vorgelegt von diplom-chemiker philipp m. grande aus...
TRANSCRIPT
"Novel bio-based catalytic strategies for the fractionation and valorization of
lignocellulose"
Von der Fakultät für Mathematik, Informatik und Naturwissenschaften der RWTH Aachen
University zur Erlangung des akademischen Grades eines Doktors der Naturwissenschaften
genehmigte Dissertation
vorgelegt von
Diplom-Chemiker
Philipp M. Grande
aus Bergheim
Berichter: Universitätsprofessor Dr. Walter Leitner
Universitätsprofessor Dr. Marcel Liauw
Tag der mündlichen Prüfung: 05.05.2014
Diese Dissertation ist auf den Internetseiten der Hochschulbibliothek online verfügbar.
iv
This work has been performed by the scientific guidance of Univ. Prof. Dr. rer. nat. Walter
Leitner and under the supervision of Dr. rer. nat. Pablo Domínguez de María. All
experimental studies have been taken place in the research group “Lehrstuhl für Technische
Chemie und Petrolchemie” of the ITMC (Institut für Technische und Makromolekulare
Chemie) at the RWTH Aachen University (Rheinisch-Westfälisch Technische Hochschule
Aachen) within the time from February, 2010 until May, 2013.
It was performed as part of the Cluster of Excellence “Tailor-Made Fuels from Biomass”
(EXC236), which is funded by the Excellence Initiative by the German federal and state
governments to promote science and research at German universities.
Affidavit
I hereby declare, that I have written this work independently and have used only the specified
sources and resourses. The work has not been presented to any examining office, but parts of
the work have already been published in scientific journals or as conference contribution (see
“List of Patents and Publications”).
Eidesstattliche Erklärung
Hiermit erkläre ich, dass ich diese Arbeit eigenständig verfasst habe und nur die angegebenen
Quellen und Hilfsmittel verwendet habe. Die Arbeit wurde bisher keiner Prüfungsbehörde
vorgelegt, wobei Teile der Arbeit in wissenschaftlichen Zeitschriften, Patenten, sowie
Konferenzbeiträgen veröffentlicht wurden (siehe “List of Patents and Publications”).
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ABSTRACT
In the coming decades replacing the depleting fossil resources will demand the development
of integrated biorefinery concepts. This study focuses on the use of biomass as starting
material to produce biomass-derived fuels and commodity chemicals.
Initially, a conceptual biomass pretreatment/fractionation strategy called 1-step OrganoCat
Process is shown and optimized. This biphasic process uses an aqueous reactive phase, with
oxalic acid as catalyst to hydrolyze hemicellulose, and 2-MeTHF as the second phase for the
in situ extraction of lignin. Different acids as catalyst as well as biomass sources showed how
versatile the system is. Recycling the liquid phases and increasing the ratio of substrate to
solvent and catalyst by fourfold made the process more economically feasible. A 3 L-scale
reaction was conducted, showing that scale-up of the process is possible while maintaining
the laboratory scale efficiency.
Second, the newly designed 2-step OrganoCat Process is presented. In this process
hemicellulose is hydrolyzed in seawater, catalyzed by oxalic acid. The resulting solid residue
(cellulose and lignin) is extracted by different bio-based solvents to extract lignin. While the
efficiency of the hydrolysis improved significantly, lignin extraction was not as efficient as
observed in the 1-step OrganoCat Process, making the variation with two steps more suitable
for biomass with low lignin content like algae.
Finally, different strategies for the valorization of the sugar-containing fractions of
lignocellulose are presented. The organic acid-catalyzed as well as the enzymatic hydrolysis
of crystalline cellulose are shown to work efficiently in (concentrated) seawater as solvent.
For further conversion of the resulting glucose, a chemo-enzymatic approach is presented. In
the first step commercially available immobilized glucose isomerase (IGI) is used for the
isomerization of glucose to fructose in seawater. In the second step dehydration of fructose to
HMF is conducted in a biphasic seawater-2-MeTHF system with oxalic acid as catalyst.
Consecutive operation of both steps afforded 64% HMF. Using the raw hemicellulose effluent
from the 1-step OrganoCat Process for fermentation with U. maydis is proven to afford
itaconic acid, not needing any further downstream processing. Also the iron(III) chloride-
catalyzed dehydration of xylose from hemicellulose to furfural is shown to work successfully.
Both reactions are examples for possible subsequent reaction steps in biorefineries.
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KURZZUSAMMENFASSUNG
Für das Ersetzen von fossilen Ressourcen werden in den kommenden Jahrzenten Verbund-
Bioraffineriekonzepte notwendig. Diese Arbeit beschäftigt sich mit der Nutzung von
Biomasse zur Herstellung biogener Kraftstoffe und Rohchemikalien.
Zunächst wurde eine konzeptuelle Strategie zur Vorbehandlung von Lignocellulose, der
1-step OrganoCat Process vorgestellt und optimiert. Dieses zweiphasige Reaktionssystem
beinhaltet eine wässrige, reaktive Phase mit Oxalsäure als Katalysator zur Hydrolyse der
Hemicellulose, und 2-MeTHF als zweite Phase zur in situ Extraktion des Lignins. Die
Vielseitigkeit des Systems wurde durch die Verwendung verschiedener Säuren als
Katalysatoren, sowie durch die Verarbeitung verschiedener Arten von Biomasse demonstriert.
Die Wirtschaftlichkeit des Prozesses wurde durch die Wiederverwendung der flüssigen
Phasen verbessert, welches eine Vervierfachung des Substrat-Lösungsmittel und des Substrat-
Katalysator-Verhältnisses ermöglicht. Die Reaktion wurde unter Beibehaltung der Effizienz
wie im Labormaßstab auf einen 3 L-Maßstab hochskaliert.
Anschließend wird der neue 2-step OrganoCat Process vorgestellt. In diesem Prozess wird
Hemicellulose mit Oxalsäure als Katalysator in Meerwasser hydrolisiert und anschließend der
feste Rückstand (Cellulose und Lignin) mit verschiedenen biogenen Lösungsmitteln extrahiert
um das Lignin zu isolieren. Während die Hemicellulose-Hydrolyse signifikant verbessert
wurde, war die Ligninextraktion weniger effizient, als beim 1-step OrganoCat Process, so
dass der neue Prozess besser für Biomasse mit geringem Ligningehalt, wie Algen geeignet ist.
Schließlich werden verschiedene Strategien für die Verwertung der Zuckerfraktionen aus
Lignocellulose vorgestellt. In (konzentriertem) Meerwasser als Lösungsmittel wird die
effektive, durch organische Säuren, oder enzymatisch katalysierte Hydrolyse von kristalliner
Cellulose gezeigt. Für die Umsetzung der Glucose wird ein chemisch-enzymatischer Ansatz
vorgestellt. Im ersten Schritt wird in Meerwasser mit handelsüblicher, immobilisierter
Glucoseisomerase (IGI) Glucose zu Fructose isomerisiert. Im zweiten Schritt wird in einem
zweiphasigen Meerwasser-2-MeTHF-System mit Oxalsäure als Katalysator Fructose zu HMF
dehydratisiert. Die konsekutive Durchführung beider Schritte resultiert in 64% HMF. Es wird
gezeigt, dass die Fermentation des unraffinierten Hemicellulose-Effluenten zu Itaconsäure mit
U. maydis möglich ist, ohne eine weitere Aufreinigung zu benötigen. Auch die Eisen(III)-
chlorid-katalysierte Dehydratisierung von Xylose aus Hemicellulose zu Furfural wird
vorgestellt. Beide Reaktionen sind Beispiele möglicher Folgeschritte in Bioraffinerien.
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ACKNOWLEDGEMENTS
First of all I would like to thank Prof. Dr. Walter Leitner for giving me the opportunity to
work on such a challenging and amazing topic.
I would like to thank Dr. Pablo Domínguez de María for being endlessly patient and
constantly encouraging during the whole process of learning the scientific work. Furthermore,
our countless fascinating and delighting discussions about my topic and science in general
improved my understanding of being a scientist. I would like to thank him for all his trust in
my abilities and the opportunities of handling also organizational tasks, apart from my
scientific work.
For frequent and fruitful discussions I would like to thank my colleagues and friends Henning
Kayser, Zaira Maugeri, Dr. César A. Urbina-Blanco, Christoph R. Müller, Dr. María Pérez
Sánchez and Thorsten vom Stein. For their fantastic work in the lab I would like to thank my
researching students and lab assistants Sophie Weber, Gent Mehmeti, Christian T. Bergs,
Anne-Christine Kick and Natascha Eickels. Moreover, I would like to thank the whole ITMC
group for the supporting working and social atmosphere. I would also like to acknowledge
Qingqi Yan, Tobias Klement and Bin Zhao for very interesting and fruitful cooperation as
well as Hannelore Eschmann and Julia Wurlitzer from the GC/HPLC-department and Ines
Bachmann-Remy from the NMR-department.
In addition I appreciate all the incredible support from my parents Dr. Silvana and Dr. Bernd
Th. Grande, my sister Sabrina C. and brother-in-law Jörg Grande as well as my close friends
Tamara and Sebastian Wissen, enduring all the good and bad times included during the past
years.
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TABLE OF CONTENTS
1. INTRODUCTION ......................................................................................................................................... 1
1.1. General composition of lignocellulose, classification and sources ............................................................. 2
1.2. Biomass processing / pretreatment. State of the art. ................................................................................... 4
1.2.1. Organosolv process ......................................................................................................................... 5
1.2.2. Steam explosion process ................................................................................................................. 6
1.2.3. Ionic liquid pretreatment ................................................................................................................. 7
1.3. Hydrolysis of polysaccharides .................................................................................................................... 8
1.3.1. Enzymatic hydrolysis ...................................................................................................................... 8
1.3.2. Chemical hydrolysis ........................................................................................................................ 9
1.4. Biomass derived chemicals ...................................................................................................................... 10
1.4.1. 2-Methyltetrahydrofuran (2-MeTHF) ........................................................................................... 10
1.4.2. 5-(Hydroxymethyl)-2-furaldehyde (HMF) .................................................................................... 11
1.4.3. Furan-2-carbaldehyde (Furfural) ................................................................................................... 13
1.4.4. 2-Methylidenebutanedioic acid (Itaconic acid) ............................................................................. 14
2. AIM OF THE THESIS ................................................................................................................................ 16
3. RESULTS & DISCUSSION ........................................................................................................................ 19
3.1. The 1-step OrganoCat-Process ................................................................................................................. 19
3.1.1. A biogenic lignocellulose fractionation concept ........................................................................... 19
3.1.2. Assessment of selectivity in the pulping process: hemicellulose vs. cellulose .............................. 21
3.1.3. Influence of the reaction vessel, pressure and preliminary scale-up ............................................. 24
3.1.4. Influence of type of acid catalyst and type of biomass .................................................................. 28
3.1.5. Influence of the reaction mixture composition .............................................................................. 31
3.1.6. Recycling of effluents ................................................................................................................... 33
3.1.7. Combination of mechanical treatment with pulping ..................................................................... 34
3.2. The 2-step OrganoCat Process ................................................................................................................. 36
3.2.1. A fractionation concept for low lignin content biomass ................................................................ 36
3.2.2. Reaction time screening of the hemicellulose hydrolysis .............................................................. 37
3.2.3. Screening of solvents for the lignin extraction .............................................................................. 38
3.2.4. Recycling of the aqueous effluent ................................................................................................. 40
3.3. Valorization strategies for the carbohydrate fractions from lignocellulose .............................................. 41
3.3.1. Introduction ................................................................................................................................... 41
3.3.2. Salt-assisted organic acid-catalyzed hydrolysis of microcrystalline cellulose .............................. 42
3.3.3. Enzymatic hydrolysis of cellulose with Accellerase® 1500 .......................................................... 45
3.3.4. Chemo-enzymatic conversion of glucose to 5-(hydroxymethyl)-2-furaldehyde ........................... 52
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3.3.5. Synthesis of furfural from xylose using the raw 1-step OrganoCat Process effluent .................... 58
3.3.6. Fermentation of raw carbohydrate effluents .................................................................................. 59
4. SUMMARY AND CONCLUSION ................................................................................................................ 62
5. EXPERIMENTAL PART ............................................................................................................................ 67
5.1. Chemicals ................................................................................................................................................. 67
5.2. General Procedures ................................................................................................................................... 67
5.2.1. Standard procedure for the 1-step OrganoCat Process .................................................................. 67
5.2.2. Standard procedure for the 2-step OrganoCat Process .................................................................. 68
5.2.3. Procedure for the organic acid-catalyzed cellulose hydrolysis in seawater ................................... 69
5.2.4. Standard procedure for the cellulose hydrolysis with Accellerase® 1500 ..................................... 69
5.2.5. Procedure for the synthesis of HMF from glucose ........................................................................ 70
5.2.6. Procedure for the synthesis of furfural from 1-step OrganoCat Process hemicellulose effluent ... 70
5.2.7. Procedure for fermentation of sugars ............................................................................................ 70
5.2.8. Procedure for the mechanical pretreatment with a screw press ..................................................... 71
5.3. Analysis .................................................................................................................................................... 71
5.3.1. High Performance Liquid Chromatography (HPLC) .................................................................... 71
5.3.2. Gas Chromatography (GC) ........................................................................................................... 72
5.3.3. Analysis with Glucose (HK) Assay Kit......................................................................................... 73
5.3.4. Analysis with PAHBAH method .................................................................................................. 74
REFERENCES ........................................................................................................................................................ 75
LIST OF PATENTS & PUBLICATIONS .................................................................................................................... 87
1
1. INTRODUCTION
The depletion of fossil resourses is fostering the research on the utilization of biomass to
produce biofuels for transportation and energy storage, as well as chemical products for
health, care, nutrition and functional applications. In this regard, lignocellulose represents a
highly attractive alternative feedstock, being the largest natural source of organic material and
additionally not competing directly with feeding purposes. Herein, efforts comprise not only
the development and cultivation of new plants, or the use of marginal lands, but also the
exploitation of readily available biomass waste streams (e.g. paper, compost, etc.) are of great
importance. Furthermore, utilizing bio-based and worldwide available chemicals for
processing may be of utmost relevance to be completely (geopolitically) independent of fossil
resourcses and financially competitive.
To achieve this transition from fossil to renewable resources, the development of biorefinery
strategies has been put forward. The integration of low cost fuel production and high value
chemicals production, using biotechnological and chemical conversion, is important to
provide energy and economic efficiency.[1–3] Furthermore, minimalizing transportation and
storage costs by locating biorefineries close to the feedstock (e.g. combination with
agriculture or algae farms) might become another important economic factor, whereby the
valorization of rural local areas may be envisaged.[4,5]
In addition to the concerns leading to the choice of biomass for its use in future biorefineries,
the envisaged enormous consumption of water for processes might become problematic,
especially in fermentative and enzymatic reaction steps. Thus, minimal pollution of waste
water streams has to be targeted by using easily biodegradable chemicals. The sources of
potable water are becoming more and more restricted, which might lead to a competition
between the chemical industry and nutrition. To tackle this issue the utilization of seawater
instead of drinking water might also prove to be a valid alternative, especially in the field of
large-scale biomass processing. For non-coastal cities the need of depurating residual water
for its further use in biorefineries appears as a promising option as well.
Along those lines, the philosophy of “Green Chemistry” can be used as a guideline to design
chemical products and processes, reducing or even eliminating the use and generation of
hazardous substances.[6] 12 principles can be used to evaluate atom and energy efficiency,
hazards, safety, abundance of feedstock, selectivity, biodegradability and pollution
2
prevention.[7] It is expected that such large-scale biorefineries will operate following these
drivers.
1.1. General composition of lignocellulose, classification and sources
Lignocellulose is generally considered to be the inedible part of plants, not suitable for food
production.[8,9] Main components of this composite material are typically cellulose (35-50%),
hemicellulose (15-30%) and lignin (15-30%) with small amounts of other substances (10%,
e.g. proteins, oils and ashes)[10–13] (Figure 1).
Figure 1. Scheme of lignocellulose fiber and the predominant monomers in cellulose, hemicellulose and lignin.
Cellulose is the most abundant biopolymer and source of fermentable sugars on earth. It is a
polydisperse linear polymer of D-anhydroglucopyranose with -1,4-glycosidic bonds,
anhydrocellobiose being the repeating unit, and a degree of polymerization from 100 to
20000. Hydrogen bonds between the polymers lead to a highly ordered supramolecular
structure.[14] The grade of regularity in hydrogen bonds between the cellulose polymers
defines the so-called crystallinity. Cleavage of the -1,4-glycosidic bonds can be achieved
3
acid-catalyzed,[15] by enzyme catalysis (e.g. cellulases)[16] or catalyzed by bases.[17] Herein the
crystallinity is of main importance for the accessibility of the catalyst to the polymer and a
crucial factor for the hydrolysis rate.[18]
Hemicellulose, the second most abundant polysaccharide in nature, is a heteropolysaccharide
consisting mainly of 1,4-linked -D-xylopyranose units (i.e. ca. 90% in beech wood) and to a
lesser part of hexoses (e.g. glucose, mannose, galactose) and other pentoses (e.g.
arabinose).[12,19] Due to its amorphous structure the hydrolysis of hemicellulose is more
straightforward than that of cellulose.[20,21]
Lignin is an amorphous macromolecule, containing the so-called monolignols (e.g. p-
coumaryl, coniferyl and sinapyl alcohol), connected with a huge variety of linkages
(-O-4-linkage being the most dominant).[22] It shows a heterogeneous composition with
primary structure, dependent on the plant and cell-wall region.[23] In contrast to the common
hypothesis of a highly cross-linked three-dimensional structure, caused by random phenoxy
radical-radical coupling, a lamella-like or linear polystyrene-like macromolecule structure
was suggested,[24,25] containing few to no cross-links.[26] However, due to strong self-
association of lignin chains[27] under relatively harsh conditions needed for the extraction,
understanding lignin structure remains a difficult task. In terms of finding applications for the
aromatic compounds in lignocellulose, investigations of phenolic lignin compounds extracted
from different plants as well as synthesized lignin model compounds have been conducted,
showing anti-oxidant activity[28–30] due to free phenolic hydroxyl groups, scavenging
radicals.[31,32] A huge variety of different molecules can be synthesized from lignin. The
production of vanillin via selective oxidation of lignin is one already successfully applied
industrial scale process.[33,34] Also, lignin gives access to the aromatics fraction (benzene,
toluene, xylene, BTX), different macromolecules (e.g. carbon fiber, polymer alloys, etc.) and
miscellaneous monomers (e.g. substituted phenols, styrenes, biphenyls, etc.).[2]
For an economically valuable utilization of lignocellulose as feedstock the selective
fractionation of this composite material into its three main components (e.g. cellulose,
hemicellulose and lignin) is of crucial importance (Figure 2). Besides avoiding problems like
catalyst inhibition, expensive downstream processing can be kept to minimum and thus lower
costs significantly. Furthermore, the separate conversion of each fraction allows application of
appropriate conditions for each substrate and product desired. By such fractionation, an array
of commodities and biofuels can be produced (Figure 2).
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Figure 2. Some possible (bio)chemical pathways for future biorefineries.
