207], shown in Figure 5.3.3-1, an forms at first an intermediate structure forms at first with lignin propylene oxide or EP (1), carbon dioxide (2) and bromide coordinated by TBA-H+ (Tetra-n-butylammonium bromide (TBAB)). A nucleophilic attack by the bromide opens the epoxide ring on the epoxide methylene. The resulting oxy-anion attacks the carbon dioxide carbon nucleophile and forms the intermediate carbonate anion (4). The final ring form of propylene carbonate develops by disconnection of the leaving bromide group.

5.3.3 Hansen Solubility Parameter (HSP) Determination

The solubility of lignin and the derivatized lignin EL and CL were tested by dissolving ca. 5 mg material in 5 ml of various solvents, see Table 6.2-5. After one day at room temperature the dissoled state was evaluated, and given an index of zero for insoluble, one for completely soluble, and zero plus for partially soluble lignins. The data were analyzed by “HSPiP 5.0.04” (UK) software to determinate the Hansen solubility parameter.

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Hansen Solubility Parameter (HSP) Determination 5.3.3

C H – 4 9 H9C4 Br + + H C N C4H9 H9C4 N C4H9 9 4 Br – O C4H9 1 O O C4H9 R O C 1 O R O O O

– Br H C O 9 4 + C4H9 H9C4 N 1 O R O O C4H9

Figure 5.3.3-1: Reaktions mechanism of carbon dioxide (2) addition to EL (1) catalyzed by TBAB (3) [207].

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Lignin and Suberin Model Compounds Modification 5.4

5.4 Lignin and Suberin Model Compounds Modification

Two model substances, one as suberin (SM) and one for lignin (LM) replacement were used for observation of the epoxidation and carbonization processes. For SM 1, 12-dodecandiol was used and for LM 4-hydroxyl-3-methoxybenzyl alcohol was used.

Epoxidation, similar to that for lignin, as described in subsection 5.3.1, in aqueous and water free conditions was performed on the models to produced epoxy suberin model (ESM) and epoxy lignin model (ELM).

Table 5.4-1: Anhydrous O-glycidylation conditions for LM

performance ELM-1 ELM-3 ELM-4

Model: 5 g 5 g 5.5 g Reaction: Lignin model (LM) (4-hydroxyl-3- methoxybenzyl alcohol) was dissolved in ECH ECH: 156 ml 150 ml 160 ml in an inert atmosphere (argon). The solution was Na2CO3: 50 g 30 g 30 g heated to 70°C in reflux. Anhydrous Na2CO3 NaOH: --- 1 g 1 g (and NaOH) was added to the solution. The suspension was stirred for 7 or 24 h at 70°C and TBAB ------1.5 g

then cooled to room temperature. Time: 7 h 22 h 24 h

Purification: The lignin epoxide solution was filtered hot, and then vacuum distilled (60 mbar at 45°C) to almost dryness. Some toluene was added and the material was distilled again. This was repeated more than three times. Finally, the material was distilled to dryness.

Additional purification: EL powder was suspended several times in toluene, centrifuged and the solvent decanted.

Drying: The solid lignin epoxide was vacuum dried at 45°C.

ELM-2 was performed following the instruction of Malutan, Nicu and Popa[181] or the same as for EL-5 (Subsection 5.3.1) using 5 g 4-hydroxyl-3-methoxybenzyl alcohol, 75 ml 20% NaOH solution and 45 ml ECH.

ESM-1 6.6 g SM (1, 12-dodecandiol) was dissolved in 150 ml ECH in inert atmosphere (argon). The

solution was heated up to 70°C in reflux, 30 g anhydrous Na2CO3 was added to the solution. After 22 h reaction a sample (a) was taken and analyzed, four hours later 5 ml of 50% NaOH was added. 19 h later a second sample (b) was taken. 3 days later the reaction was stopped.

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Lignin and Suberin Model Compounds Modification 5.4

Purification was performed on all samples taken as described in Table 5.4-1. 1.5 g NaOH (1.1 eq. of hydroxyls) was added to the yellowish viscous product, stirred and heated to 30°C for 30 min. The solution was neutralized by three times liquid-liquid extraction with ethyl acetate. The organic phase was evaporated at first without vacuum, and later under vacuum at 45°C.

ESM-2 10 g SM (1, 12-dodecandiol), 500 mg TBAB and 84 g KOH were dissolved in 27 ml ECH and 250 ml hexane under an inert atmosphere (argon) following the instruction of Karuna et al..[230] The suspension was refluxed for 6 h and then filtered. Hexane was recovered by evaporation and the mass of the residual product was measured. The mass obtained (15.7 g) corresponds to a yield of 99%, assuming that 100% of the hydroxyl groups were converted.

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NIPU Synthesis 5.5

5.5 NIPU Synthesis

Three kind of PU were prepared by addition of three different diamines under conditions adjusted to the melting temperature of the type of diamine (DA) used, see Table 5.5-1

Table 5.5-1: Reaction condition to produce PU-1, PU-2 and PU-3

Sample PU-1 PU-2 PU-3 Diamine 1, 12-dodecan diamine 1, 4-butanediamine Hexane-1, 6-diamine

Tm [°C] 67-69 25-28 39-43 Reaction temperature [°C] 80 40 50 After 5 min homogenization of the reaction material at the indicated temperatures the samples were cured at 80°C for minimum 8 h following by 24 h in vacuum.

The NIPU’s were made into films by pressing the material obtained between two Teflon films, followed by drying after removing the top layer, see Figure 5.5-1

Figure 5.5-1: Photo of NIPU film production

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Characterization Methods 5.6

5.6 Characterization Methods

The following describes the measurement parameters, the software and hardware used for the measurement, the interpretation of the data, and the sample preparation method used, which depended on the technique.

5.6.1 Fourier Transform Infrared Spectroscopy (FTIR)

The samples (bark, lignin and suberin) were characterized with an FT-IR 65 spectrometer from Perkin Elmer (Germany) in the Attenuated Total Reflectance (ATR)-mode. The cumulative 32 or 64 scans were measured with a changing resolution 1-4 cm-1 in the range of 600-4000 cm-1. The data was recorded using the “Spectrum” software (Perkin Elmer, Germany) using the automatic background correction facility. Additionally the data were processed with automatic baseline and ATR correction. The data were partially normalized to a sample dependent band before exporting the data.

−1 For the calculation of the S/G ratio the absorbance 퐴 of the typical S unit vibration band 퐴푠(1326 푐푚 ) −1 [169] was divided by the typical G unit vibration band 퐴퐺(1266 푐푚 ), see Eq. 11.

−1 퐴푠(1326 푐푚 ) −1 = 푆/퐺 Eq. 11 퐴퐺(1266 푐푚 )

5.6.2 UV-Vis Spectroscopy

Different types of measurements (see below) were performed in a 1 cm path length quartz cuvette with a UV/Vis spectrometer from Perkin-Elmer. The measurement range was 200-800 nm with a resolution of 1 nm.

Total Phenol Content (Folin-Ciocalteu-Method)

It is possible to determine the total phenol content in a solution by the method of Otto Folin and Vintila Ciocalteu [217]. A calibration curve was determined with gallic acid (GA) as standard (2-24 mg/L), as recommended by Singleton et al.[217]. Various gallic acid concentrations, solutions of the extracts and extracted suberin were used for the phenol Figure 5.6.2-1: Photo of sample with phenols (left) and blank feed (right) measurements. In each case 0.5 ml of the solution was Albert-Ludwigs-Universität Freiburg 123

Nuclear Magnetic Resonance (NMR) Study 5.6.3 added to 0.25 ml 1 mol/l Folin-Ciocalteu (CF) reagent and 1.25 ml 20% sodium carbonate solution. After 40 ± 3 min reaction time the absorbance of samples were measured at 725 nm and 747 nm [215, 216, 239]. Additionally, the absorbance spectrum was measured for various samples, and the gallic acid spectra was observed kinetically.

Indirect Determination of Tannins

For the determination of tannins 1 ml sample solution was suspended with 100 mg PVPP at 4°C for 15 min, centrifuged for 10 min at 3,000 rpm and then prepared the samples as per description in total phenol content measurements (see above) [215, 240, 241].

Determination of Condensed Tannins

For the determination of condensed tannins the Vanillin-HCl methods was used[215, 242]. A BuOH-HCl-reagent solution was made by mixing 95 ml butanol (BuOH) and 5 ml 37% HCl. Ferric- reagent was prepared by diluting 8.3 ml (37%) HCl and 1 g ammonium iron (III) sulfate dodecahydrat with water to a final Figure 5.6.2-2: Photo of diverse tannins free and samples (clear yellowish color) volume of 50 ml. The sample (0.5 ml) was added to 3 ml BuOH- and samples with tannins (reddish color) HCl reagent and 100 µl Ferric-reagent. The mixture was boiled at 95°C in a closed test tube for one hour. After cooling down to room temperature, the absorbance was determined at 550 nm. Using the molar 푙∙푐푚 attenuation coefficient for (2R, 3S, 4R)-leucocyanidin 휀1% = 460 [220] and the Beer-Lambert law: 550푛푚 푚표푙

퐴 = ε ∙ c ∙ d, Eq. 12

Where 퐴 is the absorbance, 휀 is the molar attenuation coefficient, c is the concentration and d is the path length of the beam of light through the sample.

5.6.3 Nuclear Magnetic Resonance (NMR) Study

One dimensional (1H, 13C and 31P) and two dimensional spectra (COSY, TOCSY, HSQC and HMBC) were measured using the “Bruker Avance 300” NMR spectrometer (Bruker, Germany); the device settings are listed in Table 5.6.3-1. All shifts were analyzed using the “Bruker TopSpin 3.2” software (Bruker BioSpin, US) and were expressed as parts per million (ppm) downfield from TriMethylSilyl (TMS).

