Biochemical Control Aspects
in
Lignin Polymerization
Anders Holmgren
D o c t ora l Th e s i s
R oyal Institu te of Technol ogy
S ch o ol of Ch e mi ca l S ci e n ce s a nd E ng i n e e r i ng
D e par tment of Fi br e an d Po ly mer Te ch n ol og y
D i visi o n of Wo od Ch emis try a n d Pul p Te c hn ol og y
Stockhol m 2008
Biochemical Control Aspects
in
Lignin Polymerization
Supervisors:
Professor Gunnar Henriksson
and
Dr Liming Zhang
AKADEMISK AVHANDLING
som med tillstånd av Kungliga Tekniska Högskolan i Stockholm framlägges
till offentlig granskning för avläggande av teknologie doktorsexamen fredagen
den 29 februari 2008, kl 10:00 i STFI-salen, STFI-Packforsk, Drottning
Kristinas Väg 61. Avhandlingen försvaras på engelska.
© Anders Holmgren 2008
Paper I Reproduced by permission of The Royal Society of Chemistry
TRITA-CHE-Report 2008:10
ISSN 1654-1081
ISBN 978-91-7178-864-1
Abstract
Lignins are produced by all vascular plants and they represent one of the
most abundant groups of biopolymers in nature. Lignin chemistry research,
which has been of great importance for the progress of pulping technologies,
has been plagued by the difficulties of its isolation and characterization. The
pioneering work of Karl Freudenberg in the 1950’s with synthetic models of
lignin paved the way for a detailed structural characterization of many lignin
substructures. His work with the so-called “synthetic lignins” or
dehydrogenative polymers (DHP) also laid a foundation for understanding how
different lignin substructures are formed, reinforcing the already existing
theory
of
lignin
polymerization.
However,
subsequent
structural
characterizations of DHPs and lignins have repeatedly put this theory to the
test. In the past decade, even a new radically different hypothesis for lignin
polymerization has emerged and is sustained by a few researchers in the
field.
In this work, DHPs were produced from phenolic monomers, mostly coniferyl
alcohol, a common lignin monomer, in a variety of reaction conditions. This
was done in order to establish how different chemical factors, potentially
active in the plant cell wall during lignin polymerization, influence the
polymer’s final properties. In the presence of nicotine amide adenine
dinucleotide (NADH), a quinone methide model, which is an intermediate
formed during lignin polymerization, was effectively reduced. An equivalent
reduced structure was produced during DHP synthesis in the presence of
NADH. These studies showed that reduction might take place during oxidative
polymerization, possibly explaining how reduced lignin structures are formed
in the plant cell wall. Another reductive agent, ascorbic acid, was also tested
during synthesis of DHPs. It displayed a totally different effect than NADH,
probably due to its anti-oxidant nature, by altering the final amounts of
certain inter-unit substructures, in favour of β-O-4′ structures, which are so
prominent in natural lignins. Furthermore, the new suggested model for
lignin polymerization, stating that lignin itself possesses the ability for
template replication, was tested by synthesizing DHPs in the presence of a
simple β-β′ substructure model. The DHPs produced the same amounts of β-β′
substructures as a control synthesis without the model structure, indicating
that no replication had occurred. Finally, the role of the monolignol γ-carbon
oxidation state in lignin polymerization, was studied. Hypothetically, ligninlike polymers could be produced by the plant, using monolignol biosynthetic
precursors which exhibit γ-carbonyl groups instead of an alcohol group, like
the common lignin monomer. Synthetic lignins produced with ferulic acid,
coniferaldehyde and the normal monolignol, coniferyl alcohol, displayed
important differences in chemical and physical properties. Both the ferulic
acid and coniferaldehyde polymers exhibited almost no saturated inter-unit
substructures and very few cyclic structures, both of which are very common
in coniferyl alcohol dehydrogenative polymers and natural lignins. This could
have significant implications for the formation of an important type of lignin
carbohydrate complexes (LCC). Also the hydrophobicity of the alcohol-type
polymer was lower than the other two. The biological implications of all these
findings are discussed and some suggestions are made to explain how all
these factors might affect lignin polymerization and structure in nature.
