conversion of kraft lignin into lower molecular … · conversion of kraft lignin into lower...

CONVERSION OF KRAFT

LIGNIN INTO LOWER

MOLECULAR WEIGHT

COMPOUNDS USING

ULTRASOUND

Anamarija Marinov, Matevž Mencigar,

Beryl Meg Awino Oduor, Sara Noriega Oreiro

Group: K8-K-4-F19

Conversion of Kraft lignin into lower molecular weight compounds using ultrasound

Group ID: K8-K-4-F19

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Title: Conversion of Kraft lignin into lower molecular weight compounds using ultrasound

Theme: Process Modelling

Project Period: 8th Semester 2019

Project Group: K8-K-4-F19

Supervisors: Rudi P. Nielsen

Marco Maschietti

Page Numbers: 79

Date of Completion: 26/05/2019

Participants: Anamarija Marinov

Matevž Mencigar

Beryl Meg Awino Oduor

Sara Noriega Oreiro



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Abstract

Considerable research efforts have been put into different attempts for fragmenting lignin, a

biopolymer found in wood. Wood processing is done globally for production of paper, paper products,

different kinds of biomass and biofuel. Lignin is structurally a big part of wood composition with 15-30

% of its whole mass and hence, there is need for it to be used. Usage of lignin as a wholesome material

is problematic because of its complex and large structure, so it must be pretreated into smaller

molecules for eventual use. This project reviews usage of high power ultrasound (US) for

fragmentation of lignin for said purpose. Methods researched for its fragmentation are usually

expensive and lasting, therefore US could potentially be better option for pretreatment. This project

highlights spectroscopic methods used for detecting if fragmentation with this approach is possible

and potential products gotten.



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Preface

This report is a result of a project carried out by 8th semester students of Aalborg University

(Esbjerg) in the Spring of 2019 under the course of Process Modelling. It explores fragmentation of

lignin with US. The lignin that was used is a low sulfonate content Kraft lignin and it was provided

by Aalborg University (Esbjerg). Thus, the results achieved are limited to the type of lignin provided

and not to every existing kind.

The project received guidance from the laboratory technicians who understood and provided

directions on which and how the laboratory equipment functioned. The Health, Safety and

Environment course also provided help with how to handle the laboratory work area, material and

equipment.

Finally, we would like to thank our supervisors, Rudi P. Nielsen and Marco Maschietti, who

provided a lot of support and advice as they reviewed our work and our drafts of this project. They

took the time to meet whenever a meeting was requested for consultation and to share their

knowledge. Also, we would like to thank Sergey Kucheryavskiy for his help with the spectroscopic

measurements.



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Table of contents

1. INTRODUCTION ................................................................................................................ 8

2. THEORETICAL CONSIDERATIONS ....................................................................................... 11

2.1 Ultrasound (US) ..................................................................................................................... 11

2.1.1 Main properties of US ................................................................................................... 11

2.1.2 Piezoelectric and magnetostrictive effects ................................................................... 12

2.1.3 Effects of US .................................................................................................................. 13

2.1.4 Ultrasonic equipment .................................................................................................... 15

2.1.5 Use of US in industrial applications ............................................................................... 16

2.1.6 Effects of US in polymer fragmentation ........................................................................ 17

2.2 Lignin ..................................................................................................................................... 19

2.2.1 Molecular Structure ...................................................................................................... 20

2.2.2 Lignin fragmentation ..................................................................................................... 21

2.2.3 Lignin extraction methods and their effects on the structure ...................................... 22

2.2.4 Kraft Pulping Process ..................................................................................................... 23

2.2.5 Sulfonated Kraft Lignin .................................................................................................. 25

2.3 Use of US for Lignin Fragmentation ...................................................................................... 26

2.3.1 State of the art for the use of US in lignin ..................................................................... 26

3. OBJECTIVES ...................................................................................................................... 30

4. MATERIALS AND EXPERIMENTAL METHODOLOGY ............................................................ 31

4.1 Materials ................................................................................................................................ 31

4.1.1 Chemical reagents ......................................................................................................... 31

4.1.2 Equipment ..................................................................................................................... 33

4.2 Experimental Methodology ................................................................................................... 33

4.2.1 Solubility testing and solution preparations ................................................................. 33

4.2.2 US application to the lignin solutions ............................................................................ 34

4.2.3 Determination of the degree of fragmentation of the lignin ........................................ 35



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5. RESULTS AND DISCUSSION ............................................................................................... 41

5.1 Initial characterization of the Kraft Lignin ............................................................................. 41

5.1.1 Thermal analysis ............................................................................................................ 41

5.1.2 Solubility testing and choice of concentrations ............................................................ 42

5.2 Lignin fragmentation characterization .................................................................................. 43

5.2.1 Viscosity measurements ................................................................................................ 43

5.2.2 UV/Vis results and analysis............................................................................................ 46

5.2.3 NIR spectroscopy results and analysis ........................................................................... 48

5.2.4 FTIR spectroscopy results and analysis ......................................................................... 50

5.2.5 Raman spectroscopy results and analysis ..................................................................... 55

5.2.6 pH results and analysis .................................................................................................. 58

5.2.7 HPLC results and analysis .............................................................................................. 60

5.2.8 Thermal analysis of sonicated sample ........................................................................... 63

5.2.9 Analysis of solutions of Kraft Lignin + Hexanol.............................................................. 64

6. CONCLUSIONS ................................................................................................................. 67

7. FURTHER RESEARCH ......................................................................................................... 69

8. BIBLIOGRAPHY ................................................................................................................. 70

9. APPENDIX 1 ..................................................................................................................... 77

9.1 Spectroscopic methods ......................................................................................................... 77

9.2 HPLC....................................................................................................................................... 79



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List of abbreviations and acronyms

• ATR - Attenuated Total Reflectance

• DTG - Derivative Thermogravimetry

• ESR - Electron Spin Resonance

• EU – European Union

• FTIR - Fourier-Transform Infrared

• HPLC - High Performance Liquid Chromatography

• IR – Infrared

• NIR – Near-Infrared

• SEC - Size Exclusion Chromatography

• TGA – Thermal Gravimetric Analysis

• US – Ultrasound

• UV/Vis – Ultraviolet-Visible



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1. INTRODUCTION

The use of petrochemical fuels directly affects the global climate. In order to make a transition to

a sustainable, fossil fuel free society, mere independence from oil consumption for fuel and electrical

power is not enough. Since throughout the world oil-derived products like asphalt, waxes, lubricating

oils, chemicals, plastics and synthetic materials are used every day, there is a need to find alternative

ways of producing these materials. One of the alternative paths involves the transformation of biomass

components [1].

Biomass represents a renewable feedstock for the production of fuels, chemicals and energy. In 2016,

over one quarter (27.9%) of the European Union (EU)’s primary energy production was provided by

renewable sources. Biomass and biomass derived fuels represented 66.1% of renewable energy,

surpassing the total combined contribution from wind power (13.8%), hydropower (11.4%), solar

power (6.4%) and geothermal energy (3%). In addition to energy gained from solid biofuel, biomass

represents the only renewable source for the production of liquid transportation biofuel [2].

Figure 1.1 - Production of primary energy, EU-28, 2016 (% of total, based on tonnes of oil equivalent) [3]

Since the reserves of fossil fuels are decreasing, the energy and chemicals which are currently gained

from petroleum will need to be provided from other sources. That is why the utilization of renewable

resources for producing electricity and chemicals is on the rise. This trend is widely supported by

several governments that have passed legislations mandating increases in energy and chemical

production from sustainable sources, especially biomass. In Europe, the Dutch Ministry of Economic

Affairs set goals to obtain 30% of transportation fuels from biomass and to replace 20-45% of fossil-



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based raw materials with biomass by 2040. The EU has set a target of 20% of renewable energy’s share

of total energy usage (European production + imports) by 2020 with a mandatory minimum target of

10% for biofuels for all member states. These goals supported the increased interest in the research

and development of technology for biomass processing [4].

Since the goals set by the EU contribute to higher biomass use, a particular opportunity is presented

in the development of lignin valorization processes. Lignin is an organic amorphous polymer that is a

main component in lignocellulosic biomass (15-30% by weight, 40% by energy) together with cellulose

and hemicellulose. Lignin acts as a glue that gives plants their rigidity and resistance towards decay [4].

Figure 1.2 - Schematic representation of the location and structure of lignin in lignocellulosic biomass [4]

While a lot of research has been done in utilizing cellulose and hemicellulose, lignin has received little

attention. For instance, the worldwide pulp and paper industry solely produces 70 million tonnes of

lignin annually, however the market for lignin products remains restricted to low value products such

as dispersants, adhesives and surfactants. Consequently, more than 98% of the available lignin is being

burned as a low value fuel with less than 2% being isolated and used commercially. This large difference

in commercial utilization of lignin, however, implies a large potential for different industries to gain an

edge over their competition. The pulp and paper industry could use it to diversify their product

portfolio, the bio-based industry can develop high value products such as biofuels or it can be used for

polymer formulations in the chemical industry [5].



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What makes lignin a valuable precursor for value added products is its chemical structure and

properties. Lignin is largely viewed as major source of aromatic compounds, but is not limited to them

[4]. Products like phenols, carbon fibers, activated carbon, plastic materials, monocarboxylic and

dicarboxylic acids can all be obtained from lignin as well as lignin micro- and nanoparticles which show

great promise for use as emulsion stabilizers, antioxidants and agents for UV protection coatings.

Currently petroleum gained products such as carbon fibers which are used in supercapacitors and

energy storage devices could be replaced by carbon fibers from lignin, lignin-based polymers could be

combined with petroleum-based plastics to increase the biodegradability of plastics etc [6] [7] [8].

Current research in valorizing lignin is mostly focused on catalytic lignin valorization. While lignin can

be degraded by the use of chemicals and enzymes, alternatively, ultrasound (US) can be used [4].

Whether as a sole method of degrading lignin or coupled with the use of chemicals, the use of US

provides a potentially cheaper and faster method of degrading lignin and obtaining valuable chemicals

[9].

US has been reported as a tool for lignin processing because it enhances mass transfer, dispersion

phenomena and acts as a chemical reaction initiator. During ultrasonic irradiation, solid particle’s size

gets reduced which leads to an increase in surface area. US also acts at the structural level, severing

molecular bonds, by producing free radicals through localized high pressures and temperatures. It has

been proved that by the use of ultrasound during extraction and transformation processes higher

yields are achieved within less amount of the time, at lower temperatures and with a lower

consumption chemicals required [9].

This project will focus on treatment of Kraft lignin with US with the intention of confirming the effects

US is reported to have and in order to obtain useful lignin fragmentation products with added value.



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2. THEORETICAL CONSIDERATIONS

2.1 Ultrasound (US)

US is defined as acoustic sound wave with frequencies higher than the limit of human hearing.

Frequencies that are defined for US are between 20 kHz and 10 GHz as it is shown on Figure 2.1.

Figure 2.1 - Sound spectra [10]

2.1.1 Main properties of US

US has the same type of properties as audible sound (frequency, wavelength and amplitude),

with the sole distinction from audible sound being a higher frequency.

US waves can propagate through any solid, liquid or gas medium, but they cannot travel through

vacuum because waves need particles to propagate. The wave speed depends solely on the medium

through which the wave is moving. Sound speed is, in most of the cases, bigger in solids than in liquids,

and in both bigger than in gases. There are two main factors on which speed of sound is dependent

on, elasticity and density of medium (speed=elasticity/density). So, sound speed increases with lower

density and higher elasticity [11] [12].

Figure 2.2 - US speed in dependence of media in which waves are propagating [13]

There are four different kinds of US waves: longitudinal, transverse, surface (Rayleigh) and lamb (plate)

waves. Just as speed of sound, different kinds of US waves depend on the medium they are in and



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position of particles in that medium. In liquid and gas medium, sound travels as longitudinal waves.

Particles of waves are flowing parallel with propagation, and they are reflected to the source as it is

shown on Figure 2.3. It is the most common kind of US waves that is used in science [13].

Figure 2.3 - Longitudinal US waves [14]

In solid matter transverse waves can also take place. In this case, particles vibrate perpendicular to the

wave displacement [15]. In surface waves, particle vibration is characterized by an elliptical orbit.

Waves act along the thick surface of solids penetrating into it and are mostly used to detect

imperfections of surface. Lamb waves differ from surface waves that they travel through thin and

homogeneous solid materials [16].

Another important application of US is related to the constant velocity of waves in homogenous media.

This allows the testing of properties of different materials as waves travel at one pace and are reflected

or refracted at the boundary of a different material [17].

2.1.2 Piezoelectric and magnetostrictive effects

US is generated through the use of transducers which operate by exploiting the piezoelectric

or magnetostrictive effects inherent to some materials [18].

Piezoelectric effect has an ability to change mechanical or kinetic energy into electric energy, and vice

versa, because of crystal deformation caused by mechanical stress. When voltage is applied to a

piezoelectric material, changes in its length are produced. Thus, if an alternating voltage is applied,

transducer will vibrate, inducing the vibration of the particles in the medium. When opposite reaction

takes place and returning sound vibrates, piezoelectric material produces electric pulse [19] [20] [21].