1.2. Biomass processing / pretreatment. State of the art.
Biomass processing has been investigated extensively within the last century.[35] For a
complete valorization of lignocellulose a pretreatment has to fulfill several requirements. To
start with, the formation of sugars during the pretreatment or subsequent formation of sugars
by hydrolysis has to be optimized, avoiding the loss or unselective degradation of
carbohydrates. Secondly, the formation of inhibitory byproducts for the subsequent hydrolysis
and fermentation has to be kept to a minimum, and finally the overall process has to be
necessarily cost-efficient.[13] Different physical, physicochemical, chemical, and biological
processes have been developed. In this section, three of the most important processes will be
discussed due to their effectiveness concerning fractionation and improvement of subsequent
processing steps, and due to their special relevance for this PhD thesis.
5
1.2.1. Organosolv process
In the organosolv process, breaking of the internal lignin and hemicellulose bonds of
lignocellulose by hydrolysis with inorganic acids (e.g. HCl, H2SO4, H3PO4) is conducted
typically in aqueous-organic solvent miscible mixtures (Figure 3).
Figure 3. General scheme of the organosolv process.
To avoid possible inhibition of subsequent enzymatic reaction steps a straightforward
recovery of the organic solvent is crucial, and thus low boiling point solvents are preferred
(e.g. methanol, ethanol, acetone, ethylene glycol, triethylene glycol and tetrahydrofurfuryl
alcohol).[36,37] Typical process conditions are 100-250 °C and 10-45 bar,[38] whereby at
elevated temperatures (>185 °C) due to autocatalysis no addition of acids is needed. Organic
acids (e.g. oxalic, acetylsalicylic and salicylic acid) have been studied for the process as well.
Even though the organosolv process shows to be a very promising pretreatment for
lignocellulose, certain drawbacks have to be taken into account. Due to the severe processing
conditions energy consumption is considerably high, which reduces cost-efficiency. Also high
temperature and acidic conditions might lead to considerable degradation of sugars - to
furanics, levulinic acid and humins - and condensation of the lignin. Furthermore, alcohols
used as solvents may form ethers under these severe conditions. This might lead to solvent
loss among organosolv cycles, compromising the economics of the process.[39]
The separation of the three main components in a selective way is crucial for a complete
valorization of lignocellulose. It can be achieved by extensive costly washing steps with
6
organic solvents for the removal of residual lignin. However, the separation of C5 and C6
sugar fractions is often not possible, as they remain mixed in the aqueous solution.
Additionally, the inorganic acids (e.g. HCl, H2SO4) applied cannot be efficiently recycled and
may lead to environmentally unfavorable waste water streams.
1.2.2. Steam explosion process
Another commonly applied pretreatment method for lignocellulose is steam explosion. The
macro structural decomposition of chipped biomass is achieved by treating it with high-
pressure saturated steam followed by rapid decompression (Figure 4).
Figure 4. Scheme of the steam explosion process.
Typical conditions are 160-260 °C and 7-49 bar, expanding to atmospheric pressure after
several seconds to a few minutes.[13,40,41] Addition of acids (e.g. H2SO4, SO2, H2CO3) can
improve enzymatic hydrolysis of pretreated bagasse and removal of hemicellulose. A
decrease in formation of inhibitory compounds for subsequent reaction steps can also be
achieved.[42] Compared to conventional mechanical methods (e.g. mechanical comminution)
steam explosion has low energy requirement and it is very cost efficient due to the low
recycling or environmental costs.[43,44] Additionally, an overall improvement in enzymatic
hydrolysis rate of polysaccharides compared to untreated biomass is observed after steam
explosion .[45] However, significant degradation of hemicellulose sugars and transformation of
lignin is caused due to high temperatures, leading to inhibitory materials for subsequent
7
enzymatic steps.[46] Thus, washing steps of the pretreated biomass are necessary, removing
soluble lignin fragments, hemicellulose sugars and hydrolyzed parts of the cellulose.[35,47] The
steam explosion process itself is very cost-efficient. However, it lacks selectivity and
sustainability for the complete valorization of lignocellulose.
1.2.3. Ionic liquid pretreatment
Many ionic liquids (ILs) are non-flammable,[48,49] have negligible vapor pressure[50,51] and low
melting points due to large cations and anions with low symmetry and dislocated charge.[52]
They can be designed to feature high thermal stability,[53] high conductivity[54] and low
toxicity.[55,56]
ILs have been reported to dissolve and restructure the crystalline cellulose hydrogen bond
network efficiently. This leads to an amorphous-like structure of the cellulose and thus higher
depolymerization rates by increasing the number of solvent-exposed glucan chain hydrogen
bonds with water.[18,57–59] Besides application of ILs as pretreatment, in combination with
acidic metal chlorides (e.g. CuCl2, PdCl2, FeCl3) in 1-ethyl-3-methylimidazolium chloride
([EMIM]Cl) as well as with solid catalysts (e.g. Amberlyst, Zeolite) in
1-butyl-3-methylimidazolium chloride ([BMIM]Cl), direct acidic hydrolysis of cellulose has
been published recently.[60–62] Due to the fact that ionic liquids are known to dissolve lignin
quite effectively,[63] combined with the potential of dissolving cellulose, direct dissolution of
lignocellulose in ILs could be observed, mostly using [EMIM]Cl and [BMIM]Cl and leading
to significant improvement in the subsequent hydrolysis of lignocellulosic pulp.[64–70] A
general scheme of the procedure is shown in Figure 5.
Figure 5. General scheme of the ionic liquod pretreatment of lignocellulose.
8
Even though ionic liquids show promising potential as pretreatment for lignocellulosic
biomass some potential drawbacks exist (e.g. toxicity, cost, corrosiveness, recyclability and
final disposal), reducing their applicability in industrial processes.[51] Their high cost
commands efficient recycling of the ILs which is limited due to their low volatility and thus
might involve expensive extraction with organic solvents to purify recycled ILs.
To overcome the separation issue switchable ILs have been developed, where the IL is formed
by addition of an agent (e.g. CO2) and decomposed by removing this agent.[71] Some
thermally stable ionic liquids (e.g. bis[(trifluoromethyl)sulphonyl]amide) have been shown to
be distillable under reduced pressure, facilitating separation of high boiling substances from
the IL. [72–75] Another promising group of ionic solvents can be obtained by combining two
bio-based, low cost salts (hydrogen bond acceptors, e.g. choline chloride, with hydrogen bond
donors, e.g. levulinic acid, sugar-based polyols, etc.) to form deep eutectic solvents (DES),
[56,76–80] which have been reported to solubilize cellulose as well as lignin efficiently.[76,81]
1.3. Hydrolysis of polysaccharides
Depolymerization of polysaccharides is a necessary step to produce fermentable sugars,
which can subsequently be converted into desired products. While hydrolysis of
hemicellulose is straightforward due to its amorphous structure, cellulose depolymerization
remains challenging. Due to its high crystallinity the cellulose displays a low surface area for
cleavage of -1,4-glycosidic bonds, especially, for enzymatic hydrolysis. In this section, the
most important enzymatic and chemical hydrolysis of cellulose will be presented.
1.3.1. Enzymatic hydrolysis
The depolymerization of cellulose with cellulases and cellulosomes (complexed cellulase
systems) has been investigated intensely during the last decades. The recalcitrance of
lignocellulosic biomass is one of the most challenging issues to overcome.[20,82–92] As
cellulases are relatively expensive enzymes, improving volumetric productivity, utilizing
cheaper substrates to produce the enzymes, enhancing their stability concerning temperature
and pH, raising activity on solid substrates, and higher tolerance to end-product inhibition is
of crucial interest for applying them in industrial processes.[16] Most cellulases are produced
by Trichoderma sp. and Aspergillus sp.[93–95] They contain endoglucanases to expose new
9
chain ends by hydrolyzing intramolecular -1,4-glucosidic bonds, exoglucanases to release
soluble sugar oligomers (e.g. cellobiose, glucose) by cleaving cellulose chain ends and
-D-glucosidases to hydrolyze dissolved cellobiose. Primary hydrolysis, releasing soluble
oligomers from the solid substrate is the rate-limiting step, while secondary hydrolysis takes
place in the liquid phase and produces the desired glucose in a much faster manner.[16]
Inhibition of the cellulase complex by cellobiose and glucose is another major issue to tackle
within cellulose hydrolysis.[96,97] In contrast to the separate hydrolysis and fermentation
(SHF), simultaneous saccharification and fermentation (SSF) has been put forward,
converting sugars directly to less or non-inhibitory products (e.g. ethanol). This leads to
enhanced cellulose hydrolysis rates.[98–100] Moreover, for the enzymatic conversion of
lignocellulose, simultaneous saccharification and co-fermentation (SSCF) can directly
ferment cellulose- and hemicellulose-derived sugars in one reactor with the hydrolysis.[101]
However, a major drawback of both methods may be the difference in optimal temperature for
cellulases (between 45 and 50 °C) and typical fermentative organisms (between 30 and
35 °C). Thus a compromise has to be found for each combination, lowering the respective
reaction rates and stability of the enzymes.[102] Despite these issues, reduced capital costs
(lower amount of enzyme and number of vessels needed) and higher ethanol yields than in
SHF, make SSF and SCFF preferred methods in laboratory and pilot plant studies for ethanol
production.[103,104]
1.3.2. Chemical hydrolysis
In addition to enzymatic methods mentioned above, the efficient depolymerization of
cellulose has been investigated intensively using inorganic acids (like H2SO4, H3PO4 or HCl)
as catalysts. However, due to the high crystallinity of cellulose, high temperatures (>150 °C)
have to be applied, causing degradation of the produced sugars.[90,91,105–108] To enhance the
catalytically available surface area, ionic liquids have been employed due to their ability to
reorder cellulose fibers by breaking hydrogen bond bridges leading to an amorphous-like
structure.[18,57,58,109–116] Another approach along the same lines is the utilization of inorganic
salts (e.g. LiCl, ZnCl2, CaCl2) to swell or even dissolve the cellulose slurry and thus improve
the hydrolysis rate with strong inorganic acids in an analogous way to the ionic liquid
mechanism.[15,117–123] As an alternative to the inorganic acids, and due to their biogenity
organic dicarboxylic acids (e.g. oxalic, maleic and fumaric acid) have also been studied for
cellulose hydrolysis.[62,100,124–132] As a result, dicarboxylic acids were found to perform better
10
than monocarboxylic acids due to their lower pKa values.[62,128] However, the efficient
depolymerization of highly crystalline cellulose requires very high temperatures (>160 °C),
leading again to unselective sugar degradation, lowering the attractiveness of the process.[125]
To tackle these issues, our group recently published a conceptual process, combining organic
acids (e.g. oxalic acid, maleic acid) with inorganic salts (e.g. NaCl, CaCl2 typical at brine
concentrations) under mild conditions (125 °C), leading to efficient hydrolysis of
microcrystalline cellulose (e.g. AVICEL®).[133]
1.4. Biomass derived chemicals
Producing chemicals and fuels from renewable resources as an alternative to fossil resources
and exchanging hazardous solvents and reagents by more benign components has stepped into
the focus of modern research and process design.[1,2,90,134,135] A key step is the reduction of
oxygen content in sugars, derived from lignocellulose, which can be performed by removing
small, highly oxidized carbon molecules (e.g. CO2, formaldehyde and formic acid) as in
fermentative routes (e.g. producing ethanol), via hydrogenolysis, removing oxygen and
forming water with hydrogen or by dehydration of the sugars into a variety of furan-like
compounds.[136] The U.S. Department of Energy (DOE) made a top 10 list of bio-based
chemicals, updated recently.[1,137] In the following, some important bio-based chemicals will
be introduced.
1.4.1. 2-Methyltetrahydrofuran (2-MeTHF)
In order to develop benign processes the utilization of bioderived, non-hazardous chemicals is
crucial. Especially, the choice of solvents is of utter importance as they are the biggest source
of waste in chemical processes, but will be necessarily needed for several steps within
biorefineries as well.[134,135] 2-methyltetrahydrofuran (2-MeTHF) might become an alternative
to substitute less environmentally friendly solvents (e.g. CH2Cl2). 2-MeTHF can be
synthesized from biomass-derived substrates such as furfural or levulinic acid,[138–140] by a
series of reduction-dehydration steps, also leading to other interesting chemicals (e.g.
-valero-lactone, 1,4-pentanediol) as shown in Figure 6.[141]
11
Figure 6. Example of a chemical route for the synthesis of 2-MeTHF from levulinic acid.[141]
In addition to being biogenic, 2-MeTHF also shows promising properties concerning
degradability in the environment where presumably by oxidation and ring-opening, it can be
degraded abiotically by sunlight and air.[142] Also preliminary toxicological studies enable its
use for pharmaceutical processes[143] and make 2-MeTHF a viable option for industrial pilot
plants.[144] Due to its straightforward recovery from aqueous effluents by conventional
distillation,[145] 2-MeTHF is a promising solvent for many reactions. The portfolio of possible
applications substituting hazardous solvents and even improving yields and selectivity is
already broad, including Lewis-acid-mediated ring-opening reactions, organometallic
catalyzed reactions in basic conditions, carbon-carbon and carbon-heteroatom bond
formation, organocatalysis, biocatalysis and biomass processing.[146] However, a potential
drawback of 2-MeTHF is the formation of peroxides when exposed to oxygen. This can be
overcome by addition of stabilizers (e.g. butylated hydroxyl toluene), albeit more research in
that line is necessary for large-scale practical appliactions.[147]
1.4.2. 5-(Hydroxymethyl)-2-furaldehyde (HMF)
For the production of chemicals and fuels from biomass, 5-(hydroxymethyl)-furaldehyde
(HMF) is of huge importance as a platform molecule and has been reviewed regularly
throughout the past century.[136,148–152] Due to the presence of two functional groups (the
hydroxymethyl- and the aldehyde group) many different (bio)chemical conversions of HMF
are possible, encompassing the formation of esters, ethers, halides and oxidation of the
hydroxymethyl group, reactions of the furan ring, as well as reduction, condensation and
oxidation of the formyl group (Figure 7).[152]
Apart from its variety of applications, HMF shows promising properties concerning
biodegradability, as its degradation in microorganisms takes place through oxidation and/or
reduction to the furanic alcohol and acid forms.[153]
12
Figure 7. Scheme of possible products from HMF (adapted from [136]).
The synthesis of HMF can be divided into single-phase systems, biphasic systems and ionic-
liquid-based systems. A very comprehensive overview of the different processes has been
recently published by van Putten et al.[136] Most of the reactions take place in an aqueous
phase due to the ability to dissolve the sugars, whereby homogeneous acids (e.g. HCl, H2SO4,
oxalic acid) and heterogeneous acids (e.g. Amberlyst, metal oxides) as well as metal chlorides
(e.g. CrCl2, CuCl2) are used as catalysts. Due to the tendency of HMF to form humins - that
is, oligomeric condensates of sugars and HMF -, and to prevent rehydration of HMF to
levulinic and formic acid, biphasic systems have been favored, using organic solvents (e.g.
THF, butanol and methyl isobutyl ketone) for the in situ extraction of HMF, which can be
improved by addition of salts or aprotic solvents (e.g. dimethyl sulfoxide), stabilizing
HMF.[154] The mechanism of hexose triple dehydration to HMF is not completely understood
yet. Mechanistical approaches can be divided into those based on cyclic intermediates, where
starting from ketofuranose dehydration of the hemiacetal is followed by two consecutive
-dehydrations, and those based on acyclic intermediates, where a linear 1,2-enediol is
13
formed, followed by two consecutive -dehydrations, ring closure and a final elimination of
water. Additionally, the influence of the solvent (e.g. DMSO, ILs) on the mechanism needs to
be considered.[136]
1.4.3. Furan-2-carbaldehyde (Furfural)
Furfural, another promising platform chemical, can be derived by dehydration directly from
xylose or arabinose, which are main components of the hemicellulose fraction contained in
lignocellulose.[127,155,156] It offers access to a huge variety of further products, including
2-methylfuran, 2-methyltetrahydrofuran, valerate esters, 2-(ethoxymethyl)furan and
2-(ethoxymethyl)tetrahydrofuran furan as well as various C10-C15 coupling products (Figure
8).[157]
Figure 8. Scheme of possible products from furfural.
14
Typically, strong acids (e.g. H2SO4) are used as catalyst in a temperature range from 150 to
220 °C,[158,159] which due to high energy cost and low product yield results in high
investment.[157] Quantum mechanical studies suggest the protonation of C2-OH of xylose and
subsequent ring-contraction as the most favorable mechanistic pathway to furfural in
water[160].
1.4.4. 2-Methylidenebutanedioic acid (Itaconic acid)
In the quest of options to replace petroleum-based chemicals, itaconic acid is a promising
candidate to be incorporated into polymers as an alternative for acrylic or methacrylic acids.
A broad variety of applications can be targeted, including resins, lattices, fibers, detergents,
cleaners and bioactive compounds.[161,162] While the first synthesis of itaconic acid was
conducted chemically by pyrolysing citric acid, followed up by hydrolyzing the
anhydrides,[163] fermentative processes are commercially conducted due to their higher
efficiency.[161] Herein, different organisms have been studied (e.g. Aspergillus sp., Ustilago
sp., Candida sp., Rhodotorula sp.) with A. terreus being the most frequently used organism in
commercial production.[161] Even though different biochemical pathways have been discussed,
the main route is believed to proceed with citric acid and aconitic acid as intermediates which
are then enzymatically decarboxylated to itaconic acid.[164–166] (Figure 9)
15
Figure 9. Scheme of the postulated biosynthetical pathway for itaconic acid production (adapted from [161]).
Due to high cost for refined quality starting materials needed to get reproducible and
(obviously) high productivities, the development of itaconic acid producing organisms which
are able to convert lignocellulose-derived sugars as carbon source is needed.[162]
16
2. AIM OF THE THESIS
The general aim of this thesis is the conceptual development and optimization of strategies for
environmentally-friendly and cost-efficient pretreatment methods enabling the conversion of
lignocellulosic biomass into useful raw materials following the principles of “green
chemistry”. The obtained raw materials will be evaluated for their usability in subsequent
reaction steps, demonstrating their potential application in integrated biorefinery concepts.
To this end, three major topics will be investigated in detail:
1) The 1-step OrganoCat Process (see section 3.1 for details), developed in our group will
be optimized concerning selectivity of the fractionation of beech wood, separating
cellulose and hemicellulose efficiently and avoiding sugar degradation. A comparison
of different reaction vessels and scales as well as their influence on yields of the three
products (that are cellulose, hemicellulose and lignin) will be shown, also demonstrating
a preliminary scale-up of the process. The variability of the fractionation system, using
different (acid) catalysts and processing different kinds of lignocellulose will be
investigated, as well as the possibility of economical improvement by varying the
substrate-to-solvent and -catalyst ratio. The influence of mechanical pretreatment on the
1-step OrganoCat Process will be considered as well.
2) The 2-step OrganoCat Process is a variation of the 1-step OrganoCat Process. Salty
water is used as solvent for the first step to hydrolyze hemicellulose selectively and
efficiently. In a second step lignin is extracted. The potential improvement of the
hydrolysis rate in the first step by utilizing the effect of naturally occurring salts in
seawater on the acid catalyst will be investigated. Different promising candidates (e.g.
organic solvents, water-organic solvent-mixtures, deep eutectic solvents) as solvents for
lignin will be screened for the second reaction step. An improvement of the
substrate-to-solvent and –catalyst will be shown by recycling of the aqueous effluent in
the first reaction step.