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Nuclear Magnetic Resonance (NMR) Study 5.6.3

Table 5.6.3-1: NMR measurement conditions

Number of scans Frequency [MHz] Proton Nuclear Magnetic Resonance (1H-NMR) 32 300 Carbon Nuclear Magnetic Resonance (13C-NMR) 4000 75 Phosphorus Nuclear Magnetic Resonance (31P- 4 121 NMR) Diffusion-ordered spectroscopy (DOSY): 4 0-375 Total correlation spectroscopy (Tocsy): 8 300 Heteronuclear single-quantum correlation 8 75 and 300 spectroscopy (HSQC)

Heteronuclear multiple-bond correlation 8 75 and 300 spectroscopy (HMBC)

Sample preparation: Approximately 30 mg of the samples were dissolved in about 600 µl deuterated solvent (D2O, DCCl3, d6-DMSO). Sometimes an internal (4-nitrobenzaldehyde) standard was used in a known concentration.

Derivatization with TAI

100 – 150 µl TrichloroAcetyl Isocyanate (TAI) was added to the dissolved sample under an inert gas (argon) atmosphere. The possible reactions of TAI (1) with sample groups (2) and (4) are shown in Figure 5.6.3-1.

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Nuclear Magnetic Resonance (NMR) Study 5.6.3

O O O N ROH (2) R C C CCl 3 O NH CCl 3 O (1) (3)

(4) ROOH O O

C + CO 2 R NH CCl 3 O O O (8)

C  R O NH CCl 3 (5) O O Cl 3C NH NH CCl 3 1/2 + 1/2 C C + 1/2 CO 2 R O R O O O (6) (7)

Figure 5.6.3-1: Reaction of trichloroacetyl isocyanate (TAI) with hydroxyl and carboxyl groups Derivatization with TMBP

The sample (1) was dried under vacuum at 40°C for a minimum of 24 h. A solvent solution was prepared by adding 100 mg cyclohexanol (internal standard) and 90 mg of chromium acetylacetonate (relaxation agent) in 25 mL pyridine/deuterium 1.6/1 (v/v) solvent. The samples (20-25 mg) (OL-1=28.1 mg; OL- 2=26.0 mg; EL-2=24.8 mg) were diluted in 400 µl solvent solution and 150 µl pyridine/deuterium solvent and mixed for few minutes. 70 µl 2-Chloro-4, 4, 5, 5-tetramethyl-1, 3, 2-dioxaphospholane (TMDP) (2) was add to the sample solution, shaken and transferred to an NMR tube. The reaction is shown in Figure 5.6.3-2.

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Differential Scanning Calorimetry (DSC) 5.6.4

Figure 5.6.3-2: Phosphorylating of hydroxyl functional groups with 2-chloro-4, 4, 5, 5-tetramethyl–1, 3, 2– dioxaphospholane (TMDP)

5.6.4 Differential Scanning Calorimetry (DSC)

Approximately 8 mg of sample (bark, lignin (derivative), suberin) was heated to 100°C from -30°C at a heating rate and cooling ratio of 10°C/min in two cycles with 2 min isotherm holding at the temperature extrema, using a nitrogen flow of 20 ml/min in a DSC-8500 Differential Scanning Calorimeter from

Perkin Elmer (Germany). For the determination of the glass temperature (Tg) the “Spectrum” program was used (Perkin Elmer, Germany).

5.6.5 Thermogravimetric Analysis (TGA)

Approximately 10 mg of sample (bark, lignin (derivative), suberin) was heated to 900° C from room temperature using a heating rate of 10° C/min and a nitrogen flow of 20 ml/min in a Pyris thermogravimetric analyzer from Perkin Elmer (Germany). For the determination of the degradation temperature (Td) the maximum of the first derivative was calculated with the “Pyris 1” thermos gravimetric analyser (Perkin Elmer, Germany).

5.6.6 Gas Chromatography-Mass Spectrometry (GC-MS)

GC-EI-MS measurements were made using an “Agilent 7890A/5975C” system (Agilent Technologies, Germany) with a “Gerstel MPS 2 XL” autosampler (Gerstel, Germany) and a 30 m x 0.25 mm x 0.25 µm “HP-5MS” capillary column (Agilent, Germany). The data was recorded by the “MSD ChemStation” software (Agilent Technologies, Germany). Data files were analyzed by “AMDIS Analysis” using the “NIST” data library for identification, and “MZmine 2” and “Origin Pro” for quantification. Albert-Ludwigs-Universität Freiburg 127

Elemental Analysis 5.6.7

A sample volume of 1 µl was injected in a split-splitless inlet used at 230°C in pulsed splitless mode. The inlet pressure was 9.1 and injection pulse pressure 25 psi until 0.5 min. The carrier gas helium was used at a flow rate of 1 ml/min. Temperature program of the column oven: 2 min at 80°C, then continues increase of the temperature at 5°C/min to 325°C and held for 10 min at 325°C. Therefore the total rum time was 61 min. Transfer line temperature was 280°C. The detection started after a solvent delay of 4.80 min. For the ionization and detection was used 70 eV in the full-scan made from 50 to 500 m/z at scan rate of 3.5 s-1. For the calibration, a “PFTBA auto tune” program was run approx. every 50 injections. The retention index was calibrate with n-Alkane Mix 16.

Derivatization Trimetylsilylation of alchos using N-Methyl-N-(trimethylsilyl) trifluoroacetamide (MSTFA) showed in Figure 5.6.6-1. 1 ml of samples dissolved in DCM (approximately 2 mg) and different mass of the standards used were evaporated to dryness. 20 µl Pyridine and 80 µl MSTFA (1) was added and shaken for 30 min at 37°C with 1200 rpm on a thermomixer (Eppendorf, Germany). After the reaction the derivatized samples or standards were transferred to crimp capped GC vails.

O H3C O F CH3 H3C F Si CH N 3 Si + NH F CH3 F O R R O F CH3 CH F H 3 CH3

trimethylsiloxy MTFA MSTFA (1) alcohol (2) alkane (3) N-Methyltrifluoracetamid (4)

Figure 5.6.6-1: Trimethylsilylation of alcohols using MSTFA

For the analysis of the total ion chromatogram (TIC) and the resulting Mass-spectra of the peaks diverse programs were used. Peak identification was performed with Automated Mass Spectral Deconvolution and Identification System (AMDIS) version 2.70 and National Institute of Standards and Technology (NIST) version 2.0. The Mass spectrum match (just matches greater-than 750 were admitted) and retention index (RI) was decisive for allocation of the peaks. For the quantification of TIC or single mass-to-charge ratio area was determinate with MZmine 2 and OrigenPro 9.1.

5.6.7 Elemental Analysis

Approximately 2-3 mg of the air dried sample were mixed with Tungsten oxide and measured with “vario EL” (Elementar Analysensysteme GmbH, Germany) with a thermal conductivity detector (TDC) and data recorded by the Elementar Analysesystem vario EL III (version 4.03) operating software. Oxygen was used for the oxidation and Helium as a transportation gas.

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Gel permeation Chromatography (GPC) 5.6.8

5.6.8 Gel permeation Chromatography (GPC)

An agilent 1200 series pump (Agilent Technologies, Germany) with three Polymer Standard Service (Germany) SDV 5 µm columns (100; 1,000; 10,000), a UV- and a RI-detector was used to determine the molar masses of approximately 2 mg/ml lignin and lignin derivate in THF at room temperature. Polystyrene was used as the calibration substance. The injection volume was 50 µl and the flowrate was 1 ml/min. More detailed information is shown in Figure 6.1-9 and Figure 6.1-10.

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Supporting information 6

6. Supporting information

6.1 Additional Figures

Chain class O CH 3 H3C CH3 HO O n n O 1. long chain 2. long chain carboxylic H3C alcohols acid methyl ester O OH n O O 5. -hydroxy acids H3C CH3 HO OH O O methyl ester n n 3. -dihydroxy 4. -dicarboxylic alkane acids dimethyl ester

Chain constitution

O

H3C OH O 1. unsaturated: e.g. 5:1: -hydroxyoctadec-9-enoic acid methyl ester

O O

H3C O OH Epox: e.g. 5:Epox: 9,10-epoxy-18-hydroxy-octadecanoic acid methyl ester O OH

H3C OH O OH Diol: vicinal (vic) diols: e.g. 5:Diol: 9,10,18-hydroxy octadecanoic acid methyl ester

O OH

H3C OH O O CH3 Metox. methoxy/alcohol: e.g. 5:Metox: 9,18-hydroxy-10-methoxy octadecanoic acid methyl ester

Figure 6.1-1: Expected monomers after the methanolysis

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Additional Figures 6.1

Figure 6.1-2: Expected aromatics

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Additional Figures 6.1

BE

BN

Figure 6.1-3: DSC-thermograms of BN (below) and BE (top)

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Additional Figures 6.1

Figure 6.1-4: Carbon NMR Spectrum of TAI derivatized, depolymerized Suberin monomers (TAI-SM). The observed chemical shifts are listed in Table 3.2.3-2.

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Additional Figures 6.1

m l k j i g f e d c b a

33 32 21-31 17-19

16 15

14

13

9

Figure 6.1-5:1H-13C HSQC of TAI-SM, positive correlation signals are plotted in blue, negative in green. The observed cross sections are listed in Table 3.2.3-2.

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Additional Figures 6.1

k j i g f c

A

B

7 2

1

1 13 Figure 6.1-6: H- C HMBC of TAI-SM; the Figure 6.1-7 displays an enlarged section B marked with a square in figure A; The observed cross sections are listed in Table 3.2.3-2.

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Additional Figures 6.1

k j i h g f e b-d a

32 30, 31 29 21-28 19 17

B 14

1 13 Figure 6.1-7: H- C HMBC of TAI-SM; enlarged section B of Figure 6.1-6, The observed cross sections are listed in Table 3.2.3-2.

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Additional Figures 6.1

Figure 6.1-8: FTIR Spectra of EL (red), CL in DMSO (grey), vacuum dried CL and with acetone precipitated CL (green).