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Figure 2.4 - Piezoelectric effect [22]

On another note, magnetostrictive effect has an ability to convert kinetic energy into magnetic energy

and vice versa, because magnetostrictive materials develop mechanical deformations when exposed

to magnetic field [23].

The basic approach to production of US using magnetostrictive effect is showed as a change in length

of magnetostrictive material exposed to magnetic field. An oscillator with a given frequency is

producing a magnetic field whose alternation has influence on the length of the magnetostrictive

material. Because of change in the length of material, it starts to vibrate. In the moment when vibration

made by material and vibration produced by oscillator are the same, US waves are produced [24].

Figure 2.5 - Magnetostrictive effect [25]

2.1.3 Effects of US

In general, the applications of US are divided into two categories: high and low power. Low

power US has high values of frequency (over 0.1 MHz) and low values of intensity (below 1 W/cm2),

while high power US has low values of frequency and high values of intensity [26]. Low power

applications are those where it is intended to obtain information about the environment where it is

applied but without altering it, while in high power applications, purpose is to produce permanent

effects using the ultrasonic energy [27].



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High power US is defined by the occurrence of acoustic cavitation in the system. Cavitation is a

phenomenon in which formation, growth and implosion of microscopic bubbles in the liquid is possible

because of variations of pressure produced by sound energy. Mostly, cavitation appears in the sound

range between 20 kHz and 1 MHz [28] [29].

In acoustic cavitation, as shown on Figure 2.6, sound wave is oscillating in sinusoid flow between high

pressure and low pressure regions of fluid.

During the negative pressure cycle rarefaction happens. When this negative pressure reaches the

vapor pressure exceeding the intermolecular forces (achieved with high enough intensity), distance

between molecules increases, pushing fluid apart, and creating bubbles [30]. On the other hand,

compression is the process that occurs during positive pressure cycle and bubbles are shrinking in this

part of the flow (following sinusoidal pattern on the Figure below). After multiple change of low and

high pressure, US bubble expands and finally explodes in stage of compression. Depending on the

compression and expansion of bubbles, boundary layers and thickness of bubbles itself have different

values. Because of decreasing boundary layer and increasing thickness, bubbles are exploding with

longer exposure to acoustic waves. Collapse of bubbles can result in localized temperature of at around

5300 K [31], pressure of 1000 atm and cooling and heating rates of 1010 K/s [32]. Thus, this high local

temperature and pressure are driving high power chemical reactions. The collapse leads to the

formation of radicals through separation of the molecules within and around the bubbles,

luminescence due to excited molecules formed losing energy and microjets shooting out of the bubbles

at speeds of hundreds of km per hour [29] [18].

Figure 2.6 – “Graphical summary of the event of bubble formation, bubble growth and subsequent collapse over several

acoustic cycles. A bubble oscillates in phase with the applied sound wave, contracting during compression and expanding

during rarefactions” [29]



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Through the pressure differences caused by US, physical (mechanoacoustic) and chemical

(sonochemical) processes are enhanced [18].

Mechanoacoustic effects of US are useful for their ability to enhance mixing and their erosion

capabilities. Erosion capabilities of US stem from microjets which can be used to damage cell walls and

increase the surface area in heterogeneous systems. Microjets are formed when a bubble in a solution

approaches a solid boundary, having its spherical shape distorted due to asymmetrical liquid flow

around the bubble, causing the bubble to collapse and directing the shockwave towards the solid.

Enhanced mixing occurs as a consequence of pressure differentials which are caused by the movement

of the sound wave through the solution. Mixing is enhanced through microstreaming, acoustic

streaming and shock waves. Microstreams are flow patterns which occur near bubbles or small objects

within a solution in a sound field. These streams create hydrodynamic shear stress which can

contribute for example to polymer fragmentation and enhance mass transfer. Acoustic streaming is

caused by the propagation of the pressure wave and causes a flow in the direction of the wave.

Shockwaves form after the collapse of bubbles and propel the molecules near the edge of the collapsed

bubble outwards, towards other parts of the solution [18][32].

US also enhances chemical reactions through sonochemical effects which cause the cleaving of

intramolecular bonds. Sonochemical effects are noticed as formations of radicals due to localised high

temperatures emerging from bubble collapse. For instance, when water is subjected to US it leads to

the cleaving of the hydrogen-oxygen bond in the molecule. This produces hydroxyl radicals and

hydrogen which then continue to react with other molecules. Sonochemical properties of US have the

potential to be utilised on an industrial scale since they lead to faster reactions at lower temperatures

and can reduce the quantity of chemicals required in the process [33] [34] [32].

2.1.4 Ultrasonic equipment

US process is carried out with US probe or a US bath. Sound waves are produced with the use

of external electricity source which is converted into mechanical energy in the sonicator probe (called

sonotrode) which generates sound energy. On the end of the sonotrode, as it can be seen on Figure

2.7, small bubbles are being produced in the US bath which are creditable for process of cavitation. As

explained in Section 2.1.3 cavitation causes deformations and fragmentation of samples in solution

because of the conversion of mechanical energy into heat [35].



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Figure 2.7 - Schematic display of a US probe and US bath [36]

US bath and US probe are causing cavitation, but there are certain differences between these two

techniques. Firstly, cavitation throughout the bath occurs more randomly and uncontrollably which

results in lower effect of sonication than with US probe. This instrument is not as accurate as the US

probe is, because sonication effect is spread uneven through the whole solution. On the other hand,

with usage of US probe cavitation is more arranged and solution with samples is more homogenized

which results in high efficiency and intensity of sonicator. Secondly, handling of US probe is easier

because it allows user to select most important parameter such as temperature, amplitude, pulsate

and intensity of US [37].

2.1.5 Use of US in industrial applications

Main and most common US usages are related with medical issues. It also has a great number

of large scale industrial applications, such as its use in the food industry or waste treatment [26],

although its great potential is not yet fully exploited and explored.

In the food industry low power usages of US are mainly related to non-invasive techniques used in

process and quality control, whereas high power applications are related with gas extraction from

liquid nourishment material or for enzyme and proteins treatment, among others [26].

Its usage in waste water treatment is connected with electrocoagulation for surfactants removal. Basis

of electrocoagulation is the application of electrical current to the water, generating a coagulant agent

- metal ions from electrodes form anionic and cationic complexes with hydroxyl ions (from water

US probe US bath



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splitting), and result in the coagulation of pollutants - and gas bubbles, which create turbulence and

push the produced floccules towards the surface [38]. Benefits of its combination with US are mainly

related to energy saving. Due to the radicals formed during cavitation and the gas released, electrical

conductivity is increased, and thus lower voltages are required [39].

When scaling the processes investigated at laboratory scale, it is necessary to take into account that

to process large volumes in a certain time interval it is necessary to use a greater amount of energy.

The power of industrial ultrasonic devices ranges from 500 W to 16 kW. For the most usual liquid

treatment applications, four or more units are often combined in order to increase capacity. For

example, a group of 60 kW can be used to process up to 50 m3/h of biodiesel. These equipment are

usually manufactured to be able to operate continuously [40].

Figure 2.8 – Example of ultrasonic equipment at industrial level [40]

2.1.6 Effects of US in polymer fragmentation

As shown in Section 2.1.3 the effects caused on polymers by US can be both physical or

chemical. Among the physical changes produced by US are found, for example, particle size

modification when polymers are presented as powders, or the cut of thermoplastics [31]. Chemical

changes induced by cavitation include, for example, the ultrasonically assisted polymer synthesis [41].

US can produce polymer fragmentation, being it reflected in a reduction of its intrinsic viscosity or its

molecular weight [42]. Fragmentation is understood as an irreversible shortening of the chain length

due to scission and not necessarily implying a chemical effect [31]. This mechanism basically arises

from the stretching of a sufficient long polymer chain by the solvent flow, due to the movement of this

fluid surrounding the explosive cavitation bubbles and also because of the propagation of the

corresponding shock waves generated. Thus, the strong velocity gradients are enough to break long

polymer chains into a polydisperse system of smaller size [30]. One of the more potential



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characteristics for the use of US in polymer fragmentation is that, unlike in other fragmentation -such

as thermal or photochemical- the tendency of breakage is not random and it occurs mainly in the

middle of the chain (as expected due to the stretching and cleavage mechanism) [31]. However, this

mechanism applies mostly to linear polymers and becomes more complicated with highly cross-linked

polymers, as it is the case of the lignin.

In Figure 2.9 is shown the ultrasonic fragmentation of different size polystyrene with respect to the

sonication time:

Figure 2.9 – Ultrasonic degradation of different low polydisperse polystyrenes with different starting molecular weights in

toluene. Conditions of the experiment: Volume: 50 cm3; Concentration: 0.5 wt%; Irradiation intensity: 17.4 W cm-2;

Temperature: 25ᵒC [43]

As it can be seen in the figure, the fragmentation of the polymers is faster with higher initial molecular

weights, and proceeds until a minimum value, where no matter the sonication time, or the starting

point, the molecular weight is no longer reduced. A big number of studies have shown that this

minimum molecular weight barely depends on the nature of the polymer, but on the conditions of the

US treatment [44]. So, for instance, under the conditions of Figure 2.9, a polymer with a lower

molecular mass than 30,000 u – as it can be lignin, which usually presents a molecular weight lower

than 20,000 u [45]- will not be affected by the sonication.

It has also been shown how in the presence of “weak spots” in the chain, the degradation rate is highly

increased, suggesting that chain breakage happens at these spots. However, great difference in the

relative bonding energies must exist [46].



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A compilation of the main parameters influencing polymer degradation when subjected to US is shown

below [30].

- Molecular weight of the polymer:

o Larger molecular weight → Faster degradation and higher degree of

depolymerization.

o Limiting molecular weight.

- Polymer concentration:

o More dilute solutions lead to an increase in the degradation process (as polymers

are “more free” to move around the cavitation bubbles).

- Solvents used:

o The higher the vapor pressure of the solvent, the less violent is the bubble

collapse, thus leading to a lower degradation rate.

- Temperature:

o The higher the external temperature, the lower the degradation rate (liquids vapor

pressure increases with temperature).

- Acoustic intensity:

o Higher acoustic intensity leads to an increase in degradation rate and extent (as

more and bigger bubbles are created producing stronger shear forces).

- Frequency:

o It is generally assumed that high frequency US enhances radically driven processes

(chemical effect) whereas low frequency US maximized physical effects. Anyway,

both physical and chemical effects are presented in the whole frequency range

used in US [18].

Other influences are, for example, the reactor geometry and type, and the pulse at which US is applied.

2.2 Lignin

Lignin is a highly complex aromatic heteropolymer whose biological use in plants is to increase

cell wall support and resistance to pathogen attack [47].

While lignin can be used as a base for producing materials designed for various applications as

mentioned in Chapter 1, Kraft lignin specifically can be used in foam fire extinguishers as a stabilizing

agent, a reinforcement pigment in rubber and in printing ink for high speed rotary presses [48].

Potentially, lignin usage could be much more extensive and it presents a series of associated

advantages such as [49]:



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- Renewable material

- Great potential as a raw material: it represents between 1/4 to 1/3 of all the renewable

organic carbon in the Earth

- Unique abundant renewable natural polymer of aromatic nature

- Structure with high number of hydroxyl groups: reactivity

2.2.1 Molecular Structure

The exact structure of protolignin (untreated lignin, as directly presented in plants) is

unidentified to date [50]. It is known that lignin consists of three different phenylpropane units called

p-coumaryl, coniferyl and sinapyl alcohol as shown in Figure 2.10. They result in monomer units called

guaiacyl propane, syringyl propane, p-hydroxyphenyl propane respectively. These units are linked

together by the alkyl– or aryl–ether bonds (around 60–70%), carbon–carbon (around 25–35%) and in

small quantity ester linkages at α and β positions (less than 5% in herbaceous plants). Besides these,

functional groups such as methoxy groups, phenolic and aliphatic hydroxyl groups are areas that

provide high reactivity to chemicals which makes them hydrolysable. However, the high percentage of

carbon–carbon bonds leads to the high resistance of lignin to chemical attack [51] [52].

Figure 2.10 - Lignin structure showing the three units of phenylpropane [53]

Lignin from soft wood e.g. pine, mainly comprises of approximately 90% coniferyl. However, lignin

from hardwood comprises of both coniferyl and sinaphyl at a 1:1 (w:w) ratio. Grass on the other hand

has all the monolignols present in the structure [18] [54] [52].



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The molecular weight which reflects the degree of polymerization directly correlates to the resistance

of the lignin. With the reduction in molecular weight, there is a reduction in the resistance of the lignin.

Increase in p-hydroxyphenyl monomer corresponds to a decrease in the molecular weight due to being

less cross linked [55].

Lignin is also linked to other carbohydrates such as cellulose and hemicellulose making up the cell

walls. The links of ester, ether and glycosidic bonds are believed to exist due to the difficulty in

separating the lignin from the carbohydrates [18].

Figure 2.11 - Possible lignin-carbohydrate linkages; from top to bottom; ester linkages, ether linkages, and glycosidic

linkages [18]

2.2.2 Lignin fragmentation

Fragmentation of lignin is dependent on the environment (alkaline, acidic or oxidative) it is in.

In acidic media, protonation of benzyl oxygen promotes the degradation of α- and β-ether units leading

to the formation of a benzyl carbonium ion intermediate, which can continue lignin depolymerization

[18].