17
3) Different strategies for the valorization of cellulose and hemicellulose sugars will be
developed and applied to samples from differently pretreated lignocellulose and
cellulose. Application of abundantly and naturally available seawater as solvent for
chemical (acid catalyzed) and biochemical (enzymatic) depolymerization of
microcrystalline cellulose as well as in acid catalyzed dehydration of sugars (e.g.
fructose, xylose) will be investigated, also combining enzymatic isomerization and acid
catalyzed dehydration for the synthesis of HMF. The utilization of sugar effluents from
the 1-step OrganoCat Process and cellulose depolymerization for furfural production
and fermentation to itaconic acid will be shown to be successful.
19
3. RESULTS & DISCUSSION
3.1. The 1-step OrganoCat-Process
3.1.1. A biogenic lignocellulose fractionation concept
A complete lignocellulose valorization that enables the use of the three main components -
cellulose, hemicellulose and lignin -, is of utmost importance for the set-up of future
biorefineries. Starting from lignocellulosic residues, three valuable raw materials can be
achieved for further (bio)chemical processing. Desirably, lignocellulose fractionation should
be highly selective, delivering non-degraded raw materials. Moreover, produced effluents
should be directly usable without implementing expensive downstream units operations, thus
integrating such steps in a simplified pipeline. As can be inferred from sustainability,
recycling of water, organic solvents and catalysts may become a crucial part when developing
such a system, in which optimized biomass loadings are aggregated. As a result from an ideal
fractionation set-up, a largely-delignified, crystalline cellulose pulp is achieved, which may be
more prone to cellulase-catalyzed depolymerization. Furthermore, due to its amorphous
structure the hemicellulose fraction is straightforwardly hydrolyzed by acids, affording
aqueous xylose.[129] Finally, the achievement of chemically usable lignin is rare in the
literature, due to further condensations, impurities and removal of reactive sites during
pulping processes. Herein, a fractionation system that might deliver manageable and low-
degraded lignin fractions would be an asset for future biorefineries.
Focusing on all these aspects, a novel chemical pulping procedure for lignocellulosic biomass
(the so-called 1-step OrganoCat Process) was conceptually developed in our research
group[167,168] (Figure 10).
20
Figure 10. Process scheme for one-pot fractionation of lignocellulosic biomass (1-step OrganoCat Process),
Figure adapted from [167].
In a one-pot system at 125 °C with two liquid phases (1:1 water:2-MeTHF) and a biogenic
catalyst (0.2 M oxalic acid regarding the aqueous phase) lignocellulose (i.e. beech wood) is
partially depolymerized and separated into its main components cellulose, hemicellulose
sugars (mostly xylose) and lignin. Under these conditions, hemicellulose is selectively
hydrolyzed via acid catalysis leaving cellulose untouched, while lignin is partially extracted in
situ to the organic phase. The reaction is conducted at mild conditions (125 °C, 10 bar) and at
residence times of 6 h[167]. 2-Methyltetrahydrofuran (2-MeTHF) was chosen as the (biogenic)
organic solvent which, due to its low boiling point (79 °C) can be recycled into the system via
distillation. After that distillation solid lignin is obtained. Solid cellulose pulp can be filtered
off from the aqueous phase and oxalic acid can be removed via crystallization from the
aqueous phase and be reused.
21
Taking this biogenic fractionation concept as starting point, several aspects of it were in-depth
studied and improved, with a clear focus on practical lignocellulose fractionation. In the next
sections these results will be described and discussed.
3.1.2. Assessment of selectivity in the pulping process: hemicellulose vs. cellulose
The production of pure aqueous effluents of hemicellulose sugars, whereas cellulose pulp
remained suspended, was the first aspect to be optimized, taking again beech wood as
prototypical lignocellulose. Apart from obtaining different fractions - xylose and pulp - for
their further valorization in biorefineries via fermentation or chemical conversion of sugars, a
low amount of impurities in the fractions is of great importance to overcome cumbersome and
expensive product workup. To improve selectivity for the hydrolysis of amorphous
hemicellulose without depolymerizing crystalline cellulose and thus yield mostly xylose,
different temperatures and reaction times were assessed. The yields of xylose in a temperature
range from 125 to 150 °C at two different reaction times of 3 and 6 h are compared in Figure
11. While the xylose yield after 3 h reaction time improved significantly from 125 to 140 °C,
after 6 h at 125 °C and higher temperatures the theoretical maximum of hemicellulose
contained in beech wood (15-20%[12]) was reached.
22
Figure 11. Production of xylose (wt% compared to total wood weight) in the aqueous effluent after reactions in
the range of 125-150 °C at different reaction times. Conditions: Metal-based high pressure reactor, biphasic
system water/2-MeTHF (1:1, 10 mL total volume), 10 bar CO2, oxalic acid: 0.2 M in the aqueous phase, beech
wood (0.1 mm): 100 g L-1 in the aqueous phase. Xylose and glucose production were analyzed by HPLC (see
experimental part).
Quantitative hydrolysis of hemicellulose can be achieved at 3 h at 140 °C, or for longer
reaction times at lower temperatures. Significant degradation of xylose to furfural could not
be observed in any of those experiments, enabling a broad and promising framework for
lignocellulose fractionation, at least in the case of beech wood. Depending on the biomass
applied, conditions may obviously be adapted, and a compromise yield vs. reaction time set
for each specific case.
Beside xylose, hemicellulose contains other sugars as well (e.g. pentoses, hexoses) whereby
glucose is the second largest fraction in beech wood (ca. 2-5% in relation to total wood
weight).[12] As can be seen from Figure 12, at temperatures between 125 to 140 °C, the
amount of glucose is in the range of 2-3 wt% (in relation to total wood weight). Hence, it may
be assumed that at those reaction conditions glucose originates almost exclusively from
hemicellulose. Interestingly, at higher temperatures (> 150 °C) glucose yield in the aqueous
solution enhances significantly (Figure 12). These results show that cellulose
depolymerization catalyzed by organic acids starts at temperatures of 150 °C, fully consistent
23
with studies in literature for cellulose depolymerization, under similar reaction
conditions.[127,129,131,132,169–172]
Figure 12. Production of glucose (wt% compared to total wood weight) in the aqueous effluent after reaction in
the range of 125-150 °C at different reaction times. Conditions: Metal-based high pressure reactor, biphasic
system water/2-MeTHF (1:1, 10 mL total volume), 10 bar CO2, oxalic acid: 0.2 M in the aqueous phase, beech
wood (0.1 mm): 100 g L-1 in the aqueous phase. Xylose and glucose production were analyzed by HPLC (see
experimental part).
Therefore, as a conclusion from section 3.1.2, the 1-step OrganoCat Process allows
outstanding selectivity patterns, showing that, under optimized conditions, acid
depolymerization proceeds exclusively on (amorphous) hemicellulose, whereas (more
crystalline) cellulose pulp remains suspended in the aqueous solution. Conclusively, to
achieve high selectivity of the biomass fractionation in the case of beech wood, avoiding
significant hydrolysis of cellulose pulp, 6 h at 125 °C as well as 3 h at 140 °C lead to the best
deliverables. Due to the energy efficiency of the system a shorter reaction time of 3 h at only
slightly higher temperature of 140 °C seems favorable. Yet, for further decisions on actual
reaction conditions, a more holistic perspective of a whole biorefinery process should be kept,
providing a broad heat- and mass- integration analysis.[173]
24
3.1.3. Influence of the reaction vessel, pressure and preliminary scale-up
Once the selectivity xylose-glucose was properly assessed and the processing conditions were
improved, the next step was to evaluate the influence of pressure by means of different
reaction vessels in laboratory use, as well as to provide preliminary data on the potential
scale-up of the concept and its future use at operational biorefineries.
In a first array of experiments, the 1-step OrganoCat Process was set-up using different
reaction vessels (Figure 13), reaction scales and applied pressures of CO2. Figure 14
summarizes these results. All fractionation processes were conducted under the same reaction
conditions (solvents, reaction time and temperature, catalyst and wood loadings, similar to
those of Figure 10 and Figure 11).
a) b) c) d)
Figure 13. Different reaction vessels. a) GC-vial; b) Schott-bottle; c) Metal-based high pressure reactor;
d) Glass-made high pressure reactor.
25
Figure 14. Production of xylose and glucose (wt% compared to total wood weight), detected in the aqueous
effluent after reaction for 6 h at 125 °C in different reactors and at different CO2 pressures. Conditions: Biphasic
system water/2-MeTHF (1:1), oxalic acid: 0.2 M in the aqueous phase, beech wood (0.1 mm): 100 g L-1 in the
aqueous phase. Xylose and glucose production were analyzed by HPLC (see experimental part).
Except for the pretreatments performed in GC-vials, in which xylose yields were quite low,
all other reactors led to analogous results, virtually reaching full conversion of the
hemicellulose, compared to average content in beech wood.[12] The lower outcome observed
in GC-vials can be explained by the lack of tightness of such devices, causing lower pressure
and solvent leaching, partially evaporated at reaction temperature. Remarkably, metal- and
glass-made high pressure reactors showed full conversion of hemicellulose at 125 °C after
6 h. For metal-based high-pressure reactors, CO2 was added to prevent the evaporation of the
organic phase into the upper part of the reactor. As observed previously,[167] no activity of
CO2 without addition of oxalic acid was found for lignocelluose fractionation. Albeit an effect
of dissolved CO2 in the aqueous solution lowering pH is possible, it does not appear to be
significant due to comparable results observed in the glass-made high pressure reactor, where
26
no CO2 was added (Figure 14). Therefore, due to better handling and smaller dead-volume
glass-made high-pressure reactors (10 mL and 40 mL) were chosen for further fractionation
studies at bench scale.
As observed in Figure 14, by changing from 10 to 40 mL the total reaction volume, analogous
results in xylose were achieved. Envisaging the importance of scale-up for the type of
fractionation system studied, the 1-step OrganoCat Process was further conducted in a 5 L
metal-based high pressure reactor with glass inlay. These experiments were performed in
cooperation with the Max-Planck-Institute für Kohlenforschung (MPI, Mülheim/Ruhr,
Germany). For this preliminary scale-up, 150 g of beechwood (6 mm) were suspended in
1.5 L of water containing 0.1 M oxalic acid (aq.), and 1.5 L of 2-MeTHF were added. In this
reaction only half of the oxalic acid concentration was chosen as a result from studies
showing the same high activity at both 0.2 M and 0.1 M oxalic acid in the aqueous phase (see
section 3.1.5, Table 2 for details). The high pressure reactor was tightly closed and heated to
140 °C for 3 h while magnetically stirred. Figure 15a shows the setup of the reaction. The
temperature and pressure were monitored during the course of the reaction (Figure 15b).
Heating the reactor to the desired reaction temperature as well as cooling down after reaction
took ca. 2.5 h. Pressure rose to ca. 10 bar at maximum temperature. The workup procedure is
described in the experimental part (section 5.2.1).
27
a) b)
Figure 15. Reaction-setup at the Max-Planck-Institute für Kohlenforschung. a) Filled glass inlay with substrate,
catalyst and solvents; b) Closed high pressure reactor.
After filtrating and washing 53 wt% of initial amount of beech wood (80 g) were obtained as
cellulose pulp which relates nicely to the expected amount of cellulose in beech wood
(35-50 wt%).[10] Gratifyingly, the yield in xylose, which was 24.0 wt% of initial amount of
beech wood (36 g), glucose, which was 4.33 wt% of initial amount of beech wood (6.5 g) and
lignin, which was 12.6 wt% of initial amount of beech wood (19 g) appear to be in the same
range as the small scale reactions (10 mL or 40 mL scale, Figure 16). Higher yields in sugar
fractions of the 3 L-scale experiment might be explained by the higher amount of beech wood
sample used for the experiment, leading to a higher homogeneity of the material.
28
Figure 16. Production of xylose, glucose (wt% compared to total wood weight) in the aqueous effluent and
lignin (wt% compared to total wood weight) in the organic phase, after processing at different reaction scales.
Conditions: Biphasic system water/2-MeTHF (1:1), oxalic acid: 0.1 M in the aqueous phase, beech wood:
100 g L-1 in the aqueous phase, 140 °C, 3 h. Xylose and glucose production were analyzed by HPLC (see
experimental part). Lignin production was determined gravimetrically after removing 2-MeTHF via distillation.
Overall, it can be concluded that at 3 L-scale, the 1-step OrganoCat Process worked at least
as efficiently as in the smaller scales. Based on these first results, it can be envisaged that the
scale-up to a pilot plant process might be feasible. In fact, at higher scale the fractionation
appears to be even more efficient and reproducible than when smaller volumes and loadings
are charged.
3.1.4. Influence of type of acid catalyst and type of biomass
When assessing a novel lignocellulose fractionation system, a crucial aspect is its adaptability
for different reaction conditions and scalability, as reported in previous sections. Furthermore,
another important aspect is to see whether different acid catalysts could be successfully used.
The aim of these studies is the identification of a palette of catalysts that may be recovered or
further used in subsequent reaction steps. Apart from using an easily recyclable catalyst, other
efficient, more creative ways of directly utilizing the effluent of one reaction for the next step
can be taken into account. As an example, for fermentation of aqueous xylose (and other
29
sugars) different buffer systems can be used (e.g. phosphate buffer, acetate buffer, citrate
buffer). Based on those facts, different acids were tested to substitute oxalic acid in the 1-step
OrganoCat Process, which could be subsequently neutralized and later used as buffer for
fermentation of the sugars. The results of the acid screening are shown in Figure 17.
Figure 17. Production of xylose (wt% compared to total wood weight) in the aqueous effluent, detected after
reaction for 6 h at 125 °C with different acids as catalyst. Conditions: Glass-made high pressure reactor, biphasic
system water/2-MeTHF (1:1), acid 0.1 M in the aqueous phase, pH adjusted to 1.2 (determined at RT, pH at
125 °C: ca. 1.2-1.7) with HCl, beech wood (0.1 mm): 100 g L-1 in the aqueous phase. # pH adjusted to 1.8
(determined at RT, pH at 125 °C: ca. 2.0-2.3)[174,175]. Xylose production was analyzed by HPLC (see
experimental part).
Gratifyingly, as shown in Figure 17, the xylose yield remains in the same “full conversion”
range for all different acids, as long as the same pH is employed. For organic acids the actual
pH shifts at 125 °C by approximately 0.5 units higher, as investigated recently in our
group.[175,176] A pH range of 1.2-1.7 seems to be applicable for the efficient fractionation of
beech wood. As observed, the fractionation system follows an acid-driven profile, whereby
xylose production is exclusively dependent on the proton concentration. Raising pH from 1.2-
1.7 to ca. 2.0-2.3 a significant drop in hydrolysis was observed. Analogous dependency of
organic acids and hydrolysis has been observed for cellobiose in our research group as
30
well.[175] From a practical perspective, the possibility of using different (biogenic) acids for
lignocellulose processing opens clearly a broad range of opportunities for chemical
engineering, e.g. valorization of acidic waste effluents from chemical plants, combination of
acid recovery with downstream units operations, etc.
Apart from the acid versatility, another important asset for lignocellulosic pretreatment
strategies is the possibility of adapting them to diverse biomass sources. To investigate this
the same optimized parameters applied for beech wood were evaluated for other types of
biomass. (Table 1)
Table 1. Production of xylose, glucose (wt% compared to total wood weight) in the aqueous effluent and lignin
(wt% compared to total wood weight) in the organic phase, detected after reaction for 3 h at 140 °C. Conditions:
Glass-made high pressure reactor, biphasic system water/2-MeTHF (1:1, 40 mL total volume), oxalic acid:
0.1 M in the aqueous phase, lignocellulose: 100 g L-1 in the aqueous phase. Xylose and glucose production were
analyzed by HPLC (see experimental part). Lignin production was determined gravimetrically after removing
2-MeTHF via distillation.
entry raw material xylose yield
[wt%]
glucose yield
[wt%]
lignin yield
[wt%]
1 beech wood (Fagus sp.) 17 2.3 11
2 Mate tea (Ilex paraguariensis) 5.9 2.1 10
3 reed (Phragmites australis) 19 1.2 14
4 spruce (Picea sp.) 2.6 0.3 0
Mate tea (Ilex paraguariensis) is a very popular tea in South America (e.g. Argentina).
Annually only in Argentina aproximately 280.000 tons of Mate are produced, consumed, and
then discarded.[177] For the valorization of local areas, utilizing lignocellulose should not be
restricted to plants grown for this purpose, but other biomass-based waste products may
provide economic options as well. Hence, leftovers of Mate preparation (once used for food
consumption purposes) were treated with the 1-step OrganoCat Process (Table 1, entry 2).
Hemicellulose was converted completely (content in I. paraguariensis ca. 7 wt%[178]) the
amount of recovered lignin (content in I. paraguariensis ca. 25 wt%[178]) was in the same
31
range as from beech wood (ca. 50% of theoretical content, Table 1, entry 2). The solid
cellulose pulp remained suspended in the aqueous solution. Thus, exhausted leftovers of Mate
tea might be used as a source of xylose, lignin and cellulose for further local valorization.
Obviously, such conceptual approach may be envisaged and extended for other vegetal
sources as well (e.g tea leaves).
Another substrate chosen due to its high availability was reed (Phragmites australis). Due to
its efficient cultivation reed or similar plant types could be a suitable substrate for future
provision of sugars and lignin streams. Reed yields xylose as well as lignin in the same
percentual range of amount compared to the dry biomass weight as beech wood (Table 1,
entry 3).
Spruce (Picea sp.) is a commonly used wood for paper production due to its abundance in
several northern countries. Hence, utilization as lignocellulose source appears convenient.
After conducting the 1-step OrganoCat Process, the obtained amount of xylose was relatively
low (2.6 wt% xylose yield, compared to the dry biomass weight, Table 1, entry 4) and lignin
could not be recovered at all from the organic solvent fraction. Spruce, being a so called soft
wood is characterized by low hemicellulose and high lignin content.[179] Due to the fact that
the 1-step OrganoCat Process relies on depolymerization of hemicellulose to loosen the
attached lignin and extract it into the organic phase, soft woods might not be the appropriate
substrate for a pulping procedure under the mild reaction conditions employed. Applying
more severe conditions (e.g. temperature, pressure) might lead to depolymerization of
cellulose and thus extraction of lignin. In that case aspects to be carefully optimized would be
the degradation of resulting sugars and eventual formation of humins - due to harsher reaction
conditions -, that could decrease the product purity and efficiency of separation. A
compromise between efficient processing conditions and low degradation patterns appears
feasible once optimizations would be conducted.
3.1.5. Influence of the reaction mixture composition
An application-oriented lignocellulose fractionation system must be straightforward and
economically feasible. An important aspect is to provide high selectivity in fractionation
(xylose-cellulose-lignin). Another crucial step is the optimization of the biomass and catalyst
loadings, together with reduction and recycling of water and solvent effluents. Due to the
physical feasibility of stirring a suspension of wood chips in water-2-MeTHF mixtures, there
32
is a limitation for substrate loadings at ca. 100 g L-1 (referring to the concentration only in the
aqueous phase, Table 2, entry 2). Consequently, increasing the biomass loadings up to
200 g L-1 resulted in a reduction in xylose and lignin yield (Table 2, entry 3). Albeit a
compromise may be found, other strategies to increase the amount of biomass processed per
volume would be clearly desirable.
For further improvements, the amount of catalyst used for the reaction was reduced to half.
That resulted in a concentration of 0.1 M instead of 0.2 M referring to the concentration only
in the aqueous phase (Table 2, entry 1). Xylose and lignin were obtained in the same range of
amount as with higher concentration of oxalic acid. Even though oxalic acid could be recycled
via crystallization, decreasing the amount needed for the fragmentation may economically
enhance the process significantly.
