Preliminary Study on Enzymatic Degradation of Organosolv Lignin

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Bachelor Thesis Chemistry

Preliminary Study on Enzymatic Degradation

of Organosolv Lignin

by

Laura de Wal

26-06-2017

Student number

10756426

Research institute

Supervisor

Van ’t Hoff Institute for Molecular

Sciences

Prof. dr. ir. Peter Schoenmakers

Research group

Daily supervisor

Analytical Chemistry

Dr. Bert Wouters

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Abstract

Our dependence on fossil fuel to generate energy is causing an increasing concern as the fossil fuel stock is limited and its use is hazardous for our own environment due to its relation with global warming. Since the introduction of new technologies such as bio refineries, biomass-‐ based fuels present a very promising alternative for fossil fuels. Lignin was chosen as the subject of interest due to its use for bio fuels. Additionally it represents one of the richest sources of aromatics, which gives a promising perspective in future bio refinery. The exact structure of lignin is hard to determine, and the depolymerisation of lignin remains a challenge. This research project is focussed on exploring possibilities for enzymatic degradation of Organosolv lignin, and the study of both undegraded and degraded Organosolv lignin through the use of size-‐exclusion (SEC) and reversed-‐phase (RP) liquid chromatography. The size-‐exclusion analysis shows that Organosolv lignin used in this research are relatively small polymers with a majority of a mass of 526,9 Da. In a later stage of this project the goal is to make an online method to combine the degradation with two-‐dimensional LC analysis, for instance SEC×RP. For this reason the effect of tetrahydrofuran (THF), a common mobile phase for size-‐exclusion chromatography, on the enzymatic activity of laccase was assessed. Laccase was used for the enzymatic degradation of Organosolv lignin, both trough in-‐solution degradation and through the use of an immobilized-‐enzyme reactor. For in-‐solution degradation, it was observed that the laccase enzyme is compatible with a high content of THF, with the enzymatic degradation still successful at 70 vol.-‐% THF. At least 15 vol.-‐% of THF was necessary to fully dissolve Organosolv lignin. Unfortunately, these THF contents were shown to be incompatible with the immobilized-‐ enzyme reactor.

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Samenvatting

Tegenwoordig zijn we als mensen bevoorrecht om energie te gebruiken voor alle gewenste doeleinden. Een wereld zonder een grote energieconsumptie is moeilijk voor te stellen. Als de energiebehoefte in dezelfde lijn blijft groeien, wordt dit zorgwekkend omdat de fossiele brandstoffen niet oneindig zijn, maar deze wel een gevaar vormen voor onze eigen omgeving door bijvoorbeeld de opwarming van de aarde. Door de opkomst van nieuwe technologieën voor bio-‐zuiveringen zullen op biomassa gebaseerde brandstoffen een goed alternatief vormen voor fossiele brandstoffen. In dit project is er gekeken naar lignine als biobrandstof vanwege de grote beschikbaarheid en lignine heeft een van de grootste opslag aan aromaten, wat een veelbelovend vooruitzicht is voor de

bio-‐raffinaderij. De exacte structuur van lignine is lastig vast te stellen, daarnaast is de depolymerisatie van lignine ook nog steeds een uitdaging. De structuur van lignine is weergeven in het figuur rechts.

Bij dit project lag de focus op het onderzoeken van mogelijkheden voor enzymatische degradatie van Organosolv lignine, zowel de niet-‐ gedegradeerde als de gedegradeerde vorm van Organosolv lignine werden

onderzocht door middel van size-‐exclusion (SEC) en reversed-‐phase (RP) vloeistofchromatografie. De SEC analyse liet zien dat Organosolv lignine, die gebruikt is in dit project, een relatief klein polymeer is, met een gewichtsverdeling rond de 526,9 Da. In een later stadium van dit onderzoek is het doel om een online methode te combineren met de degradatie met een twee-‐ dimensionele LC analyse, bijvoorbeeld met SECxRP. Vanwege deze reden was het effect van tetrahydrofuran (THF), een veel gebruikte mobiele fase voor SEC, op de enzymatische activiteit onderzocht. Laccase was gebruikt voor de enzymatische degradatie van Organosolv lignine, doormiddel van in-‐solution degradatie en door het gebruik van een immobilized-‐enzyme reactor (IMER). De resultaten van de in-‐solution degradatie lieten zien dat het laccase enzym geschikt is voor een hoge concentratie THF, de enzymatische degradatie was nog steeds geslaagd bij 70 vol.-‐% THF. Tenminste 15 vol.-‐% THF is nodig om Organosolv lignine volledig op te lossen. Helaas zijn deze THF hoeveelheden nog niet geschikt voor de immobilized-‐enzyme reactor.

Lignine wordt gevonden in de celwand van een plantencel, waarbij het dient als een soort lijm voor (hemi)cellulose. Lignine is een willekeurig gevormde structuur door polymerisatie van de drie monolignols.