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Additional Figures 6.1

Figure 6.1-9: GPC-data of Organosolv lignin

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Additional Figures 6.1

Figure 6.1-10: GPC-data of EL

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Additional Figures 6.1

Figure 6.1-12: DSC thermogram of OL

EL-1

EL-2

EL-3

Figure 6.1-12: DSC EL-1 (red), EL-2 (blue), and EL-3 (green)

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Additional Figures 6.1

Figure 6.1-13: FTIR spectra of the NIPUs in adequate concentration (colored) and in excess of diamines (grey).

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Additional Figures 6.1

NIPU-1

NIPU-2

NIPU-3

Figure 6.1-14: DSC thermogram of NIPU-1 (top), NIPU-2(center) and NIPU-3 (below)

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Additional Tables 6.2

6.2 Additional Tables

Table 6.2-1 contains gravimetric triple determined water content of the pre-dried, fresh fine milled bark samples with appendant standard deviation and the mass of the solvent extractable material related of the dry bark weight.

Sample name water content DCM-soluble MeOH-soluble water- soluble [wt.%] [wt.%] of dry bark

BS 1 41.7 ± 3.5

Fresh BS 2 43.0 ± 2.5

BS 3-IB 27.2 ± 1.3

Frozen BS 3-OB 27.2 ± 3.4

BN "130930" 3.7 ± 0.1 1.90 7.91 3.75

BN "140218" 3.9 ± 0.1 1.42 5.38 3.64 0.81 4.44 3.02 1.42 4.19 4.16

BN "141216" 3.3 ± 0.1 1.03 3.09 2.0 ± 0.4 1.19 4.92 1.20 4.94 0.77 5.13 2.64

BN "150413" 3.2 ± 0.1 1.11 5.75 2.04

Pre dried anddried milled Pre BN "150414" 5.5 ± 0.1 1.49 4.74 3.57

BN "151029" 2.2 ± 0.0 1.61 5.63 3.07 1.36 5.12 2.73 average 3.42 ± 0.12 1.28 5.10 3.18 SD 1.17 ± 0.13 0.33 1.14 0.66

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Additional Tables 6.2

Table 6.2-2: Typical IR transmissions bands for suberin, bark, and bark extracts

풗̃ [cm-1] Bonds vibrations of functional groups

3400 O-H of alcohols[40, 107, 114, 162, 163, 181, 182, 195] or water

[32, 37, 40, 163, 181, 195, 230, 243] [157] 2958-2950 C-H asymmetric vibration aliphatic CH3 (C-O stretching)

[37, 40, 163, 181, 195, 202, 230, 243] 2917-2916 C-H asymmetric vibration of aliphatic CH2

[32, 37, 40, 163, 181, 202, 230, 243] 2850-2840 C-H symmetric vibration of aliphatic CH2 1736-1733 C=O carbonyl vibration of ester group or carboxylic acids[40, 162, 168, 194, 195, 244]

1715 C=O stretch vibration of non-conjugated carbonyl[40, 157, 168, 169, 243, 245]

1637

1619-1598 C=C stretch vibration of aromatic rings[32, 40, 95, 157, 163, 168, 169, 181, 182, 243–245] (C=O carbonyl vibration)[40] 1513-1512 C-H vibrations of aromatic rings[157, 163, 168, 169, 181, 194, 244, 245], C=C of alkene in lignin[107]

[157, 168, 169, 195, 244, 245] 1461 asymmetric deformations vibration of aliphatic CH2 and HH3

[95] [163] 1436-1422 C-H bending of CH2 , vibration of methoxy groups , aromatic ring C-H in plan deformation stretch[182, 202] 1379-1364 phenolic OH stretch vibration[163, 181] 1321-1318 phenolic OH stretch vibration[157, 168, 169, 244, 245] 1262-1180 C=O stretch vibration of carbonyls[40, 157], stretch vibration of C-O-C[40, 181, 194] saccaride or epoxy groups, C-O and C-C stretch vibration of aromatic rings [40, 157, 168, 169, 244, 245], oxirane ring (epoxy group)[181, 231] 1171-1162 C=O stretch vibration of (lignin) ester groups[168, 169, 244] 1139-1113 C-O-C asymmetric vibrations of saccharide[37, 40, 231], aromatic C-H stretch vabration[157, 168, 169, 244, 245], C-O vibration of secondary alcohols[163, 181, 195] 1093 1075 asymmetric C-O-C vibration[40] 1023-1032 O-H vibration of primary alcohol, symmetric C-O-C vibration[40] and C-H, C-O deformation in lignin and saccharide like cellulose[32] 894-890 C-H deformation of saccharide 820-817 Vibration typical for saccharide, aromatic C-H vibration[168]

796 C-O-C of glucosides bond in cellulose and C-H of CH3 and CH2 781

725 Deformations vibration of aliphatic CH2 718 C-O ester, C=C alkene chain[95, 129]

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Additional Tables 6.2

Table 6.2-3: Summarized determination values produced by EnBW

wet bark BN (dry) BR method ash content [wt.%] 8.3 7.4 16.0 ASTM D-5142 04 colorific value [kJ/kg] 10279 16592 14548 DIN 51900 volatile material (900°C) [wt.%] 75.4 74.4 71.5 DIN EN 15148 K [wt.%] 0.3 0.3 0.17 EN ISO 11885 N [wt.%] 0.6 0.6 0.53 E DIN 51732 Ca [wt.%] 3.0 2.5 2.54 EN ISO 11885 Na [wt.%] trace trace 2.99 EN ISO 11885 Phenol index in the eluent [mg/l] 92.3 89.6 28.1 DIN 38409-H16-1

Table 6.2-4: Observed band vibration of OL, EL, CL and NIPU 1-3 by FTIR.

풗̃ [cm-1] Bonds vibrations of functional groups 3400 O-H of alcohols[40, 107, 114, 162, 163, 181, 182, 195] or urethanes [25, 27, 229, 230]N-H vibration of amines [40, 228] 3267 (C-O stretching), NH amide[228, 229]

(Epoxy ring)

[32, 37, 40, 163, 181, 195, 202, 230, 3000-2840 C-H asymmetric and symmetric vibration aliphatic CH2 and CH3

243] [157] (C-O stretching) 2255 it is assumed that the vibration belongs to a cyclic carbonate 2127 it is assumed that the vibration belongs to a cyclic carbonate 1793 C=O vibration of cyclic carbonates[228] 1714 C=O stretch vibration of non-conjugated carbonyl[40, 157, 168, 169, 243–245], C=O vibration of urethanes[24, 27, 230] 1670 C=O vibration of conjugated carbonyl [157, 168, 169, 184, 245], of urethane[27, 228, 230], of amide and amine[228] 1600-1593 C=C stretch vibration of (lignin) aromatic rings[32, 40, 95, 157, 163, 168, 169, 181, 182, 243–245] (C=O carbonyl vibration)[40], N-H deformations vibration of amines[228] 1530-1545 N-H deformation urethane and amide[25, 27, 40, 228–230] 1513-1507 C-H vibrations of aromatic rings[157, 163, 168, 169, 181, 194, 244, 245], C=C of alkene in lignin[107]

[40, 157, 168, 169, 181, 1461 C-H asymmetric deformations vibration of methyl (CH3), methylene (CH2)

195, 244, 245] [163] and methoxy (OCH3)

[95] [163] 1425 C-H bending of CH2 , vibration of methoxy groups , aromatic ring C-H in plan deformation stretch[182, 202]

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Additional Tables 6.2

풗̃ [cm-1] Bonds vibrations of functional groups 1330-1325 C-O stretch vibration of syringyl ring[157, 168, 169, 243–245], N-H and C-N deformations vibration of amide[40] 1269-1259 C-O stretch vibration of guaiacyl ring[157, 168, 169, 243–245] 1254 it is assumed that the vibration belongs to oxirane[181, 231] 1223-1213 C-C, C-O, C=O stretch vibration of guaiacyl ring[40, 157, 168, 169, 181, 194, 231, 245] 1150 C=O stretch vibration in ester of HGS lignin[168, 169, 244], or C-O-C stretch vibration[40] 1145-1140 C-H stretch vibration of aromatic guaiacyl ring[40, 168, 169, 244, 245], C-O vibration of secondary alcohols[181] 1125-1113 aromatic C-H deformation vibration of syringyl and guaiacyl ring[169, 244], or aliphatic chlorides[232] 1092 alkyl aryl ether vibration [37] 1031 C-O stretch vibration of primary alcohols[157, 168, 169, 181, 195, 244, 245] 1000 it is assumed that the vibration belongs to a cyclic carbonate 960 vibration of oxirane[202] or amide[40] 925 oxirane ring vibration[24], aromatic ring[168, 169, 244, 245] or N-H deformation vibration of amide[40] 910 oxirane ring vibration,[163, 180–184, 202] aromatic ring 880 oxirane ring vibration, C-H vibration of aromatic ring[40, 232] 851 oxirane ring vibration [25, 163, 180, 181], C-H vibration belonging to guaiacyl units[12, 37, 157, 232] 830 C-H bending of S units or oxirane ring vibration[12, 24, 25], C-H vibration belonging to syringyl units[157] 757 alkyl ether of epoxide[12, 163] 732 it is assumed that the vibration belongs to aliphatic chlorides[232] 695 it is assumed that the vibration belongs to aliphatic chlorides[232]

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Additional Tables 6.2

Table 6.2-5: Solvents used for the calculation of the EL HSP with their specific HSP such as applied in the HSPiP program. A score of one signal indicates complete solubility of the sample in the specific solvent, a score of zero indicates insolubility and partial solubility is indicated by 0+.