In addition, the amount of 2-MeTHF was reduced to assess the optimal solvent-to-water ratio.
Thus, when half of the amount of 2-MeTHF (Table 2, entry 4) was added, xylose yields in the
aqueous solutions remained complete. Albeit the lignin yield dropped slightly (from 13 to
10 wt%) the fractionation of beech wood shows to be still possible with half the amount of
2-MeTHF (water:2-MeTHF 2:1 v/v).
Table 2. Variation of reaction mixture composition. Production of xylose, glucose (wt% compared to total wood
weight) in the aqueous effluent and lignin (wt% compared to total wood weight) in the organic effluent, detected
after reaction for 3 h at 140 °C. Conditions: Glass-made high pressure reactor, biphasic system water/2-MeTHF
(10 mL total volume), oxalic acid: different concentrations in the aqueous phase, beech wood (6 mm): 100 g L-1
in the aqueous phase. Xylose and glucose production were analyzed by HPLC (see experimental part). Lignin
production was determined gravimetrically after removing 2-MeTHF via distillation.
entry catalyst loading
[mol L-1]
water:
2-MeTHF
wood loading
[g L-1]
xylose yield
[wt%]
glucose yield
[wt%]
lignin yield
[wt%]
1 0.1 1:1 100 17 1.2 14
2 0.2 1:1 100 18 1.2 13
3 0.2 1:1 200 15 2.0 9
4 0.1 1:0.5 100 18 1.2 10
33
3.1.6. Recycling of effluents
Apart from decreasing (thus optimizing) the amount of 2-MeTHF, another way to further
improve the solvent- and catalyst-substrate ratio is the recycling of the liquid phases by
adding new biomass, once the first (solid) loading has been processed. As stated above, there
is a physical limitation to the addition of solid biomass, which can be assumed to be in the
range of 100-150 g L-1 (referring to the aqueous phase for beech wood, 6 mm, Table 2). Thus,
to improve this, after conducting the reaction for 3 h at 140 °C the liquid effluents (that is, the
aqueous phase with hemicellulose sugars and oxalic acid, as well as the organic 2-MeTHF
phase with lignin) were recovered via filtration of the reaction mixture - removing solid
pulp -, and then a new beech wood loading (100 g L-1, referring to the aqueous phase) was
added (Figure 18).
Figure 18. Concept for recycling liquid phases in the 1-step OrganoCat Process.
The envisaged filtration-addition approach was performed up to four cycles (with a total wood
addition of 400 g L-1, referring to the aqueous phase). Gratifyingly, it led to a remarkable
improvement in hemicellulose sugar concentrations, producing an aqueous solution of ca.
34
60 g xylose L-1 (Figure 19). An almost linear increase in xylose concentration shows that,
applying state of the art filtration and distillation techniques, the direct recycling of the
solvents can yield much higher concentrations in product streams, resulting highly beneficial
for further reaction steps (e.g. fermentation,[180] dehydration[155]).
In addition to xylose deliverables, an increase of lignin in the organic phase was observed as
well (Figure 19). It must be noted that due to the experimental setup a loss of liquid phase
during filtrations could not be avoided. Consequently, the improvement of lignin yield, which
was determined gravimetrically, was only about twofold. However, this aspect can easily be
circumvented by optimization of the recovery process.
Figure 19. Recycling of liquid phases. Concentration of xylose, glucose in the aqueous effluent and lignin in the
organic effluent, detected after reaction for 3 h at 140 °C. Conditions (1st cycle): Glass-made high pressure
reactor, biphasic system water/2-MeTHF (1:1, 10 mL total volume), oxalic acid 0.1 M in the aqueous phase,
beech wood (6 mm): 100 g L-1 in the aqueous phase. following cycles: filtration (paper filter, MN615), addition
of new beech wood (6 mm): 100 g L-1 in the aqueous phase, 140 °C, 3 h. Xylose and glucose concentrations
were analyzed by HPLC (see experimental part); lignin production was determined gravimetrically after
removing 2-MeTHF via distillation and concentration calculated.
3.1.7. Combination of mechanical treatment with pulping
Prior to chemical pretreatment, the particle size has to be reduced by mechanical processing.
Phragmites australis was mechanically prepared at the Aachener Verfahrenstechnik (AVT,
RWTH Aachen University). One sample was processed only applying the cutting mill and
35
one was subsequently treated with the screw press. The screw press applies a shearing force,
decreasing moisture, physically changing the fibers and increasing the specific surface
area.[181]
Figure 20 shows the yields of xylose, glucose and lignin after cutting mill, chopping reed into
particles of 10 mm length, in comparison to additional screw-pressing after cutting mill.
Under optimized conditions of the 1-step OrganoCat Process xylose yield was about 18 to
20% and lignin yield about 12%. Here, no significant change was observable, comparing
cutting-mill reed to screw-press reed.
Figure 20. Impact of mechanical pretreatment on 1-step OrganoCat Process. Production of xylose, glucose
(wt% compared to total wood weight) in the aqueous effluent and lignin (wt% compared to total wood weight) in
the organic phase, detected after reaction for 3 h at 140 °C. Conditions: Glass-made high pressure reactor,
biphasic system water/2-MeTHF (1:1, 40 mL total volume), oxalic acid: 0.1 M in the aqueous phase, reed
(10 mm): 100 g L-1 in the aqueous phase. Xylose and glucose production were analyzed by HPLC (see
experimental part). Lignin production was determined gravimetrically after removing 2-MeTHF via distillation.
Further investigations at lower temperatures or shorter reaction times might show stronger
impact on separation of the lignocellulose components and thus lead to lower overall energy
costs. Likewise, the set-up of mechanical steps might contribute to homogenize the size of
biomass, and thus to enable more reproducible results during the subsequent (bio)chemical
steps. An investigation of the subsequent enzymatic hydrolysis of the resulting pulp after the
combination of cutting mill or screw press and 1-step OrganoCat Process will be discussed in
section 3.3.3.
cutting mill cutting mill + screw press
36
3.2. The 2-step OrganoCat Process
3.2.1. A fractionation concept for low lignin content biomass
For certain (bio)chemical segments contamination with organic solvents may become
unacceptable. Additionally, contamination of waste water effluents with organic solvents
represents an important ecological issue. Waste must be completely removed before recycled
water is placed back into the ecosystem. Albeit 2-MeTHF shows preliminary promising
features in biodegradability,[143] avoiding contamination completely or reducing it to a
minimum might become a desirable option in specific applications whereby sugars (e.g.
xylose) are envisaged for food industry or analogous “sensitive” markets.
To meet the requirements for those applications, the 2-step OrganoCat Process was designed
(Figure 21). In the first step, 100 g L-1 of beech wood are stirred at 140 °C for 3 h in seawater
as the sole liquid phase with 0.1 M oxalic acid as catalyst.
In a second step, after cellulose pulp filtration to achieve aqueous xylose, the obtained solid
cellulose-lignin mixture was extracted with a broad range of different organic solvents and
deep-eutectic-solvents (DES) which are known to dissolve lignin to a significant extent.[76]
Figure 21. Process scheme of the "2-Step-OrganoCat-Process".
37
For biorefineries a large consumption of water will be inevitable, due to its use as solvent in a
number of (bio)chemical conversions, and especially in enzymatic and fermentative reaction
steps. Whereas ca. 70% of the earth is covered with water, only 3% of that water is potable
water,[182] leading to an unfavorable competition between drinking water and usage for
industrial processes. To avoid this competition, utilization of water-depurated effluents from
municipal wastes for biorefineries could open up interesting opportunities. In parallel to that,
seawater is abundantly available and easily accessible, especially in coastal regions. As
described in literature seawater contains different salts (mostly NaCl, ca. 3-4 wt%),[183] which
may improve acid catalyzed cellulose hydrolysis rate.[133]
3.2.2. Reaction time screening of the hemicellulose hydrolysis
In a first step, reaction time screening of the hydrolysis of hemicellulose was assessed. As can
be seen in Figure 22, xylose yield reached a maximum already after 1 hour. Such fast results
in depolymerization are consistent with results observed in depolymerization of crystalline
cellulose in seawater and salty effluents catalyzed by organic acids (see also section
3.3.2).[133] Conducting the fractionation process at longer times (up to 3 hours) did not result
in higher content of xylose, aligned with the assumption that full hydrolysis of hemicelluloses
was reached after 1 hour already.
Figure 22. Reaction time screening of the 2-step OrganoCat Process. Production of xylose and glucose (wt%
compared to total wood weight) in the aqueous effluent. Conditions: Monophasic system (seawater, 10 mL total
volume), oxalic acid: 0.1 M, beech wood (6 mm): 100 g L-1, 140 °C. Xylose and glucose production were
analyzed by HPLC (see experimental part).
38
3.2.3. Screening of solvents for the lignin extraction
The solid cellulose pulp from the first step was separated from the aqueous phase by filtration
and washed with water until neutral pH. Then, lignin was extracted from the solid residue
(containing cellulose pulp and lignin) by adding 2-MeTHF and stirring for 3 h at room
temperature. Then, the organic solvent was removed via distillation. As observed, lignin yield
in the second step of the 2-step OrganoCat Process was not as high as in the 1-step
OrganoCat Process (Figure 23). To investigate this further, different mixtures of water and 2-
MeTHF, as well as other organic solvents were screened to see whether lignin yields could be
improved (Figure 23).
Figure 23. Screening of different solvents for lignin extraction after the first step. Conditions first step:
Monophasic system (seawater, 10 mL total volume), oxalic acid: 0.1 M, beech wood (6 mm): 100 g L-1, 140 °C,
3 h. Lignin extraction was conducted with different solvents at different reaction times and temperatures; yield
was determined gravimetrically after removing organic solvent via distillation. The amount of lignin was
determined gravimetrically and is given in wt% compared to total wood weight. *Lignin was precipitated by
addition of water to the DES, then filtered and redesolved in 2-MeTHF; organic solvent was removed via
distillation.
39
Ethanol and 1-butanol are known as good solvents for lignin from the organosolv process,[184]
as well as 2-MeTHF from the 1-step OrganoCat Process. The mixture of lactic acid with
betaine (2:1 mol:mol) forms a DES and has also been reported as a highly efficient solvent for
lignin dissolution.[76] As can be seen, all extractions led to analogous lignin yields (4-6 wt%).
Yet, similar yields to those observed with the in situ extraction from 1-step OrganoCat
Process could not be achieved in any of the experiments performed (Figure 23). This leads to
the conclusion that the set-up of in situ biphasic systems is crucial for achieving high lignin
yields. It could be hypothesized that in the absence of organic solvent lignin remains in acidic
aqueous solutions over longer reaction times, thus being prone to further irreversible
(re)polymerizations. More research in that direction is needed to validate these considerations,
which might bring a competitive advantage for the 1-step OrganoCat Process compared to
more classic organosolv treatments (typically performed in monophasic solutions under
strong acidic conditions).
In any case, the xylose production in this monophasic system remains outstanding, making it
especially appealing for biomass containing low lignin proportions (e.g. grasses), or for other
promising raw materials (e.g. algae). In fact, the improvement of hydrolytic efficiency when
using seawater shows great opportunity for processing seawater-based algae, not needing to
remove their salty-water content, and thus simplifying treatment processing. Additionally, the
lack of contamination with organic solvents in the first step shows high potential for
application of the sugars in food industry. Also in the subsequent waste (sea)water treatment
the absence of organic solvents is beneficial.
40
3.2.4. Recycling of the aqueous effluent
Based on previous results considering quantitative xylose yields, further improvements in the
solvent- and catalyst-to-biomass ratio were considered, by recycling the liquid phase in the
first step, analogous to the procedure discussed in section 3.1.5 for the 2-step OrganoCat
Process. A similar benefit for further biomass processing, like in the 1-step OrganoCat
Process (section 3.1.5), was targeted. For this purpose, after conducting the reaction for 3 h at
140 °C, the liquid effluent (that is, the 0.1 M oxalic acid solution in seawater) was recovered
via filtration of the reaction mixture – removing solid pulp –, and then new beech wood
(100 g L-1) was added. The filtration-addition approach was performed up to four cycles.
As can be seen from Figure 24, the xylose concentration increases almost linearly with every
cycle, leading to a remarkable xylose concentration of ca. 90 g L-1 after 4 cycles. This nicely
shows that also with this monophasic variant, higher xylose concentrations can be achieved
straightforwardly, significantly improving economics of this and following processing steps.
Figure 24. Recycling in 2-step OrganoCat Process. Concentration of xylose, glucose in the aqueous effluent,
detected after reaction for 3 h at 140 °C. Conditions (1st cycle): monophasic system (seawater, 10 mL total
volume), oxalic acid: 0.1 M, beech wood (6 mm): 100 g L-1, following cycles: filtration (paper filter, MN615),
addition of new beech wood (6 mm): 100 g L-1, 140 °C, 3 h. Xylose and glucose concentrations were analyzed
by HPLC (see experimental part).
41
3.3. Valorization strategies for the carbohydrate fractions from lignocellulose
3.3.1. Introduction
The valorization of all components of lignocellulose (e.g. cellulose, hemicellulose and lignin)
might become important for future biorefinery concepts. To simplify subsequent reaction
steps it is crucial to minimize downstream processing, as well as using abundantly available
and thus economic solvents and catalysts. Thus, conducting subsequent process steps after the
fractionation of lignocellulose with the not further purified effluents could demonstrate further
benefits of this process.
Based on these considerations, in this section some valorization strategies for cellulose and
xylose will be presented (Figure 25). Herein, the focus is on valorization of the carbohydrate
fractions lignin will remain out of the scope of this work. Processing of cellulose can be
achieved via enzymatic hydrolysis[104] as well as by setting up acid-catalyzed
depolymerizations.[185] Obtained sugar effluents can subsequently be converted chemically or
fermentative to synthesize so called platform molecules, such as HMF, furfural, itaconic acid,
etc. These molecules are then used to produce desired products, such as fuels, fine chemicals,
etc. (see section 1.4 for details).[141]
42
Figure 25. Flow scheme envisaged for the valorization of the 1-step OrganoCat Process product streams
3.3.2. Salt-assisted organic acid-catalyzed hydrolysis of microcrystalline cellulose
Based on findings showing that organic acids (e.g. oxalic acid, maleic acid) are able to
depolymerize amorphous cellulose[129] and the beneficial effect of adding salts, improving the
hydrolysis rate of acid-catalyzed (1-4)glycosidic bond cleavage[15] a proof-of-concept
process was designed for more crystalline celluloses.[133,167] Herein, microcrystalline cellulose
(AVICEL®) was hydrolyzed efficiently under mild conditions, combining organic acids with
NaCl.
43
Figure 26. Reaction scheme of the salt-assisted organic acid-catalyzed hydrolysis of microcrystalline cellulose.
Organic-acid catalyzed hydrolysis of cellulose in (concentrated) seawater
Abundantly available seawater was introduced as a natural source of NaCl (ca. 3-4%). Figure
27 compares hydrolysis of AVICEL® in different reaction media. Without the addition of salt
(white bars), the activity was close to zero under the mild reaction conditions applied (125 °C,
6 h). However, using seawater instead of distilled water improved hydrolysis rate significantly
(green bars). In drinking water production, seawater is commonly desalinated, leading to
waste water with high salt content,[186] which could be beneficial for the hydrolysis rate of this
reaction. To simulate this high-salt-content-seawater, seawater was concentrated fivefold
(blue bars) by distillation, reaching about the same NaCl loading as in brine. Gratifyingly, in
concentrated seawater the reaction rate is in the same range as with brine (grey bars). This
opens up a great opportunity to utilize this waste stream for sugar production out of
microcrystalline cellulose.
44
Figure 27. Impact of salt content in seawater on hydrolysis rate of organic acids in crystalline cellulose.
Conditions: AVICEL® cellulose: 20 g L-1, 0.1 M maleic acid, 125 °C, 6 h. Samples were taken after 0, 1, 4 and
6 h. Different solvents were used: distilled water, seawater, 5-fold concentrated seawater, brine. Concentrations
of reducing sugars were determined with PAHBAH method (see experimental part).
Organic-acid catalyzed hydrolysis of ionic liquid-pretreated cellulose
Another way to lower crystallinity in cellulose, enabling straightforward hydrolysis, is
dissolution and re-precipitation in ionic liquids (ILs).[187] To analyze this effect, the hydrolysis
rates of differently pretreated samples, provided by the Institut für Technische und
Makromolekulare Chemie (ITMC, RWTH Aachen University),[187,188] were compared (Figure
28). Samples of -cellulose were dissolved in different ILs and then precipitated by addition
of water. Cellulose was recovered by filtration and then dried. In comparison with only dried
-cellulose an improvement in initial reaction rate of hydrolysis with 0.1 M oxalic acid was
observed for [DMIm]Butyrate and even more after pretreatment with [EMIM]Butyrate.
Due to preliminary results, showing a positive effect in the hydrolysis rate caused by the
addition of NaCl,[133] also the impact of IL pretreatment under those conditions was
investigated. The initial reaction rate of oxalic acid with NaCl to hydrolyze untreated -
45
cellulose is already higher than without NaCl. Interestingly, after dissolution and
reprecipitation in [DMIm]Acetate the initial hydrolysis rate was doubled.
Figure 28. Effect of pretreating -cellulose with different ionic liquids. Conditions: pretreated -cellulose:
20 g L-1, 0.1 M oxalic acid, 125 °C, 6 h; # addition of 30 wt% NaCl compared to the aqueous phase. Initial
reaction rate was calculated from reducing sugar concentration, determined with PAHBAH after reaction (see
experimental part).
3.3.3. Enzymatic hydrolysis of cellulose with Accellerase® 1500
Another very efficient and selective way to hydrolyze cellulose to glucose is performed by
cellulases (see section 1.3.1). Herein, crystallinity[18] and content of associated material to
cellulose (i.e. lignin)[189] are of major importance, as well as capillary structure of cellulose
fibers, molecular arrangement and degree of water swelling of the substrate.[190] Moreover,
another important factor is product inhibition of cellulases and -glucosidases, typically
caused by increasing (soluble) concentrations of cellobiose and glucose[97]. These inhibitory
patterns can be overcome by applying simultaneous saccharification and fermentation
(SSF),[99] converting glucose in-situ through fermentation, thus removing the inhibitor (see
section 1.3.1 for details).[13]
46
The commercially available cellulase Accellerase® 1500 (kindly donated from Genencor®
International Corporation) was used to determine hydrolysis activity of different cellulose
substrates. It is actually an enzyme “cocktail”, intended specifically for lignocellulosic
biomass processing industries. The enzyme mixture is produced with a recombinant strain of
Trichoderma reesei and includes genetically improved exoglucanases, endoglucanases,
hemicellulases as well as -glucosidases at optimized proportions.
Influence of substrate and catalyst loading
In order to define the optimal conditions for hydrolysis of cellulose catalyzed by Accellerase®
1500 (see experimental part) different substrate and catalyst loadings were firstly screened.
Table 3 compares initial reaction rates at substrate loadings from 10 to 30 g L-1 and catalyst
loadings from 0.2 to 5 vol% (v/v). Best results were achieved at substrate loadings of 20 g L-1
(Table 3, entry 2). Higher cellulose loadings lead to lower reaction rates, presumably due to
substrate inhibition.[191] Likewise, the addition of more enzyme per solvent shows higher
activity but reaches maximum activity at about 1 vol% of Accellerase® 1500 (v/v) (Table 3,
entries 4-6). Hence, for further reactions 20 g L-1 AVICEL® and 1 vol% Accellerase® 1500
were chosen as standard conditions for further uses with cellulose samples.