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1. Introduction

1.1 General introduction

In modern society we have the privilege to use energy for all kinds of purposes in almost limitless amounts. A world without large energy consumption is hard to imagine. As our energy consumption continues to increase, some major concerns arise that cannot be ignored. Our dependence on fossil fuel to generate energy is causing an increasing concern1_{as
the
fossil
fuel}

stock is limited and its use is hazardous for our own environment due to its relation with global warming. Barreto et al. shows that there is an ongoing energy transition from fossil fuels towards renewable and biomass energy.2_{In
Figure
1,
it
is
shown
that
the
concerns
about}

dependence on fossil fuel result in a shift towards renewables, biomass and nuclear energy3_{.
In}

the beginning of the 20th_{century
the
energy
production
was
mostly
based
on
coal.
Fifty
years}

later, around 1950-‐1970, coal became less predominant and oil and gas filled up a large part of energy production (grey line). Forty years later, around 2010, a new interest in renewable energies arises. The predicted developments show that within the next century, around 2100, renewables and biomass-‐based energy will have the majority in energy production (blue and green line).3

Figure 1: The movement of interest for different kinds of energy in time. It started around 1900 with mostly coal. Around 1950-‐1970 this made a shift towards a majority of oil and gas. The prediction is that in de future the demand for renewables, biomass and nuclear energy becomes larger and larger. Figure is based on Barreto et al. 2_{and
courtesy
of
Dr.
Wim}

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Since the introduction of new technologies such as bio refineries, biomass-‐based fuels present a very promising alternative for fossil fuels.1_{Bio
fuels
can
be
divided
into
first
and
second-‐}

generation products. First-‐generation bio fuels are made out of sugar, starch and vegetable oil. Its disadvantages include competition with the food industry for raw materials, and the large energy input needed to make bio fuels yielding a small net reduction of carbon dioxide.4_Second-‐

generation bio fuels can solve the biggest problem of first-‐generation fuels because they don’t compete with the food industry for raw materials, which can grow next to or even together with the raw materials from the food industry. For example, lignocellulosic materials are used which are residues from agriculture forestry.

Lignocellulosic materials consist of cellulose, hemicellulose and lignin, and can be found in the cell wall of plants. The main compound of lignocellulose is cellulose, which is a beta-‐linked chain of glucose molecules. Hemicellulose is composed out of various 5-‐ and 6-‐carbon sugars. Lignin can be seen as the cellular ‘glue’ that holds cellulose and hemicellulose together and the ratio of lignin to (hemi) cellulose varies between different plants. Lignin is composed out of three phenolic compounds; coumaryl alcohol, coniferyl alcohol and sinapyl alcohol.5

The exact structure of lignin is still unknown, but if the biomass-‐based fuels are to be generated from lignin, it is very important to determine its exact structure. This structure can for instance be elucidated using liquid chromatography, which can provide information about the molar mass distribution of lignin and about the connection between the size of degraded polymers and its hydrophobicity or hydrophilicity.

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2. Theory

2.1 Lignin

Lignin is a polymer with a three-‐dimensional structure composed of three main building blocks; coumaryl alcohol, conifernyl alcohol and synapyl alcohol.5_{These
building
blocks
are
also
called}

monolignols. The structure of lignin is given in Figure 2.

Despite the fact that the main building blocks are known, the structure of the lignin polymer has remained difficult to characterize6_{,
in
part
because
the
ratio
of
monolignols,
molar
weight
and}

availability varies by plant source. Chakar et al. have shown that the availability of lignin decreases from softwood (27-‐33%) to hardwoods (18-‐25%) to grasses (17-‐24%). Lignin of softwood mostly consist of coniferyl alcohol (90%), whereas hardwood lignin has an equal ratio of coniferyl alcohol and sinapyl alcohol. Grass lignins mostly contain coniferyl and sinapyl alcohol, but also 10-‐20% of p-‐coumaryl alcohol.7_{The
polymerization
process
is
initiated
by
the}

oxidation of the hydroxyl phenolic groups of the monolignols. This oxidation is catalysed via an enzymatic route, this route is initiated by an electron transfer that yields monolignols with free radicals, which can couple with each other.6_{The
polymerization
of
lignin
is
done
by
different}

kind of couplings. Crestini et al. shows that the first coupling is from head to tail, where the head (the phenyl group) of one monolignols is coupled to the tail of monolignols. This can occur in two ways, the first way is with the oxygen bonded to the head of the monolignol, the second Figure 2: Lignin can be found in the cell wall of a plant, here it functions as the ‘glue’ for (hemi)cellulose. Lignin is a randomly formed structure by polymerisation of the three monolignols, shown in the upper part of this figure. Courtesy of Edward M. Rubin

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option is with the carbon next to a double-‐bonded oxygen on the head of the monolignol (Figure 3 and Figure 4).