EL- CL- solvent δD δP δH Score Score Acetic Anhydride 16.00 11.70 10.20 1 0 Acetone 15.50 10.40 7.00 0+ 0 Acetonitrile 15.30 18.00 6.10 0 0 Acetophenone 18.80 9.00 4.00 1 0 Anisole 17.80 4.40 6.90 0 0 Benzaldehyde 19.40 7.40 5.30 1 0 1-Butanol 16.00 5.70 15.80 0 0 Butyl Lactate 15.80 6.50 10.20 1 0 Chlorobenzene 19.00 4.30 2.00 0 0 Chloroform 17.80 3.10 5.70 0+ 0+ Cyclohexanol 17.40 4.10 13.50 0 0 Cyclohexanone 17.80 8.40 5.10 1 0 Diacetone Alcohol 15.80 8.20 10.80 1 0 o-Dichlorobenzene 19.20 6.30 3.30 0 0 Diethyl Ether 14.50 2.90 4.60 0 0 Diethylene Glycol Monobutyl Ether 16.00 7.00 10.60 1 0 Dimethyl Sulfoxide (DMSO) 18.40 16.40 10.20 1 0 1, 4-Dioxane 17.50 1.80 9.00 1 0 Ethanol 15.80 8.80 19.40 0 0 Ethyl Acetate 15.80 5.30 7.20 0+ 0 Ethyl Benzene 17.80 0.60 1.40 0 0 2-Ethyl-1-Butanol 15.80 4.30 13.50 0 0 Furfuryl Alcohol 17.40 7.60 15.10 1 0 Hexane 14.90 0.00 0.00 0 0 Methanol 14.70 12.30 22.30 0+ 0 N, N-Dimethyl Acetamide 16.80 11.50 9.40 1 0 2-Propanol 15.80 6.10 16.40 0 0 Propylene Carbonate 20.00 18.00 4.10 1 0+ Propylene Glycol 16.80 10.40 21.30 0 0 Tetrahydrofuran (THF) 16.80 5.70 8.00 1 0 Water 15.50 16.00 42.30 0 0 p-Xylene 17.80 1.00 3.10 0 0

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Table 6.2-6: GC-MS peak identification via NIST-MS database analyses of the DCM-extract

GC-MS analyses of DCM-extracts UPAC-name [min] ± ± [%] ± [%] No. propane-1, 3-diol 5.350 860 42 1082.3 0.3 0.143 0.091 4 2-hydroxypropanoic acid 5.650 889 99 1093.7 9.8 0.296 0.168 4 hexanoic acid 5.758 918 38 1098.1 6.6 1.365 1.469 7 1-phenylethan-1-one 5.795 843 1099.9 0.001 0.001 1 2-hydroxyacetic acid 5.891 859 45 1103.3 7.8 0.069 0.054 3 heptanoic acid 7.780 893 24 1125.0 114.2 0.099 0.088 5 benzoic acid 9.411 897 9 1242.6 1.4 0.028 0.016 4 octanoic acid 9.962 872 34 1264.2 7.8 0.153 0.079 7 propane-1, 2, 3-triol (glycerol) 10.503 884 19 1288.0 13.2 0.175 0.059 7 oct-2-enoic acid 11.243 859 34 1317.7 14.2 0.337 0.383 6 deca-2, 4-dienal 11.118 873 1309.9 0.295 0.511 1 3-butylcyclohex-1-en-1-ol 11.439 740 17 1322.6 7.5 0.352 0.291 5 (2E)-but-2-enedioic acid (fumaric acid) 12.330 715 1357.8 0.059 0.102 1 Nonanoic acid 12.366 877 40 1362.7 16.0 0.137 0.080 5 2-hydroxyheptanoic acid 12.672 809 35 1368.6 9.1 0.090 0.069 5 decanoic acid 14.675 810 38 1456.9 16.8 0.073 0.054 6 3-hydroxyoctanoic acid 15.245 736 36 1477.5 10.4 0.177 0.129 4 4(3)-hydroxy-3(4)-methoxybenzaldehyde 16.453 906 44 1531.1 13.1 0.536 0.359 7 3(4)-hydroxy-4(3)-methoxybenzaldehyde 16.851 940 1 1554.4 13.6 1.910 2.571 2 3-phenylprop-2-enoic acid (cinnamic acid) 16.859 753 76 1551.8 20.7 0.023 0.035 3 4-(2-hydroxyethyl)phenol 17.501 872 17 1582.4 29.3 0.037 0.052 2 5-(hydroxymethyl)-2-methoxyphenol 19.060 865 61 1645.9 21.0 0.215 0.160 5 Dodecanoic acid 19.336 781 11 1661.6 23.4 0.018 0.015 3 2, 6-dimethoxybenzene-1, 4-diol 19.953 765 42 1685.9 18.4 0.876 0.495 7 methyl 4-hydroxy-3-methoxybenzoate 20.490 829 1729.6 0.022 0.038 1 octanedioic acid (suberic acid) 20.134 848 7 1691.1 0.6 0.197 0.174 2 4-hydroxy-3, 5-dimethoxybenzaldehyde (syringic aldehyde) 20.303 896 26 1702.0 18.1 1.059 0.759 7 5-(2-hydroxyethyl)-2-methoxyphenol (homovanillic alcohol) 20.545 797 19 1710.0 14.4 0.479 0.420 3 4(3)-hydroxy-3(4)-methoxybenzoic acid 21.685 915 24 1759.6 10.2 0.615 0.313 7 nonanedioic acid (azelaic acid) 22.361 905 23 1797.6 19.8 0.796 0.389 7 4-(3-hydroxypropyl)-2-methoxyphenol 22.873 728 40 1824.0 23.9 0.123 0.024 5

GC-MS analyses of DCM-extracts UPAC-name [min] ± ± [%] ± [%] No. 3, 4-dihydroxybenzoic acid 22.762 770 1813.5 0.000 0.000 1 2, 5-bis(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (fructofuranose) 22.978 788 41 1808.1 22.6 0.012 0.012 2 5-allyl-3-methoxybenzene-1, 2-diol 23.189 699 9 1839.2 21.2 8.327 2.762 6 3-(4-hydroxy-3-methoxyphenyl)acrylaldehyde 23.240 799 53 1841.1 19.2 2.386 0.965 7 tetradecanoic acid 23.109 796 1831.1 0.093 0.132 1 2-hydroxy-2-(4-hydroxy-3-methoxyphenyl)acetic acid 23.577 853 24 1860.9 25.1 0.072 0.041 4 4-hydroxy-3, 5-dimethoxybenzoic acid 24.471 863 41 1903.5 20.6 0.196 0.075 7 2, 3, 4, 5 ,6-pentahydroxyhexanal (mannose) 24.629 830 1908.1 0.013 0.023 1 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (glucopyranose) 24.633 777 123 1908.2 0.2 0.064 0.074 3 2, 3, 4, 5, 6-pentahydroxyhexanal (glucose) 24.646 831 1908.9 0.013 0.023 1 3-(3(4)-hydroxy-4(3)-methoxyphenyl)acrylic acid (cinnamic acid) 24.701 703 1911.7 0.025 0.035 1 2-hydroxy-2-(4-hydroxy-3-methoxyphenyl)acetic acid 25.237 824 47 1947.0 25.8 0.026 0.025 3 2-ethoxy-6-(hydroxymethyl)tetrahydro-2H-pyran-3, 4, 5-triol 25.721 826 15 1967.6 22.3 0.199 0.134 6 3-(4-hydroxy-3-methoxyphenyl)propane-1, 2-diol 26.006 791 72 1983.1 24.4 0.092 0.092 5 3-(4-hydroxy-3, 5-dimethoxyphenyl)acrylaldehyde 26.477 845 40 2009.2 26.2 0.212 0.177 4 9-hexadecenoic acid (palmitelaidic acid) 26.731 790 58 2021.3 26.2 0.112 0.065 5 hexadecanoic acid 27.136 916 71 2042.2 21.4 6.789 1.665 7 Heptadecanoic acid 28.852 863 36 2137.3 23.0 0.374 0.077 7 3-(3, 4-dihydroxyphenyl)acrylic acid 29.030 863 49 2156.9 41.4 0.003 0.006 2 octadecan-1-ol 28.904 812 2136.2 0.021 0.036 1 9, 12-octadecadienoic acid 30.04 917 13.9 2203.7 24.0 2.737 2.029 7 octadec-9-enoic acid (oleic acid) 30.150 886 58 2210.0 23.7 3.384 2.664 7 octadec-11-enoic acid 30.240 882 29 2215.3 24.5 0.930 0.511 7 octadecanoic acid 30.571 916 17 2235.1 25.0 0.903 0.291 7 nonadecanoic acid 32.598 852 0 2367.5 20.6 0.060 0.070 2 icosan-1-ol 32.786 746 2394.0 0.043 0.075 1 eicosanoic acid 33.590 851 52 2433.8 29.0 0.389 0.234 7 hexadecandioic acid 34.453 729 45 2477.6 34.6 0.128 0.076 4 heneicosanoic acid 35.316 861 77 2530.5 27.5 0.430 0.161 7 1-docosanol 35.531 935 13 2545.2 27.8 4.018 2.266 6 hexadecanoic acid, 2, 3-bis(oxy)propyl ester 36.243 856 29 2591.3 28.8 0.511 0.324 7 docosanoic acid 36.780 912 28 2433.8 29.0 2.027 1.125 7 tricosan-1-ol 36.766 933 39 2625.8 6.7 0.636 0.262 3