Table 3. Initial reaction rates with different substrate and catalyst loadings. AVICEL® cellulose, 0.1 M citrate
buffer (pH = 4.5), 50 °C, 3 h. Initial reaction rate was calculated from glucose concentrations, determined with
Glucose (HK) Assay Kit after reaction (see experimental part).
entry substrate loading
[g L-1]
Accellerase® 1500
loading* [vol%]
initial reaction rate
[g L-1 h-1]
1 10 0.2 1.6
2 20 0.2 3.7
3 30 0.2 1.7
4 20 0.6 5.7
5 20 1.0 5.0
6 20 5.0 6.3
* related to the amount of solvent
47
Multistep-hydrolysis of AVICEL® with Accellerase® 1500
Due to product inhibition during enzymatic hydrolysis of cellulose, glucose has to be
continuously removed to reach a complete cellulose depolymerization. To assess whether
commercial cellulases would hydrolyze microcrystalline cellulose completely, a sample of
AVICEL® was hydrolyzed in multiple steps. After each step the aqueous glucose solution was
removed via filtration, the residual cellulose was washed with water and new Accellerase®
1500 and solvent were added. Figure 29 shows initial reaction rates of the different cycles.
Reaction rates diminish constantly over hydrolytic cycles. This could be explained by
different grades of crystallinity of the substrate.[192,193] First, less crystalline regions (of a
microcrystalline cellulose) are hydrolyzed. Herein, depolymerizing the solid cellulose is the
rate-determining step while hydrolyzing soluble oligomers is relatively fast. In the following
cycles, only the higher crystalline regions of cellulose are left so reaction rate decreases. In
the 5th cycle, cellulases showed almost no activity on the residue of cellulose particles with
highest crystallinity. Finally, in the 6th cycle no glucose could be determined anymore,
supporting that no further hydrolysis of the cellulose particles was possible upon such highly
recalcitrant cellulosic materials.
Figure 29. Initial reaction rates observed along the multistep hydrolysis of AVICEL® catalyzed by cellulases.
Conditions: AVICEL® cellulose: 20 g L-1, 0.1 M citrate buffer (pH=4.5), 1 vol% Accellerase® 1500, 50 °C, 3 h.
Initial reaction rate was calculated from glucose concentrations, determined with Glucose (HK) Assay Kit after
reaction (see experimental part).
48
Hydrolysis of cellulose pulps obtained from the 1-step OrganoCat Process
The cellulose pulp of the 1-step OrganoCat Process was hydrolyzed with Accellerase® 1500.
Figure 30 compares produced sugar concentration of Accellerase® 1500 with different
cellulose pulps. As benchmark commercially available AVICEL® cellulose was used.
Figure 30. Kinetics of the hydrolysis with Accellerase® 1500 of AVICEL® cellulose, beech wood (6 mm) and
beech wood pulp (after 1-step OrganoCat Process). Conditions: substrate loading: 20 g L-1, 0.1 M citrate buffer
(pH=4.5), 1 vol% Accellerase® 1500, 50 °C. Reducing sugar concentrations were determined with PAHBAH
after reaction (see experimental part).
The enzymatic cocktail converts about half of the microcrystalline cellulose (compared to
theoretical full conversion of 20 g L-1 cellulose yielding ca. 22 g L-1 glucose) within 3 h. At
about 10 g L-1 product inhibition takes place, consistent with the literature[97] and practically
avoided by simultaneous saccharification and fermentation (SSF) processes.[99] Conversely,
when using non-pretreated beech wood as substrate virtually no conversion was observable.
After applying the 1-step OrganoCat Process cellulase reaction rate increased significantly.
Compared to AVICEL®, the activity is slightly lower which might be explained by residual
49
lignin in the cellulose pulp which may inhibit the cellulases. However, hydrolysis rate was
greatly improved compared to the untreated beech wood.
Furthermore, the hydrolysis rates of the differently pretreated substrates (see section 3.1.7)
after conducting the 1-step OrganoCat Process were investigated. Herein, the reaction rate of
screw-pressed reed pulp after 1-step OrganoCat Process was significantly higher than that of
cutting-mill reed pulp (Figure 31).
Figure 31. Kinetics of hydrolysis with Accellerase® 1500 of cut and screw pressed reed and only cut reed before
and after 1-step OrganoCat Process. Conditions: cellulose pulp: 20 g L-1, 0.1 M citrate buffer (pH = 4.5), 1 vol%
Accellerase® 1500, 50 °C. Glucose concentrations were determined with Glucose (HK) Assay Kit after reaction
(see experimental part).
A possible explanation for this improvement could be the enhancement of the available
surface area and the decrease in cellulose crystallinity by screw press processing, presumed to
be analogous to ball milling treatment.[190] Even though without chemical pretreatment
compared to cutting-mill reed no change in hydrolysis rate was observable, after conducting
the 1-step OrganoCat Process, the higher surface area enhanced the hydrolysis rate
50
significantly. Overall, it appears that the combination of mechanical pre-steps with further
1-step OrganoCat Process may deliver promising synergies for a more adequate biomass
processing and valorization.
Enzyme catalyzed hydrolysis of cellulose in seawater
Scientific literature offers a broad number of examples of enzymes and microorganisms (e.g.
halofilic), able to live in high concentration of salt (e.g. NaCl), even up to saturated
solutions.[180,194] Some biotechnological applications have been reported,[180] and recently also
fermentative applications producing succinic acid from glucose in seawater were
published.[183] With regard to cellulases, there are some examples of these enzymes dealing
with high saline concentrations or ionic liquids.[195–197] Yet, surprisingly, the direct use of
cellulases in (concentrated) seawater for depolymerization purposes had never been assessed
or reported in the literature.
With such state of the art in mind seawater was used instead of distilled water to prepare the
buffer for the hydrolysis of cellulose with commercial glycosidic “cocktail”
Accellerase® 1500 (see experimental part). The first step was to evaluate whether cellulase
activity in seawater was analogous to that observed in distilled-water-based standard buffer.
Figure 32 compares the kinetics of hydrolysis of AVICEL® cellulose in distilled water (white
bars) and in seawater (grey bars). Gratifyingly, no significant difference was observable when
seawater was used instead of distilled water. Therefore, some commercially available
cellulase “cocktails” might be directly used in seawater as reaction media for the provision of
fermentable sugars.
Furthermore different concentrations of seawater were compared, simulating waste stream
from e.g. desalination plants. Higher concentrations were achieved by distillation of seawater
to the desired salt concentration. At twofold concentrated seawater (blue bars), the enzymatic
“cocktail” displayed a slightly lower performance. Furthermore, with fourfold concentrated
seawater (green bars) a significant reduction of hydrolysis rate was observed, presumably due
to the higher osmolarity of the solvent. However, cellulose was still hydrolyzed efficiently,
even at this high salt concentration. Accellerase® 1500 is a cellulase “cocktail”, composed of
several glycosidases, optimized in performance and proportion. Thus further investigations
with isolated enzymes are needed to understand which components are affected by the salt. In
combination with the knowledge about halofilic cellulases this might open up a possibility for
51
genetic evolution of cellulases, to support occurring high salt concentrations in concentrated
seawater, and thus to integrate this reaction media into future biorefinery plants.
Figure 32. Effect of higher concentration of seawater on cellulose hydrolysis. Conditions: cellulose: 20 g L-1,
0.1 M citrate buffer (pH=4.5), 1 vol% Accellerase® 1500, 50 °C. Glucose concentrations were determined with
Glucose (HK) Assay Kit after reaction (see experimental part).
52
3.3.4. Chemo-enzymatic conversion of glucose to 5-(hydroxymethyl)-2-furaldehyde
In the scope of utilizing biomass-derived chemicals and fuels, several molecules have gained
importance over the past decades. In this respect, 5-(hydroxymethyl)-2-furaldehyde (HMF) is
considered a platform chemical[1,137] due to its broad number of applications for further
conversions (see also section 1.4.2).[139,141,152,198–208] It can be directly derived from glucose or
fructose, upon a triple dehydration, and a variety of syntheses have been published using
homogeneous, heterogeneous, acid or metal catalysts[202].
Effluents of biomass-derived glucose are mostly aqueous, as cellulose hydrolysis must be
obviously conducted in aqueous solutions. Hence, further water removal as work-up step
might be considered energetically not efficient for this chemical segment. Thus, most of the
reported HMF syntheses take place in aqueous media. However, straightforward production is
challenging due to the instability of HMF,[209] especially in high concentrations and in
presence of sugars, where the so-called humins (e.g. oligomeric condensates of sugars and
HMF) are formed. In addition, in water media at severe conditions there is the possibility of
HMF rehydration, leading to formic and levulinic acid formation. To overcome this, several
strategies have been investigated: a) removal of HMF from the reactive phase into a second
organic phase, such as MIBK, 2-butanol or THF, b) addition of cosolvents, such as DMSO,
stabilizing sugars and HMF, c) utilization of non-aqueous solvents, such as ionic liquids.[136]
HMF can be produced either from glucose or fructose. The synthesis using glucose as raw
material that is obviously preferred for lignocellulose-based biorefineries, typically requires
more severe conditions (e.g. high temperature, pressure, etc.).[202] However, this enhances side
reactions and thus reduces selectivity for HMF. Consequently, new approaches, avoiding high
concentration (and thus degradation) of HMF in the reaction mixture but still converting the
complete sugar substrate selectively, have to be developed.
Herein a combined chemo-enzymatic synthesis of HMF from D-glucose is presented (Figure
33). In a first step, D-glucose is enzymatically converted to D-fructose by means of a
commercially available immobilized glucose isomerase (IGI, see Experimental Part).
Isomerization takes place in seawater where the immobilized (suspended) enzyme can be
easily recycled via filtration. In a second step, dehydration of D-fructose, catalyzed by oxalic
acid dissolved in seawater leads to HMF, which is in situ extracted into a 2-MeTHF phase and
can easily be isolated via distillation, while 2-MeTHF is recycled into the system. Oxalic acid
53
in the aqueous phase can be recycled and residual sugars can be either led back into the
enzyme tank or fermented.
Figure 33. Reaction scheme of the chemo-enzymatic conversion of D-glucose to HMF.
Conversion of D-fructose to 5-(hydroxymethyl)-2-furfural
At first, the oxalic-acid-catalyzed dehydration of D-fructose to HMF in a biphasic system was
investigated. The substrate and the catalyst were dissolved in water and a second organic
phase of 2-MeTHF was added for the in situ extraction of HMF. Table 4 compares HMF
yields after synthesis in different solvents. Adding salts to the acid-catalyzed dehydration of
sugars has been reported to be beneficial by decreasing solubility of organic species in
water.[210] Consequently, addition of NaCl (20 wt%, referring to the aqueous phase) in the
synthesis of HMF led to higher yields (Table 4, entry 2) than that observed for reactions
conducted without NaCl (Table 4, entry 1). In general, results are fully consistent with
previous studies using similar NaCl-water biphasic systems for HMF production.[154] Inspired
by the results shown above, pure seawater (Table 4, entry 3) was tested as a natural source of
NaCl in water as well as two- and fourfold concentrated seawater (Table 4, entries 4-5). In
case of pure seawater, the HMF yield is slightly lower than with the addition of NaCl
54
(20 wt%, referring to the aqueous phase), as consequence of the lower salt content (3-4%
NaCl in seawater[183]). However, at twofold as well as fourfold concentration of seawater,
yields of HMF were in the same range as observed for water-NaCl mixtures (Table 4, entry
2). In all cases a brownish coloring of both phases (that are the aqueous phase and 2-MeTHF)
was observed, suggesting some by-product formation, typically observed in these HMF
syntheses [211,212]. However, no solid humins were observed in the applied reaction conditions.
This low degradation of the reagents in combination with recyclability of solvents and catalyst
as well as the possibility to further use the glucose containing seawater for fermentation
without the need for workup[183] compensates the relatively low HMF yields and presents
great opportunity for a selective and complete valorization of the sugars.
These remarkable results, implementing seawater to enhance HMF production, keeping
selectivity at considerably high level, encourage further utilization of this solvent for the
dehydration of sugars. For instance, the dehydration of algae-derived saccharides[5] to
synthesize HMF, which has been reported to be straightforward (compared to that of
glucose),[213] could be combined with desalination plants and lead to a promising approach for
an integrated HMF production.
Table 4. Production of HMF (% compared to theoretical yield) in 2-MeTHF after reaction in different solvents.
Conditions: Biphasic system water/2-MeTHF (1:1, 10 mL total volume), oxalic acid: 0.1 M in the aqueous
phase, D-fructose: 1 mmol, 140 °C, 1 h. HMF production was analyzed by HPLC (see experimental part).
entry solvent NaCl* [wt%] HMF yield [%]
1 water – 27
2 water - NaCl 20 52
3 seawater ca. 4 45
4 2x conc. seawater ca. 8 54
5 4x conc. seawater ca. 16 57
* wt% related to the aqueous phase
55
Isomerization of D-glucose to D-fructose with immobilized glucose isomerase
Whereas conversion of fructose to HMF was shown to work efficiently in the seawater-
2-MeTHF setup presented above, glucose is not converted at the same conditions as also
shown in section 3.1.2, where no significant degradation of glucose or xylose was observed.
In order to not apply harsher conditions (e.g. higher temperature, lower pH) and consequently
lose selectivity by degradation of HMF, D-glucose can be isomerized to D-fructose by the so-
called Lobry deBruyn-Alberda van Ekenstein (LBAE) transformation.[214,215] For food-
industry-related applications, glucose isomerase (GI, D-xylose ketol isomerase; EC 5.3.1.5) is
commonly applied to isomerize D-glucose into D-fructose[216]. For instance, it is used in food
industry to produce high fructose corn syrup (HFCS, via saccharification of starch),[216] and it
has also been shown to be active in ionic liquids[217] and with boron-based systems in the
synthesis of HMF[218]. However, application of the GI in seawater has not been investigated
previously and will be presented in this work. Due to its straightforward recyclability and
improved stability over soluble forms,[219] a commercially available immobilized glucose
isomerase (IGI, Sweetzyme IT Extra, 5.1.3.5; ≥300 µg-1), was chosen.
The isomerization has an equilibrium at about 50-60% fructose content,[220] which makes
multiple treatment of the aqueous phase after conversion of D-fructose to HMF necessary.
Hence, the activity of the IGI in seawater (saturated with 2-MeTHF, analogue to the
envisaged aqueous effluent of the second step, Figure 33) was compared to the activity in
phosphate buffer (Figure 34). Impressively, under these non-optimized conditions, the activity
in seawater (saturated with 2-MeTHF) was only slightly lower than in buffer and reached
equilibrium in ca. 5 h (data not shown). This is consistent with previous findings reported in
the literature, where the robustness of the enzyme is also shown by reaction in ionic liquids or
in ethanol-aqueous systems.[217,221]
56
Figure 34. Kinetics of the isomerization of D-glucose to D-fructose in phosphate buffer and in seawater
(saturated with 2-MeTHF, no addition of buffer). Conditions: D-glucose: 140 g L-1, pH 8.2, IGI: 3 wt% (referring
to D-glucose), 60 °C. Glucose concentrations in samples were determined with Glucose (HK) Assay Kit after the
reaction (see experimental part).
Following these promising results, stability and activity of the immobilized enzyme using the
raw effluent from the former step was investigated over multiple cycles. After conducting the
isomerization of 140 g L-1 D-glucose for 40 min at 60 °C with 3 wt% IGI (referring to
D-glucose), the immobilized enzyme was recovered via filtration, added to new solution of
140 g L-1 D-glucose in seawater (saturated with 2-MeTHF) and tested again (Figure 35).
Gratifyingly, the enzyme remained stable during all of the cycles, which clearly emphasizes
the robust nature of the IGI as stated above, and provides promising prognoses on the use of
these enzymes in raw salty effluents to be used in future biorefineries.
57
Figure 35. Recycling of immobilized glucose isomerase (IGI) in 2-MeTHF-saturated seawater. Conditions:
D-glucose: 140 g L-1, seawater (saturated with 2-MeTHF, no buffer added, pH 8.2), IGI: 3 wt% (referring to
D-glucose), 60 °C, 40 min. After each cycle recovery of IGI by filtration and addition to new substrate solution.
Glucose concentration was determined with Glucose (HK) Assay Kit after the reaction and enzyme activity
standardized to remaining glucose in the first cycle (see experimental part).
From D-glucose to HMF by combining the isomerization and dehydration
Since both steps (enzymatic and chemical) showed good performance in the seawater media,
the combination of them was the subsequent experiment. For this purpose, 3 wt% IGI
(referring to D-glucose) was added to a solution of 140 g L-1 D-glucose in 5 mL seawater
(saturated with 2-MeTHF). It was stirred for 2 h at 60 °C, forming ca. 90 g L-1 of D-fructose
(calculated as difference from determined D-glucose concentration of 52 g L-1). After
removing the enzyme via filtration, oxalic acid was added to achieve a 0.1 M solution. Then,
5 mL 2-MeTHF was added and the mixture was stirred for 1 h at 140 °C. Analysis of the
organic phase showed 40 g L-1 HMF, equivalent to 64% yield (relative to the theoretical HMF
yield from D-fructose).
The organic phase was removed and the pH was adjusted to 8 (with solid NaOH). To start a
second cycle D-glucose was added to achieve a total concentration of ca. 140 g L-1. 3 wt% IGI
(referring to theoretical concentration of 140 g L-1 D-glucose) was added and reaction
58
conducted for 2 h at 60 °C, leading to 60 g L-1 of D-fructose (calculated as difference from
determined D-glucose concentration of 77 g L-1).
These results show nicely that the two steps, the isomerization of D-glucose to D-fructose and
the conversion of D-fructose to HMF, can be combined straightforwardly. Since immobilized
glucose isomerase proved to be very stable under the applied conditions, the aqueous effluent
after dehydration of D-fructose can be conversely recycled back into the enzyme reactor.
Alternatively, the remnant seawater with D-glucose (ca. 50%) could be used for further
fermentative processes. Optimization of the concept may provide useful and sustainable
synergies for future biorefineries combining enzymes and bio-based acids, and using seawater
as readily available and non-potable water (and salt) source.
3.3.5. Synthesis of furfural from xylose using the raw 1-step OrganoCat Process effluent
The platform chemical furfural, which is of great importance due to its numerous
applications[139,141,151,222] can be derived from xylose, main component of hemicellulose, by
triple dehydration of the monomeric C5 sugar. For the furfural synthesis our research group
developed a concept using FeCl3 as acidic catalyst in aqueous media and addition of NaCl to
improve furfural production rate, while the product is in situ extracted into a second
2-MeTHF phase[155] (Figure 36).
Figure 36. Scheme of the iron-catalyzed xylose dehydration. Figure adapted from[155]
59
The non-purified aqueous hemicellulose sugars effluent of the 1-step OrganoCat Process,
conducted with beech wood particles (0.5-0.1 mm, Figure 16) was directly used after removal
of oxalic acid and the 2-MeTHF phase (containing lignin). Furfural was synthesized upon
xylose dehydration with FeCl3 ∙ 6 H2O as catalyst. Xylose concentration in the effluent was
ca. 30 g L-1 (determined in the aqueous phase together with small amounts of other
carbohydrates after the 1-step OrganoCat Process).