The second coupling is from tail to tail, this can also occur in two ways, where one route can cause two 5-‐membered rings (Figure 5 and Figure 6). These couplings give lignin a random three-‐dimensional structure without any repetitive units or bonding patterns. 8

O HO OCH₃ + O OCH₃ Lignin O HO OCH₃ O CH₃ Lignin HO O HO OCH3 + OCH3 Lignin O OCH3 O Lignin OCH3 HO OH OCH3 O HO OCH3 Lignin OCH3 O HO H3CO O OH + OCH3 O HO H3CO O OH OCH3 OH O O H3CO OH OCH3 O HO + OCH3 O HO OCH3 O HO OCH3 O HO O OCH3 O OH OH HO OCH3 HO OH OCH3 HO HO OCH3 OH

Figure 3: Head to tail coupling of monolignols. This coupling can take place because of the two radicals. The radical on the ß-‐place reacts with the radical on the oxygen, this forms a ß-‐ O-‐4 coupling.

Figure 4: Head to tail coupling of monolignols. This coupling also takes place because of the two radicals. Only the location is different. This forms a ß-‐5 coupling.

Figure 5: Tail to tail coupling of monolignols. The OH-‐tails (drawn in blue) form within two steps two 5-‐membered rings, called ß-‐ß coupling

Figure 6: Tail to tail coupling of monolignols. The OH-‐tails (drawn in blue) react and form a coupling without a final 5-‐membered ring. This is called a ß-‐1 coupling.

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Unlike cellulose and hemicellulose, lignin is not fermentable but it can provide energy or undergo thermochemical modifications to produce biofuels.1_{Before
lignin
can
function
as
a}

biomass-‐based fuel, it has to undergo a production process. The first and very essential step in this production process is the pre-‐treatment of lignin. In this step the three main components of the cell wall, cellulose, hemicellulose and lignin are separated from each other.6_{This
is
followed}

by the pulping process. The most used pulping process is the Kraft process, in which lignin is fragmented into smaller water/alkali-‐soluble components. This happens in a high-‐pressure vessel called a digester, with aqueous sodium hydroxide and sodium sulphide, where the anions of those molecules cause the fragmentation.6_{Alternatively,
the
process
of
Organosolv
pulping}

can be used, which is a pulping process based on low-‐boiling point solvents. Organosolv pulping is one of the routes that can produce high-‐quality cellulose bio fuel with a lignin purity of <1% wt. of residual carbohydrate content.9_{The
most
well-‐known
kind
of
Organosolv
is
the
Allcel}

process, which uses ethanol as solvent. With Organosolv lignin the average molecular weight (Mn) is less than 1000, while with Kraft pulping the Mn value is much higher, between 1000-‐ 3000.8_{In
this
research
project,
the
lignin
was
obtained
using
the
Organosolv
pulping
method.}

2.2 Liquid Chromatography

(Ultra) High-‐Pressure Liquid Chromatography (UHPLC) is an analytical tool widely used for the separation of both small and large molecules.10_{When
a
sample
is
injected,
it
is
carried
by
a
fluid}

from one end through the whole column to the other end. The column is usually packed with a particulate layer, such as silica modified with octadecyl alkyl (C18) chains.11 Separation is based

on a difference in distribution of sample molecules between the stationary and mobile phase. This difference in distribution can be described by the partitioning ratio (K), which is the ratio between the concentration of the sample molecules on the stationary phase and in the mobile phase. Different components will elute from the column at different times, depending on the difference in their partitioning ratios. The most important types of liquid chromatography include reversed-‐phase (RP), normal phase (NP), ion-‐exchange (IEX) and size-‐exclusion (SEC). These types are based on different interactions between the stationary phase and the analyte molecules.

2.2.1 Size-‐Exclusion Chromatography: SEC

Size-‐Exclusion Chromatography (SEC) is a type of liquid chromatography in which separation is based on partial exclusion of the analyte through the pores of the stationary phase. Molecules with a smaller hydrodynamic volume will penetrate further into smaller pores than larger molecules, thus small molecules experience a larger accessible pore volume than larger molecules. Therefore, the small molecules have a longer pathway and elute later. All molecules that are very small compared to the pore size will spend the same amount of time in the column

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and therefore will have the same elution time. At the same time all large molecules that are too large to penetrate the pores will also have the same elution time. 10_{Hence,
the
range
of}

separation of polymers is related to the pore size of the SEC column. In SEC, analyte molecules ideally do not interact with the surface of the stationary phase. For this purpose, Tetrahydrofuran (THF) can be used as mobile phase.

2.2.2 Reversed-‐Phase Liquid Chromatography

Despite the fact that it is called reversed-‐phase liquid chromatography (RPLC), this is the more common kind of liquid chromatography compared to normal-‐phase liquid chromatography (NPLC). RPLC is based on a difference in hydrophobicity and uses an hydrophobic stationary phase, e.g., silica-‐based bonded phases such as C18 chains, combined with a gradient of mobile

phase that goes from a hydrophilic mobile phase, e.g. water to a hydrophobic mobile phase, e.g., aqueous acetonitrile or methanol.

2.2.3 Hydrophilic Interactions Liquid Chromatography: HILIC

Hydrophilic interactions liquid chromatography (HILIC) is a valuable alternative for reversed-‐ phase liquid chromatography for the separation of polar and weakly acidic of basic samples.12

HILIC is a type of normal-‐phase chromatography. A HILIC column contains a polar stationary phase, which usually is silica, without any C18 chains. Irgum et al. suggests that the mechanism of

HILIC is building up a water layer on the stationary phase, which has interactions with the polar compounds in the injected sample. This water layer can be built up by adding small amounts of water to the mobile phase. If the ratio of water is too high, then the water layer will break down.13_{For
every
run
with
HILIC
the
water
layer
has
to
be
built
up
at
the
beginning
and
flushed}

out at the end.

