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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
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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
Sigma-Aldrich 64-18-6 46.03
Glycolic acid
Sigma-Aldrich 79-14-1 76.05
Lactic acid
Sigma-Aldrich 50-21-5 90.08
Acetic acid
Sigma-Aldrich 64-19-7 60.05
Glutaric acid
Sigma-Aldrich 110-94-1 132.11
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
VWR Chemicals 110-15-6 118.09
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Methanol
VWR Chemicals 67-56-1 32.04
Potassium
dihydrogen
phosphate
Sigma-Aldrich 7778-88-0 136.08
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)
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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.
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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
Peak, max. int.: 1.42
Apparently
condensation with time
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but without clear
pattern
420 cm-1
Peak, max. int.: 5.50
Apparently
condensation with time
but without clear
pattern
135 -
Peak, max. int.: 10.8
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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8. BIBLIOGRAPHY
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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].