To 2.5 mL of this raw solution, FeCl3 6 H2O (0.24 M) was added together with NaCl
(30 wt%, related to the aqueous phase). Then 2.5 mL of 2-MeTHF were added and the
reaction mixture was stirred for 2 h at 140 °C. Remarkably ca. 7 g L-1 of furfural were
observed in the organic phase, indicating a rate of formation of ca. 3.5 g L-1 h-1 and
corresponding to an initial yield of 37%, still under these largely non-optimized process
conditions.
Figure 37. Conversion of xylose into furfural mediated by FeCl3 6 H2O, using aqueous non-purified xylose
effluent obtained from the 1-step OrganoCat Process. Reaction conditions: ca. 30 g L-1 xylose, 0.12 M
FeCl3 6 H2O, 30 wt% NaCl with respect to aqueous phase, 2.5 mL 2-MeTHF, 2.5 mL of aqueous phase
produced in 1-step OrganoCat Process, 140 °C, 2 h.
3.3.6. Fermentation of raw carbohydrate effluents
Itaconic acid may be used as a building block for polymerization (e.g. plastics, adhesives,
elastomers and coatings),[161,162] as well as for the synthesis of potential biofuels or biogenic
60
solvents (e.g. 3-MeTHF) (see also section 1.4.4).[141] It can be synthesized via chemical
multistep reactions[223] or via biotransformations.[161] To keep downstream processing of sugar
effluents at a minimum, research with the goal of demonstrating compatibility of raw
effluents with subsequent processes is of high interest. In cooperation with
Bioverfahrenstechnik (AVT, RWTH Aachen University) different effluents of the 1-step
OrganoCat Process (section 3.1) and enzymatically hydrolyzed cellulose in seawater (section
3.3.3) were fermented by U. maydis to itaconic acid[180] (Figure 38).
Figure 38. Fermentation of raw sugar effluents with U. maydis.
Due to the fact that the organism preferes glucose over xylose as substrate, especially in the
initial growing phase, the sole fermentation of the hemicellulose effluent (13 g L-1 xylose in
reaction medium after preparation of the Tabuchi medium) yielded very low amounts of
itaconic acid. However, by adding 30 g L-1 glucose to that hemicellulose effluent (10 g L-1
xylose in reaction medium) a final itaconic acid concentration of 4.1 g L-1 was found after
36 h, consuming both xylose and glucose. In a reference experiment using a solution of
61
30 g L-1 xylose and 90 g L-1 glucose a concentration of 13.1 g L-1 itaconic acid was achieved.
The product concentration is much lower in the case of the hemicellulose sugars due to the
overall lower concentrations.[180] However, fermentation of the hemicellulose effluent of the
1-step OrganoCat Process was successfully achieved, even though higher sugar
concentrations in the reaction media as well as genetically improving acceptance of xylose as
substrate for the organism could further improve production.
The effluent of enzymatically hydrolyzed cellulose in seawater was produced by stirring
100 g L-1 amorphous cellulose (Sigmacell®101) with 1 vol% Accellerase® 1500 at 50 °C for
seven days in 0.1 M citric buffer (pH 4.5) prepared with seawater. The raw glucose solution
(ca. 60 g L-1 glucose, determined with HPLC) was used to prepare the Tabuchi medium
(without further addition of glucose) for fermentation. Remarkably, a slight increase of the
overall production of itaconic acid, compared to a reference cultivation using regular Tabuchi
medium with 60 g L-1 glucose, was observed. After 96 h, a final itaconic acid concentration of
6.4 g L-1 was obtained, demonstrating excellent results for the fermentation of this raw
sample.
62
4. SUMMARY AND CONCLUSION
Figure 39. Summary scheme.
63
In this thesis development and optimization of conversion strategies for lignocellulosic
biomass was targeted. It was desired to yield value-added raw materials following the
principles of “green chemistry”. Evaluating usability of the effluents originating from single
reaction steps in subsequent reactions was conducted to demonstrate a potential application in
integrated biorefinery concepts.
The 1-step OrganoCat Process [224] (a biphasic, biogenic lignocellulose fractionation strategy)
was in-depth studied in terms of selectivity and optimized parameters, identifying the best
conditions for an energy efficient, quantitative depolymerization of hemicellulose, leaving
cellulose untouched (3 hours at 140 °C). As a result, three non-degraded raw materials are
obtained from beech wood (xylose, cellulose and lignin). Moreover, a glass-made high
pressure reactor setup was shown to display the best results in laboratory-scale, regarding
yields of product streams and handling at 10 mL as well as 40 mL reaction scale (referring to
total solvent volume). Furthermore, the reaction was conducted at a 3 L scale (referring to
total solvent volume), proceeding as efficiently as in the smaller lab-scales, showing that a
scale-up for practical purposes can be realized straightforwardly. Additionally, the versatility
of catalysts used in the process was successfully demonstrated, showing full conversion of
hemicellulose, independently of the kind of acid, as well as the adaptability to a broad range
of different biomass sources with moderate-to-high hemicellulosic content, whereas
softwoods (e.g. spruce) still have some room for improvement due to their low hemicellulose
content, which might be tackled by applying harsher conditions (e.g. higher temperature,
pressure or addition of salt), thus extracting lignin at the cost of some cellulose degradation
extent. Furthermore, to improve solvent-to-substrate and -catalyst ratio, pretreatments at high
substrate loadings of 200 g L-1 were shown, as well as the reduction of organic solvent within
the process, yet leading to slightly lower yields. This issue was successfully overcome by
recycling the liquid effluent and adding new lignocellulose in multiple steps, leading to
remarkably enhanced hemicellulose sugar concentrations (ca. 60 g L-1 after four pretreatment
cycles) as well as recovered increased amounts of lignin. Finally, the combination of a
mechanical screw press pretreatment with chemical pulping resulted in comparable results
considering the product streams. Herein, reduction of reaction time and temperature, and thus
working at less optimized conditions might show higher impact of the mechanical
pretreatment on the fractionation of lignocellulose.
64
The 2-step OrganoCat Process, a variation conducted in two steps has been put forward as
well. First hemicellulose was hydrolyzed selectively in natural and abundantly available
seawater as reaction medium with bioderived and recyclable oxalic acid as catalyst. In a
second step lignin was extracted from the solid residue in different organic solvents, water-
organic solvent mixtures and deep eutectic solvent (DES). The utilization of salty water
proved to enhance the hydrolysis rate in the first step significantly (full hydrolysis of the
hemicellulose fraction after 1 hour at 140 °C). The lignin yield obtained was lower than with
the original 1-step OrganoCat Process (ca. 3-6 wt% compared to ca. 12 wt% in the 1-step
OrganoCat Process, in relation to total wood weight), independent on the extraction solvent
used, which might be explained by lignin being exposed to the acidic medium longer than in
the biphasic system, and thus being prone to further irreversible (re)polymerizations. Finally,
the straightforward recycling of the reactive phase in the first step led to significantly
improved substrate-to-solvent and substrate-to-catalyst ratios (90 g L-1 xylose after total
addition of 400 g L-1 beech wood to 0.1 M oxalic acid in seawater within 4 cycles). Those
promising results display great opportunities for lignocellulose containing low amounts of
lignin (e.g. grasses, algae, etc.), efficiently and selectively hydrolyzing hemicellulose and
leaving cellulose for subsequent hydrolysis. Furthermore, sugar effluents not being
contaminated with organic solvents might favor application in nutritional and pharmaceutical
processes.
After showing two promising concepts for the fractionation of lignocellulose various
strategies for the valorization of the effluents cellulose and hemicellulose sugars were
presented, focusing on the application of integrated biorefinery concepts. For the salt-assisted
organic-acid catalyzed depolymerization of microcrystalline cellulose[133] seawater can be
used as natural source of salt, improving the hydrolysis rate and substituting more exhaustable
potable water reservoirs. As another method to reduce crystallinity of cellulose, pretreatment
(e.g. dissolution and reprecipitation) with ionic liquids (e.g. [DMIm]Acetate) was shown to
significantly improve the hydrolysis rate of the organic-acid as well as the salt-assisted
organic-acid catalyzed depolymerization of -cellulose.
As an example for the enzymatic hydrolysis of cellulose, the commercially available cellulase
cocktail Accellerase® 1500 was characterized and shown to efficiently depolymerize
microcrystalline cellulose (Avicel®) at optimum conditions (e.g. 20 g L-1 substrate loading,
0.1 vol% enzyme compared to solvent, 50 °C). Furthermore, hydrolysis of Avicel® cellulose
65
was performed removing the liquid product mixture and adding new enzyme solution in
multiple cycles, thus avoiding product inhibition of the catalyst. As a result, particles of high
crystallinity were obtained which could not be hydrolyzed anymore, and might find
application as defined-size cellulose particles, when controlling crystallinity appropriately. To
further evaluate the effect of the different pretreatments hydrolysis of cellulose samples from
the 1-step OrganoCat Process and the combination with mechanical pretreatment was
compared to the untreated biomass. In case of beech wood, after the 1-step OrganoCat
Process the enzymatic activity arose significantly compared to the non-pretreated sample,
which virtually showed no activity at all. Compared to Avicel® cellulose the activity was
slightly lower, presumably due to inhibition by residual lignin. However, based on those good
results, evolving cellulase and adapting it to the specific kind of biomass, hydrolysis
effectivity could be further improved and optimized. When combined with mechanical screw
press pretreatment of reed hydrolysis rate of the cellulose pulp was shown to be improved
greatly. The effect of the screw press on the 1-step OrganoCat Process is not yet well
understood and has to be investigated further. However, considerations involving an extended
surface area and a decrease in cellulose crystallinity, analogous to ball milling treatment,[190]
are probable and might deliver promising synergies of mechanical pretreatment and the 1-step
OrganoCat Process. Furthermore, application of commercial cellulase (e.g.
Accellerase® 1500) using seawater to prepare the buffer was successfully conducted, showing
the same high activity as when distilled water was used. Also in higher concentrated seawater
decent activity was observed.
To synthesize HMF from glucose a chemo-enzymatic concept conducted in seawater was put
forward, using IGI for the isomerization of glucose to fructose, followed by the oxalic acid
catalyzed triple-dehydration of fructose to HMF in a biphasic system. Herein, seawater was
used as source of salt, boosting the activity of the dehydration reaction while keeping
selectivity at a considerably high level. IGI proved to be very robust, efficiently working in
2-MeTHF saturated seawater over several cycles, and thus enabling the combination of both
reaction steps for multiple cycles, adding new glucose for the isomerization and yielding
HMF in the dehydration. Those examples indicate great opportunity for integrated processes
at coastal areas, using salt-water based biomass (e.g. algae) and converting it directly in their
natural medium seawater. Additionally, utilization of the waste stream of drinking water
production (e.g. concentrated seawater) might further enhance cost-effectiveness of both
processes and make them more benign and integrated.
66
Finally, the direct utilization of the hemicellulose effluent from the 1-step OrganoCat Process
for subsequent conversion into platform chemicals was shown. In the iron-catalyzed
dehydration of xylose to furfural in a bio-based biphasic system[155] the crude effluent
afforded 3.5 g L-1 h-1 under non-optimized conditions, which could be straightforward
improved by higher xylose concentration in the effluent. Also the fermentation with U.
maydis showed first promising results in the production of itaconic acid from glucose,
obtained from enzymatic hydrolysis of cellulose in seawater, and from xylose, derived from
beech wood in the 1-step OrganoCat Process.[180] Avoiding an expensive downstream
processing unit for the sugars, both chemical and biochemical approaches reveal promising
success in the utilization of the crude effluents coming from the pretreatment units.[155,180]
67
5. EXPERIMENTAL PART
5.1. Chemicals
Avicel® (PH-101), -cellulose, Amorphous cellulose (phosphoric-acid-swollen-cellulose,
PASC), fructose, glucose, xylose, oxalic acid, phosphoric acid, acetic acid, citric acid, maleic
acid, hydrochloric acid, lactic acid, betaine, sodium chloride, magnesium sulfate, iron(III)
chloride hexahydrate, calcium chloride, sodium hydroxide, dipotassium hydrogen phosphate,
potassium dihydrogen phosphate, trisodium citrate, glucose isomerase Sweetzyme IT Extra
(5.1.3.5; ≥300 mg-1), Glucose (HK) Assay Kit (GAHK20), reagents for PAHBAH method,
1-butanol, ethanol, and 2-MeTHF were purchased from Sigma–Aldrich. Beech wood, reed
and spruce were obtained from local suppliers and dried until constant weight. Mate tea leaves
were obtained in supermarket. After tea consumption, leaves were dried for several weeks and
used for fractionation systems. Seawater was collected in Bandol (France), and stored at 4 °C
until use. Accellerase® 1500 was kindly donated by Genencor International Corporation. It is
an enzyme cocktail containing several enzymes with endoglucanase activity
(2200-2800 CMC U g-1; CMC: carboxymethylcellulose), and -glucosidase activity
(525-775 pNPG U g-1; pNPG: p-nitrophenyl glucoside), with an optimum pH of 4.5.
5.2. General Procedures
5.2.1. Standard procedure for the 1-step OrganoCat Process
Glass-made high pressure reactor
In a glass-made high pressure reactor (Ace pressure tube, 15 mL or 80 mL) 2-MeTHF (5 mL
or 20 mL) and the same volume of aqueous oxalic acid solution with the indicated
concentration (0.1-0.2 M, referring to the aqueous phase) were introduced. 100 g L-1
(referring to the aqueous phase, e.g. 500 mg or 2 g) lignocellulose and a stirring bar were
added and the reactor was closed tight. The suspension was stirred at room temperature for
20 min and then heated to the indicated reaction temperature for the indicated time.
After reaching desired reaction time the reactor was cooled down to room temperature and
opened. The two phases were separated, 2-MeTHF was removed over reduced pressure and
lignin was weighed and analyzed via NMR. The aqueous phase with solid residue was
68
centrifuged at 4000 rpm for 5 min. Then, 2 mL of the aqueous phase were filtered with a
syringe filter (Multoclear-13 PTFE, 0.45 µm) and analyzed in HPLC. The solid cellulose pulp
was filtered off with a filter paper (MN 615), washed neutral with distilled water and then
dried.
Standard procedure for the metal-based high pressure reactor
In a stainless steel high pressure reactor (10 mL or 20 mL) 2-MeTHF (5 mL or 10 mL) and
the same volume of aqueous oxalic acid solution with the indicated concentration (0.1-0.2 M,
referring to the aqueous phase) were introduced. 100 g L-1 (referring to the aqueous phase,
e.g. 500 mg or 1 g) lignocellulose and a stirring bar were added. The indicated amount of CO2
was pressed on and the reactor was closed tight. The suspension was stirred for 20 min at
room temperature and then heated to the indicated reaction temperature for the indicated time.
After reaching desired reaction time the reactor was cooled down to room temperature and the
high pressure reactor was vented carefully to release CO2. The workup procedure was
analogous to the reaction in the glass-made high pressure reactor.
Procedure for the 5L metal-based high pressure reactor
In a glass inlay 100 g L-1 beech wood lignocellulose (150 g) was suspended in water with
0.1 M oxalic acid (1.5 L). As the organic phase, 2-MeTHF (1.5 L) was added, the glass inlay
was inserted into the metal-based high-pressure reactor and the reactor was closed. The
reactor was heated to 140 °C for 3 h. After cooling of the reactor, the organic phase was
separated by decantation. The aqueous phase was filtered to isolate the cellulose-pulp, and
xylose and glucose concentrations were determined in the aqueous phase. The solid residue
was washed with distilled water until neutral pH, dried until constant weight, extracted with
2-MeTHF (1.5 L) at room temperature for 24 h and then filtrated and the solid residue dried
again until constant weight. The organic phases were collected, and 2-MeTHF was evaporated
to obtain the lignin fraction.
5.2.2. Standard procedure for the 2-step OrganoCat Process
In a glass-made high pressure reactor (Ace pressure tube, 15 mL or 80 mL) 100 g L-1
lignocellulose was suspended in seawater with oxalic acid (0.1 M). The temperature was set at
140 °C for 3 h reaction time. After cooling off the reactor, the aqueous phase was filtered to
isolate the cellulose-pulp, and xylose and glucose concentrations were determined in the
69
aqueous phase via HPLC. The solid residue was washed with distilled water until neutral pH
and dried until constant weight.
Dried pulp obtained was suspended (final concentration 100 g L-1) in the respective solvent
(2-MeTHF / distilled water, 1:1 v/v; 2-MeTHF / seawater, 1:1 v/v; 2-MeTHF / 0.1 M oxalic
acid, 1:1 v/v; 2-MeTHF; EtOH; 1-BuOH; DES, lactic acid / betaine, 2:1 mol/mol), and stirred
at the indicated temperature for the indicated time. After cooling and filtration the organic
phase was decanted (in case of mixtures with water), and the organic solvent was removed via
distillation to obtain the lignin fraction. In case of the DES after filtration lignin was
precipitated by addition of water, filtrated and redissolved in 2-MeTHF. The organic solvent
was then removed via distillation to obtain the lignin fraction.
5.2.3. Procedure for the organic acid-catalyzed cellulose hydrolysis in seawater
Slurries of Avicel® cellulose (20 g L-1) were suspended in distilled water, brine, seawater and
5-fold concentrated seawater, in which 0.1 M maleic acid or oxalic acid were aggregated.
Reactions were magnetically stirred at 125 °C for 6 h in a refluxing system (106 °C as
effective temperature). At 0, 1, 4 and 6 h samples of 0.3 mL were taken and analyzed with the
PAHBAH method.
5.2.4. Standard procedure for the cellulose hydrolysis with Accellerase® 1500
Preparation of citrate-buffer in distilled water
44 mmol citric acid and 52 mmol of trisodium citrate were added to 900 mL of distilled water,
leading to a pH of 4.5 (0.1 M buffer). The buffer solution was stored at 4 °C until use.
Preparation of citrate-buffer in seawater
12 mmol of citric acid and 14 mmol of trisodium citrate were added to 250 mL of seawater,
leading to a pH of 3.8. Trisodium citrate was further added (ca. 1 mmol) until pH was
equilibrated at 4.5 (0.1 M buffer). The buffer solution was stored at 4 °C until use.
Hydrolysis procedure
Cellulose samples (15-40 g L-1) were suspended in citrate buffer in a round flask with a
stirring bar. The suspension was heated to 50 °C and the indicated amount of
Accellerase® 1500 was introduced. 0.3 mL samples were taken at indicated times, heated to
100 °C for 5 min and then analyzed with the Glucose (HK) Assay Kit.
70
5.2.5. Procedure for the synthesis of HMF from glucose
Preparation of phosphate buffer
In 1 L distilled water 7.4897 g K2HPO4, 0.9526 g KH2PO4, and 0.3000 mg MgSO4 were
dissolved under stirring at room temperature. The solution was stored at 4 °C until use.
Standard procedure for the isomerization of D-glucose to D-fructose with IGI
Solutions of glucose (140 g L-1) were prepared in phosphate buffer and in 2-MeTHF-saturated
seawater. In a closed glass reactor 5 mL reaction phase and 3 wt% (referring to glucose) of
immobilized glucose isomerase (IGI) were added. The reaction mixture was shaken in an
Eppendorf Thermomixer Comfort at 600 rpm and 60 °C for the indicated time. After cooling
the reactor was opened, the liquid phase was decanted and analyzed with Glucose (HK) Assay
Kit.