GC-MS analyses of DCM-extracts UPAC-name [min] ± ± [%] ± [%] No. heptadecanoic acid, glycerine-monoester 37.481 787 2673.8 0.028 0.040 1 Alizarine, bis-TMS 38.007 819 0 2727.6 22.3 0.024 0.037 2 tetracosan-1-ol 38.349 923 40 2740.7 27.8 2.201 1.723 7 disaccharide 39.235 652 2 2815.8 21.4 0.139 0.164 2 disaccharide 39.313 817 0 2821.5 21.6 0.123 0.204 2 tetracosanoic acid 39.635 755 28 2834.7 29.9 2.454 0.928 6 tricosan-1-ol 40.700 789 2906.4 0.253 0.234 1 pentacosanoic acid 41.302 816 1 2967.6 25.0 0.051 0.049 2 hexacosan-1-ol 41.211 905 12 2955.2 38.3 0.196 0.208 3 disaccharide 41.805 677 1 3004.5 25.9 0.074 0.118 2 22-hydroxydocosanoic acid 41.649 823 28 2984.1 38.1 0.130 0.152 4 tetracosan-1-ol (probably longer) 41.690 792 28 2977.9 29.5 0.193 0.160 2 hexacosanoic acid 42.405 799 75 3022.7 29.0 0.189 0.089 3 tetracosanoic acid (probably longer) 43.106 813 0.252 0.223 1 (8S, 9S, 10R, 13R, 14S, 17R)-17-((2R, 5S)-5, 6-dimethylheptan-2-yl)-10, 13- dimethyl-2, 3, 4, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17-tetradecahydro-1H- 45.079 828 12 3200.1 0.253 0.223 2 cyclopenta[a]phenanthren-3-ol (campesterol) (3S, 8S, 9S, 10R, 13R, 14S, 17R)-17-((2R, 5S)-5-ethyl-6-methylheptan-2-yl)- 10, 13-dimethyl-2, 3, 4, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17-tetradecahydro- 46.053 933 25 3288.0 17.5 1.947 0.602 7 1H-cyclopenta[a]phenanthren-3-ol (β-sitosterol) (3S, 4aR, 6aR, 6bS, 8aR, 12aR, 14aR, 14bR)-4, 4, 6a, 6b, 8a, 11, 11, 14b- octamethyl-1, 20, 3, 4, 4a, 5, 6, 6a, 6b, 7, 8, 8a, 9, 10, 11, 12, 12a, 14, 14a, 14b- 46.571 846 29 3330.5 0.0 0.282 0.039 2 icosahydropicen-3-ol (β-Amyrin) (8R, 9S, 10S, 13R, 14S, 17R)-17-((2R, 5R)-5-ethyl-6-methylheptan-2-yl)- 10, 13-dimethylhexadecahydro-1H-cyclopenta[a]phenanthren-3-ol 46.598 806 75 3332.5 0.0 0.282 0.039 2 (stigmastanol) (3S, 4aR, 6aR, 6bS, 8aR, 11R, 12S, 12aR, 14aR, 14bR)- 4, 4, 6a, 6b, 8a, 11, 12, 14b-octamethyl-1, 2, 3, 4, 4a, 5, 6, 6a, 47.586 619 82 3323.2 0.8 1.269 1.020 3 6b, 7, 8, 8a, 9, 10, 11, 12, 12a, 14, 14a, 14b-icosahydropicen-3-ol (α-Amyrin) Total identification rea 49.996 8.172

Table 6.2-7: GC-MS peak identification via NIST-MS database analyses of the methanol-extract

GC-MS analyses of MeOH-extracts UPAC-name [min] ± ± [%] ± [%] No. propane-1, 2, 3-triol (glycerol) 10.382 887 8 1280.9 0.3 0.661 0.757 3 3(4)-hydroxy-4(3)-methoxybenzaldehyde 16.329 918 5 1523.1 0.1 0.037 0.007 3 2, 3, 4, 5-tetrahydroxypentanal (Arabinose, Xylose) 18.757 865 12 1627.6 0.2 0.026 0.003 3 2, 3, 4, 5-tetrahydroxypentanoic acid (Xylonic acid or Arabinonic acid) 18.903 789 18 1634.3 0.1 0.019 0.003 3 2, 3, 4, 5, 6-pentahydroxyhexanal (Manose) 19.042 848 30 1640.7 0.1 0.016 0.002 3 2, 3, 4, 5-tetrahydroxypentanal (Lyxose, Ribose, Xylose) 19.432 898 14 1658.7 0.2 0.031 0.003 3 2, 3, 4, 5-tetrahydroxypentanal (Xylose, Arabinose) 19.496 847 15 1661.7 0.1 0.012 0.007 3 2, 6-dimethoxybenzene-1, 4-diol 19.786 762 11 1675.0 0.1 0.015 0.021 2 4-hydroxy-3, 5-dimethoxybenzaldehyde (syringic aldehyde) 20.129 879 11 1690.9 0.2 0.040 0.004 3 5-(2-hydroxyethyl)-2-methoxyphenol (homovanillic alcohol) 20.376 612 1702.3 0.026 1 tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (Xylopyranose or Lyxopyranose) 20.729 899 7 1718.5 0.2 0.117 0.006 3 pentane-1, 2, 3, 4, 5-pentaol (arabitol) 21.154 909 1738.1 0.025 0.043 1 Ribitol, Adonitol 21.242 860 1742.2 0.029 0.051 1 4(3)-hydroxy-3(4)-methoxybenzoic acid 21.530 905 13 1755.4 0.1 0.070 0.010 3 tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (Xylopyranose or Lyxopyranose) 21.907 842 28 1772.8 0.1 0.115 0.002 3 nonanedioic acid (azelaic acid) 22.213 739 1786.9 0.004 0.007 1 Glucofuranoside, Allofuranose 22.783 774 25 1814.5 0.4 0.419 0.381 3 2, 5-bis(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (fructofuranose) 23.071 933 8 1829.1 1.3 0.284 0.279 2 2, 5-bis(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (fructofuranose) 23.251 927 14 1838.1 0.8 11.646 10.171 3 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (fructopyranose, 23.281 953 1 1839.8 0.2 11.194 6.063 2 psicopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (psicopyranose, 23.689 812 1 1860.4 0.0 0.750 0.751 2 fructopyranose, tagatopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (mannopyranose, 24.779 917 28 1915.7 1.4 14.091 12.102 3 glucopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (talopyranose, 24.891 916 10 1921.3 3.3 8.289 13.289 2 mannopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (talopyranose, 24.943 902 12 1924.0 1.3 0.894 0.805 2 mannopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (glucopyranose, 26.587 907 20 2007.9 3.0 8.262 13.933 3 mannopyranose, talopyranose)

GC-MS analyses of MeOH-extracts UPAC-name [min] ± ± [%] ± [%] No. 2, 3, 4, 5, 6-pentahydroxyhexanal (talose) 26.634 9047 2010.5 8.224 14.245 1 cyclohexane-1, 2, 3, 4, 5, 6-hexaol (myo-Inositol) 28.315 869 28 2103.6 0.4 0.067 0.030 3 glyceryl-pyranosside 32.429 843 18 2342.7 0.1 0.047 0.003 3 disaccharide 35.861 857 9 2561.1 0.3 0.093 0.026 3 disaccharide 36.813 795 11 2625.4 0.4 0.055 0.019 3 disaccharide (sucrose) 37.616 939 4 2683.5 0.9 1.843 0.833 3 disaccharide (cellobiose, lactose, mannobiose, maltose) 38.178 849 28 2724.1 1.0 0.042 0.013 3 disaccharide (cellobiose, maltose) 38.795 838 52 2768.7 0.3 0.255 0.133 3 disaccharide (mannobiose) 39.554 834 93 2823.5 0.3 0.127 0.021 3 catechine 40.643 808 4 2902.2 0.7 0.728 0.015 3 Total identification area 64.553 12.963

Table 6.2-8: GC-MS peak identification via NIST-MS database analyses of the water-extract

GC-MS analyses of water-extracts UPAC-name [min] [%] No. propane-1, 3-diol 5.369 868 25 1083.1 0.7 0.097 0.022 6 2-hydroxypropanoic acid 5.524 900 18 1089.2 0.4 0.092 0.057 5 hexanoic acid 5.664 754 1094.7 0.002 0.005 1 2-hydroxyacetic acid 5.791 906 20 1099.7 0.5 0.120 0.096 6 benzoic acid 9.400 815 15 1242.1 0.1 0.004 0.007 2 propane-1, 2, 3-triol (glycerol) 10.376 877 12 1280.7 0.3 0.586 0.702 6 ethane-1, 2-dicarboxylic acid 11.151 822 42 1311.2 0.2 0.137 0.217 6 2, 3-dihydroxypropanoic acid 11.764 871 8 1335.1 0.8 0.141 0.042 6 2-hydroxybutanedioic acid (malic acid) 15.620 892 11 1493.3 0.3 0.278 0.137 6 butane-1, 2, 3, 4-tetraol (threitol) 16.259 873 27 1520.2 0.2 0.106 0.134 4 4(3)-hydroxy-3(4)-methoxybenzaldehyde 16.246 762 1523.8 0.003 0.007 1 2, 3, 4-trihydroxybutanoic acid (threonic acid) 17.090 850 32 1555.1 0.5 0.086 0.043 6 2,3,4-trihydroxybutyric acid (erythronic acid) 17.519 871 15 1573.1 0.1 0.129 0.032 6 2, 3, 4, 5-tetrahydroxypentanal (Lyxose) 18.751 805 32 1627.3 0.0 0.009 0.015 2 2, 3, 4, 5-tetrahydroxypentanoic acid (Xylonic acid or Arabinonic acid) 18.898 748 27 1634.1 0.3 0.015 0.019 3