In alkaline media, lignin is degraded by the breakage of α- and β-aryl ether bonds.



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In oxidative environments, degradation arises from the cleavage of carbon-carbon linkages between

phenolic units and α-β-carbon bonds, resulting in the formation of carboxylic acids. Displacement of

side chains to a non-substituted ring position may also occur [18].

However, problems sometimes arise from these chemical methods as they may produce a loss of

carbohydrates [18].

2.2.3 Lignin extraction methods and their effects on the structure

Lignin causes a significant resistance to the degradation of biomass because of its difficulty to

break down. Therefore, extraction is normally done to remove it from cellulose. However, these

extraction methods result in altered forms of the lignin due to the heterogeneous nature of the raw

material. There is no available method for the isolation of lignin without the risk of structurally

modifying it during the process [56].

These methods (mechanical and / or chemical) can be grouped into two main routes. The first group

includes methods in which cellulose and hemicellulose are released by solubilization, leaving lignin as

an insoluble residue. The second group includes methods that involve the dissolution of lignin, leaving

cellulose and hemicellulose as insoluble residues, followed by the recovery of lignin from the liquid

phase [56].

Lignin composition and yield is affected by several factors such as the extraction method, solvent type,

time, and temperature, being all important variables. The choice of extraction method also depends

on the type of starting material. Categorization of lignin depends on the extraction method which can

be used to identify the lignin such as Brauns lignin, lignosulfonate and Kraft lignin [54].



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2.2.4 Kraft Pulping Process

Figure 2.12 - Kraft process for production of paper pulp [57]

The Kraft process (Figure 2.12) for the manufacture of cellulose pulp is one of the most

prevalent in the market (85% of world production), but the recovery of Kraft lignin for chemical uses

is not widely practiced at this time [56].

Kraft pulping is accomplished by dissolving the lignin in hot alkaline sodium sulfide solution (“white

liquor”). Preparation of Kraft white liquor is done by dissolving sodium sulfide in 1 M NaOH, which is

used break lignin, hemicellulose and cellulose bonds indicated in Figure 2.11. The selected wood chips,

with size about 8 mm, and white liquor are fed to the digester. The objective of the digester is to

disintegrate the wood to result in a fiber product. Operating temperature is 160-180° C at a pressure

of 800 kPa, being the retention time needed of a maximum of 3 hours [58] [54].

The pulp is washed with water in the pulp washing system and separated from the combined liquids

(black liquor), mainly composed of the white liquor, lignin and some of the carbohydrates that break

down from the hemicellulose. The washing water flows counter current to the pulp [58][59]. The black

liquor is then sent to the recovery system, a part of the pulping process that consists of the evaporator,

the recovery boiler, the causticizer and the lime kiln. The recovery system functions to reduce the

waste from the pulping process, recycling the NaOH and Na2S for the pulping process, and also to

produce steam and power [60] [61].

Recovery of the chemicals in the pulping process starts by the evaporators which are arranged in series.

The steam that is used to heat the evaporators runs co current to the to the black liquor. This is done

to avoid fouling in the pipe which would occur because of the increased viscosity of the black liquor as



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it becomes more concentrated. The temperatures of the long tube vertical evaporators start from the

first effect at 150 ͦC with the last effect having a temperature of 46 ͦC with pressures of 340 kPa to 90

kPa (vacuum) respectively [58].

This concentrated black liquor with approximately 65% solids, is fed into the recovery boiler. The

recovery boiler operates at an oxygen deficient environment and it is where the black liquor is burnt

to form Na2S and Na2CO3 which is recovered as a smelt. To prevent carbon monoxide emissions, the

upper region of the recovery boiler is the oxidative ensuring complete combustion of the organic

materials. The combustion temperatures are at 1100-1300 ᵒC for 65% solids concentrated black liquor

[58].

𝑁𝑎2𝑆𝑂4 + 2𝐶 → 𝑁𝑎2𝑆 + 2𝐶𝑂2

The processing water is used to dissolve the smelt and results in a solution known as green liquor which

contains Na2S and Na2CO3. The solution is then reacted with calcium oxide to produce calcium

hydroxide in a reactor called slaker which is between the causticizer and the kiln. The operating

temperature is 99-109 ᵒC with a retention time 10-15 minutes for the causticizing reaction. However,

the bulk of the causticizing reactions occur at the causticizers which are a series of 2-4 continuous

stirred reactors. They are operated by a turbine revolving at 70-80 rpm and a retention time of 1.5-2.5

hours [58], [59].

𝑁𝑎2𝑆 + 𝑁𝑎2𝐶𝑂3 + 𝐶𝑎(𝑂𝐻)2 ↔ 𝑁𝑎2𝑆 + 2𝑁𝑎𝑂𝐻 + 𝐶𝑎𝐶𝑂3

The lime kiln is a chemical reactor that is used for calcination of the calcium carbonate to produce

calcium oxide. The retention time required by the kiln is 2-3 hours. Temperatures of 1200 ᵒC are

supplied from the combustion of natural gas which runs counter-clockwise to the lime.

𝐶𝑎𝐶𝑂3 → 𝐶𝑎𝑂 + 𝐶𝑂2

The CaO from the kiln is reacted to water to produce Ca(OH)2 and the solution is directed back to the

caustisizer to be used again in the process.

𝐶𝑎𝑂 + 𝐻2𝑂 → 𝐶𝑎(𝑂𝐻)2

The recovery boiler is also the unit where steam is produced to generate electricity when passed

through a steam turbine. Heat is produced when the black liquor undergoes combustion resulting in a

high temperature and pressure super-steam. The black liquor contains the bulk of the energy content

in wood [61] [60].

However, the Kraft process present a series of inconveniences [49]:



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- Limitation of the thermal capacity of the recovery boilers

- High investment: the boiler of recovery in an industry increases the capital in

approximately 19%

- A waste of a renewable and eco-sustainable product with high industrial added value

When purpose is Kraft lignin extraction from the black liquor, it is done by precipitation. Precipitation

occurs when the pH is decreased to 5.0 or lower though acidification [56]. The precipitated lignin is

washed thoroughly with distilled water and freeze-dried [54].

Kraft lignin recovered direct from the pulping process is characterized by a relatively high degree of

purity. It has a molecular weight between 2,500 and 39,000 u, with a hydroxyl group content of 1.2 to

1.27 groups per phenylpropane unit [56].

2.2.5 Sulfonated Kraft Lignin

Kraft lignin is not soluble in water and thus, presents a series of limitations [62]. At the moment

most of the lignin production is coming from the pulp and paper industry - sulfite process and Kraft

process – so, lignosulfonates are obtained from them: as a direct by-product in the first case, or by

sulfonation of Kraft lignin [63][64]. In the sulfite process, liquor consists in a solution of sulfur dioxide

(obtained from sulfur burning) in an alkaline solution (usually limestone). Sulfur dioxide reacts with

water and sulfurous acid is formed, which in turn reacts with the limestone, producing calcium

bisulfite, that dissolves the lignin, and lignosulfonates are obtained as process byproducts [65] [66]

[67].

When lignin is sulfonated, it can become water soluble as anionic charge density is provided. Kraft

lignin sulfonation can be achieved thought different methods: via sulfuric acid or via sodium sulfide

treatment. Pre-treatment is first applied in both cases in order to improve the reactivity to sulfonation,

with phenolation and hydroxymethylation [62].

Figure 2.13 – Sulfonated lignin molecule structure [68]



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2.3 Use of US for Lignin Fragmentation

2.3.1 State of the art for the use of US in lignin

As it has been explained in Section 2.2, lignin is a type of complex organic polymer, so, a priori

its treatment with US seems as a viable option in order to revalue this side-product. However, to date,

few studies have been conducted and/or reported, and focus and study has been placed on different

chemical and biological treatments. The thermal fragmentation attracts a lot of interest, and a scheme

of thermochemical processes for the transformation of lignin and their potential products is shown in

Figure 2.14.

Figure 2.14 – Scheme of thermochemical processes for lignin fractionation and their products and application [56]

However, despite the remarkable perspectives for the lignin revaluation, the methodologies used have

multiple technical limitations, which is why it is necessary to develop alternative processes that

operate with a high efficiency and are competitive from the industrial point of view [56].

Below, as starting point to this project is collected main available literature about experimental use of

US with lignin.

● Bussemaker et al. (2013) [18] studied the use of US in lignocellulose as a pretreatment

option. The purpose of pretreatment is to separate cellulose, hemicellulose and lignin in order to

increase and facilitate the fermentation of sugar monomers. After the ultrasonic pretreatment, both

the enzymatic and acid hydrolyses were improved, increasing the yields of glucose and xylose, as well



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as reducing treatment times, being it attributed to a better accessibility and delignification.

Delignification was achieved in different solvents, although in most of the cases it was also observed a

loss of carbohydrates, not desirable as the main purpose is its conversion into biofuels.

Purpose was not focused on lignin fragmentation but on the capacity of US for its extraction. However,

the extracted lignin was analyzed and although in most of the cases the molecular structure was

maintained, isolated cases were also observed, where the molecular weight was both decreased and

increased, inferring this that certain chemical reactions of condensation were taken place. Reason for

this to happen is that, depending on the solvent, conditions may induce species accumulation in the

bubble interface, causing a transfer of protons or increase radical scavenger, that could lead to a

recondensation. Low frequency US was used, and the power was fixed at a value of 120 W (volume

treated and acoustic intensity are not specified). It is pointed that lignin polymerization occurs in

insignificant amounts under the conditions of this experiment. Anyway this fragmentation was studied

by Electron Spin Resonance (ESR) revealing that occurs due to the homolytic cleavage (breakage of a

covalent bond in such a way that each fragment gets one of the shared electrons [69]) of the phenyl

ether α-O-4 and mainly β-O-4 bonds. Also, hydroxyl attack on lignin compounds (happening mainly in

the aromatic ring), led to hydroxylated, demethoxylated and elimination of side chains. Aqueous,

oxidizing and alkaline media were studied, achieving, in general, a higher delignification with the last

one, although in some cases it also led to condensation reactions. Higher treatment times did not

always give better results.

● Araceli García et al. (2012) [9] studied the effect of US on alkaline lignin coming from olive

tree pruning residues. Main goal was the removal of impurities presented in the lignin, both

hemicellulose and inorganic content. Solute was dispersed in aqueous media (1:20 weight ratio) and

different sonication times were applied at 40 kHz and 200 W (volume treated and acoustic intensity

are not specified). Once ultrasonic treatment was applied, the liquid fraction was analyzed with High

Performance Liquid Chromatography (HPLC), both before and after acid post-hydrolysis (to release the

monomeric sugars from the polysaccharides), determining the concentration of xylose, glucose and

arabinose. It is stated that short sonication times (15-30 min) allows the obtaining of the largest

carbohydrates content, whereas longer treatments lead a new rearrangement of these carbohydrates.

Also, with long US irradiation (120 min) it was observed how bigger amounts of lignin were dissolved

(19%), as the higher shear stresses could be promoting chain breakage of the polymer.

The solid part was also analyzed through Attenuated Total Reflectance (ATR), infrared (IR)

spectroscopy (chemical structure study), Thermal Gravimetric Analysis (TGA) and Size Exclusion

Chromatography (SEC), to analyze molecular weight.



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Figure 2.15 – Thermal behavior of the samples before (L1) and after US treatment (L2-30 (30 min) and L2-120 (120 min)).

Figure (a) shows TGA and Figure (b) DTG (Derivative Thermogravimetry) curves (inert atmosphere)

From the thermal analysis (Figure 2.15), some conclusions were made about lignin depolymerization

and fragmentation, based on the comparison between samples before and after US treatment. While

observing DTG curve, first peak at 223 o C (for initial sample) and 265 o C (for samples after US exposure),

and last peak at 706 ᵒC are showing that degradation profiles differ, meaning that lignin fragmentation

is taken place and that lignin has suffered some kind of modification in its chemical structure due to

US. From 300 ᵒC thermal degradation displays degradation of aromatic compounds. As the peak at 360

ᵒC is the same for all samples, its shows trivial or none degradation of aromatic structure of lignin.

It is also suggested that the low degradation of lignin in this case is due to the low power used during

its treatment with US, being in principle possible a greater depolymerization if the applied power is

increased.

● Finch et al. (2012) [70] extracted lignin in both acid (Formic Acid Lignin) and basic medium

(Ammonia Lignin) from a Miscanthus x giganteus crop. Lignin was subjected to catalytic

depolymerization under ultrasonic activation, using different nickel catalysts. This type of lignin is

insoluble in water, so the amount of lignin that was converted into lower molecular weight compounds



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was calculated as the difference in mass from the solid lignin before and after sonication. Lignin

extracted from the acidic pretreatment led to 86.5% delignification and basic one to a 36.3% (being

methanol the solvent in which better results were obtained). Thermal and Fourier-Transform Infrared

(FTIR) analysis were also performed for the characterization. However, experiments were carried out

under the same conditions without the use of catalysts, and the lignin was not depolymerized.

Available literature on lignin depolymerization with US is scarce, being it applied in most of the cases

as a purification method. However, from the results obtained it seems possible its use as a

fractionation stage, varying the conditions in which sonication is carried out.