Standard procedure for the synthesis of HMF from fructose
Solutions of D-fructose (72 g L-1) were prepared in indicated aqueous solvents, in which
oxalic acid (0.2 M) was aggregated. In a closed glass reactor a mixture of 2.5 mL reaction
phase and 2.5 mL 2-MeTHF phase was magnetically stirred for 1 h at 140 °C. After cooling
the reactor was opened, the organic phase was decanted and analyzed via HPLC.
5.2.6. Procedure for the synthesis of furfural from 1-step OrganoCat Process hemicellulose
effluent
The aqueous effluent from the 1-step OrganoCat Process (see section 0 for details) contained
ca. 30 g L-1 xylose (determined in the aqueous phase together with small amounts of other
carbohydrates after 1-step OrganoCat Process). Oxalic acid was removed by adding 2 mmol
CaCl2 to 20 mL of the effluent, stirring for 1 h at room temperature, and filtrating. To 2.5 mL
of this solution, FeCl3 6 H2O (0.12 M, related to the total volume) was added together with
NaCl (30 wt%, related to the aqueous phase). Then 2.5 mL 2-MeTHF was added and the
reaction mixture was stirred for 2 h at 140 °C. After cooling the reactor was opened, the
organic phase was decanted and analyzed via GC.
5.2.7. Procedure for fermentation of sugars
The experiments were conducted by Tobias Klement at the Bioverfahrenstechnik (AVT, Prof.
Dr. J. Büchs, RWTH Aachen University), following the protocol of Klement et al.[180]
71
5.2.8. Procedure for the mechanical pretreatment with a screw press
The experiments were conducted by Qingqi Yan at the Aachener Verfahrenstechnik (AVT,
Prof. Dr. M. Modigell, RWTH Aachen University), following the protocol of Yan et al.[181]
5.3. Analysis
5.3.1. High Performance Liquid Chromatography (HPLC)
HPLC measurements of glucose and xylose were carried out on a Jasco HPLC equipped with
a SUGARSH1011 column with a 0.01 wt% aqueous acid solution as the eluent. The flow rate
was set to 0.6 mL min-1 and samples of 30 µL were injected. Amounts of xylose and glucose
present in the hemicellulose fraction were determined based on calibration curves built using
commercially available authentic substrates.
Figure 40. Calibration curves of xylose and glucose for HPLC measurement. Retention times: xylose 13.2 min,
glucose 12.2 min.
HPLC measurements of HMF were carried out on a Jasco HPLC equipped with a
Kromasil 100 C18 column with a 10 wt% aqueous methanol solution as the eluent. The flow
rate was set to 0.8 mL min-1 and samples of 50 µL were injected. The amount of HMF present
in 2-MeTHF was determined based on calibration curves built using commercially available
authentic substrates.
72
Figure 41. Calibration curve of HMF for HPLC measurement. Retention time HMF 9.8 min.
5.3.2. Gas Chromatography (GC)
GC measurements to quantify the concentration of furfural in the 2-MeTHF phase were
conducted with a 50 m OV1-IVA column, nitrogen as carrier gas, a split ratio of 33:1, and a
flame-ionization detector. The initial temperature was 50 °C, raised at 8 °C min-1 to 250 °C.
Quantification was done using n-decane and n-dodecane as internal standards.
73
5.3.3. Analysis with Glucose (HK) Assay Kit
10 µL of the sample were diluted with 990 µL distilled water. After shaking for 1 min 50 µL
of the diluted sample were transferred with an Eppendorf pipette into a microtiter plate well.
Then 200 µL of Glucose (HK) Assay Kit were added and absorption at 340 nm was measured
with a UV/VIS spectrometer (BioTek PowerWave HT) until absorption was constant.
Figure 42. Calibration curve for the Glucose (HK) Assay Kit.
74
5.3.4. Analysis with PAHBAH method
5 g p-hydroxy benzoic acid hydrazide (PAHBAH) were added to 30 mL distilled water and
slurried. Then 5 mL of concentrated HCl were added and the volume filled to 100 mL with
distilled water (solution A). In a second flask 12.45 g trisodium citrate were dissolved in
500 mL distilled water, 1.10 g CaCl2 and 20.00 g NaOH were added. Then the volume was
adjusted to 1 L with distilled water (solution B). 10 mL of solution A were added to 90 mL of
solution B, stirred and stored on ice freshly for every analysis (PAHBAH-reagent).[225]
50 µL sample were mixed with 100 µL PAHBAH-reagent, heated to 100 °C for 10 min and
then absorption at 410 nm was measured with a UV/VIS spectrometer (BioTek PowerWave
HT).
Figure 43. Calibration curve for the PAHBAH-method.
75
REFERENCES
[1] T. Werpy, G. Petersen, A. Aden, J. Bozell, Top Value Added Chemicals From Biomass.
Volume I-Results of Screening for Potential Candidates From Sugars and Synthesis
Gas, 2004.
[2] J. Bozell, J. Holladay, D. Johnson, J. White, Top Value Added Chemicals from
Biomass-Volume II: Results of Screening for Potential Candidates from Biorefinery
Lignin., 2007.
[3] J. J. Bozell, Clean Soil Air Water 2008, 36, 641–647.
[4] J. Leboreiro, A. K. Hilaly, Bioresour. Technol. 2011, 102, 2712–2723.
[5] P. M. Foley, E. S. Beach, J. B. Zimmerman, Green Chem. 2011, 13, 1399–1405.
[6] P. T. Anastas, Chem.rev. 2007, 107, 2167–2168.
[7] P. Anastas, N. Eghbali, Chem. Soc. Rev. 2010, 39, 301–312.
[8] K. Sanderson, Nature 2011, 474, S12–S14.
[9] B. Kamm, M. Kamm, M. Schmidt, T. Hirth, M. Schulze, in Biorefineries-Industrial
Process. Prod. Status Quo Futur. Dir. (Eds.: B. Kamm, P.R. Gruber, M. Kamm),
Wiley-VCH Verlag GmbH, Weinheim, Germany, 2008.
[10] C. E. Wyman, B. Yang, Calif. Agr. 2009, 64, 185–190.
[11] C. Somerville, H. Youngs, C. Taylor, S. C. Davis, S. P. Long, Science (80-. ). 2010,
329, 790–792.
[12] A. Wisogel, S. Tyson, D. Johnson, in Handb. Bioethanol Prod. Util. (Ed.: Ch.E.
Wyman), Taylor & Francis, Washington D.C., 1996, pp. 105–116.
[13] Y. Sun, J. Cheng, Bioresour. Technol. 2002, 83, 1–11.
[14] V. L. Finkenstadt, R. P. Millane, Macromolecules 1998, 31, 7776–7783.
[15] R. Rinaldi, F. Schüth, ChemSusChem 2009, 2, 1096–1107.
[16] Y.-H. Percival Zhang, M. E. Himmel, J. R. Mielenz, Biotechnol. Adv. 2006, 24, 452–
481.
[17] F. Hu, A. Ragauskas, Bioenerg. Res. 2012, 5, 1043–1066.
[18] S. P. S. Chundawat, G. Bellesia, N. Uppugundla, L. da Costa Sousa, D. Gao, A. M.
Cheh, U. P. Agarwal, C. M. Bianchetti, G. N. Phillips, P. Langan, et al., J. Am. Chem.
Soc. 2011, 133, 11163–11174.
76
[19] B. C. Saha, J. Ind. Microbiol. Biot. 2003, 30, 279–291.
[20] C. E. Wyman, Annu. Rev. Energ. Env. 1999, 24, 189–226.
[21] A. Ebringerová, Z. Hromádková, T. Heinze, Adv. Polym. Sci. 2005, 186, 1–67.
[22] J. Zakzeski, P. C. A. Bruijnincx, A. L. Jongerius, B. M. Weckhuysen, Chem.rev. 2010,
110, 3552–3599.
[23] L. B. Davin, N. G. Lewis, Curr. Opin. Biotech. 2005, 16, 407–415.
[24] K. Ruel, F. Barnoud, D. A. I. Goring, Wood Sci. Technol. 1978, 12, 287–291.
[25] T. Hatakeyama, T. Yoshida, H. Hatakeyama, Tenth Int. Symp. Wood Pulping Chem.
1999, 1, 478–481.
[26] M. Dolk, F. Pla, J. F. Yan, J. L. McCarthy, Macromolecules 1986, 19, 1464–1470.
[27] W. J. Connors, S. Sarkanen, J. L. McCarthy, Holzforschung 1980, 34, 80–85.
[28] J. M. Cruz, H. Domı́nguez, J. C. Parajó, Food Chem. 2005, 90, 503–511.
[29] C. Pouteau, P. Dole, B. Cathala, L. Averous, N. Boquillon, Polym. Degrad. Stabil.
2003, 81, 9–18.
[30] L. R. C. Barclay, F. Xi, J. Q. Norris, J. Wood Chem.Technol. 1997, 17, 73–90.
[31] C. Rice-Evans, N. Miller, G. Paganga, Trends iPlant Sci. 1997, 2, 152–159.
[32] X. Pan, J. F. Kadla, K. Ehara, N. Gilkes, J. N. Saddler, J. Agric. Food. Chem. 2006, 54,
5806–5813.
[33] M. B. Hocking, J. Chem. Educ. 1997, 74, 1055–1059.
[34] S. Ramachandra Rao, G. Ravishankar, J. Sci. Food Agr. 2000, 80, 289–304.
[35] J. McMillan, in Pretreat. Lignocellul. Biomass Enzym. Convers. Biomass Fuels Prod.
ACS Symp. Ser. (Eds.: M.E. Himmel, J.O. Baker, R.P. Overend), American Chemical
Society, Washington, DC, 1994, pp. 292–324.
[36] H. L. Chum, D. K. Johnson, S. Black, J. Baker, K. Grohmann, K. V Sarkanen, K.
Wallace, H. a Schroeder, Biotechnol. Bioeng. 1988, 31, 643–649.
[37] R. W. Thring, E. Chornet, R. P. Overend, Biomass 1990, 23, 289–305.
[38] X. Zhao, K. Cheng, D. Liu, Appl. Microbiol. Biot. 2009, 82, 815–827.
[39] M. Mascal, E. B. Nikitin, ChemSusChem 2010, 3, 1349–1351.
[40] J. Li, G. Henriksson, G. Gellerstedt, Bioresour. Technol. 2007, 98, 3061–3068.
77
[41] S. M. Ewanick, R. Bura, J. N. Saddler, Biotechnol. Bioeng. 2007, 98, 737–746.
[42] P. J. Morjanoff, P. P. Gray, Biotechnol. Bioeng. 1987, 29, 733–741.
[43] M. T. Holtzapple, a E. Humphrey, J. D. Taylor, Biotechnol. Bioeng. 1989, 33, 207–
210.
[44] T. A. Clark, K. L. Mackie, J. Wood Chem.Technol. 1987, 7, 373–403.
[45] W. R. Grous, A. O. Converse, H. E. Grethlein, Enzym. Microb. Tech. 1986, 8, 274–
280.
[46] K. L. Mackie, H. H. Brownell, K. L. West, J. N. Saddler, J. Wood Chem.Technol.
1985, 5, 405–425.
[47] M. Mes-Hartree, B. E. Dale, W. K. Craig, Appl. Microbiol. … 1988, 29, 462–468.
[48] D. M. Fox, J. W. Gilman, A. B. Morgan, J. R. Shields, P. H. Maupin, R. E. Lyon, H. C.
De Long, P. C. Trulove, Ind. Eng. Chem. Res. 2008, 47, 6327–6332.
[49] M. Smiglak, W. M. Reichert, J. D. Holbrey, J. S. Wilkes, L. Sun, J. S. Thrasher, K.
Kirichenko, S. Singh, A. R. Katritzky, R. D. Rogers, Chem. Commun. 2006, 2554–
2556.
[50] P. Wasserscheid, T. Welton, Eds., Ionic Liquids in Synthesis, Wiley-VCH Verlag
GmbH & Co. KGaA, Weinheim, Germany, 2007.
[51] S. Tan, D. MacFarlane, Top. Curr. Chem. 2010, 290, 311–339.
[52] P. Wasserscheid, W. Keim, Angew. Chem. Int. Ed. 2000, 39, 3772–3789.
[53] A. Bösmann, P. S. Schulz, P. Wasserscheid, Monatshefte für Chemie - Chem. Mon.
2007, 138, 1159–1161.
[54] P. Bonhôte, A.-P. Dias, N. Papageorgiou, K. Kalyanasundaram, M. Grätzel, Inorg.
Chem. 1996, 35, 1168–1178.
[55] E. B. Carter, S. L. Culver, P. a Fox, R. D. Goode, I. Ntai, M. D. Tickell, R. K. Traylor,
N. W. Hoffman, J. H. Davis, Chem. Commun. 2004, 630–631.
[56] P. Nockemann, B. Thijs, K. Driesen, C. R. Janssen, K. Van Hecke, L. Van Meervelt, S.
Kossmann, B. Kirchner, K. Binnemans, J. Phys. Chem. B 2007, 111, 5254–5263.
[57] A. P. Dadi, C. a Schall, S. Varanasi, Appl. Biochem. Biotech. 2007, 137-140, 407–421.
[58] R. P. Swatloski, S. K. Spear, J. D. Holbrey, R. D. Rogers, J. Am. Chem. Soc. 2002,
124, 4974–4975.
[59] C. Graenacher, Cellulose Solution, 1934, US1943176.
78
[60] Y. Su, H. M. Brown, G. Li, X. Zhou, J. E. Amonette, J. L. Fulton, D. M. Camaioni, Z.
C. Zhang, Appl. Catal. A-Gen. 2011, 391, 436–442.
[61] R. Rinaldi, R. Palkovits, F. Schüth, Angew. Chem. Int. Ed. 2008, 47, 8047–8050.
[62] L. Vanoye, M. Fanselow, J. D. Holbrey, M. P. Atkins, K. R. Seddon, Green Chem.
2009, 11, 390–396.
[63] Y. Pu, N. Jiang, A. J. Ragauskas, J. Wood Chem.Technol. 2007, 27, 23–33.
[64] I. Kilpeläinen, H. Xie, A. King, M. Granstrom, S. Heikkinen, D. S. Argyropoulos, J.
Agric. Food. Chem. 2007, 55, 9142–9148.
[65] D. a. Fort, R. C. Remsing, R. P. Swatloski, P. Moyna, G. Moyna, R. D. Rogers, Green
Chem. 2007, 9, 63–69.
[66] J. Viell, W. Marquardt, Chem. Ing. Tech. 2009, 81, 1213–1213.
[67] J. Viell, W. Marquardt, Holzforschung 2011, 65, 519–525.
[68] S. H. Lee, T. V Doherty, R. J. Linhardt, J. S. Dordick, Biotechnol. Bioeng. 2009, 102,
1368–1376.
[69] S. S. Y. Tan, D. R. MacFarlane, J. Upfal, L. a. Edye, W. O. S. Doherty, A. F. Patti, J.
M. Pringle, J. L. Scott, Green Chem. 2009, 11, 339–345.
[70] J. B. Binder, R. T. Raines, P. Natl. Acad. Sci. USA 2010, 107, 4516–4521.
[71] P. G. Jessop, D. J. Heldebrant, X. Li, C. A. Eckert, C. L. Liotta, Nature 2005, 436,
1102.
[72] M. J. Earle, J. M. S. S. Esperança, M. a Gilea, J. N. C. Lopes, L. P. N. Rebelo, J. W.
Magee, K. R. Seddon, J. a Widegren, Nature 2006, 439, 831–834.
[73] P. Wasserscheid, Nature 2006, 439, 797.
[74] A. E. Rosamilia, C. R. Strauss, J. L. Scott, Pure. Appl. Chem. 2007, 79, 1869–1877.
[75] U. P. Kreher, A. E. Rosamilia, C. L. Raston, J. L. Scott, C. R. Strauss, Molecules 2004,
9, 387–393.
[76] M. Francisco, A. van den Bruinhorst, M. C. Kroon, Green Chem. 2012, 14, 2153–2157.
[77] Z. Maugeri, P. Domínguez de María, RSC Adv. 2012, 2, 421–425.
[78] P. Domínguez de María, Z. Maugeri, Curr. Opin. Chem. Biol. 2011, 15, 220–225.
[79] Z. Maugeri, W. Leitner, P. Domínguez de María, Tetrahedron Lett. 2012, 53, 6968–
6971.
79
[80] C. Ruß, B. König, Green Chem. 2012, 14, 2969–2982.
[81] R. M. Hertel, A. S. Bommarius, M. J. Realff, Y. Kang, Deep Eutectic Solvent System
Comprising Betaine Monohydrate and Hydrogen Bond Donor, 2012, WO2012145522.
[82] M. M. Chang, T. Y. C. Chou, G. T. Tsao, Adv. Biochem. Eng. Biot. 1981, 20, 15–42.
[83] A. L. Demain, M. Newcomb, J. H. D. Wu, Microbiol. Mol. Biol. R. 2005, 69, 124–154.
[84] L. T. Fan, Y.-H. Lee, M. M. Gharpuray, Adv. Biochem. Eng. Biot. 1982, 23, 157–187.
[85] H. E. Grethlein, Biotechnol. Adv. 1984, 2, 43–62.
[86] K. W. Lin, M. R. Ladisch, D. M. Schaeffer, C. H. Noller, V. Lechtenberg, G. T. Tsao,
AIChE Symp. Ser. 1981, 203, 102–106.
[87] J. D. McMillan, ACS Sym. Ser. 1994, 566, 234–292.
[88] M. A. Millett, A. J. Baker, L. D. Satter, Biotechnol. Bioeng. Symp. 1976, 6, 125–153.
[89] N. Moreira, Sci. News. Online 2005, 168, 209–224.
[90] N. Mosier, C. Wyman, B. Dale, R. Elander, Y. Y. Lee, M. Holtzapple, M. Ladisch,
Bioresour. Technol. 2005, 96, 673–86.
[91] J. Weil, P. Westgate, K. Kohlmann, M. R. Ladisch, Enzym. Microb. Tech. 1994, 16,
1002–1004.
[92] C. E. Wyman, B. E. Dale, R. T. Elander, M. Holtzapple, M. R. Ladisch, Y. Y. Lee,
Bioresour. Technol. 2005, 96, 1959–1966.
[93] J. R. Cherry, A. L. Fidantsef, Curr. Opin. Biotech. 2003, 14, 438–443.
[94] H. Esterbauer, W. Steiner, I. Labudova, A. Hermann, M. Hayn, Bioresour. Technol.
1991, 36, 51–65.
[95] O. Kirk, T. V. Borchert, C. C. Fuglsang, Curr. Opin. Biotech. 2002, 13, 345–351.
[96] M. Holtzapple, M. Cognata, Y. Shu, C. Hendrickson, Biotechnol. Bioeng. 1990, 36,
275–287.
[97] A. V Gusakov, A. P. Sinitsyn, Biotechnol. Bioeng. 1992, 40, 663–671.
[98] P. Ghosh, N. B. Pamment, W. R. B. Martin, Enzym. Microb. Tech. 1982, 4, 425–430.
[99] J. D. Wright, C. E. Wyman, K. Grohmann, Appl. Biochem. Biotech. 1988, 18, 75–90.
[100] J.-W. Lee, R. C. L. B. Rodrigues, T. W. Jeffries, Bioresour. Technol. 2009, 100, 6307–
6311.
80
[101] L. C. Teixeira, J. C. Linden, H. A. Schroeder, Appl. Biochem. Biotech. 2000, 84-86,
111–128.
[102] C. Tengborg, M. Galbe, G. Zacchi, Biotechnol. Progr. 2001, 17, 110–117.