Sammanfattning
Lignin tillverkas av alla kärlväxter och är därmed en av de vanligaste
naturligt förekommande polymererna på jorden. Forskningen kring ligninets
kemi, som har varit viktig för utvecklingen av massatillverkningsprocesser,
har länge lidit av svårigheter vad gäller dess isolering och karaktärisering.
Karl Freudenbergs arbete med syntetiska ligninmodeller (DHP) på 1950-talet
lade grunden för den kemiska strukturbestämningen av många av ligninets
viktiga bindningstyper. Genom dessa upptäckter förstärktes den dåvarande,
och idag allmänt accepterade, teorin för ligninpolymeriseringen. Efterföljande
forskning kring DHP- och ligninstrukturen har vid upprepade tillfällen satt
denna teori på prov. Några forskare har även utvecklat ytterligare en hypotes,
som skiljer sig markant från den befintliga teorin.
I föreliggande arbete har ligninmodeller (DHP) framställts av olika fenoliska
monomerer och i olika reaktionsförhållanden med syftet att försöka bestämma
hur och vilka kemiska faktorer, som kan vara aktiva i växternas cellväggar,
påverkar ligninpolymeriseringen och därmed ligninets egenskaper. Studier
med två biologiskt viktiga reduceringsmedel visade att sådana ämnen kan
spela en roll i ligninpolymeriseringen och påverka dess struktur. För det
första påvisades hur en kinonmetidmodell, som föreställer en vanlig
ligninpolymeriseringsintermedjär, kunde reduceras av nikotinamid adenin
dinukleotid (NADH) och bilda en så kallad reducerad ligninstruktur.
Dessutom användes samma ämne, NADH, under polymerisering av DHP,
vilket producerade bl a motsvarande reducerade strukturer. Dessa studier
påvisade hur reduktion kan äga rum under oxidativ polymerisering och
därmed ge en förklaring till hur reducerade ligninstrukturer bildas i
cellväggen. För det andra visades att bildningen av vissa bindningstyper (β-O4′, den vanligaste bindningstypen i lignin) gynnades framför andra när DHP
syntetiserades i närvaro av en antioxidant, askorbinsyra, som har
rapporterats förekomma i cellväggen. En del av arbetet uppmärksammar den
nya modellen som framhåller att ligninpolymeriseringen står under strikt
biokemisk kontroll genom att påstå bl a, att lignin kopierar sig självt under
polymeriseringen (eller replikerar sig) genom ”template” polymerisering. Ett
experiment där DHP syntetiserades i närvaro av en enkel modell, en β-β′
struktur som är en vanligt förekommande bindningstruktur i lignin, visade
att mängden β-β′ strukturer inte ökade överhuvudtaget i modellens närvaro
jämfört med en motsvarande syntes utan modellen. Därigenom kunde den
enligt hypotesens förväntade kopiering inte påvisas i detta fall. Slutligen har
också den roll studerats, som monolignolernas (ligninmonomererna) kemiska
struktur spelar i ligninpolymeriseringen med avseende på γ-kolets
oxidationstal. I teorin skulle någon ligninlik polymer kunna bildas i
cellväggen med vissa av monolignolernas biosyntetiska föregångare som har
en aldehyd- eller karboxylsyragrupp i γ-kolet i stället för en alkoholgrupp,
som de vanliga monolignolerna har. DHP har syntetiserats med ferulsyra,
koniferylaldehyd och koniferylalkohol (en vanlig monolignol) var för sig och
polymerernas kemiska och fysiska egenskaper har jämförts med varandra. De
mer oxiderade monomererna, ferulsyra och koniferylaldehyd (båda med
karbonylgrupper) bildade mera hydrofobiska polymerer och dessutom förekom
i stort sett inga saturerade bindningstyper, vilka annars är vanliga i
koniferylalkohol-DHP och i naturligt lignin. Betydelsen för bildningen av en
viktig typ av lignin-kolhydrat komplex (LCC) behandlas därefter. Alla resultat
diskuteras i ett biologiskt sammanhang och förslag lämnas på hur alla dessa
faktorer påverkar ligninpolymeriseringen och därigenom också dess struktur.
If we knew what it was we were doing,
it would not be called research, would it?