Besides, the type of lignin treated in the different studies was not Kraft lignin, so different properties

are expected to be found in the development of this project.



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3. OBJECTIVES

Lack of studies about this topic are found in the literature and none of them is focused on Kraft

lignin. So, the challenges in the study are presented from the initial preparation and US treatment of

the solutions, to the methodology used in order to quantify the possible lignin conversion into lower

molecular weight compounds.

In general terms, this project’s focus will lay on several spectroscopic methods, which will be used in

order to prove eventual fragmentation of Kraft lignin.

The use of different solvents will be researched, with water being one of them (as used Kraft lignin is

soluble in it). Sodium hydroxide solutions will be also analyzed in order to check if and how, lignin

fractionation can take place in alkaline media, and 1-hexanol will be used because of its low vapor

pressure, beneficial for US treatment. However, main focus will be on oxidative environment, using

hydrogen peroxide. With HPLC it will be checked if it is possible the obtention of carboxylic acids by

the application of ultrasonic treatment instead of through its wet partial oxidation, which entails the

usage of high pressure and temperatures.



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4. MATERIALS AND EXPERIMENTAL METHODOLOGY

4.1 Materials

Chemical reagents and equipment used in order to achieve the set objectives (Chapter 3) are

shown in the next section.

4.1.1 Chemical reagents

Below are introduced the main chemicals used in this project, which were used without any

further purification.

Table 4.1 – Main chemicals used and their specifications

Reagents Structure Supplier CAS number Molecular

weight (u)

Kraft lignin, low

sulfonate content

*

Sigma-Aldrich 8068-05-1 ~ 10,000

1 – Hexanol (98%)

Sigma-Aldrich 111-27-3 102.17

Hydrogen

peroxide (33%) VWR Chemicals 7722-84-1 34.04

Sodium Hydroxide

VWR Chemicals 1310-73-2 40.00

* Aggregation state is powder and it presents a brownish appearance.

o Specifications:

- Solubility: H2O soluble

- pH in H2O solution: 10.5 (3 wt. %)

- Impurities: 4% sulfur

Also, different compounds were needed to carry out HPLC procedure, including 10 carboxylic acids:



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Table 4.2 – Chemical regents used for HPLC methodology

Chemical

compound

Chemical structure Supplier CAS number

Molecular

weight (u)

Formic acid


Glycolic acid


Lactic acid


Acetic acid


Glutaric acid


Oxalic acid

MERCK 144-62-7 90.03

Malonic acid

MERCK 141-82-2 104.06

Maleic acid

MERCK 110-16-7 116.07

Fumaric acid

MERCK 110-17-8 116.07

Succinic acid




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Methanol


Potassium

dihydrogen

phosphate


Demineralized water was used in the preparation of all solutions.

4.1.2 Equipment

Main equipment used corresponding to the different analysis carried out, is specified in their

description in next section, 4.2 Experimental Methodology. To carry out the different operations, other

laboratory equipment such as pH meters, heating chambers, balances, etc. were used. Also, the

different Software associated with each equipment was used and spectroscopic data was treated with

MATLAB software.

4.2 Experimental Methodology

4.2.1 Solubility testing and solution preparations

As a first step, lignin solubility in 4 different solvents was tested: Demineralized water, 1 M

hydrogen peroxide (H2O2), 1 M sodium hydroxide (NaOH) and 1-hexanol (C6H14O). It was mainly based

on visual inspection, but in some of the higher concentrations, solution was passed through a vacuum

filter for confirmation.

From there, in most of the cases presented in this project, concentrations of 1 g/L, 5 g/L and

10 g/L of lignin were dissolved and magnetically stirred before sonication.



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4.2.2 US application to the lignin solutions

Direct sonication was applied to 400 mL of solution with the equipment shown in Figure 4.1

(Vibra-Cell Ultrasonic Liquid Processor (Sonics & Materials, Inc.) Model: VCX 750). Manual cooling was

carried out in the samples, every 20 minutes, in order to avoid temperature rising, that would lower

the degradation rate.

Figure 4.1 – US equipment

Frequency used was 20 kHz (low frequency), being this a fixed parameter of the equipment used, but

several other parameters can be modified:

- Amplitude: The equipment is designed to deliver constant amplitude. This magnitude can

be modified as a percentage of the maximum one. Amplitude is directly proportional to

the power, so, in order to achieve highest power possible, tests were run with 100%

amplitude. However, the maximum power that is possible to be delivered may not be

750 W, as to achieve this quantity, the resistance to the movement of the probe should be

high enough to draw 750 W [71]. In this specific case, once ultrasonic treatment was run,

amplitude was automatically reduced to around a 22 ± 2% percent. Thus, the actual power

being applied was around 165 W.

- Pulse: It allows the application of US discontinuously. In this case, solutions were measured

with a pulse of 2” ON / 2” OFF, and 4” ON / 2” OFF. US was not applied in a continuous

way, as due to the high power used, and the amount of time that it was run, it could have

led to overheating and possible damage of the equipment.

- Time: All samples were treated for 120 minutes, except in one of the cases (1 g/L lignin in

H2O – 2” ON / 2” OFF), where US was carried out for 240 minutes in order to see if greater

time would allow a bigger fragmentation.



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Except in the solutions prepared for specific methods described below, during the two hours of the

experiment, samples were taken every 20’ for the measurement of Ultraviolet-Visible (UV/Vis) (3 mL)

and at 0’, 60’ and 120’ (40 mL) for FTIR, as due to the low concentrations of lignin studied, big amounts

of solution are required.

It should be noticed that power density is variable with time, beginning for example in this case with a

value of 0.41 W/cm3 and ending up with a value of 0.64 W/cm3, as samples are being extracted (volume

variations). Diameter of the probe used was 18 mm, thus applied intensity is around 65.8 W/cm2.

4.2.3 Determination of the degree of fragmentation of the lignin

Apart from the achievement of the fragmentation of lignin, one of the biggest challenges of this

project is the determination of whether this fragmentation is taking place as well as its

characterization.

The two main methods used in polymer science for this purpose are SEC and capillary viscometry,

which would allow an easy measurement of a decrease in molecular weight of the lignin molecules.

However, these two techniques are not available at AAU Esbjerg laboratory, so other different

techniques were performed in order to try to accomplish this goal.

• TGA

TGA is a technique that measures the quantity and the speed of the weight change of a sample

as a function of the temperature and/or the time in a controlled atmosphere. Generally, it allows

measurements to determine the composition of the materials and to predict their stability at

temperature of up to 1500 ᵒC [72] (in this case temperature range was 50-700 ᵒC at a heating rate of

10 ᵒC/min). This technique can, therefore, characterize materials that present weight loss or gain due

to decomposition, oxidation and dehydration [73].

It consists of a sample pan that is supported by a precision balance. The pan resides in a furnace and

is heated or cooled during the experiment [74]. The mass of the sample is monitored, and a sample

purge gas controls the sample environment, which may be inert (as in this case, N2 at a flow rate of 25

mL/min) or reactive [75].

TGA has been widely used for the investigation of the decomposition and thermal stability of organic

polymers [76], and in this project, as an initial characterization of the commercially available lignin,

TGA was carried out. Device used is TGA 550 (TA Instruments). Also, 1 g/L solution in 1 M H2O2 was

analyzed for comparison of the lignin structure (sample was dried at 85 ᵒC during 12 hours before

analysis), being possible an observation of changes in chemical structure and release of simpler



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components by comparing both thermal degradation profiles. Approximately 7 mg of each sample

were used.

TGA and DTG curves are obtained from the same result, as DTG is the first derivative of the TGA curve,

which represents the weight loss of the sample. DTG curves allow the identification of the range where

weight loss occurs more rapidly, as the first derivative of a signal is the rate of change of y with x,

interpreted as the slope of the tangent to the signal at each point.

• Viscosity

Due to the relationship between viscosity and molecular

weight, viscosity measurements could allow the identification of a

lignin fragmentation, being rheometry one of the experimental

techniques used for its measurement.

Rotational viscometer shown on Figure 4.2 (Programmable

rheometer Model DV-III (Brookfield)) is an instrument which

measures the resistance of the solution by applying a certain shear

rate (as viscosity is defined as 𝑆ℎ𝑒𝑎𝑟 𝑠𝑡𝑟𝑒𝑠𝑠

𝑆ℎ𝑒𝑎𝑟 𝑟𝑎𝑡𝑒 [Pa·s]). Around 8 mL of

solution was tested in each case with a SC4-18 spindle at 250 rpm,

resulting in an applied shear rate of 330 s-1. Temperature was kept

constant at 23 ᵒC and measurements were repeated 6 times in each case.

• Spectroscopic methods

Different spectroscopic methods were carried out in this project, including ultraviolet, visible

and IR radiation: UV/Vis, Near-Infrared (NIR), FTIR and Raman, in order to analyze chemical structure

of the lignin samples before and after sonication in the different solvents. Their main differences and

the mode of application in this project are shown below, and some theoretical extra information can

be found in Appendix 1.

- UV/Vis spectroscopy

UV/Vis refers to absorption spectroscopy or reflectance spectroscopy in the ultraviolet and

the visible spectral regions (200 – 800 nm). In these regions the energies of electromagnetic

radiation at different wavelengths match the excitation energies needed for non-bonding n

electrons or bonding π electrons to transition to the π* excited state of an unsaturated molecular

bond. Saturated molecular bonds cannot be excited in this region since the photon energy of the

incident beam is not high enough [77]. Therefore, UV/Vis spectroscopy is, although not exclusively,

Figure 4.2 – Rheometer



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used for quantitative analysis of unsaturated organic compounds in solutions [78]. For lignin

specifically, phenolic units can be detected and thus provide a quantitative lignin analysis [79].

Apparatus used was Cary 60-UV-Vis Spectrophotometer (Agilent Technologies). During the initial

analysis the scanning was conducted by measuring the absorbance in the electromagnetic spectrum

from 200 nm – 800 nm. From the obtained spectral lines, the wavelength of 279 / 280 nm proved to

be the local maximum for all measurements providing a reference point for future measurements.

The energy of photons at this wavelength was assigned to the excitation of non-conjugated phenolic

units present in lignin [80]. Afterwards, the analysis was conducted by measuring the absorption

values solely at 280 nm.

- FTIR spectroscopy

FTIR is a vibrational spectroscopic technique that identifies the information on the chemical

composition of samples based on the absorption of electromagnetic radiation through a wavelength

range of 2,500 nm to 25,000 nm [81] [82].

The FTIR obtained spectrum can be divided into 2 regions; the functional group region (4000 - 1500

cm-1), where individual functional groups can be identified and the fingerprint region (1500 - 400

cm-1), characteristic of the molecule.

FTIR is most sensitive to hetero-nuclear functional group vibrations and polar bonds, like C-H

stretch/bend or O-H stretching in aromatic and aliphatic alcohols in lignin and water [81] [82].

Because of high absorbance of water which leads to many overlapping absorption bands against the

molecule of interest, aqueous solutions are not well suited for FTIR analysis, which is why the

measurements of solid samples were conducted in this project. More detailed description of

detectable functional groups and their approximate wavenumber region is noted in the results and

analysis part in Section 5.2.4.

FTIR analysis was performed with Nicolet FTIR Spectrometer (Thermo Scientific) by measuring the

samples before sonication, after 60 minutes of treatment and after 120 minutes. To carry out this

procedure, 40 mL of each solution was dried in the oven at 85ᵒC, for 12 hours. After that, the

obtained solids were grounded and measured.

- Raman spectroscopy

Raman spectroscopy is a spectroscopic technique based on scattering phenomenon of

molecules. The nearly monochromatic electromagnetic radiation (laser), irradiated in order to

acquire the spectra, is usually in the visible or NIR region (850 nm in this case). Raman spectroscopy



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can be applied as a qualitative or quantitative method of analysis but was only used for qualitative

purpose in this project [78].

Raman and FTIR are referred to as complementary spectroscopic techniques since a different subset

of the full vibrational spectrum can be detected with one or the other method. Similarly to FTIR

spectra, Raman spectra have a functional group region (2000 to 3500 cm-1) and a fingerprint region

(500 to 2000 cm-1). Raman spectra though, are more informative about certain types of organic

compounds compared to IR spectra. For instance, homo-nuclear molecular bonds like C=C can be

more visible as well as aromatic carbon rings due to the expanding/contracting movement of the

ring [78] [83] [81] [82].

The use of this method has an advantage over IR spectroscopy in that it is free from interference

due to water bands so samples can be measured in aqueous solutions. Additionally, since the laser

probe of the Raman spectrometer used was not embedded in the instrument it allowed for in-situ

measurements of the sample. Raman spectroscopy’s main disadvantage is that some samples show

fluorescence, which can mask the Raman signal and make measurements difficult [81].

For this analysis, 400 mL solution of 1 g/L lignin in water was used, and spectrum was recorded every

minute while US was applied with RamanRXN1 Research Raman Instrument (Kaiser Optical Systems,

Inc).

- NIR spectroscopy

As FTIR and Raman, NIR is a type of vibrational spectroscopy. NIR is based on the absorption

of electromagnetic radiation ranging from wavelengths of 750 nm to 2500 nm. Energy of the

radiation provided in this range is lower than the one needed to promote electrons to an excitation

level but higher than the one required for molecules to achieve their lowest excited vibrational state

[84].