[103] N. Mosier, C. Wyman, B. Dale, R. Elander, Y. Y. Lee, M. Holtzapple, M. Ladisch,
Bioresour. Technol. 2005, 96, 673–686.
[104] M. J. Taherzadeh, K. Karimi, Bioresources 2007, 2, 707–738.
[105] R. Rinaldi, F. Schüth, ChemSusChem 2009, 2, 1096–1107.
[106] R. P. Chandra, R. Bura, W. E. Mabee, A. Berlin, X. Pan, J. N. Saddler, Adv. Biochem.
Eng. Biot. 2007, 108, 67–93.
[107] D. Klemm, B. Heublein, H.-P. Fink, A. Bohn, Angew. Chem. Int. Ed. 2005, 44, 3358–
3393.
[108] J. Tollefson, Nature 2008, 451, 880–883.
[109] J. B. Binder, J. J. Blank, A. V. Cefali, R. T. Raines, ChemSusChem 2010, 3, 1268–
1272.
[110] F. Schüth, R. Rinaldi, Method for the Depolymerization of Cellulose, 2009, WO
2009/115075 A1.
[111] T. Balensiefer, J. Brodersen, G. D’Andola, K. Massonne, S. Freyer, V. Stegmann,
Method for Producing Glucose by Hydrolysis of Cellulose That Can Be Pretreated with
an Ionic Liquid Containing a Polyatomic Anion., 2008, WO2008090155.
[112] M. Klemens, G. D’Andola, V. Stegmann, W. Mormann, M. Wezstein, W. Leng,
Method for Breaking down Cellulose in Solution, 2009, US2009062524.
[113] R. Rinaldi, R. Palkovits, F. Schüth, Angew. Chem. Int. Ed. 2008, 47, 8047–8050.
[114] P. Domínguez de María, A. Martinsson, Analyst 2009, 134, 493–496.
[115] A. Pinkert, K. N. Marsh, S. Pang, M. P. Staiger, Chem.rev. 2009, 109, 6712–6728.
[116] C. Graenacher, Cellulose Solution, 1934, US1943176.
[117] T. Heinze, A. Koschella, Polim. Cienc. Tecnol. 2005, 15, 84–90.
[118] P. L. Ragg, P. R. Fields, P. B. Tinker, Phil. Trans. R. Soc. Lond. A 1987, 321, 537–547.
[119] L. F. Chen, C.-M. Yang, Quantitative Hydrolysis of Cellulose to Glucose Using Zinc
Chloride, 1984, US4452640.
[120] S. A. Barker, P. J. Somers, A. J. Beardsmore, B. L. F. Rodgers, Solubilisation and
Hydrolysis of Cellulose-Containing Materials, 1983, EP 0096497.
81
[121] R. A. Penque, Method of Hydrolyzing Cellulose to Monosaccharides, 1977,
US4018620.
[122] E. C. Sherrard, W. H. Gauger, Ind. Eng. Chem. 1923, 15, 63–64.
[123] H. G. Deming, J. Am. Chem. Soc. 1911, 33, 1515–1525.
[124] U. Wongsiriwan, Y. Noda, C. Song, P. Prasassarakich, Y. Yeboah, Energ. Fuel. 2010,
24, 3232–3238.
[125] A. M. J. Kootstra, H. H. Beeftink, E. L. Scott, J. P. Sanders, Biotechnol. Biofuels 2009,
2, 31.
[126] J. P. M. Sanders, P. H. M. De Bot, A. M. J. Kootstra, Method for Treating Vegetable
Material with Acid as Well as Products Obtained with This Method, 2009,
WO2009145617.
[127] A. M. J. Kootstra, N. S. Mosier, E. L. Scott, H. H. Beeftink, J. P. M. Sanders, Biochem.
Eng. J. 2009, 43, 92–97.
[128] J.-W. Lee, R. C. L. B. Rodrigues, H. J. Kim, I.-G. Choi, T. W. Jeffries, Bioresour.
Technol. 2010, 101, 4379–4385.
[129] Y. Lu, N. S. Mosier, Biotechnol. Progr. 2007, 23, 116–123.
[130] N. S. Mosier, J. J. Wilker, M. R. Ladisch, Biotechnol. Bioeng. 2004, 86, 756–764.
[131] N. S. Mosier, C. M. Ladisch, M. R. Ladisch, Biotechnol. Bioeng. 2002, 79, 610–618.
[132] N. S. Mosier, A. Sarikaya, C. M. Ladisch, M. R. Ladisch, Biotechnol. Progr. 2001, 17,
474–480.
[133] T. vom Stein, P. M. Grande, F. Sibilla, U. Commandeur, R. Fischer, W. Leitner, P.
Domínguez de María, Green Chem. 2010, 12, 1844–1849.
[134] C. Jiménez-González, P. Poechlauer, Q. B. Broxterman, B.-S. Yang, D. am Ende, J.
Baird, C. Bertsch, R. E. Hannah, P. Dell’Orco, H. Noorman, et al., Org. Process Res.
Dev. 2011, 15, 900–911.
[135] D. J. C. Constable, C. Jimenez-Gonzalez, R. K. Henderson, Org. Process Res. Dev.
2007, 11, 133–137.
[136] R.-J. van Putten, J. C. van der Waal, E. de Jong, C. B. Rasrendra, H. J. Heeres, J. G. de
Vries, Chem.rev. 2013, 113, 1499–1597.
[137] J. J. Bozell, G. R. Petersen, Green Chem. 2010, 12, 539–554.
[138] G. W. Huber, S. Iborra, A. Corma, Chem.rev. 2006, 106, 4044–4098.
[139] A. Corma, S. Iborra, A. Velty, Chem.rev. 2007, 107, 2411–2502.
82
[140] R. Palkovits, Angew. Chem. Int. Ed. 2010, 49, 4336–4338.
[141] F. M. A. Geilen, B. Engendahl, A. Harwardt, W. Marquardt, J. Klankermayer, W.
Leitner, Angew. Chem. Int. Ed. 2010, 49, 5510–5514.
[142] Sigma-Aldrich, 2-methyltetrahydrofuran Mater. Saf. Data Sheet [MSDS] 2012,
Retrived from http://www.sigmaaldrich.com.
[143] V. Antonucci, J. Coleman, J. B. Ferry, N. Johnson, M. Mathe, J. P. Scott, J. Xu, Org.
Process Res. Dev. 2011, 15, 939–941.
[144] R. K. Henderson, C. Jiménez-González, D. J. C. Constable, S. R. Alston, G. G. a.
Inglis, G. Fisher, J. Sherwood, S. P. Binks, A. D. Curzons, Green Chem. 2011, 13,
854–862.
[145] B. Comanita, S. Chemicals, Spec. Chem. Mag. 2006.
[146] V. Pace, P. Hoyos, L. Castoldi, P. Domínguez de María, A. R. Alcántara,
ChemSusChem 2012, 5, 1369–1379.
[147] D. F. Aycock, Org. Process Res. Dev. 2007, 11, 156–159.
[148] F. H. Newth, Adv. Carbohyd. Chem. 1951, 6, 83–106.
[149] C. J. Moye, Rev. Pure Appl. Chem. 1964, 14, 161–170.
[150] M. S. Feather, J. F. Harris, Adv. Carbohyd. Chem. Bi. 1973, 28, 161–224.
[151] B. F. M. Kuster, Starch - Stärke 1990, 42, 314–321.
[152] J. Lewkowski, ARKIVOC 2001, i, 17–54.
[153] N. Wierckx, F. Koopman, H. J. Ruijssenaars, J. H. de Winde, Appl. Microbiol. Biot.
2011, 92, 1095–1105.
[154] D. M. Alonso, J. Q. Bond, J. A. Dumesic, Green Chem. 2010, 12, 1493–1513.
[155] T. vom Stein, P. M. Grande, W. Leitner, P. Domínguez de María, ChemSusChem 2011,
4, 1592–1594.
[156] F. Carvalheiro, G. Garrote, J. C. Parajó, H. Pereira, F. M. Gírio, Biotechnol. Progr.
2005, 21, 233–243.
[157] J.-P. Lange, E. van der Heide, J. van Buijtenen, R. Price, ChemSusChem 2012, 5, 150–
166.
[158] A. P. Dunlop, Ind. Eng. Chem. 1948, 40, 204–209.
[159] D. L. Williams, A. P. Dunlop, Ind. Eng. Chem. 1948, 40, 239–241.
83
[160] M. R. Nimlos, X. Qian, M. Davis, M. E. Himmel, D. K. Johnson, J. Phys. Chem. A
2006, 110, 11824–11838.
[161] T. Willke, K.-D. Vorlop, Appl. Microbiol. Biot. 2001, 56, 289–295.
[162] M. Okabe, D. Lies, S. Kanamasa, E. Y. Park, Appl. Microbiol. Biot. 2009, 84, 597–
606.
[163] S. Baup, Ann. Pharm. 1836, 19, 29–38.
[164] R. Bentley, C. Thiessen, J. Biol. Chem. 1957, 226, 673–687.
[165] N. Winskillm, Microbiology 1983, 129, 2877–2883.
[166] P. Bonnarme, B. Gillet, a M. Sepulchre, C. Role, J. C. Beloeil, C. Ducrocq, J.
Bacteriol. 1995, 177, 3573–3578.
[167] T. vom Stein, Organic-Acid-Catalysed Selective Fractionation of Lignocellulose,
Diplomarbeit, 2010.
[168] P. Domínguez de María, T. vom Stein, P. M. Grande, F. Sibilla, W. Leitner, Integrated
Process for the Selective Fractionation and Separation of Lignocellulose in Its Main
Components., 2011, EP 11154705.5.
[169] A. M. J. Kootstra, H. H. Beeftink, E. L. Scott, J. P. M. Sanders, Biochem. Eng. J. 2009,
46, 126–131.
[170] A. M. J. Kootstra, H. H. Beeftink, E. L. Scott, J. P. Sanders, Biotechnol. Biofuels 2009,
2, 31.
[171] N. S. Mosier, J. J. Wilker, M. R. Ladisch, Biotechnol. Bioeng. 2004, 86, 756–764.
[172] N. Li, G. a. Tompsett, T. Zhang, J. Shi, C. E. Wyman, G. W. Huber, Green Chem.
2011, 13, 91–101.
[173] J. A. Vaselenak, I. E. Grossmann, A. W. Westerberg, Ind. Eng. Chem. Process Des.
Dev. 1986, 25, 357–366.
[174] P. Dominguez de Maria, H. Kayser, F. Rodríguez-Ropero, W. Leitner, M. Fioroni, RSC
Adv. 2013, DOI 10.1039/c3ra41307a.
[175] H. Kayser, Mechanistic Studies on Organic Acid Catalyzed Hydrolysis of Glycosidic
Bonds, Diplomarbeit, 2010.
[176] E. L. Shock, Am. J. Sci. 1995, 295, 496–580.
[177] C. I. Heck, E. G. de Mejia, J. Food Sci. 2007, 72, R138–R151.
84
[178] C. M. Pagliosa, K. N. de Simas, R. D. M. C. Amboni, A. N. N. Murakami, C. L. O.
Petkowicz, J. de D. Medeiros, A. C. Rodrigues, E. R. Amante, Ind. Crop. Prod. 2010,
32, 428–433.
[179] T. E. Timell, Wood Sci. Technol. 1967, 1, 45–70.
[180] T. Klement, S. Milker, G. Jäger, P. M. Grande, P. Domínguez de María, J. Büchs,
Microb. Cell Fact. 2012, 11, 43.
[181] Q. Yan, M. Modigell, Chem. Eng. Trans. 2012, 29, 601–606.
[182] M. Pidwirny, “Introduction to the Oceans”. Fundamentals of Physical Geography,
2006.
[183] C. S. K. Lin, R. Luque, J. H. Clark, C. Webb, C. Du, Energ. Environ. Sci. 2011, 4,
1471–1479.
[184] A. Johansson, O. Aaltonen, P. Ylinen, Biomass 1987, 13, 45–65.
[185] M. Taherzadeh, K. Karimi, Bioresources 2007, 2, 472–499.
[186] B. Van der Bruggen, C. Vandecasteele, Desalination 2002, 143, 207–218.
[187] B. Zhao, L. Greiner, W. Leitner, RSC Adv. 2012, 2, 2476–2479.
[188] B. Zhao, Syntheses and Applications of Ionic Liquids as Solvents and Reactants, PhD
Thesis, 2012.
[189] A. Berlin, M. Balakshin, N. Gilkes, J. Kadla, V. Maximenko, S. Kubo, J. Saddler, J.
Biotechnol. 2006, 125, 198–209.
[190] L. T. Fan, Y. Lee, D. H. Beardmore, Biotechnol. Bioeng. 1980, 22, 177–199.
[191] P. Väljamäe, G. Pettersson, G. Johansson, Eur. J. Biochem. 2001, 268, 4520–4526.
[192] M. Hall, P. Bansal, J. H. Lee, M. J. Realff, A. S. Bommarius, FEBS J. 2010, 277,
1571–1582.
[193] P. Bansal, B. J. Vowell, M. Hall, M. J. Realff, J. H. Lee, A. S. Bommarius, Bioresour.
Technol. 2012, 107, 243–250.
[194] A. Oren, J. Ind. Microbiol. Biot. 2002, 28, 56–63.
[195] S. Grant, D. Y. Sorokin, W. D. Grant, B. E. Jones, S. Heaphy, Extremophiles 2004, 8,
421–429.
[196] C. Liang, Y. Xue, M. Fioroni, F. Rodríguez-Ropero, C. Zhou, U. Schwaneberg, Y. Ma,
Appl. Microbiol. Biot. 2011, 89, 315–326.
85
[197] J. Pottkämper, P. Barthen, N. Ilmberger, U. Schwaneberg, A. Schenk, M. Schulte, N.
Ignatiev, W. R. Streit, Green Chem. 2009, 11, 957–965.
[198] D. M. Alonso, J. Q. Bond, J. A. Dumesic, Green Chem. 2010, 12, 1493–1513.
[199] P. Domínguez de María, ChemSusChem 2011, 4, 327–329.
[200] G. W. Huber, J. N. Chheda, C. J. Barrett, J. a Dumesic, Science (80-. ). 2005, 308,
1446–1450.
[201] B. Kamm, P. R. Gruber, M. Kamm, Biorefineries — Industrial Processes and
Products. Status Quo and Future Directions, Wiley-VCH, Weinheim, 2010.
[202] A. A. Rosatella, S. P. Simeonov, R. F. M. Frade, C. A. M. Afonso, Green Chem. 2011,
13, 754–793.
[203] X. Tong, Y. Ma, Y. Li, Appl. Catal. A-Gen. 2010, 385, 1–13.
[204] F. M. A. Geilen, T. vom Stein, B. Engendahl, S. Winterle, M. A. Liauw, J.
Klankermayer, W. Leitner, Angew. Chem. Int. Ed. 2011, 50, 6831–6834.
[205] W. Xu, Q. Xia, Y. Zhang, Y. Guo, Y. Wang, G. Lu, ChemSusChem 2011, 4, 1758–
1761.
[206] J. Julis, M. Hölscher, W. Leitner, Green Chem. 2010, 12, 1634–1639.
[207] S. Lima, M. M. Antunes, M. Pillinger, A. a. Valente, ChemCatChem. 2011, 3, 1686–
1706.
[208] M. E. Zakrzewska, E. Bogel-Łukasik, R. Bogel-Łukasik, Chem.rev. 2011, 111, 397–
417.
[209] A. D. Patel, J. C. Serrano-Ruiz, J. A. Dumesic, R. P. Anex, Chem. Eng. J. 2010, 160,
311–321.
[210] Y. Román-Leshkov, C. J. Barrett, Z. Y. Liu, J. a Dumesic, Nature 2007, 447, 982–985.
[211] H. C. Silberman, J. Org. Chem. 1961, 26, 1967–1969.
[212] T. J. Christian, M. Manley-Harris, R. J. Field, B. a Parker, J. Agric. Food. Chem. 2000,
48, 1823–1837.
[213] B. Kim, J. Jeong, S. Shin, D. Lee, S. Kim, H.-J. Yoon, J. K. Cho, ChemSusChem 2010,
3, 1273–1275.
[214] J. Speck, Adv. Carbohyd. Chem. 1958, 13, 63–103.
[215] S. Angyal, Glycoscience 2001, 215, 1–14.
[216] S. H. Bhosale, M. B. Rao, V. V Deshpande, Microbiol. Rev. 1996, 60, 280–300.
86
[217] T. Ståhlberg, J. M. Woodley, A. Riisager, Catal. Sci. Technol. 2012, 2, 291.
[218] R. Huang, W. Qi, R. Su, Z. He, Chem. Commun. 2010, 46, 1115–1117.
[219] R. Dicosimo, J. McAuliffe, A. J. Poulose, G. Bohlmann, Chem. Soc. Rev. 2013, DOI:
10.1039/c3cs35506c.
[220] Y. B. Tewari, Appl. Biochem. Biotech. 1990, 23, 187–203.
[221] K. Visuri, A. M. Klibanov, Biotechnol. Bioeng. 1987, 30, 917–920.
[222] A. S. Mamman, J.-M. Lee, Y.-C. Kim, I. T. Hwang, N.-J. Park, Y. K. Hwang, J.-S.
Chang, J.-S. Hwang, Biofuels, Bioprod. Bioref. 2008, 2, 438–454.
[223] G. P. Chiusoli, US3025320: Process for Preparing Itaconic Acid, and 2, 3-Butadienoic
Acid, 1962.
[224] T. vom Stein, P. M. Grande, H. Kayser, F. Sibilla, W. Leitner, P. Domínguez de María,
Green Chem. 2011, 13, 1772–1777.
[225] M. Lever, Anal. Biochem. 1972, 47, 273–279.
87
LIST OF PATENTS & PUBLICATIONS
[1] P. Domínguez de María, T. vom Stein, P.M. Grande, F. Sibilla, W. Leitner,
EP 11154705.5, 2011, Integrated process for the selective fractionation and
separation of lignocellulose in its main components.
[2] T. Klement, S. Milker, G. Jäger, P.M. Grande, P. Domínguez de María, J. Büchs,
Microb. Cell Fact. 2012, 11, 43, Biomass pretreatment affects Ustilago maydis in
producing itaconic acid.
[3] P.M. Grande, C. Bergs, P. Domínguez de María, ChemSusChem 2012, 5, 1203-1206,
Chemo-enzymatic conversion of glucose into 5-hydroxymethylfurfural in seawater.
[4] P.M. Grande, P. Domínguez de María, Bioresource Technol. 2012, 104, 799-802,
Enzymatic hydrolysis of microcrystalline cellulose in concentrated seawater.
[5] T. vom Stein, P.M. Grande, W. Leitner, P. Domínguez de María, ChemSusChem 2011,
4, 1592-1594, Iron-catalyzed furfural production in biobased biphasic systems: from
pure sugars to direct use of crude xylose effluents as feedstock.
[6] T. vom Stein, P.M. Grande, H. Kayser, F. Sibilla, W. Leitner, P. Domínguez de María,
Green Chem. 2011, 13, 1772-1777, From biomass to feedstock: one-step fractionation
of lignocellulose components by the selective organic acid-catalyzed depolymerization
of hemicellulose in a biphasic system.
[7] T. vom Stein, P. Grande, F. Sibilla, U. Commandeur, R. Fischer, W. Leitner,
P. Domínguez de María, Green Chem. 2010, 12, 1844-1849, Salt-assisted
organic-acid-catalyzed depolymerization of cellulose.