Albert Einstein
List of Papers
This thesis is based on the following works which will be referred to by their
numbers:
Paper I
A. Holmgren, G. Brunow, G. Henriksson, L. Zhang and J. Ralph. “NonEnzymatic Reduction of Quinone Methides during Oxidative Coupling of
Monolignols: Implications for the Origin of Benzyl Structures in Lignins.”
Organic & Biomolecular Chemistry, 2006, 4, 3456-3461.
Paper II
A. Holmgren, G. Henriksson and L. Zhang. “Effects of a Biologically Relevant
Anti-oxidant on the Dehydrogenative Polymerization of Coniferyl Alcohol.”
Manuscript
Paper III
A. Holmgren, L. Zhang and G. Henriksson. “In vitro Monolignol
Dehydrogenative Polymerization in the Presence of a Candidate Dimer
Template Model.” Manuscript
Paper IV
A. Holmgren, M. Norgren, L. Zhang and G. Henriksson. “On the Role of the
Monolignol γ-carbon in Lignin Biosynthesis.” Manuscript
The author’s contribution to each paper:
Papers I-III
Principal author, performed all experimental work.
Paper IV
Principal author, performed all experimental work except for the preparation
of spin-coated surfaces and their characterization.
Related Material:
A. Holmgren, L. Zhang, G. Brunow and G. Henriksson. “Origin of Reduced
Lignin Structures: Quinone Methides” 13 t h ISWFPC Proceedings, Auckland
NZ, 2005, 3, 147-15
Table of contents
INTRODUCTION
1
O BJECTIVES
1
BACKGROUND
2
B IOLOGICAL P ERSPECTIVES
L IGNINS S TRUCTURE AND C HEMISTRY
L IGNIN B IOSYNTHESIS
L IGNIN AMONG O THER P OLYCYCLIC B IOPOLYMERS
2
5
9
15
METHODS
19
RESULTS AND DISCUSSION
22
R EDUCTION D URING O XIDATIVE C OUPLING (P APER I)
E FFECTS OF AN ANTI - OXIDANT DURING L IGNIN P OLYMERIZATION (P APER II)
T ESTING THE L IGNIN T EMPLATE P OLYMERIZATION M ODEL (P APER III)
O N THE R OLE OF THE M ONOLIGNOL γ- CARBON (P APER IV)
22
26
30
33
CONCLUSIONS
42
ACKNOWLEDGEMENTS
44
REFERENCES
45
GLOSSARY
59
Introduction
In spite of more than 100 years of research in the field of wood chemistry,
neither the structure of lignins * nor their polymerization is fully understood.
Much of the research on lignins has been driven by the development of
pulping technologies and it has therefore been focused on their degradation
and chemical modification to improve pulp properties. So a considerable
amount of fundamental knowledge has been obtained over the years making
lignins the best known biopolymers of their kind. Nevertheless, in comparison
with the knowledge collected on other common biopolymers such as cellulose,
proteins and DNA, the understanding of lignins is vague. Merely comparing
the available structural data on lignins with these other biopolymers makes
them look like chaotic and complex compounds. Yet the plant, which deposits
lignin into the walls of its cells, must somehow be able to control its
structure and consequently its properties.
Lignins’ particular chemical and physical properties are believed to
depend on the polymerization process, which leads to the lignification of the
plant cell wall. The polymerization step in lignin biosynthesis has been a
matter of discussion and speculation for a long time and recently a debate
has arisen over the control mechanisms at work in the plant cell wall. This is
the main subject of the present work.
This thesis starts with a background which describes lignins from two
different but related contexts: biology and chemistry. The biosynthesis of
lignins is also considered in the light of our knowledge to date and a special
focus is placed on the polymerization step by presenting the currently
accepted theory and its challenges. In addition, lignins and their
biopolymerization are compared to other natural polycyclic polymers:
melanins. Subsequently there is a general description of the experimental
methods applied during the research that has led to the present thesis,
followed by a summary of the results obtained and some discussion. The
conclusions in summary form are presented at the end, and additional
detailed information can be found in the appended articles.
Objectives
It is the aim of this work to shed more light into how plants control lignin
polymerization and structure. Understanding what roles the different
biochemical factors play in building lignin is useful, e.g., when “engineering”
a lignin structure that will improve pulping technologies. Based on existing
models for lignin polymerization, experiments have been performed to gain
understanding on the potential biochemical factors influencing the
polymerization process. Factors such as reductive agents, possible template
effects and the importance of the monomeric chemical structure were
assessed for their potential influence upon the structure of the lignin
polymer.