Since NIR spectroscopy’s range is in between the visible light and mid-IR wavelength range it is not

as useful for qualitative analyzes, as FTIR or Raman spectroscopy, but more useful for quantitative

analyzes, as UV/Vis spectroscopy [78]. The absorption regions of NIR spectroscopy are divided into

overtones of fundamental vibrational frequencies (harmonics) and combination bands [85]. The

bands that are mainly observed arise mainly from stretching of O-H, C-H, and N-H junctions and are

much wider in relation to what is observed in FTIR [81]. For the purpose of this project NIR

spectroscopy can be used to detect the quantity of phenols, alcohols and organic acids based on the

O-H vibrational stretch and of esters and carboxylic acids based on the overtone of carbonyl stretch

vibrations [78].



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Six different solutions of the same composition (1 g/L lignin in 1M H2O2) were prepared and analyzed

with NIR (at 0’, 60’ and 120’) and each of them was measured in triplicate in order to avoid

measurement and random errors. The equipment used was Quant NIR Analyzer (Q-interline).

• HPLC

HPLC is an analytical method utilizing the principles of column chromatography in order to

detect the amount of predetermined molecular species in a sample solution. The output data is

shown in chromatographic peaks at different retention times. Based on the area under the peaks

and the retention time the concentration and the type of the species is determined, respectively

[86].

The analysis was inspired by Maschietti et al. (2018) [6] where the presence of 10 organic acids was

investigated and detected after wet partial oxidation of guaiacol (a primary lignin product) in

hydrogen peroxide solution at temperatures of 150 – 200°C and pressure 100 bar. Since lignin is

comprised of guaiacyl units among others and US produces high localized temperatures and

pressures at bubble collapse as mentioned in Section 2.1.3, similar results were expected especially

for hydrogen peroxide dissolved samples.

For each acid a standard solution of 1 g/L was prepared and 0.1 g/L for fumaric and maleic acid, in

order to measure the retention times of the acids [6].

The analysis was done using a HPLC 1260 Infinity (Agilent Technologies). The columns consisted of

5 µm sized particles packed in a Hypersil GOLD C18 selectivity column (150mm x 4.6mm ID). During

the analysis 2 eluents were used as the mobile phase in a gradient mode of analysis (0.025M

KH2PO4(aq) solution at pH 2.5 and a methanol-water solution with a 9:1 ratio of methanol to water).

The flow of the mobile phase was 0.7 mL/min. For the detection of separated species a UV detector

was used with the incident beam wavelength set at 210 nm, where the absorption maximum for the

carboxylic group is known [78]. The analysis ran at ambient temperatures.

Table 4.3 – HPLC gradient program (A=MeOH-H2O, B=KH2PO4)

Time (min) Eluent composition

0-14 A=0% B=100%

18-20 A=50% B=50%

22-30 A=0% B=100%

As a summary of the experimental methodology and clarification for next chapter (Chapter 5- Results

and discussion), a diagram is included below with the different operations carried out in this project.

Solubility testing in

H2O, NaOH, H2O2 and 1-hexanol

Thermal analysis

US

H2O

1 g/L 1 M H2O2

1 M NaOH

H2O

1 M H2O2

H2O

1 M H2O2

20 g/L H2O 2”2”

Treatment time: 120’ (all) + 240 ‘ (1 g/L H2O 2”2”)

Power = 165 W Freq = 20 kHz

Dynamic viscosity

20 g/L H2O 2”2”

(biggest concentration for apparatus accuracy)

250 rpm / 6 repetitions

pH measurements

HPLC

1 g/L H2O 4”2”

1 g/L H2O2 4”2”

Spectroscopic techniques

UV/Vis

(All sonicated samples)

FTIR

(All sonicated samples-

dried)

Raman

1 g/L 1 M H2O

4”2”

(Continuous analysis while

sonication)

NIR

1 g/L 1 M H2O2

4”2”

(6 solutions sonicated and

analyzed)

Thermal analysis

1 g/L 1 M H2O2 4”2” (dried)

US

1 g/L 1-hexanol 4”2”

Treatment time: 120 ‘ Power = 165 W Freq = 20 kHz

UV/Vis HPLC

KRAFT

LIGNIN

(POWDER)

+ + +

2”2”

4”2”

2”2” 5 g/L

10 g/L 2”2”

All analyzes shown were performed at different time intervals during the ultrasonic treatment



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5. RESULTS AND DISCUSSION

5.1 Initial characterization of the Kraft Lignin

As a first step of this project a characterization of the Kraft Lignin presented as a powder was

performed, in order to study, among others, its humidity content or its behavior in different solvents.

5.1.1 Thermal analysis

Figure 5.1 – TGA and DTG (Derivative Thermogravimetry) curves of commercial low sulfonate content Kraft Lignin supplied

by Sigma-Aldrich

Figure 5.1 shows the TGA and DTG curves as a function of the temperature, for the Kraft Lignin

used as the solute in this project, performed following the procedure explained in Section 4.2.3. In this

case DTG curve is not very useful, as it can be seen how in the range between 500 and 600 ᵒC, weight

loss was not “smooth”, so that this curve interval with peaks - and thus large slopes - has altered the

overall DTG shape.

Below, in Table 5.1 is collected a summary of weight losses from Figure 5.1:



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Table 5.1 – Weight losses of Kraft Lignin in the analyzed temperature range using TGA

Temperature range Range weight loss (%) Cumulative weight loss (%)

0 - 150 ᵒC 9.14 9.14

150 – 200 ᵒC 1.14 10.28

200 – 350 ᵒC 20.01 30.29

350 – 500 ᵒC 7.91 40.00

500 – 575 ᵒC 8.57 48.57

575 – 660 ᵒC 0.57 49.14

660 – 700 ᵒC 2.29 51.43

Total weight loss (%) (0 - 700 ᵒC) 51.43

When examining TGA curve, it can be seen that weight decreases around a 9% between the 0 and 150

ᵒC, which corresponds mainly to water evaporation. So, it can be concluded that humidity content of

the samples is around 9% of the total weight (although this could be a bit overestimated if there is

presence of some volatile gases coming out at this range of temperature, such as carbon dioxide or

carbon monoxide [87]). After that, degradation continues at a slower pace. Biggest weight loss occurs

between 200 and 575 ᵒC. Reason for this wide range in which lignin degradation occurs can be due to

the complex molecular structure of lignin [76]. In this region, monomeric phenols such as guaiacol are

released, due to fragmentation of inter-unit linkages [88].

Between 500 and 575 ᵒC a deep and abrupted decrease is observed, that could be related to the

decomposition of some of the aromatic rings [88] although slower decrease was expected, as both

degradation and recondensation could be happening [80].

In the 575 to 660 ᵒC range, a plateau can be seen, decreasing weight slightly afterwards. Within the

range of temperatures studied (0 - 700 ᵒC) a total weight loss of 51.43% was achieved. According to

Tejado et al. [89] at 800 ᵒC around half of the sample remains non-volatilized because of the formation

of highly condensed aromatic structures [90] [91]. Reason for this is that at temperatures above

700 ᵒC polycyclic aromatic hydrocarbons begin to form quickly, which have a very stable nature [92].

5.1.2 Solubility testing and choice of concentrations

As explained in Section 4.2.1, the solubility of lignin was observed in different solvents. When

lignin was dissolved in aqueous solutions (including demineralized water, 1 M NaOH and 1 M H2O2)

complete solubility was achieved up to values around 50 g/L. The solubility was confirmed by passing

the solution with the dissolved solute through a vacuum filter which did not show any filtered out



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solids on the filter paper. However, Kraft lignin was not soluble in the organic solvent (1-Hexanol), even

at the lowest concentrations.

A few tests with US were initially carried out at the highest concentrations (50 g/L - for aqueous

solutions) where complete solubility was achieved. As the final consideration of this project would be

to optimize the performance of fractionating lignin by increasing the amount that can be treated at

the same time, working with larger concentration would have proved necessary. However, as it will be

further explained in detail, US treatment did not achieve the expected lignin fragmentation and the

project was therefore focused mainly on 1 g/L, 5 g/L and 10 g/L concentrations since it has been

explained in Section 2.1.6 that more dilute solutions lead to an increase in the fragmentation process.

5.2 Lignin fragmentation characterization

5.2.1 Viscosity measurements

As explained in Section 4.2.3, one of the methods used for the study of a possible lignin

fractionation was the measurement of the dynamic viscosity with the help of a rheometer. In this case,

if fragmentation is occurring a decrease in viscosity would be expected.

First, measurements were made with different concentrations of lignin in water (before US treatment)

and the results obtained are shown in Table 5.2 and plotted in Figure 5.2.

Table 5.2 - Viscosity values for lignin samples with different concentrations at 23 ᵒ C (standard deviations based on 6

measurements)

Concentration of lignin (g/L) Viscosity (mPa · s)

1 1.81 ± 0.15

5 1.85 ± 0.13

30 1.83 ± 0.11

40 2.03 ± 0.15

50 2.05 ± 0.08

75 2.13 ± 0.07

100 2.17 ± 0.07



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Figure 5.2 – Average dynamic viscosity vs lignin concentration at 23 ᵒC showing standard deviations

From Figure 5.2 it can be seen how from the average values the expected trend is obtained –

increase in viscosity with increase in concentration – except for the 30 g/L solution. However, despite

the large range of concentrations analyzed, the viscosity variation is not very large (0.4 mPa·s),

deducing that a small lignin degradation could not be identified with this method. Besides, due to the

characteristics of the equipment, maximum available angular velocity applied is 250 rpm. When

measuring an initial blank sample of water, a value of viscosity of 1.67 mPa·s was obtained, while the

real viscosity of the water at 23 ᵒC (temperature at which the experiment was carried out) is 0.933

mPa·s [93]. This is due to the small viscosity of the water, that cannot be measured with the instrument

used, since with spindle used, minimum viscosity that can be measured is 3 mPa·s, and maximum

10000 mPa·s [94]. Therefore, it is intuited that in this case measurements are also overcalculated and

this is the explanation of the high variations obtained for each measurement. Thus, obtained

calibration line would be erroneous.

However, as a check, viscosity measurements were also done for one of the solutions treated with US:

20 g/L lignin dissolved in water and sonicated for 120 minutes, with samples taken every 20 minutes.

Although this project is mainly focused on solutions of smaller concentration due to their greater ease



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for fragmentation, in this case a higher concentration was chosen to try to minimize the error arising

from a very small viscosity that equipment could not measure correctly.

Results obtained are collected in Table 5.3 and plotted in Figure 5.3.

Table 5.3 - Viscosity values for sonicated sample of lignin with concentration of 20 g/L and time of US treatment at 23 ᵒC

(standard deviations based in 6 measurements)

Time of US treatment (min) Viscosity (mPa · s)

0 1.66 ± 0.11

20 1.78 ± 0.09

40 1.76 ± 0.11

60 1.91 ± 0.12

80 1.92 ± 0.08

100 1.98 ± 0.07

120 1.88 ± 0.09

Figure 5.3 – Average dynamic viscosity vs time of US treatment for an initial solution of 20 g/L lignin in water at 23 ᵒC

showing standard deviations



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It can be seen that an increasing trend with sonication time is obtained, which does not match the

expectations. Results suggest that instead of a fragmentation, a lignin condensation is happening.

However, results are not accurate as it can be seen in the big standard deviations obtained, due, as

explained before, to the low viscosity of the solution, being lower than the minimum that the

rheometer is capable of measuring. Thus, this method is inconclusive.

Capillary viscometer appears to be the technique that would work for our specific solutions, as it is the

technique usually used when working with polymers. It is based on the measurement of the time it

takes to a solution to travel through a capillary of a given length and diameter.

However, as this method was not available, several spectroscopic techniques were carried out and

results are collected below.

5.2.2 UV/Vis results and analysis

UV/Vis spectroscopy was done with the intention of detecting the quantity of phenolic units

present in lignin and to observe any changes caused by prolonged sonication effects.

Firstly, samples with known concentrations were analyzed. Since the spectra produced a lot of noise

when concentrations were higher than 0.1 g/L, lower concentrations were used and sonicated samples

were diluted before measurements. From the Figure 5.4 it was noted that the most important

wavelength in order to determine lignin content was at 280 nm, where the local maximum can be seen

for all spectra. As mentioned in Section 4.2.3 (UV/Vis spectroscopy) the wavelength of 280 nm

corresponds to the excitation of non-conjugated phenolic units present in lignin [80].

Figure 5.4 – UV/Vis spectra of different concentrations of lignin



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By studying the lignin polymer structure it was noted that lignin monomers, which are conjugated

phenolic molecules, transform into non-conjugated phenolic units during polymerization. Most often

polymerization leads to the formation of a hydroxyl group on the alpha position and ether linkage to

another lignin monomer on the beta position. In some cases, the ether linkage to a monomer forms

on the alpha position and a carbon-carbon bond to the same monomer on the beta position. It is also

possible that an ether linkage to the same monomer forms on both the alpha and beta positions or for

ester linkages to form.

Therefore, the hypothesis was made that the decrease of non-conjugated phenolic units is caused by

fragmentation. Through the decrease of non-conjugated phenolic units, UV/Vis spectroscopy could

thus show us whether lignin fragmentation occurs.