GC-MS analyses of water-extracts UPAC-name [min] [%] No. 2, 3, 4, 5, 6-pentahydroxyhexanal (Manose) 19.038 805 34 1640.5 0.1 0.012 0.011 4 2, 3, 4, 5-tetrahydroxypentanal (Lyxose, Ribose, Xylose) 19.424 831 48 1658.3 0.1 0.030 0.020 5 2, 3, 4, 5-tetrahydroxypentanal (Xylose, Arabinose) 19.486 782 13 1661.2 0.1 0.013 0.019 2 2, 6-dimethoxybenzene-1, 4-diol 19.775 740 44 1674.5 0.2 0.033 0.054 4 4-hydroxy-3, 5-dimethoxybenzaldehyde (syringic aldehyde) 20.139 770 33 1691.3 0.2 0.029 0.016 5 tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (Xylopyranose or Lyxopyranose) 20.717 864 25 1718.0 0.3 0.078 0.054 5 pentane-1, 2, 3, 4, 5-pentaol (arabitol) 21.135 905 24 1737.2 0.2 0.087 0.084 4 Ribitol, Adonitol 21.229 843 16 1741.6 0.2 0.084 0.093 3 4(3)-hydroxy-3(4)-methoxybenzoic acid 21.519 869 21 1754.9 0.1 0.093 0.024 6 tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (Xylopyranose or Lyxopyranose) 21.889 833 50 1772.0 0.2 0.084 0.074 4 2, 3, 4 ,5-tetrahydroxypentanoic acid (ribonic acid) 22.263 810 63 1789.3 6.6 0.140 0.149 6 nonanedioic acid (azelaic acid) 22.225 685 87 1787.6 1.6 0.013 0.020 4 5-(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (ribofuranose) 22.701 786 3 1810.4 0.1 0.048 0.079 2 3, 4-dihydroxybenzoic acid 22.794 883 34 1815.1 3.8 0.828 1.822 5 2, 5-bis(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (fructofuranose) 22.979 897 33 1824.5 3.0 4.828 1.349 6 2, 5-bis(hydroxymethyl)tetrahydrofuran-2, 3, 4-triol (fructofuranose) 23.127 894 59 1832.2 0.9 8.650 7.954 3 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (psicopyranose, 23.154 910 23 1833.3 0.0 1.244 1.650 2 fructopyranose, tagatopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (psicopyranose, 23.617 771 7 1856.8 0.2 0.199 0.175 4 fructopyranose, tagatopyranose) 2, 3, 4, 5, 6-pentahydroxyhexanal (fructose) 23.770 874 12 1864.5 0.3 0.726 0.253 6 2, 3, 4, 5, 6-pentahydroxyhexanal (glucose) 24.017 820 54 1877.1 0.2 0.099 0.110 3 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (mannopyranose, 24.686 922 7 1910.9 1.0 13.710 4.915 6 glucopyranose) 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (glucopyranose) 24.753 899 1914.3 0.586 1.309 1 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (talopyranose, 24.880 880 11 1920.8 0.4 0.345 0.125 6 mannopyranose) hexane-1, 2, 3, 4, 5, 6-hexaol (mannitol, sorbitol) 25.424 836 95 1948.4 0.3 0.179 0.167 4 hexane-1, 2, 3, 4, 5, 6-hexaol (mannitol, sorbitol) 25.568 888 24 1955.6 0.2 0.461 0.357 6 cyclohexane-1, 2, 3, 4, 5, 6-hexaol (myo-Inositol, inositol) 25.857 868 7 1970.2 0.2 0.218 0.058 6 2-(hydroxymethyl)tetrahydro-2H-pyran-2, 3, 4, 5-tetraol (glucopyranose, 26.538 940 4 2005.2 1.1 18.417 10.670 6 mannopyranose, talopyranose) 2, 3, 4, 5, 6-pentahydroxyhexanoic acid (gluconic acid) 26.813 909 8 2020.4 0.2 5.890 11.152 4

GC-MS analyses of water-extracts UPAC-name [min] [%] No. cyclohexane-1, 2, 3, 4, 5, 6-hexaol (myo-Inositol) 28.304 895 6 2102.9 0.1 0.581 0.563 6 octadecanoic acid 30.386 824 9 2219.8 0.6 0.016 0.026 2 glyceryl-pyranosside 32.426 802 17 2342.6 0.3 0.023 0.031 3 glycerol-pyranoside 33.253 809 2392.3 0.019 0.043 1 disaccharide 35.852 796 2560.5 0.027 0.059 1 disaccharide 36.800 755 46 2624.6 0.3 0.095 0.073 6 disaccharide (sucrose) 37.607 939 7 2682.9 1.6 8.780 9.607 6 disaccharide (cellobiose, lactose, mannobiose, maltose) 38.169 831 39 2723.4 0.2 0.056 0.062 3 disaccharide (cellobiose, maltose) 38.785 820 47 2768.0 0.3 0.152 0.086 5 disaccharide (mannobiose) 39.547 842 67 2823.0 0.4 0.255 0.032 6 disaccharide (mannobiose) 39.965 828 34 2853.2 0.2 0.079 0.059 5 disaccharide (mannobiose) 40.230 826 28 2672.3 447.3 0.314 0.284 5 catechine 40.612 802 6 2900.0 0.4 0.614 0.478 6 disaccharide (melibiose) 40.905 841 28 2921.1 0.1 0.041 0.065 2 Total identification area 68.678 8.759

Table 6.2-9: GC-MS peak identification via NIST-MS database analyses of SM

GC-MS analyses of water-extracts UPAC-Name [min] ± ± [%] ± [%] No. 1 benzoic acid 9.394 873 38 1258.3 0.2 0.013 0.003 6 2 propane-1, 2, 3-triol (glycerol) 10.371 858 19 1297.3 0.1 0.089 0.052 6 3 9-oxononanoic acid 13.893 744 11 1439.9 0.1 0.024 0.004 5 4 octanedioic acid 14.203 831 11 1453.0 0.1 0.049 0.033 4 5 4-hydroxybenzoic acid 15.338 840 17 1501.3 1.0 0.027 0.013 6 4-hydroxy-3-methoxybenzaldehyde or 3-hydroxy-4- 6 16.348 934 6 1543.6 0.4 0.471 0.347 6 methoxybenzaldehyde 7 nonanedioic acid 16.600 875 47 1554.3 0.5 0.402 0.240 6 8 4-hydroxy-3-methoxybenzoic acid 18.976 922 8 1659.6 0.2 0.204 0.093 6 9 2, 6-dimethoxybenzene-1, 4-diol 19.808 777 7 1698.0 0.3 0.080 0.054 6

GC-MS analyses of water-extracts UPAC-Name [min] ± ± [%] ± [%] No. 10 4-hydroxy-3, 5-dimethoxybenzaldehyde 20.164 908 4 1714.5 0.6 0.425 0.264 6 11 3,4-dihydroxybenzoic acid 20.606 822 17 1734.9 0.3 0.138 0.094 6 12 4-hydroxy-3-methoxybenzoic acid 21.524 869 23 1777.3 0.2 0.112 0.035 5 13 3-(4-hydroxy-3-methoxyphenyl)acrylic acid 22.719 799 0 1835.7 0.0 0.111 0.000 1 14 4-(Trimethylsilyloxy)-3-methoxycinnamaldehyde 23.039 739 131 1851.9 0.3 0.158 0.057 4 9, 10-dihydroxyoctadecanedioic acid (probably 23.425 698 22 1871.5 0.3 0.173 0.068 6 dihydroxybutadecanedioic acid) 15 4-hydroxy-3, 5-dimethoxybenzoic acid 24.308 804 0 1917.4 0.0 0.080 0.025 2 16 hexadecanoic acid 24.613 892 33 1931.8 0.9 0.908 0.486 6 3-(3-hydroxy-4-methoxyphenyl)acrylic acid or 4-(4-hydroxy-3- 17 25.129 809 30 1958.0 0.2 0.141 0.096 6 methoxyphenyl)acrylic acid 9, 10-dihydroxyoctadecanedioic acid (probably 25.165 701 36 1959.9 0.1 0.162 0.052 6 dihydroxyhexadecanedioic acid) 18 heptadecanoic acid 26.479 814 53 2028.9 0.4 0.108 0.033 6 19 hexadecanoic acid 26.915 890 18 2053.2 0.3 0.291 0.068 6 20 9, 12-octadecadienoic acid 27.731 838 22 2098.5 0.3 0.275 0.081 6 21 9-octadecenoic acid or 11-octadecenoic acid 27.867 889 12 2106.1 0.5 0.451 0.159 6 22 13-octadecenoic acid or 11-octadecenoic acid 27.955 850 29 2111.0 0.6 0.215 0.079 6 23 octadecanoic acid (steric acid) 28.313 868 16 2130.8 0.4 0.146 0.040 6 24 octadecan-1-ol 28.896 819 8 2163.2 0.2 0.049 0.026 3 25 9, 12-octadecadienoic acid 29.830 639 104 2203.1 19.9 0.099 0.026 4 26 9-octadec-9-enoic acid (oleic acid) 29.834 733 0 2216.5 0.0 0.269 0.000 1 27 octadecanoic acid (steric acid) 30.403 825 32 2250.8 0.3 0.220 0.121 5 28 hexadecanedioic acid 30.609 918 6 2263.2 0.6 1.406 0.299 6 29 16-hydroxyhexadecanoic acid 31.103 846 5 2292.9 0.3 0.729 0.070 6 30 eicosanoic acid 31.761 823 27 2332.5 0.3 0.147 0.072 6 icosan-1-ol 32.223 821 0 2360.2 0.0 0.037 0.000 1 18-hydroxyoctadec-9-enoic acid 33.829 706 57 2461.5 0.5 2.075 1.036 4

GC-MS analyses of water-extracts UPAC-Name [min] ± ± [%] ± [%] No. octadecanedioic acid 33.891 735 9 2465.8 0.7 0.060 0.024 5 31 docosanoic acid 34.997 901 12 2538.1 0.8 1.258 0.357 6 32 1-docosanol, trimethylsilyl ether 35.360 899 5 2561.6 0.4 1.461 0.546 6 33 8-(3-(8-hydroxyoctyl)oxiran-2-yl)octanoic acid 36.457 716 22 2635.9 36.5 2.512 1.392 6 eicosanebioic acid 36.936 749 14 2668.9 0.4 0.883 0.579 5 34 tetracosanoic acid 37.976 861 2 2741.2 1.6 1.881 0.676 6 35 tetracosan-1-ol 38.213 891 40 2757.7 0.4 1.139 0.263 6 36 9, 10-dihydroxyoctadecanedioic acid 38.446 863 10 2773.9 0.4 1.079 0.225 6 37 9, 10, 18-trihydroxyoctadecanoic acid 38.724 847 19 2800.4 1.4 44.228 7.935 6 38 22-hydroxydocosanoic acid 40.049 781 11 2891.7 0.7 2.122 0.559 6 39 hexacosanoic acid 40.652 748 40 2936.7 0.4 0.206 0.082 5 40 22-hydroxydocosanoic acid 41.382 781 36 2991.3 0.2 0.098 0.042 6 41 tetracosan-1-ol (probably longer chain) 41.417 797 8 2993.9 0.2 0.120 0.044 2 42 β-sitosterol 45.849 900 17 3324.9 0.6 0.282 0.053 6 Total identification area 66.065