2.3 Enzymatic degradation with Laccase

The depolymerisation of lignin is still a challenge. Especially compared to polysaccharides, the degradation of lignin is not as fully developed. Difficulties with the depolymerisation of lignin in catalytic processing are caused by the difference in inter-‐molecular linkages. 14_{In
this
research}

project the degradation of lignin is focused on the use of the enzyme laccase.

Bourbounnais et al. has shown that laccase is a multi-‐copper oxidase complex, which is produced by certain trees and fungi. It is found that fungal laccase plays a role in the biodegradation of lignin. Laccase oxidizes phenols and polyphenols by one electron oxidation with the reduction of O2 into H2O. Laccase from the Coriolus (Trumetes) versicolor, which is used

in this research project, is found most effective for degradation of lignin.15_{The
enzymatic}

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reaction equation of laccase is given. Figure 7B is an example of the linkage monolignols, the bond marked in red is most likely the one broken by laccase.14

The most straightforward method for degradation of lignin with laccase is by in solution-‐ degradation. The solution can be brought to ideal circumstances for the enzyme, which consists of a pH of 4-‐5 and a temperature of 65 ˚C. Typically, these digestions are carried out overnight. Alternatively, an immobilized-‐enzyme reactor (IMER) can be used. Girelli et al. shows that immobilized enzymes are widely used in biocatalysts and for bio-‐specific detection. Enzymes are immobilised in a confined space, resulting in shorter diffusion distances, which leads to shorter digestion times. Additionally, immobilized enzymes are typically more stable than free enzymes to changes in temperature, organic solvents and pH. Enzymes can be immobilised through matrix entrapment, adsorption, or covalent bonding. As a substrate for enzyme immobilisation, the surface of an open microfluidic channel, membranes, particles or porous monolithic structures can be used.17_{Figure
8
shows
an
example
of
covalent
attachment
of
an
enzyme
to
a}

polymer monolithic substrate inside a micro channel of a cyclic-‐olefin-‐copolymer-‐based microfluidic device.

O2 + 4 OH OH laccase O OH 4 + 2 H₂O O O R O O HO A B

Figure 7A: Laccase has an enzymatic activity within lignin, because it oxidizes phenol-‐groups by one electron oxidation with the reduction of O2 into H2O. Figure 7B: The location marked in red

is where the depolymerisation takes place. With the depolymerisation there is an increase in aliphatic/phenolic OH.

Figure 8: Production process of an immobilized enzyme reactor (IMER). In several steps, as shown in the figure, the enzyme is immobilized at the surface of the channel. Courtesy of Dr. Bert Wouters.

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2.4 Aim of this research project

Lignin was chosen as the subject of interest for its use for bio fuels because of its large availability on earth and additionally it represents one of the richest sources of aromatics which gives a promising perspective in the future bio refinery.18_{As
mentioned
before,
the
exact}

structure of lignin is hard to determine, and the depolymerisation of lignin remains a challenge.9

In this research project the focus was placed on the analysis of the structure of Organosolv lignin and the analysis of the degradation products of lignin by the enzyme laccase. Degradation of lignin in combination with two-‐dimensional LC may give information about the subcomponents of the polymer. These subcomponents can contain hydrophilic components, for example OH-‐ groups, which were hidden in the three-‐dimensional structure of the polymer.

To tackle these subjects, this research project was divided in two main parts. The first part of this research project focussed on the optimization of the liquid chromatography separation for Organosolv lignin. First the Organosolv lignin was separated by size-‐exclusion chromatography (SEC) and reversed-‐phase liquid chromatography (RPLC). For SEC, polystyrene standards were used to set up a calibration curve in order to predict a molar mass distribution of Organosolv lignin. For the optimization of SEC, the influence of water present in the sample was studied. This is important because water is almost unavoidable if a multi-‐dimensional analysis is used.

The second part of this research project focuses on connecting the analysis of both lignin and the degradation products with the degradation process itself. For the separation and analysis of the enzymatic degradation products of Organosolv lignin, RPLC and SEC were used. SEC is used because it is expected that the degradation products are smaller than the original lignin structure. Additionally, degradation products of Organosolv lignin likely have a different hydrophobicity because hidden OH-‐groups may become available for interaction with stationary phase, which is why the degradation products were also studied with RPLC. The enzymatic degradation process can be done in two ways: in-‐solution or with immobilized-‐enzymes degradation. Lignin degradation takes place with the laccase enzyme, which cuts the lignin molecules by oxygen reduction into smaller pieces. Degradation with immobilized enzymes is a more advanced way for lignin degradation. The enzyme and microfluidic reactor are not compatible with (a high ratio of) THF, but the lignin will not dissolve in (exclusively) TRIS buffer. Therefore, to combine the analysis with the degradation, different ratios of tetrahydrofuran (THF) and aqueous buffer were tested to find an optimum for both the analysis and degradation. If a compromise can be found the lignin samples can be degraded in the immobilized-‐enzyme reactor (IMER), which can be used in between the first and second dimension of the LC analysis.