The analyzes were followed as described in the beginning of this section. Firstly, different

concentrations of lignin at different ultrasonic treatment times, were tested to see if any changes were

present. The results are shown in Figure 5.5.

Figure 5.5 - Relative absorbance for samples of different concentrations in water at 2"2" pulsation time

From the above figure no obvious difference between the sonicated samples at different

concentrations was noted. If our hypothesis about the decrease of non-conjugated phenolic units

indicating lignin fragmentation are true, the results point to random reaction of lignin fragmentation

and condensation occurring with no dominant trend in either way. The occurrence of these results

could as well be attributed, at least in part, to dilution error.

0,84

0,86

0,88

0,9

0,92

0,94

0,96

0,98

1

1,02

1,04

1,06

0 20 40 60 80 100 120

Rel

ativ

e ab

sorb

ance

(A

t/A

t 0)

Time [min]

1g/L

5g/L

20g/L



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Furthermore, samples of 1 g/L concentration were dissolved in different solvents, sonicated at

different pulsation intervals and sampled at various sonication times. The obtained results are

presented in Figure 5.6.

Figure 5.6 – Relative absorbance for samples in different solvents with different pulsations vs time of sonication

From the figure it was noted that there is no clear decreasing or increasing trend for each type of

experiment nor is there any correlation among the different experiments. Although, for pulsation

times of 4’’ ON 2” OFF additional observation was noted. The results show that for water and NaOH

almost all the measurements, with the exception of sonication in NaOH at 120’, had a higher

concentration of non-conjugated phenolic units than before sonication, indicating the occurrence of

lignin condensation. Secondly, the results from hydrogen peroxide are all lower than the pre-sonicated

sample, meaning that fragmentation of lignin occurred. These results indicate that at 4” ON 2” OFF

pulsation times some results were obtained and should be further investigated, while the 2” ON 2”

OFF pulsation times samples seemed to change due to random reactions taking place during sonication

(condensation/fragmentation) or there was an error made during sample dilutions.

5.2.3 NIR spectroscopy results and analysis

From the UV/Vis analysis most promising results arise from the 1 g/L solution in 1 M H2O2 (4”

ON 2” OFF), so, as explained in Section 4.2.3, 6 new solutions with this concentration were prepared

0,8

0,85

0,9

0,95

1

1,05

1,1

1,15

0 20 40 60 80 100 120

Rel

ativ

e ab

sorb

ance

(A

t/A

t0)

Time [min]

H2O 2"2"

H2O 4"2"

H2O2 2"2"

H2O2 4"2"

NaOH 2"2"

NaOH 4"2"



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and sonicated, being samples taken at 0’, 60’ and 120’, and each analysis was repeated 3 times for

each case.

The NIR spectra obtained is shown in Figure 5.7.

Figure 5.7 – Spectra of 1g/L Kraft lignin in 1 M H2O2 with color legend indicating sonication time

Spectra was preprocessed with Matlab Sofware for noise reduction, distortions correction, etc.

Besides, wavelength range was reduced to the NIR spectrum range (up to 2500 nm), removing thus

the noise presented in the tail.

Figure 5.8 - NIR spectra of 1 g/L Kraft lignin solution in 1 M H2O2 with color legend indicating sonication time, pre-processed



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It can be seen how all spectra corresponding to the different US time lies in the same transmittance

range. Even after preprocessing, up peaks (high transmittance) present noise and distortion and

besides, there is no a clear pattern between all lines.

Two main peaks are seen (in this case, attention should be paid to the local minima, as results are

based on transmittance):

Table 5.4 – Peaks observed in NIR of Kraft lignin in 1M H2O2 and their theoretically band contribution

Wavelength (nm) Band contribution

1685 Signal connected to C-H vibration [95]

2100 Combination of C-H and O-H stretching

vibration [96][95]

Smaller peaks are observed at a wavelength of 945 nm (that could correspond to the third overtone of

C-H, C-H2 and C-H3), 1130 nm and 1285 nm (both could correspond to the second overtone region of

these same bonds) [97]. Peak around 1830 nm may be associated to the first overtone of C-H

stretching, as according Jaya et al. (2015) it is found between 1670 and 1830 nm [96]. However, by

comparison with other NIR spectra of lignin found in the literature [95] [96] many other peaks should

have been obtained, for example those ascribed to C=O or different overtones related with the O-H

stretching. Besides, lignin is dissolved in a 1 M H2O2 solution and it is stated that O-H have a very strong

influence on the vibration overtones through the whole NIR spectra and are influenced by the

hydrogen bonding.

Thus, no valuable information can be obtained from this method. In addition to the no difference in

observance between the samples subjected to different ultrasonic treatments, spectra obtained do

not correspond to the one expected from a lignin solution.

5.2.4 FTIR spectroscopy results and analysis

Lignin spectrum from the FTIR showed the various peaks caused by the absorption of the IR

radiation that correlated with the various functional groups that are present in the lignin molecule.

The molecule has a complex structure, and as a result there are several peaks that highlight the various

bonds. In the Table 5.5 below, the key wavelengths that represent some of the functional groups that

are present in the lignin molecule and their molecular motions are shown.

Table 5.5 -IR Absorption for representative functional groups [98]



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Functional group Molecular motion Wavenumber (cm-1)

Alcohols O - H stretch 3650 or 3400-3300

C - O stretch 1260-1000

Ethers C-O-C stretch (dialkyl) 1300-1000

C-O-C stretch (diaryl) 1250 & 1120

Aromatics C-H stretch 3020-3000

C=C stretch 1600 &1475

Esters

C=O stretch 1750-1735

C-C(O)-C stretch (acetates) 1260-1230

C-C(O)-C stretch (all others) 1210-1160

Alkanes C–H stretch 2950-2800

Alkenes =CH stretch 3100-3010

The most defined peak in the spectrum of lignin is the one that occurs at the wavelength of about

1000-1300 cm-1. This falls in the area that shows the presence of ester and ether groups. Additionally

there are C-H bonds displayed at the wavenumber of 2930 cm-1 [99]. The aromatic groups were

represented by the grouped peaks between 1600 & 1475 cm-1. According to Araceli et al. (2012) the

peaks at 1265, 1215 and 1115 cm-1 are characteristics of peaks associated to syringyl and guaiacyl units

in alkaline lignin [9].



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Figure 5.9 - Spectrum of Kraft lignin

Spectra of lignin dissolved in the different solvents and dried again without any sonication taking place

was compared to ascertain the changes caused by the solvents within the lignin molecule. The spectra

are presented in Figure 5.10.

Figure 5.10 - Lignin 1 g/L non-sonicated samples dissolved in the different solvents vs pure lignin (undissolved)

The expectation was that all the spectra would be the same, but this does not match the results, as it

can be seen on Figure 5.10. The spectrum for lignin dissolved in 1M NaOH is probably different because

some NaOH residue was left on the dried lignin sample (NaOH boiling point is 1388 ᵒC [100]), which

showed up in the analysis. The changes observed in the spectrum of lignin dissolved in water showed

a decrease in all the notable bonds wavelength peaks, as well as a large baseline shift which was a

surprising result. At first the assumption as to why the water dissolved sample showed this kind of

spectrum was that the sample was not dry enough before measurement and contained some more

water which affected the measurement, but since the shape of it is practically the same as the shape

of undissolved lignin spectrum and the spectrum of lignin dissolved in hydrogen peroxide, but more

stretched, this anomaly was finally attributed to a measurement or program error.

Since the NaOH was interfering with the measurement and the spectra, further analysis for those

samples were omitted.

As the dissolved lignin was sonicated for a period and samples were withdrawn, the spectra of the

different intervals were compared. These comparisons were done in terms of time interval of exposure

to sonication. First, an analysis of 1 g/L of lignin in water at sonication time 4” ON 2” OFF was done

with sampling at 0’, 60’ and 120’ intervals of sonication. The spectra are shown in Figure 5.11.



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Figure 5.11 - Lignin 1 g/L in water at 4" ON 2" OFF sonication time

The overall result of the sonication proved hard to interpret as it remained unknown whether the

resulting shifts between the individual spectra were caused by measurement or program error or they

reflected the actual state of the samples. Assuming that the spectra reflects the actual situation, by

looking at the functional group region (4000 - 1500 cm-1) the biggest change that can be noticed is the

difference in the peaks around 2900-3000 nm which correspond to C-H bond stretch of the aromatic

and aliphatic structures. Furthermore, since the transmittance is increasing with sonication time it

should mean that the amount of C-H bonds in the samples is decreasing and that after 120’ of US

exposure, 2% of all the aromatic and aliphatic C-H bonds were gone and should be replaced by some

other functional group, or the ratio between the aliphatic and aromatic C-H bond peaks should change

(each peak corresponds to either aromatic (~ 3000 nm) or aliphatic C-H (~ 2900 nm) bonds). Since the

120’ as well as the 60’ spectral lines continually stay at a higher transmittance than the non-sonicated

sample spectrum in the functional group region and since the 2 peaks which correspond to C-H bonds

stay in the same ratio this means that no new functional groups are being formed and no aliphatic

molecules are made from the aromatic molecules or the products formed from fragmentation are

evaporated during drying of the samples. Furthermore, when comparing the FTIR spectra to UV/Vis

results obtained at the same sonication times do not match, UV/Vis results would indicate that more

non-conjugated phenolic units are formed meaning that the amount of C-H bonds should not be

decreasing. The comparison between FTIR and UV/Vis results indicates further that the spectral

differences obtained from FTIR are due to measurement or program error.

90

91

92

93

94

95

96

97

98

99

%T

1000 1500 2000 2500 3000 3500 4000

Wav enumbers (cm-1)



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The measurement or program error seems to be confirmed by looking at the spectra for lignin samples

of 5 g/L in water at the pulsation time 2” ON 2” OFF in Figure 5.12. The baseline shift is in this case

shifted in the other way having the pre-sonicated sample a continuously higher transmittance,

suggesting that lignin condensation was occurring with US exposure. That is why, at least for water

solvent, the FTIR analysis method does not contribute to any worthy results.

Figure 1.12 - Lignin spectra of 5 g /L in water for the pulsation time 2” ON 2”OFF

Despite the results for water solvent, the analysis for 1 g/L lignin sample in hydrogen peroxide at the

pulsation time of 4” ON 2” OFF were done as well. The spectra are shown in Figure 5.13.

Figure 5.13 - Lignin 1 g/L in H2O2 solvent with 4" ON 2" OFF sonication time

74

76

78

80

82

84

86

88

90

92

94

96

%T

1000 1500 2000 2500 3000 3500 4000

Wav enumbers (cm-1)

90

91

92

93

94

95

96

97

98

99

100

%T

1000 1500 2000 2500 3000 3500 4000

Wav enumbers (cm-1)



of 79

From looking at the region around 3000 nm and 1000-1300 nm it seems that C-H aromatic and aliphatic

groups have decreased while ether, ester and C-O alcohol functional groups have increased. The band

from 1000-1300 nm is thus pointing to the result that polymer condensation is occurring with

sonication which again does not match the results obtained from UV/Vis spectroscopic analysis for the

same samples where the results point to less non-conjugated phenolic units being present in the 60

min and 120 min sonicated samples. To further investigate if the FTIR obtained results are just

fluctuation in the spectra that cannot be interpreted, several runs of the same sample of 1 g/L in

hydrogen peroxide without sonication was done. The difference in each of the spectrum was too large

to be able to make any stable conclusions, thus leading to the conclusion that although, the FTIR can

be used to make a qualitative analysis of lignin molecule, an accurate quantitative measurement

cannot be reliably done.

5.2.5 Raman spectroscopy results and analysis

Below, in Figure 5.14, is shown the Raman spectra collected while a 1 g/L solution in water was

being subjected to US (4”ON 2”OFF pulse for 2 hours).

Figure 5.14 – Raman spectra of 1 g/L Kraft lignin solution in water while US is being applied. Color legend represents the

sonication time in minutes

When observing the raw spectra, a phenomenon of fluorescence (light absorption) can be seen,

resulting in noise that makes difficult the peaks observance. Fluorescence can be usually identified in

the Raman spectra as a curvature of the baseline [101] and it is one of the main challenges when

commercial lignin is studied using Raman spectroscopy (it emerges from the “intense” color of the

solutions) [102].



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Spectra was treated with Matlab Sofware in order to try to eliminate this fluorescence and other

sources of noise such as shot noise (result of the random nature of the light), instrumentation noise,

etc.

In Figure 5.15 is shown the spectra after different kinds of pre-processing were applied to the data,

including ALS baseline correction and smoothing (Savitzky-Golay), among others.

Figure 5.15 – Raman spectra of 1 g/L Kraft lignin solution in water while US is being applied pre-processed

Spectra of the first 4 minutes were considered outliers and removed for further analysis. US was set to

work after these initial measurements and probably mixing influenced this baseline deviation.