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List of Tables

List of Tables

Table 2.2.1-1: The beech bark composition and ash content refer to dry bark (db). IB: inner bark; OB: outer bark; rA: number refers to relative peak area ...... 9

Table 2.2.1-2: Yield of beech bark extractable materials refers to dry bark and their composition refers to the extract amount...... 10

Table 2.3.2-1: Suberin monomer types fond in beech (Fagus sylvatica) via GC-MS, cf. Figure 6.1-1. The values are given in areas found in total ion chromatogram (TIC)...... 28

Table 2.4.1-1: Comparison between commercial lignin types and organosolv lignin, their economic interest and properties[159] ...... 32

Table 2.4.5-1: OL of beech bark produced and analyzed by Over and Maier[41]...... 39

Table 3.1.1-1: Mean degradation temperatures, onset and end temperature of the maximal degradation step and the mass loss after heating to 900°C of four milled bark samples detected by TGA. SD is the standard deviation. The glass temperature and glass temperature onset was detected by DSC...... 49

Table 3.1.2-1: Summary of the photometric analysis. Total phenols, PVPP non-precipitated phenol content and condensed tannins content in percent by weight of the extract mass for the different solvents...... 54

Table 3.1.2-2: Heavily summarized presentation of identified substances groups found as areas in TIC chromatogram of the solvent extracts GC-MS measurements...... 56

Table 3.2.2-1: Photometric analyzed total, PVPP non precipitate phenol content and condensed tannins content in percent by weight referred to the extract mass of the samples. The calculated weight percent of total and PVPP non precipitate phenols rest on the gallic acid calibration, condensed tannins based on the molar attenuation coefficient for (2R, 3S, 4R)-leucocyanidin [220]...... 62

Table 3.2.3-1: Observed peak shifts using proton NMR and TCSY with the observed kind of splitting and correlations signals by spin-spin coupling (SSC). The monitored shifts are given in the range which was used for the integration and the TAI-SM spectrum signals are labelled alphabetically. Consequentially the most probable kind of protons are described and marked in red in the formula. ... 67

Table 3.2.3-2: Generated NMR data observed peaks from 13C-NMR spectra and correlation peaks detected by HSQC and HMBC. Proton signals are labeled as in Paragraph “Proton NMR” or in Table 3.2.3-1. Carbon signals observed in 13C-NMR are labelled numerically. The non-numbered carbon or not labeled proton signals were not visible in HSQC or HMBC, but were observed in the carbon or proton NMR (Figure 6.1-4; Figure 3.2.3-1; Figure 3.2.3-2)...... 69

Albert-Ludwigs-Universität Freiburg 170

List of Tables

Table 3.2.3-3: Calculated average chain length and percentage portion of functional groups in chains for the TAI derivatized Suberin monomers...... 72

Table 3.2.4-1: Area proportion in percentage of the different substance classes for the six MSTFA-SM measured samples, illustrated in Figure 3.2.4-3 ...... 76

Table 3.2.4-2: Determinate calibration curve slope of specific m/z standards ...... 78

Table 3.2.5-1: Elemental analysis of SM. Carbon, hydrogen, nitrogen and sulfur were determined experimentally, with oxygen being estimated. Values are in percent by weight [wt.%]...... 79

Table 3.2.5-2: Theoretical molecular formula for 18 carbon long chain molecules, which are found in a high amount in GC-MS measurements...... 81

Table 3.3.2-1: Proton NMR (300 MHz, DMSO) chemical shifts observed for OL, EL and CL...... 91

Table 3.3.2-2: Calculated EI for all EL batches using an IS. EL-1, EL-2 and EL-3 were measured without an additional purification step. EL-2 and EL-3 were purified and measured afterwards, EL-4 directly after the reaction...... 92

Table 3.3.2-3: 31P-NMR samples spectra of OL and two EL batches. The area was calculated relative to the internal standard, cyclohexanol. The EL sample was measured twice, from which the sample mean and covariance was calculated. Additionally, the increase or decrease in signals of EL and OL were calculated and compared...... 94

Table 3.3.3-1: Degradation temperature with the mass loss of OL, EL and CL by TGA and glass transition temperatures of OL and EL samples (EL-1, EL-2, EL-3) by DSC...... 95

Table 3.3.4-1: SEC data of OL and EL in THF detected by UV- and RI- detector...... 97

Table 3.3.5-1: Generic Algorithm used to calculate dispersion (D), polar (P), hydrogen bonding (H) and total cohesion Hansen solubility parameter δ. The calculated radius of the green sphere (R) and fit quality (Fit) of OL and EL...... 98

Table 3.4.1-1: Determination of CI by using 1, 4-butanediamine (NIPU-2); and 1, 6- hexanediamine(NIPU-3); and 1, 12-dodecan diamine (NIPU-1) as curing agent of CL...... 103

Table 3.4.3-1: Degradations temperatures Td and the residual mass after heating to 900°C of NIPU 1-3 observed by TGA measurements. The glass temperature Tg was determined by the first heating cycle of the DSC measurements...... 104

Table 5.1-1: Lignin data determined by Fraunhofer CBP and SEC by University of Hamburg...... 109

Table 5.1-2: Alphabetical list of chemical used with source specification ...... 110

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Table 5.3.1-1: Procedure and water- free using conditions for O-glycidylation of lignin to produce lignin epoxide (EL)...... 117

Table 5.3.2-1: Using conditions for addition of carbon dioxide...... 118

Table 5.4-1: Anhydrous O-glycidylation conditions for LM ...... 120

Table 5.5-1: Reaction condition to produce PU-1, PU-2 and PU-3 ...... 122

Table 5.6.3-1: NMR measurement conditions ...... 125

Table 6.2-1 contains gravimetric triple determined water content of the pre-dried, fresh fine milled bark samples with appendant standard deviation and the mass of the solvent extractable material related of the dry bark weight...... 143

Table 6.2-2: Typical IR transmissions bands for suberin, bark, and bark extracts ...... 144

Table 6.2-3: Summarized determination values produced by EnBW ...... 145

Table 6.2-4: Observed band vibration of OL, EL, CL and NIPU 1-3 by FTIR...... 145

Table 6.2-5: Solvents used for the calculation of the EL HSP with their specific HSP such as applied in the HSPiP program. A score of one signal indicates complete solubility of the sample in the specific solvent, a score of zero indicates insolubility and partial solubility is indicated by 0+...... 147

Table 6.2-6: GC-MS peak identification via NIST-MS database analyses of the DCM-extract ...... 148

Table 6.2-7: GC-MS peak identification via NIST-MS database analyses of the methanol-extract .... 151

Table 6.2-8: GC-MS peak identification via NIST-MS database analyses of the water-extract ...... 152

Table 6.2-9: GC-MS peak identification via NIST-MS database analyses of SM ...... 154

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List of Figures

List of Figures

Figure 1.3-1: Diagram of proposed strategy ...... 4

Figure 2.1.1-1: Relative presence probability of European beech [44] ...... 7

Figure 2.1.1-1: Scheme of bark with the various tissues [70]...... 7

Figure 2.2.1-1: Structure of the cambium zone (CZ), early and late phloem (EPh and LPh) of beech bark observed via light microscopy [59]...... 11

Figure 2.2.1-2: Structure of beech bark observed via light microscopy [59]...... 11

Figure 2.2.1-3: Structure of beech bark periderm [59]...... 12

Figure 2.3-1: Isolated periderm of potato tuber fixed with OsO4. [73] ...... 13

Figure 2.3.1-1: Proposed suberin organization in cork periderm. [95] ...... 14

Figure 2.3.1-2: 1980 proposed aliphatic-aromoatic polymer structure of suberin [72, 94, 106]...... 15

Figure 2.3.1-3: Suggested suberin structure of potato periderm with C: carbohydrate, P: phenolic; S, suberin (phenolic or aliphatic)[91]...... 17

Figure 2.3.2-1: Famous suberin depolymerization methods and their products ...... 19

Figure 2.3.3-1: Cell model with the tree lamellea proposed by von Höhnel 1877 with m: meddle lamella, s: suberin lamella, c: cellulose lamella (tube) [115] ...... 24

Figure 2.3.3-2: Schematic illustration of suberized cell wall: medial lamella (M); primary wall (P); secondary wall (S); suberin (SU); wax films (W); tertiary wall (T); pores (PO) [140] ...... 25

Figure 2.4-1: Determination of most common inter-unit linkages in lignin macromolecules: R1 and R2 are protons or methoxy groups depending on the type of monolignol hydrogen or methoxy group ...... 29

Figure 2.4.1-1: Purity as a function of kappa (top) spruce (left) and pine (right) pulps and resulting paper strength properties of this two species (below) [156]. The numerated lines represent the various solvents used during the organosolv pulping: 1. peroxoic acid, 2. methanol, 3. cresol (spruce)/phenol (pine), 4 propanol (spruce)/ amine (pine), 5. and 6 ethanol, 7 acetic acid, 8. phenol...... 31

Figure 2.4.2-1: First derivative of the TGA thermogram: L1 is Kraft lignin; L2 soda/anthraquinine lignin; and L3 organosolv lignin. [157] ...... 34