* In the present work, the term “lignins” is used to refer to the wide variety of naturally
occurring lignin-type polymers that fulfil similar functions in the cell walls of plants.
1
Background
Biological Perspectives
The Appearance and Roles of Lignins
In plant history, the appearance of vascular tissues such as xylem and
phloem implied a cellular specialization and a dedicated infrastructure for
water and nutrient transport. The emergence of the first vascular plants,
around 430 million years ago, is associated with the appearance of lignins [13]. Lignins are therefore believed to have been crucial for the development of
large land plants [4]. Today, there is a great variety of vascular plants
including club mosses, horsetails, ferns and the seed-based gymnosperms
and angiosperms. Woody plants, called softwoods and hardwoods, are part of
the gymnosperm and angiosperm groups respectively. Vascular plants have
the ability to grow taller than non-vascular plants, a factor which is
important in the competition for sunlight. Their rigid structures have
efficiently overcome the challenge of gravity; as is evident in certain redwoods
and eucalypts which reach heights of over 100 meters. The ability to grow tall
is also dependent on an effective system for the distribution of water to the
whole plant. Lignins are believed to play a role which is essential for both
rigidity and water transport.
Lignins can be found mostly in the cell walls and middle lamella of tissues
involved in mechanical support and water transport like phloem fibres,
schlerenchyma cells and mainly in the xylem tissue [1,5]. The xylem exhibits
a remarkable morphological variation and specialization at the cellular level.
In some cases the same cell type, tracheids, is responsible for both support
and water conduction. Then there are parenchyma cells, which are for
storage. Another type of diversification and specialization can be observed in
the difference between earlywood and latewood tracheids in the xylem of
softwoods.
Earlywood
tracheids,
specialized
for
water
transport,
display a large lumen (Figure 1) and
thin cell wall whereas the opposite
relation is observed in latewood
tracheids, where a thick cell wall is
favoured to provide structural rigidity
while the lumen is practically absent
in many cases. Angiosperms have
developed a particular cell type for
water transport called vessel elements
while the structural rigidity function
is taken care of by other cell types
such as libriform fibres.
Figure 1 Sof twood xylem tissue displaying
late wood and earlywood tracheids (crosssection photomicrograph of spruce, bar:
100 µm).
2
As mentioned above, the first lignins appeared together with the
development of the first vascular plants. Lignins are believed to be part of the
product of the evolution of phenol metabolism and more specifically the
phenylpropanoid pathway. This secondary metabolic pathway is thought to be
important in the adaptation of plants to land life providing, among other
things, protection against UV-light and pathogens [6-8]. Other phenolic
compounds derived from this pathway are lignans, the phenolic portion of
suberin, flavonoids, coumarins, styrylpyrones and stillbenes, some of which
are present in bryophytes, which are non-vascular. Lignins have commonly
been attributed not only the roles of increasing cell wall rigidity and
hydrophobicity but also of improving the plant’s pathogenic defence [5,9-11].
Lignins fix cells to each other and harden the cell wall. A lignified cell wall is
considered necessary for effective water conduction and the hydrophobic
nature of lignins enhances cell wall impermeability. Furthermore, lignins’
chemical properties are believed to improve the cell wall’s efficiency as a
barrier against microbial attack [12]. Looking at the enormous diversity of
plants that produce lignins, it is interesting to notice their chemical
variations as well. Since diversification can mean gain or loss of functions, to
what extent does the lignin structure need to be conserved to retain the
necessary properties? In other words, to what extent does the plant need to
control its lignin structure and chemistry?
Figure 2 a. Wheat leaf schlerenchyma cells (cross-section TEM)
showing middle lamella (ML), primary wall (PW) and secondary wall
(SW). Bar: 1µm. From Brett et al. 1996 [5] with kind permission of
Springer Science and Business Media. b. Model of wood cell wall
displaying Middle lamella (ML), Primary wall (P) and Secondary wall
(S1, S2 and S3). Adapted f rom Côté 1967 [13].With permission of
The University of Washington Press.