Figure 5.16– Preprocessed Raman spectra of 1 g/L Kraft lignin solution in water while US is being applied (Raman shift

range: 0-2000 cm-1 – Zoom in the peaks)

Main peaks are observed at 135 cm-1, 420 cm-1, 578 cm-1, 750 cm-1 and 1860 cm-1.



of 79

As lignin is a big and heterogeneous molecule, different spectral data was expected. Obviating the fact

that fragmentation occurs or not, many different interlinkages and functional groups are presented in

the lignin: ether linkages (C-O-C), carbon-carbon bonds (C-C), hydroxyl groups (OH), carbonyl groups

(C=O), aldehydes (CHO), methyls (CH3), etc. [103] and the little presence of peaks results unexpected.

For organic compounds, there are several bond types of C, H and O with a very characteristic

frequency, being the vibrational transitions presented in a local mode. Here, are included bands

associated with C-H, O-H, C=C and C=O stretching vibrations. On the other hand, bonds such as C-C

and C-O, when presented closed to similar bonds may be coupled and thus presented in a broad range

of Raman shift [102].

Agarwal et al. (2005)[103] reported that as H-bonding are formed with some of the groups, when lignin

is in solution, some band shifting occurs, depending on the type of lignin analyzed. However, main

peaks reported in the literature do not correspond with the ones observed in this study in a close

range.

Table 5.6 – Comparison between expected and observed Raman spectra

Raman shift (cm-1) Theoretical contribution [102] Observed (Figure 5.16)

3100 - 2800 Aromatic and aliphatic C-H

stretch Noise - No peaks

1860 -

Peak, max int.: 4.5

Fragmentation over time but without

clear pattern

1800 - 1500 Mainly aromatic rings, ethlylenic

C=C and γ-C=O No peaks

1500 - 1100

Mixed vibrations: coupled modes

(O-CH3, CH, phenolic and aliphatic

O-H…)

No peaks

1000 - 350

Difficult to identify – Skeletal

deformation of aromatic rings,

substituent groups and side

chains

750 cm-1

Peak, max. int.: 2.27

Apparently

condensation with time

but without clear

pattern

578 cm-1


Apparently




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but without clear

pattern

420 cm-1


Apparently


but without clear

pattern

135 -


Apparently condensation with time but

without clear pattern

As it can be seen the results obtained do not match the expected ones and those recorded in the

literature. Thus, conclusions related to US effect cannot be drawn either from this experiment as,

although changes are not being observed, the main linkages of the lignin molecule are not being

represented.

The analysis was carried out with lignin from other containers (same type and brand) to verify that the

lignin used was not contaminated. In all cases the same peaks were observed. Also, analyzes were

performed with diluted samples, in order to decrease fluorescence, that could be influencing and

“hiding” certain band contributions. Again, in this case, same peaks were observed.

5.2.6 pH results and analysis

pH was measured in the different solutions and results obtained are collected in Table 5.7 and

plotted in Figure 5.17.

Table 5.7 - pH values of lignin solutions with different concentration, pulse and solvent (1 month after preparation)

Concentration 1 g/L 5 g/L 10 g/L

Time H2O

2" 2"

H2O

4" 2"

H2O2

2" 2"

H2O2

4" 2"

NaOH

2" 2"

NaOH

4" 2"

H2O

2" 2"

H2O2

2" 2"

H2O

2" 2"

H2O2

2" 2"

0 8.10 8.47 5.22 5.96 12.50 12.86 8.32 5.42 7.92 5.68

20 7.63 7.36 4.92 4.56 12.49 12.63 8.31 5.15 7.48 5.46

40 7.71 7.34 4.10 4.21 12.50 12.24 8.23 5.05 7.43 5.23

60 7.58 7.16 4.02 4.04 12.50 12.51 8.15 4.50 7.38 5.12

80 7.53 7.23 3.91 3.82 12.52 12.88 8.09 4.35 7.25 5.08

100 7.39 7.07 3.88 3.73 12.48 12.80 8.08 4.36 7.24 4.99



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120 7.35 7.25 3.68 3.49 12.42 12.92 8.06 4.28 7.18 4.86

Figure 5.17 – Graphical representation of the pH variations with time in the different solutions

It can be seen how in most of the cases pH is decreasing, at least slightly, with sonication time. Kraft

lignin has an alkaline nature, thus increasing the pH of the solvents. From the information provided by

the supplier (Sigma-Aldrich) it is known that the pH of a solution of lignin in water (3 wt. %) should be

around 10.5. However, it can be seen how initial pH of samples diluted in water is around 8 in all cases.

pH of the samples was measured one month after their preparation. Some of the samples were made

again and pH measurements carried immediately after sonication, and the results obtained are

collected in Table 5.8:

Table 5.8 – pH measurements at the moment of solutions preparation

1 g/L lignin in H2O (4”2”) 1 g/L lignin in H2O2 (4”2”)

0 min 9.88 9.57

60 min 8.97 8.87

120 min 8.54 8.23

It can be seen how in both cases pH is higher than the pH measured after one month. The decrease of

pH with time was partly attributed to CO2 dissolving in the samples and forming carbonic acid which

in turn lead to the decrease of pH.

2

4

6

8

10

12

14

0 20 40 60 80 100 120

pH

US time (min)

1 g/L H2O 2"2"

1 g/L H2O 4"2"

1 g/L H2O2 2"2"

1 g/L H2O2 4"2"

1 g/L NaOH 2"2"

1 g/L NaOH 4"2"

5 g/L H2O 2"2"

5 g/L H2O2 2"2"

10 g/L H2O 2"2"

10 g/L H2O2 2"2"



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For the solutions with NaOH there is no changes in the pH with sonication time. Small variations may

be due to the accuracy of the pH-meter.

In the case of solutions with H2O, for the 1 g/L solution, a difference can be seen depending on the

pulse applied. When 2” ON 2” OFF was used, pH decrease is less obvious whereas for 4” ON 2” OFF the

decrease is more evident with a loss of 1.22 in pH, being the biggest drop during the first 20 minutes

of sonication.

The drop of pH was attributed to the decrease of hydroxide ions in the solution. UV/Vis spectroscopy

analysis showed the condensation of lignin occurring for 4” ON 2” OFF. As written in Section 5.2.2 in

some cases during polymerization a hydroxyl group forms on the alpha position. The formation of

hydroxyl groups means the hydroxide ions floating freely in the solution get used up during

polymerization, consequently decreasing the pH of the solution.

Solutions prepared with H2O2 are the ones showing the biggest pH change, except in the case of

solutions with 10 g/L concentration. The bigger the concentration, the harder is the fragmentation,

thus this could explain the slight changes observed in this sample. For the 1 g/L solution, again, the

biggest decrease is obtained during the first 20 minutes of ultrasonic treatment. Total pH variation is

2.2. As it has been explained in Theoretical Considerations Section 2.2.5, in an oxidative environment

the fragmentation of lignin is driven mostly by the cleavage of carbon-carbon linkages from which the

formation of acidic groups occurs. Thus, this variation in pH could be due to the occurrence of lignin

fragmentation, releasing these acidic compounds.

Also, it was noticed that although the original samples (0’ US treatment) were completely soluble, a

precipitate appeared when samples were sonicated (for water and hydrogen peroxide solutions). This

could mean that sulfuric acid is being formed, since the dissolved lignin had a low sulfonate content,

making it water soluble. If sulfur was removed from lignin during sonication it would make lignin

insoluble in water and form sulfuric acid. The decrease in pH would thus be due to carbonic and sulfuric

acid being formed.

All these assumptions were further tested in HPLC and results are shown below.

5.2.7 HPLC results and analysis

The intention of HPLC analysis was to investigate the presence of acids in the sonicated

samples. These experiments were done as further research based on the finding that in some of the

solutions the pH was decreasing with longer sonication time as described in Section 5.2.6.



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The analysis focused on the detection of 4 monocarboxylic (formic, acetic, glycolic and lactic acid) and

6 dicarboxylic acids (oxalic, malonic, maleic, succinic, fumaric and glutaric acid).

The samples of 1 g/L of lignin solution in water, hydrogen peroxide and hexanol at 4” ON 2” OFF

pulsation times were tested. The analysis showed some results for samples in hydrogen peroxide,

while no presence of investigated organic acids for samples dissolved in water and hexanol.

The results for samples dissolved in hydrogen peroxide are presented in Table 5.9, where

coincidentally the presence of hydrogen peroxide was detected at the retention time of oxalic acid,

which is why the written concentrations of hydrogen peroxide do not correspond to the actual

concentration of the solvent in the sample.

Table 5.9 – Results from HPLC analysis of 1 g/L lignin in 1 M H2O2

Time (min) Hydrogen peroxide (mg/L) Formic acid (mg/L) Maleic acid (mg/L)

0 6,911 84 0.547

20 6,922 120 /

40 6,824 120 /

60 6,821 136 0.514

80 6,724 133 0.5

100 6,619 106 0.562

120 6,536 123 0.67

From the analysis only the presence of 2 organic acids, formic and maleic, was detected. The

“surprising” result was that the acids were already present in the non-sonicated sample at 0 min. This

indicates that hydrogen peroxide is reacting with lignin even without sonication and to confirm this

hypothesis further tests were done. The measurements at longer sonication times proved that the

concentration of formic acid was higher than with the non-sonicated sample but did not show a

continuous increasing trend. Maleic acid was detected in trace concentrations only in 5 out of 7

samples which do not show any obvious correlation with sonication time. The non-monotone trend

for both acids indicates further oxidation of maleic to formic acid and formic acid to carbon dioxide as

hypothesized by Maschietti et al. (2018) [6].

As already mentioned above, the presence of hydrogen peroxide was coincidentally detected and

measured as well. A clear decreasing trend of its concentration was noticed, showing that hydrogen

peroxide is being used throughout the sonication. This finding, combined with the decreasing pH of



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samples and an increasing amount of precipitate with longer sonication time, indicates that during

sonication some chemical reactions are taking place within the samples. Part of the hydrogen peroxide

is probably used up by the oxidation of carboxylic acids while the other part by the formation of sulfuric

acid as mentioned in Section 5.2.6 [6]. To test the hypothesis of the formation of sulfuric acid, its

presence would need to be detected directly. Since no methods of sulfuric acid detection were found

in literature this hypothesis was not verified.

Further analysis with non-sonicated samples of 1 g/L lignin in hydrogen peroxide was done. The

intention of this test was to investigate whether hydrogen peroxide was reacting with lignin even

without sonication. A new solution of 1 g/L lignin in 1 M H2O2 was made, and HPLC analysis was

performed in the moment of its preparation (without sonication). The results only showed a peak

corresponding to hydrogen peroxide, but no formic or maleic acids were detected. When the same

sample was measured again after two weeks formic acid was detected as well as a drop in hydrogen

peroxide concentration, confirming the results obtained from the previous measurements.

As it has been explained in the previous section, the difference in pH between non-sonicated samples

measured in the moment of their preparation and one month later, was due to the double influence

of sulfuric and carbonic acid forming. However, the difference is bigger for samples dissolved in

hydrogen peroxide (~ 4 pH reduction) than in water (~ 1.8 pH reduction), and this is explained by the

results achieved, showing that lignin is reacting with hydrogen peroxide, releasing carboxylic acids

spontaneously without the need of ultrasonic treatment, matching the results of reduction in non-

conjugated phenolic groups obtained from UV/Vis. The HPLC results also showed that sonication acted

as a kind of catalyst, speeding up the process of lignin oxidation by hydrogen peroxide.

In the study realized by Maschietti et al. (2018) [6] about wet partial oxidation of guaiacol (product

from lignin hydrothermal decomposition) an obtention of carbon-based yield of monocarboxylic and

dicarboxylic acids in a range of 4% to 19% was reported, depending on the different operation

parameters (temperature was ranged between 150 and 300 ᵒC and retention times in the reactor

between 1 and 25 min). From our experimental results, the carbon-based yield for carboxylic acids

range (for the sonicated samples that were one month in solution before measurement), is between

10.6 % and 13.7 % (wt. % carboxylic acid per lignin), corresponding the highest value to 60 minutes of

treatment. So, results obtained from both methodologies are comparable. However, as explained

before, it should be noticed that an 8.5 % was obtained without the need of US and just by reaction

between hydrogen peroxide and lignin at ambient temperature.



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5.2.8 Thermal analysis of sonicated sample

Thermal analysis was also performed to a dried sample of 1 g/L lignin in 1 M H2O2 after 120‘

(4” ON 2” OFF pulse) of US treatment, for comparison with the analysis of the pure lignin (Section

5.1.1). TGA and DTG curves are presented on Figures 5.18 and 5.19.

Figure 5.18 – TGA and DTG of dried 1 g/L lignin 1 M H2O2 solution after 120’ of US treatment

Figure 5.19 – Untreated Kraft lignin TGA vs sonicated Kraft lignin TGA

The first observation made is that for the untreated lignin the weight loss in the temperature range

studied is 51.43 %, whereas for the sonicated one is 59.31 %. Thus, almost an 8 % difference in weight

loss was obtained.