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3. Experimental

3.1 Reagents & chemicals

Tetrahydrofuran (THF, stabilized/BHT, AR grade, ≥99.8%), acetonitrile (ACN, LC-‐MS grade), Acetone (99%) and methanol (MeOH, absolute, LC-‐MS grade) were purchased from Biosolve (Valkenswaard, The Netherlands). Aluminium Oxide (Al2O3) butyl methacrylate (BMA, 99%),

ethylene glycol dimethacrylate (EDMA), ethylene glycol diacrylate (EDA, 90%, technical grade), methyl methacrylate (MMA, 99%), 1-‐propanol (>99.8%), 1,4-‐butanediol (ReagentPlus quality, ≥99%). 2,2-‐dimethoxy-‐2-‐phenylacetonephenone (DMPA, ≥99%), 4,4-‐bis(diethylamino)-‐ benzophenone (DEBP, ≥99%), poly (ethylene glycol) methacrylate (PEGMA, number-‐average molar mass 500 g/mole), ethanolamine (≥98%), tert-‐butanol (American Chemical Society reagent, ≥99%), tris(hydroxymethyl)aminomethane hydrochloride (Tris–HCl), trifluoroacetic acid (TFA, ≥99%), Laccase (from Trametes Versicolor), Calcium Chloride (CaCl2) and toluene

(Chromasolve grade) were purchased from Sigma Aldrich (Zwijndrecht, The Netherlands). 2-‐ Vinyl-‐4,4-‐dimethylazlactone was purchased from Pure Chemistry Scientific (Watertown, MA, USA). Polystyrene standards (PS; MW=580 to 7450000). Water was purified in-‐house using a Milli-‐Q Q-‐POD water-‐purification system (Millipore). Organosolv lignin (OSL, product no. 37, 101-‐7) was provided by Shell B.V.

3.2 Instruments

Size-‐exclusion chromatography (SEC) experiments were performed on the Shimadzu 2 system (Table 1) and the reversed-‐phase liquid chromatography (RPLC) experiments were performed on the Agilent system, shown in Table 1. The column used for SEC was PLRP-‐S (7.5x300 mm, 3-‐ µm particles, 100Å) from Polymer Laboratories England, UK). The column used for RPLC was an Agilent Zorbax Eclipse Plus C18 (Rapid Resolution HT 4.6x50 mm, 1.8-‐µm particles). The column used for HILIC was a Phenomenex Kinetex column (50x3 mm, 1.7-‐µm particles, 100 Å). For the immobilization of laccase a microfluidic reactor was used based on the prototype of Wouters et

al.17_{For
UV-‐light
exposure,
Spectrolinker
XL-‐1500
UV
Crosslinker
(Spectronics
Corporation,}

365nm) was used. A Hamilton syringe (81320, 1001TL, 1.0 mL) and Spark Holland HPLC Column Thermostat oven (SPH99) were used for the immobilized enzyme degradation.

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Table 2: Preparation of lignin samples

3.3 Method

3.3.1 Calibration with polymer standards

The SEC column was calibrated with polystyrene (PS) standards. The PS-‐standards were dissolved in tetrahydrofuran (THF, stabilized/BHT). Toluene (500 ppm) was used as internal standard. The calibration was performed on system Shimadzu 2 and measured with an isocratic flow for 12 minutes. The mobile phase was THF (stabilized/BHT, AR-‐grade).

3.3.2 Sample preparation

For the analysis of lignin and its degradation products, Organosolv lignin samples were prepared with different ratios THF/aqueous buffer. The preparation of the aqueous buffer is done with TRIS HCl (394 mg), CaCl2 (148,5 mg) and water (MilliQ, 50 mL), based on the protocol

for the immobilized-‐enzyme reactor. The sample preparation is shown in Table 2. Organosolv lignin with a concentration of 250 ppm was used, diluted from a stock-‐solution of 50000 ppm (dissolved in THF).

Shimadzu 2 Agilent

Pump/ Degasser LC-‐10AD/ DGU-‐14A 1290 BinPump G42290A Detector SPD-‐M20A (1,5 Hz) 1290 DAD G4212A (160 Hz) Injection volume 20 µL 5 µL

Software Lab solutions 1.21 SP1 Agilent OpenLAB CDS Chemstation edition Rev. C01.04 [35]

Sampler -‐ 1290 Sampler G4226A

Ratio THF/buffer (%) Lignin dissolved in THF (µL) THF (µL) Aqueous buffer (µL)

90/10 5 895 100 80/20 5 795 200 75/25 5 745 250 70/30 5 695 300 60/40 5 595 400 50/50 5 495 500 45/55 5 445 550 40/60 5 395 600 30/70 5 295 700

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3.3.3 SEC analysis of lignin

The SEC analysis of Organosolv lignin and its degradation products was performed on system Shimadzu 2 and measured in isocratic mode with a flow rate of 1.0 mL/min with an analysis time of 12 minutes. The mobile phase was THF (stabilized/BHT, AR-‐grade). For the effect of water on the chromatogram of lignin the same LC system and mobile phase was used. In the sample vial, Organosolv lignin was dissolved in different ratios of THF/water, with the following compositions: 90%, 80%, 75%, 70%, 65% and 60% THF.