Figure 2.4.4-1: Conventional PU synthesis (left) and the isocyanate-free way producing NIPUS (right)[10] ...... 38

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Figure PART I-1: Schematic representation of the processes done in Part I. The blue-labeled steps are analyzed in Section 0 and the green-marked product is examined in Section 3.2. (s: solid, l: liquid) and a photo of the three solvent extracts...... 42

Figure 3.1.1-1: ATR-FTIR spectrum of native milled bark (BN) is black, of Soxhlet extracted bark (BE) is red and of bark residue after methanolysis (BR) is blue. Observed transmittance band are marked with dotted lines and labeled with corresponding wavenumber and listed in Table 6.2-2...... 45

Figure 3.1.1-2: Solid-state 13C-NMR of solvent extracted bark BE (red) and rest bark after methanolysis BR (blue). The red arrows mark the increasing signals, the blue arrows the decreasing signals after the methanolysis of the BE. The signals are normalized at δ=105.2 ppm, which supposed to correspond to polysaccharides (cellulose) carbons...... 47

Figure 3.1.1-3: TGA thermograms of different treated bark sampels are on the left side, and the first derivatives of the thermograms are on the right side. On the top (black) is untreated bark, in the middle (red) is extracted bark and below (blue) remaining bark after the methanolysis...... 48

Figure 3.1.2-1: ATR-FTIR measurements of the DCM-extract (green), methanol-extract (blue) and water (red). Observed transmittance bands are marked with black dotted lines and labeled with corresponding wavenumber, the grey dotted lines are already found in Figure 3.1.1-1 and listed in Table 6.2-2...... 51

Figure 3.1.2-2: Measurements for application of the Folin-Ciocalteu-Method: a-b. Kinetically observation of gallic acid spectra, c. spectra of different substances containing phenol structures, d. linear calibration curve with confidence band and forecast ribbon. The red vertical lines in a. and c. mark the wavelength recommended in the literature [108, 109] and the dashed red line was the one used in this work. The blue vertical line in b. marks the time that elapsed before the samples were measured...... 53

Figure 3.1.2-3: 300 MHz proton NMR of all solvent extract fractions: green is the DCM-extract in

CDCl3; blue the MeOH-extract in d6-DMSO; and red the water-extract in deuterium oxide...... 55

Figure 3.1.2-4: Graphical representation of Table 3.1.2-2...... 56

Figure 3.2.1-1: ATR-FTIR spectrum of suberin monomers ...... 60

Figure 3.2.3-1: Proton NMR (300 MHz) spectra of SM (red) and derivatized TAI-SM (blue) dissolved in

CDCl3...... 63

Figure 3.2.3-2: Enlarged section of the 300 MHz proton NMR spectra presented in Figure 3.2.3-1. The red spectrum belongs to the SM sample and the blue spectrum belongs to TAI-SM...... 64

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Figure 3.2.3-3: TOCSY spectrum of TAI-SM sample in CDCl3, measured in deuterated chloroform. The by Proton NMR applied letters for labeling the signals see Figure 3.2.3-1 or Table 3.2.3-1 are used to describe the two dimension spectrum...... 66

Figure 3.2.3-4: DOSY spectra of SM (red) and TAI-SM (blue) in CDCl3 stacked atop each other. The bars mark the main diffusion area (D-area)...... 74

Figure 3.2.4-1-: Example of total ion chromatogram (TIC) of the GC-MS measurement of MSTFA-SM ...... 75

Figure 3.2.4-2: TIC spectrum versus fragments spectrum, a screen shot of AMDIS ...... 77

Figure 3.2.4-3: Bar charts of analyzed six MSTFA-SM samples bear on the various substance classes refer to the monitored TIC peak area (left side) and refer to identify components (right). Additional a bar of Fagus sylvatica L. roots SM composition is showed (right) found by Spielvogel et al. [77]...... 77

Figure 3.2.4-4: Calibration diagram for different carboxylic acid methyl esters using characteristic m/z ratio 74...... 79

Figure PART II-1: Schematic representation of the processes done in Part II. The derivatization steps (marked red) are investigated in Section 3.3 and the polymer synthesis is examined in Section 3.4 (marked yellow)...... 84

Figure 3.3.1-1: FTIR spectra of OL (black), EL (red), ECH (pink), CL– vacuum dried (blue), CL - dissolved in DMSO (Cyan deshed), and pure DMSO (grey). For OL, EL and CL (vac.) the reference band at 1597 cm-1 was used. All important bands are marked with dotted lines and labeld on the top. New obands are labeld with the coresonding colour of the spectrum observed. Observed transmission bands are marked with dotted lines and labeled Newly identified bands are indicated by the color of the label at the top: black in the OL spectrum; red in the EL spectrum; and blue in CL (vac.) spectrum. ... 87

Figure 3.3.1-2: FTIR spectra of CL-1, CL-2 and a mix of CL-2 and CL-3 (high pressure) and CL-4 (normal pressure)...... 88

Figure 3.3.2-1: Ethylene oxide derivative of EL (R=lignin unit) ...... 89

Figure 3.3.2-2: 300 MHz proton NMR spectra of OL (black), EL (red) and CL (blue) in DMSO. The typical region for epoxide ring is shown in red and for cyclic carbonate in blue...... 90

Figure 3.3.2-3: Ethylene carbonate derivative of CL (R=Lignin unit)...... 92

Figure 3.3.2-4: 121 MHz 31P-NMR of OL-1 (black) and EL-2 (red) ...... 93

Figure 3.3.3-1: TGA termograms of OL (black), EL (red) and CL (blue) left side; first derivatives (right side)...... 95

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Figure 3.3.5-1: 3D-application of OL (left) EL (right) using the dispersion (D), polar (P), and hydrogen bond (H) cohesive Hansen-solubility Paramers. Blue spheres are selvents desolving EL, red cubes mark the solvents non-disolving the sample. The green speres show mark the parameter with are necessery to desolve EL...... 98

Figure 3.4.1-1: FTIR spectra of the educts: CL (black) and the relevant diamine 1,12-dodecan diamine (blue spotted); 1,4-butanediamine (red spotted); and 1,6-hexanediamine (green spotted) and spectra of the resulting PU colored with the right ratio for reacting all groups...... 101

Figure 3.4.1-2: FTIR cyclic carbonate band high monitoring of δ=1793 cm-1 normed by band of δ=1597 cm-1 (60%) after reaction with the three diamines: 1, 12-dodecan diamine (blue); 1,4- butanediamine (red); and 1,6-hexanediamine (green) in different proportional with CL...... 102

Figure 3.4.1-3: Schematically illustration of endcapped lignin NIPU (left) and cross-linked lignin NIPU (right). Substituents of epoxidation are marked red; of carbon ´dioxide are blue; and the diamines green...... 103

Figure 3.4.2-1: Photo of the solubility test of NIPU's in ethanolamine after two weeks at room temperature...... 104

Figure 3.4.3-1: TGA thermogram of NIPU-1 (blue), NIPU-2 (red) and NIPU-3 (green) ...... 105

Figure 5.2.2-1: Reaction mechanism of methanolysis...... 114

Figure 5.2.3-1: Process scheme of the suberin monomer (SM) extraction from bark...... 114

Figure 5.3-1: Process scheme of lignin O-glycidylation and CO2 addition to produce EL and LC. .... 115

Figure 5.3.1-1: Reactions mechanism of epichlorohydrin reaction ...... 116

Figure 5.3.3-1: Reaktions mechanism of carbon dioxide (2) addition to EL (1) catalyzed by TBAB (3) [207]...... 119

Figure 5.5-1: Photo of NIPU film production ...... 122

Figure 5.6.2-1: Photo of sample with phenols (left) and blank feed (right) ...... 123

Figure 5.6.2-2: Photo of diverse tannins free and samples (clear yellowish color) and samples with tannins (reddish color) ...... 124

Figure 5.6.3-1: Reaction of trichloroacetyl isocyanate (TAI) with hydroxyl and carboxyl groups ...... 126

Figure 5.6.3-2: Phosphorylating of hydroxyl functional groups with 2-chloro-4, 4, 5, 5-tetramethyl– 1, 3, 2–dioxaphospholane (TMDP) ...... 127

Figure 5.6.6-1: Trimethylsilylation of alcohols using MSTFA...... 128

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Figure 6.1-1: Expected monomers after the methanolysis ...... 130

Figure 6.1-2: Expected aromatics ...... 131

Figure 6.1-3: DSC-thermograms of BN (below) and BE (top) ...... 132

Figure 6.1-4: Carbon NMR Spectrum of TAI derivatized, depolymerized Suberin monomers (TAI-SM). The observed chemical shifts are listed in Table 3.2.3-2...... 133

Figure 6.1-5:1H-13C HSQC of TAI-SM, positive correlation signals are plotted in blue, negative in green. The observed cross sections are listed in Table 3.2.3-2...... 134

1 13 Figure 6.1-6: H- C HMBC of TAI-SM; the Figure 6.1-7 displays an enlarged section B marked with a square in figure A; The observed cross sections are listed in Table 3.2.3-2...... 135

1 13 Figure 6.1-7: H- C HMBC of TAI-SM; enlarged section B of Figure 6.1-6, The observed cross sections are listed in Table 3.2.3-2...... 136

Figure 6.1-8: FTIR Spectra of EL (red), CL in DMSO (grey), vacuum dried CL and with acetone precipitated CL (green)...... 137

Figure 6.1-9: GPC-data of Organosolv lignin ...... 138

Figure 6.1-10: GPC-data of EL ...... 139

Figure 6.1-12: DSC thermogram of OL ...... 140

Figure 6.1-12: DSC EL-1 (red), EL-2 (blue), and EL-3 (green) ...... 140

Figure 6.1-13: FTIR spectra of the NIPUs in adequate concentration (colored) and in excess of diamines (grey)...... 141

Figure 6.1-14: DSC thermogram of NIPU-1 (top), NIPU-2(center) and NIPU-3 (below) ...... 142

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