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In this case a weight reduction of a 2.8 % is obtained up to a temperature of 125 ᵒC. The difference in

weight loss compared with the pure sample makes sense, since sample was dried prior to its analysis,

thus reducing its water and H2O2 content. However, this weight loss may correspond to some residues

of peroxide left, or because thermal analysis of the samples was performed one month after its drying

and therefore some humidity could have been gained by ambient contact. After 125 ᵒC weight is

reduced with approximately same slope until a temperature of 400 ᵒC. In this region weight is reduced

around an 17 % more than in the untreated sample, that could be due to a simpler lignin molecule

structure. From 500 ᵒC. Poletto (2017) stated that degradation of aromatic rings begins. In the first

case, an abrupt change was seen in the region to 500-575 ᵒC, that here is not presented. Thus, this

could indicate - as concluded from UV/Vis results and somehow corroborated by FTIR and HPLC - that

a depolymerization is occurring due to the transformation of the non-conjugated phenol groups to the

conjugated monomers, being thus carboxylic acids released, which are easily degraded at lower

temperatures. Although a comparable weight decrease is observed from 575 ᵒC, that may also be

related to aromatic rings degradation.

As a final conclusion, by comparison between both profiles, differences can be noticed, highlighting

that in the sonicated samples bigger weight loss occurred (and water content was less, so the actual

degradation of lignin was even bigger). This could mean that some kind of fragmentation was achieved

with the ultrasonic treatment, or at least that some chemical changes in the structure have taken place,

that favors the thermal degradation.

5.2.9 Analysis of solutions of Kraft Lignin + Hexanol

The analysis of the samples in hexanol is collected separately, since Kraft lignin is not soluble

in this solvent, and thus the procedure carried out for its analysis was a bit different.

Hexanol was chosen because of its low vapor pressure (1 mm Hg at 25.6 ᵒC [104]), which favors the US

treatment and degradation process. Solution consisted of 1 g/L lignin in 1-hexanol and was sonicated

during 120’ with a pulsation time of 4” ON 2” OFF. After US was applied, samples looked like shown in

Figure 5.20.



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Figure 5.20 – Solutions of Kraft lignin in hexanol with different sonication time

At first glance it might seem that with US, solute begins to be partially soluble, meaning this that some

kind of fragmentation was taking place, releasing smaller molecules. However, after some time, the

precipitate was deposited on the bottom, and the initial cloudy appearance may be due to a very small

particle size (caused by US).

In order to check both assumptions, different tests were carried out: UV/Vis and HPLC. Since lignin was

only partially soluble in hexanol, the samples were centrifugated and the supernatant was measured

without prior dilution in both cases.

● UV/Vis - The results are presented in Figure 5.21.

Figure 5.21 – Relative absorbance for samples in hexanol with 4” ON 2” OFF pulsations vs time of sonication

0

0,5

1

1,5

2

2,5

3

0 30 60 90 120

Rel

ativ

e ab

sorb

ance

(A

t/A

t0)

Time [min]

0’ 30’ 60’ 90’ 120’



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The obtained results are surprising. The results show that the absorbance is decreasing till 60 min of

sonication and later jumps up to almost 2.5 times the value of the non-sonicated measurement, at 90

min and 120 min of sonication. The sudden jump in relative absorbance could not be attributed to

lignin condensation since the difference is too big. The most probable reason for the occurrence of

higher absorbance is that through sonication lignin got more soluble in hexanol. This assumption would

mean that fragmentation was occurring, making the lignin molecules smaller and thus more likely to

dissolve. Interestingly, the effect of higher lignin solubility is noticed only after 60 min of sonication

and not before. Perhaps the lignin molecule was getting fragmented into still large enough pieces of

molecules, at 30 and 60 min, to not be soluble in hexanol but afterwards, at 90 min, those pieces

became small enough, dissolved and increased the absorbance. If this is the case, that would mean

that 50% of the dissolved lignin got fragmented from 0 till 30 min of sonication, afterwards some of

the lignin molecules got small enough to dissolve, slightly increasing the absorbance at 60 min, while

later fragmentation to smaller molecules on a larger scale took place increasing the absorbance to the

value seen at 90 min. The absorbance measured at 120 min is smaller than at 90 min meaning that

maybe further conversion of the smaller, already dissolved, molecules took place, but no more small

pieces of the bulk lignin molecule got fragmented and dissolved. To further investigate whether these

assumptions were true and that the absorbance obtained at 90 min indeed is the maximum

absorbance possible for lignin in hexanol, this sample would need to be sonicated for a longer time

and later analyzed using multiple methods.

● From the analysis in HPLC of the liquid part of the samples, no carboxylic acids were detected.

So, it is assumed that the apparent fragmentation of the lignin molecule observed in UV/Vis did not

take place through the degradation of aromatic rings.



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6. CONCLUSIONS

In this project, experimental part was conducted as US treatment of different Kraft lignin of low

sulfonate content solutions, followed by methods for determination of the degree of fragmentation of

lignin itself. This project focused on spectroscopic methods, TGA, pH tests and HPLC.

Based on the conducted test and obtained results it is conclusive that with instruments provided

fragmentation of lignin with US was not accomplished for water, sodium hydroxide and 1-hexanol

solvents while US enabled faster fragmentation by hydrogen peroxide.

Rheometry was used in this project for observation of possible changes in dynamic viscosity of the

sample before and after sonication, however due to equipment limitations and its accuracy with small

values of viscosity, this method could not be used for the indication of any changes in molecular

structure.

The pH values of sonicated samples were measured next. For all samples the pH decreased (except the

ones in NaOH), being the biggest decrease in pH before and after exposure to US, noted for 1 g/L lignin

in hydrogen peroxide with 4” ON 2” OFF. Two possibilities were considered: 1) That carboxylic acids

were released as a consequence of reactions taking place after the carbon-carbon bond cleavage and

aromatic ring breakage and 2) that sulfonates presented in Kraft lignin used were released generating

sulfuric acid and precipitating some lignin from the solution.

To check the nature of the acids released HPLC was conducted for the lignin solution in water, hexanol

and hydrogen peroxide. Only the samples in hydrogen peroxide showed formic and maleic acids were

present in sample both before and after exposure to US in small amounts. Thus, explaining why the pH

was lowest for hydrogen peroxide dissolved samples. The decrease of pH for samples in other solvents

was attributed to the formation of sulfuric acid. The presence of sulfuric acid, however, was not

experimentally analyzed and proven. Comparing the carbon-based yields of carboxylic acids obtained

by the ultrasonic treatment of lignin samples in hydrogen peroxide and those recorded in the literature

from its wet partial oxidation, results are comparable (US: 10.6 % -13.7 % vs oxidation: 4 % - 19%)

In UV/Vis, samples were measured at a wavelength of 280 nm, which showed the presence of non-

conjugated phenolic units. From all solutions, the only one showing a decreasing trend was 1 g/L lignin

in H2O2 with 4” ON 2” OFF pulsation time, indicating that fragmentation has taken place, which was in

accordance with the results from HPLC.



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FTIR and NIR were the next spectroscopic methods preformed in this project. Samples of multiple

concentrations and with different solvents were analyzed but the obtained results were mixed. These

spectroscopic methods were deemed inaccurate for a proper qualitative or quantitative analysis.

Raman spectra for water and lignin solution showed few main peaks which do not have clear pattern

(with respect to time) and were not corresponding to theoretical values for bonds presented in lignin,

which should have shown a different spectrum. Overall, this experiment did not prove the

fragmentation of lignin with usage of US as results were hard to interpret.

When 1-hexanol was used as a solvent, due to its advantage of low vapor pressure, no meaningful

results were obtained.

In conclusion, the objective studied, lignin conversion into smaller molecular weight compounds with

US, was not achieved. Only one sample (1 g/l in H2O2 4” ON 2” OFF) showed in some of the performed

analyzes that a small fragmentation could be taken place in the aromatic rings, but it cannot be

assured.

Main explanation for the failure in the achievement of the objective is believed to be the low power

(165 W) and frequency (20 kHz) that was applied to the samples.



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7. FURTHER RESEARCH

Further research could be undertaken to better accomplish and complete the objectives studied

in this project.

Due to resource limitations, the project is based on the Kraft lignin with low sulfonate content, one

available at Aalborg University (Esbjerg). However, the analysis of different kinds of lignin (without

sulfonate content) could be researched for eventual different results.

Furthermore, solubility could be tested in acidic solvents, as in the current project acidic environments

were not studied. Since solubility of lignin in 1-hexanol is not total but partial in small concentrations

of lignin (1 g/L), it is recommended to find a solvent with a lower vapor pressure, thus ensuring

degradation rate, but where lignin is soluble and facilitating the process. In order to increase solubility,

it is proposed to carry out tests with the addition and combination with other solvents.

Due to detection problems of lignin fragmentation, methods and instruments with ability to measure

molecular weight distribution (SEC) or intrinsic viscosity (capillary viscometer), could precisely

determinate the influence of US on lignin.

As it was explained throughout the project, because of probe resistance, ultrasonic device performed

with lower power, so sonicator with higher power or smaller probe should be used. Also, US setup

could be improved with flow cooling system which can ensure constant temperature of the solution

exposed to US, since higher external temperature lowers degradation rate.

Connected to the flow cooling system is another proposal for further research involving higher pulse

of sonicator. US exposure to solution in this project was limited to the pulses 2’’ ON 2’’ OFF and 4’’ ON

2’’ OFF since with pulse 6’’ ON 2’’ OFF there were overheating problems. Higher US exposure time

would possibly help fragmentation if lower external temperature can be delivered.



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9. APPENDIX 1

9.1 Spectroscopic methods

Figure 8.1 – Electromagnetic spectrum

Spectroscopic methods are based on the principle that when electromagnetic radiation interacts and

is absorbed by matter, there is a change in energy in the sample. When spectroscopy is based on

absorption most of the radiation passes through the sample without any interaction, but at certain

wavelength, its intensity is decreased (absorbed). However, this mechanism differs from visible and

ultraviolet radiation to the IR radiation. For the first two, the photon absorption modifies the energy

of the valence electrons of the sample (so atoms and molecules undergo electronic transitions,

measuring transitions from the ground state to the excited state), whereas for IR it is the bond

vibrational energy that is modified [105] [106].

An absorption spectrophotometer is an instrument that measures the intensity of the incident light

transmitted through a specimen. Comparing the intensity measured through a sample solution to the

intensity of the initial background (solvent), the amount of light absorbed by the sample is measured

indirectly [107]. The absorption is expressed through Beer-Lambert equation (1).

A = −log (I

𝐼0) (1)

Where A represents the absorbance of the sample, I the intensity of light measured through the sample

and I0 the intensity measured through the background (solvent) [107].

λ(m)



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FTIR

The FTIR obtained spectrum corresponds to the fundamental modes of molecular bond

vibrations in the sample, which arise due to the changes in the dipole moment of the bonds.

The spectrum can be divided into 2 regions; the functional group region (4000 - 1500 cm-1) and the

fingerprint region (1500 - 400 cm-1). The functional group region includes peaks that indicate a specific

kind of bond and are used to identify individual functional groups. The fingerprint region is made up

of many overlapping signals from multiple bonds deforming at the same time and is therefore less

suited for qualitative analysis. On the other hand, the fingerprint region is useful to identify the

measured molecule and note any changes in its constitution with consecutive measurements, since

each organic molecule produces its own unique spectrum and even small changes in the molecule’s

structure result in significant changes of the spectra. To some degree the fingerprint and functional

group regions overlap [108] [78].

A single beam of IR light is produced by the instrument and passed through the ATR crystal onto its

surface. Hence, when a sample is brought in contact with the crystal, the beam interacts with it, travels

through the detector and thus a spectrum can be obtained. The IR light undergoes several reflections

inside the crystal to increase the interaction with the sample [109].

In this project, the FTIR is set on the iD7 ATR-Diamond program and set to take 10 scans at a

wavenumber range of 500 - 4000 cm-1 and a resolution of 4 cm-1. An empty beam background (no

sample in the light path) is recorded first. This spectrum shows the instrument energy profile, which is

affected by the characteristics of the source, the beam splitter (KBr in this case), the absorption by the

air (mainly due to CO2 and water vapor) in the beam path, and the sensitivity of the detector at

different wavelengths. The sample is placed on the crystal and the arm is placed down on it before the

sample spectrum is collected.

Raman

The main principle behind Raman spectroscopy consists in the shift of the energy state of the

photons of the excitation laser beam (850 nm in this case – NIR region) with which the sample is

irradiated. Like FTIR, Raman also investigates fundamental vibration modes of molecules, although

these are not based on the change of the dipole moment, but instead, on the difference in polarizability

of a molecule as it vibrates [83] [81] [82].



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9.2 HPLC

Column chromatography uses a stationary phase (solid) and a mobile phase (liquid or gas). The

sample mixture is dissolved in the mobile phase which passes through the stationary phase. Different

molecules travel through the stationary phase at different times and are thus separated and identified

at the detector. With the instrument used the amount of passed molecules is detected using a UV

detector and the concentration calculated through Beer-Lambert’s Law (1) and calibration [86].

HPLC uses a high pressure pump (up to 400 bar) to push the mobile phase through tightly packed

particles in the columns of the stationary phase. The high pressure forces the different species to travel

faster through the stationary phase than in the gravitationally-driven column chromatography, making

HPLC a more convenient method. The output data is shown in spectral peaks at different retention

times. Based on the area under the peaks and the retention time the concentration and the type of

the species is determined. Even low concentration of species (down to hundreds of µg/L) can be

reliably detected by noting the retention time of the corresponding peaks [86] [110].

conversion of kraft lignin into lower molecular … · conversion of kraft lignin into lower...

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