3.3.4 RPLC analysis of lignin

The RPLC analysis of lignin and its degradation products were performed on the Agilent system measured with the gradient in Table 3. This gradient works with two different mobile phases. The first mobile phase, A, consist of 95% water and 5% ACN. The second mobile phase, B, consist of 5% water and 95% ACN. Both the original and degraded lignin with different ratios THF/aqueous buffer were analysed with RP.

3.3.5 In-‐solution degradation of lignin

The in-‐solution degradation of lignin by laccase is done in five steps. The first step is to sonicate both lignin as laccase for 10 minutes. Laccase (1µL, 1000 ppm) was then added to lignin (500µL, 250 ppm). The following step was to lower the pH to 3/4 with HCl (5 µL, 1M). The mixture is now set in an oven (50˚C and 65˚C). After 24-‐hour trifluoroacetic acid (TFA, 2,5 µL) is added to lower the pH and stop the degradation.

3.3.6 Immobilized enzyme degradation of lignin 3.3.6.1 creating an immobilized-‐enzyme reactor

The procedure of creating an immobilized-‐enzyme reactor is based on the protocol of Wouters

et al. (2017) and consists of three steps. The first step consist of a surface modification of cyclic

olefin copolymer (COC) and in-‐situ polymerization. For the surface modification, a pre-‐treatment mixture of DEBP (30.0 mg), EDA (485.0 mg) and MMA (485.8 mg) and a monolith mixture of

Time (minutes) A% B% Flow rate (mL/min)

0 100 0 1.3

3.00 0 100 1.5

3.50 0 100 1.5

5.00 100 0 1.5

6.00 100 0 1.5

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DMPA (4,1 mg), BMA (160,3 mg), EDMA (239,4 mg), 1-‐propanol (281,0 mg) and 1,4-‐butanediol (321,2 mg) were prepared, sonicated (10 minutes) and degassed (10 minutes). Then the chip was flushed with acetone (2 syringe volumes) and dried with nitrogen. The pre-‐treatment mixture was injected into the chip. The chip was then exposed to UV light (365 nm, 4 minutes) and flushed with acetone (2 syringe volumes) again. For the in-‐situ polymerization the monolith mixture was injected into the chip. The chip was exposed to UV light (365 nm, 10 minutes) and flushed with methanol (2 hours, 1 µL/min)

The second step consist of photografting of poly(ethylene glycol) methacrylate (PEGMA) and 2-‐vinyl-‐4-‐4-‐dimethylazlactone onto the polymer monolith. A mixture of DEBP (20.1 mg) in methanol (0.5 mL), a mixture of PEGMA (30.2 mg) in water (MilliQ, 574 µL) and a mixture of DEBP (2.3 mg), 2-‐vinyl-‐4,4-‐dimethylazlactone (150.4 mg), tert-‐butanol (635.3 mg) and water (MilliQ; 211.0 mg) were prepared and sonicated (10 minutes). Then the DEBP mixture was pumped through the reactor (5µL/min for 2 minutes, then 0.5 µL/min for 30 minutes). The reactor was exposed to UV light (365 nm, 2 minutes) and flushed with methanol (5µL/min for 2 minutes, then 0.5 µL/min for 30 minutes). The PEGMA mixture was pumped through the solution (5µL/min for 2 minutes, then 0.5 µL/min for 30 minutes). The reactor was exposed to UV light (365 nm, 2 minutes) and flushed with water (5 µL/min for 2 minutes, then 0.5 µL/min overnight). The azlactone mixture was pumped through the reactor (5 µL/min for 2 minutes, then 0,5 µL/min for 1 hour). The reactor was exposed to UV light (365 nm, 5 minutes) and flushed with acetone (5 µL/min for 2 minutes, then 0,5 µL/min for 1 hour).

The third and last step consists the laccase immobilization and quenching of the unreacted azlactone groups. A mixture of laccase (1.5 mg) and TRIS buffer (0.75 mL) and a mixture of ethanolamine (62.0 mg) and water (MilliQ, 1015 µL) were prepared and sonicated (10 minutes). The enzyme solution was pumped through the reactor (5 µL/min for 2 minutes, then 0.5 µL/min for 2 hour). For quenching of the unreacted azlactone groups the ethanolamine mixture was pumped through the reactor (5 µL/min for 2 minutes, then 0.5 µL/min for 1 hour). The reactor was last flushed with TRIS buffer (5 µL/min for 2 minutes, then 0.5 µL/min for 1 hour) and stored at 4˚C.

3.3.6.2 Degradation of lignin with an IMER

For the degradation of lignin with immobilized laccase with a residence time of 3.8 minutes, a mixture of TRIS buffer and THF (80:20 vol.-‐%) was flushed through the reactor (2µL/min, 30 minutes, 65˚C). Then for the sample introduction a lignin sample in a TRIS buffer/THF solution (80:20 vol.-‐%) was pumped through the reactor (0.5 µL/min, 20 minutes, 65˚C). The sample was then collected in a vial (0.5 µL/min, 100 minutes, 65˚C). The reactor is flushed with an 80:20

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Different ratios of TRIS buffer/ THF (15/20/30 % THF) and different concentrations lignin (125 ppm and 250 ppm) were tested.

4. Results and discussion

4.1 Calibration curve of polystyrene standards

The calibration curve of the polystyrene (PS) standards is shown in Figure 9. A calibration curve is used to relate retention time to molar weight and shows the optimum molar weight range for the column. The optimum lies in the most horizontal part of the curve. The flow rate was fixed at 1.00 mL/min. The slalom effect is observed for PS standards from a molar mass of 2x106_Da.

Slalom effects occur when molecules are so large they get a higher retention time again. This is due to the fact that large polymers won’t go past the pores and stick randomly in the column and therefore elute later. When slalom effects are observed, the column is not compatible for that range of molar mass.

4.1.1 Molar weight distribution of lignin

Lignin is not a polymer with just one single molar mass, but has a wide distribution of molar weight. The molar weight distribution of Organosolv lignin is shown in Figure 6. This figure was created using the calibration curve with a chromatogram of lignin. The polynomial function of the calibration curve (y= -‐0.0487x3_{+
0.8944x}2 _{-‐5,918x
+18,839)
is
used
for
the
relation
between}

the retention time and molar weight. Together with the lignin chromatogram the retention time is converted to molar mass. Figure 10 shows that Organosolv lignin has a majority of molar mass of 526.9 Da.

Figure 9: Calibration curve of column PLRP-‐s. This calibration curve shows that this column has its optimum around the 15.000 Da. System Shimadzu 2. Mobile phase THF stabilized

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4.2 Effect of water on SEC of lignin

The effect of water on the chromatogram with SEC analysis is shown in Figure 11. When water was added to the lignin sample, a new peak was observed around 10 minutes. Generally, when the concentration of water is higher, the peak at 10 minutes was also higher, but this is not consistent. The lignin peak is normalized. Remarkably the shape of the lignin peak stays the same, so there can be concluded that water doesn’t affect lignin on the chromatogram.

Figure 10: Molar weight distribution of lignin. This distribution is built up from the polynomial function of the calibration curve (y= -‐0.0487x3_{+
0.8944x}2 _{-‐5,918x
+18,839)
and}

the lignin Chromatogram

Figure 11: Lignin chromatogram with different ratios of THF/water. With a higher ratio of water, the peak at almost 10 minutes gives a higher intensity. The shape of the lignin peak (at 8,5 minutes) stays constant. Peaks are normalized. System: Shimadzu 2. Mobile phase:

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4.3 SEC analysis of lignin and degradation products

4.3.1 In-‐solution degradation with 50˚C and 65˚C

Lignin is degraded with the enzyme laccase with two temperatures, 50˚C and 65˚C. First 65˚C was chosen because this is the optimum temperature for enzymatic activity, but this temperature also has a disadvantage as it is close to the boiling point of THF (65-‐67˚C). Therefore, the mixtures with a high ratio of THF were (partly or fully) evaporated after 24 hours. Because of this observation a lower temperature of 50˚C was chosen. There was much more of the mixture still present after 24 hours of degradation. Figure 13 shows that also with a temperature of 50˚C lignin is still degraded with laccase. Furthermore, from Figure 12, 13 and Appendix 1 it can be concluded that the enzyme activity is related to the THF content of the mixture. With the ratio of 90/10 THF/aqueous buffer a small shift is observed towards lower molar mass (Appendix 1). But from the ratio 70/30 (75/25 with 50˚C) a clear shift to the lower molar mass is observed, as shown in Figure 12,13 and Appendix 1.

Figure 12: Lignin chromatogram with and without laccase. The blue chromatogram is an original lignin sample and the red chromatogram is lignin degraded with laccase. Degradation is observed because there is a shift towards the lower molar masses. Peaks are normalized. Temperature degradation 65˚C. System: Shimadzu 2. Mobile phase: THF stabilized

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To establish with certainty that the degradation is due to the enzymatic activity of laccase and not due to the heat or other changes (for example the change in pH), a lignin sample was kept under the same conditions as for in-‐solution degradation, except for adding the laccase enzyme. Figure 14 shows a corresponding peak for the original lignin and the lignin that was treated with the in-‐solution method (without the laccase). This shows that under the conditions for in-‐ solution degradation lignin is not degraded by heat only, but the enzyme laccase is essential for degradation of lignin.

Figure 13: Lignin chromatogram with and without laccase. The blue chromatogram is the original lignin and the red chromatogram is lignin degraded with laccase. The shift towards the lower molar masses (to the right) indicates that lignin is degraded by laccase. Peaks are normalized. Temperature degradation 50˚C. System: Shimadzu 2. Mobile phase: THF stabilized

Figure 14: Lignin chromatogram with and without laccase with an additional lignin chromatogram, which is only exposed at 50˚C without any laccase (chromatogram in green). This chromatogram shows that lignin with only heat (green graph) has the same retention time and shape as the original lignin (blue chromatogram). This indicates that lignin is not degraded only by heat. Peaks are normalized. Temperature degradation 50˚C. System:

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4.3.2 Immobilized enzyme degradation

The degradation of lignin can also be done with immobilized enzymes using an IMER. It is challenging to make lignin compatible with the enzymes and reactor. For the enzymes, THF with a ratio of 70 vol-‐% or lower does not seem to be a problem, this is also shown in paragraph 4.3.1. The reactor is less compatible with THF, small ratios of THF (30 vol-‐%) seems to damage the monolith layer of the reactor. Therefore, a sample with a THF ratio of 15 vol-‐% was pumped through the reactor and analysed with SEC, shown in Figure 15. The problem arises with lignin that is dissolved in a low ratio of THF, because its solubility goes down. It is expected that very little to no lignin went through the reactor, as a brown residue was found at the inlet of the reactor and only one peak at 11 minutes is observed which is most probably caused by water.

To improve to solubility of lignin, the ratio of THF is increased to 20 vol-‐%. Furthermore, other changes are made, the concentration lignin was lowered (125 ppm) and the temperature was increased (65˚C). This degradation is also analysed with SEC, and shown in Figure 16. Because of the lower concentration of lignin, the intensity of the signal is low, therefore no clear peaks are observed. Moreover, there is no clear degradation pattern observed compared with in-‐solution degradation, as shown in paragraph 4.3.1. A minimum concentration of 250 ppm is required for a clear detection with this SEC column (PLRP-‐S). In addition, damage in the reactor was observed after one day of degradation with 20 vol.-‐% of THF. After ± 6 hours it was clearly visible by eye.

Figure 15: Chromatogram of lignin (blue) with both in-‐solution as immobilized degradation (red and green, respectively). Only a peak caused by water is observed with the immobilized enzyme degradation. Peaks are normalized. Temperature degradation: 50˚C. System: Shimadzu 2. Mobile phase: THF stabilized.

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4.4 RPLC analysis of in-‐solution degradation of lignin with 65˚C

Besides size-‐exclusion chromatography, lignin was also analysed with reversed-‐phase chromatography. RP was only used for the first batch of degradation products (in-‐solution degradation at 65˚C). The RP chromatograms (Figure 15 and Appendix 2) are more difficult to interpret and it is more difficult to identify every single peak. There is a small pattern that is recognisable in the different chromatograms, which is a shift towards earlier elution. This may indicate that the degraded form of lignin is more hydrophilic. A possible explanation is that OH-‐ groups previously shielded in the 3D structure have become available for interaction after the degradation.

Figure 16: Lignin chromatogram with and without laccase. The blue chromatogram is the original lignin sample and the red chromatogram is the lignin sample after degradation with the IMER. There is no clear degradation pattern observed. Concentration lignin: 150 ppm. Temperature degradation: 65˚C. System: Shimadzu 2. Mobile phase: THF stabilized

Figure 15: Lignin RP chromatogram with and without laccase. The blue chromatogram is the original lignin sample and the red chromatogram is the lignin after its degradation with laccase. A shift towards less hydrophobicity is observed. Temperature degradation: 65˚C.

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When analysing lignin with RPLC, one has to take into account that THF is used as solvent for the first dimension with SEC. THF is such a strong solvent that lignin would rather stay solvated in THF than interact with ACN, the mobile phase of reversed phase. This can be a problem because this will cause less retention and less separation. This is observed in Appendix 2, where the original lignin of a high ratio THF (for example 90% or 80% THF) gives rise to less peaks and therefore has less separation.

5. Conclusion

A SEC and RP separation method for lignin was investigated. The results of the SEC experiments suggested that Organosolv lignin is a relatively small polymer with a majority of a mass of 526,9 Da. This SEC method is also compatible for analysing degradation products, favourably with a high ratio of THF as solvent. A high ratio of THF is favourable because lignin is easily dissolved in THF and the SEC method works well with strong solvents, like THF. From the RP method, chromatograms were difficult to interpret, and more optimization is necessary to analyse lignin and its degradation products. The in-‐solution degradation of lignin with the enzyme laccase was assessed. The laccase enzyme is compatible with a high ratio of THF, up to 70 vol.-‐% THF the enzymatic degradation was still successful. The immobilized-‐enzyme degradation with the IMER is not yet compatible for lignin degradation because of the solvent THF. > 15 vol.-‐% of THF is necessary to fully dissolve Organosolv lignin, because it has a low solubility for other solvents. Unfortunately, prolonged exposure of the polymer monolith in the IMER to 20 vol.-‐% or more THF content leads to irreversible damage.

Preliminary Study on Enzymatic Degradation of Organosolv Lignin

Bachelor Thesis Chemistry