The Plant Cell Wall and the Lignification Process
The cell wall acts as a skeleton in plants, bringing shape and strength to
the cell and rigidity to the whole plant. In woody plants, over 95% of the dry
weight is attributed to the cell wall. It is a dense material, limiting the
movement of large molecules, such as proteins or nucleic acids, in and out of
the cell. Consequently, it also works as an effective barrier against
pathogenic organisms, blocking even viruses. During cell growth, the cell wall
is deposited in a series of layers (Figure 2). The first layer formed is the one
between two neighbouring cells, the generally very thin middle lamella (0.1-1
3
µm). The primary cell wall then follows, varying between 0.1 and 0.3 µm in
thickness. Certain specialized cells, e.g., tracheids, fibres and vessels deposit
a secondary cell wall which is normally thicker than the primary cell wall
(1-5 µm). For cells that do lignify their cell walls, lignification means a nonreversible process. In the xylem the completion and lignification of the
secondary cell wall is followed by cell death. Accordingly, this process implies
a terminal step for the cell and tissue, determining the final ultrastructure of
the cell wall and its properties. The morphological distribution of lignin in
cell walls of woody plant xylem has been studied with many methods but the
best results have been obtained by UV and electron microscopy of woody
plants [14,15]. It has generally been observed that the concentration of lignin
is very high in the middle lamella and primary cell wall, ranging between 40
and 100% whereas the secondary wall displays a lignin concentration of
about 20%.
The material of the cell wall is a composite of mainly three types of
biopolymers: cellulose, hemicelluloses, and lignins. The cell wall is thus a
highly heterogeneous material that can be divided into two phases: the
microfibrillar phase composed of cellulose in a more or less homogenous
phase with a high degree of crystallinity and a heterogeneous phase, the
matrix, composed of different polysaccharides, proteins and phenolics (e.g.,
lignins) [5]. The cell wall synthesis, i.e., the deposition of the different
components, generally follows an order. Thus lignins are never deposited in
the absence of carbohydrates [1] and they are deposited from the outside and
inwards [16,17]. Lignification follows three stages in xylem tissue. It is
believed to start in the cell corner and middle lamella when the deposition of
carbohydrates in the first layer (S1) of the secondary cell wall, begins. The
second stage starts when the second layer (S2) of carbohydrates is deposited
in the secondary cell wall, and the main lignification stage starts when
cellulose fibrils are deposited in the third layer (S3) of the secondary wall
(Figure 2b) [18,19]. Lignification proceeds, as described, towards the inside of
the cell through the secondary cell wall, incrusting it by filling the pores of
the polysaccharide matrix [20,21]. The carbohydrate contribution to the
matrix varies considerably in composition within the cell wall. The middle
lamella is mainly composed of pectins while the primary and secondary cell
walls are made of different hemicelluloses. In addition, hemicelluloses vary
between cell types and plant species. The physical and chemical conditions
such as charge, hydrophobicity/hydrophilicity and pH are affected by the
polysaccharides’ own properties. As a result, the local environment into
which lignin is deposited varies. This raises questions about what influence,
if any, the environment, and more specifically each type of carbohydrate,
exerts on the deposition of lignins [22-25].
So far, the understanding of the organization of the different components
in the plant cell wall and how these interact with each other is very limited.
This is especially important when ascribing the general properties of the cell
wall to the different components. The role(s) of each component should be
regarded in the context of the cell wall as a whole [11]. Maybe the most
studied type of interactions is the covalent linkages of lignin to
carbohydrates, called lignin carbohydrate complexes (LCC) [26-30]. How LCCs
are actually formed in the cell wall is a matter of debate, and it is further
complicated by the fact that there are several types of LCCs. It is nonetheless
generally accepted that LCCs exist naturally in the plant cell wall and they
4
may even cross-link the matrix polysaccharides to each other [30]. Noncovalent interactions between lignins and carbohydrates have been less
investigated but they could involve, e.g., high interfacial affinity of lignin to
hemicelluloses rather than cellulose [31] and to hydrophobic regions of the
cell wall matrix [32]. Several tentative models describing possible
organizations of the different components of the lignified cell wall have been
proposed [33-37].
Lignins Structure and Chemistry
Description and Characterization
Chemically, lignins can be described as aromatic polymers of methoxylated
phenylpropanoid units, connected by both ether and carbon-carbon bonds.
They are optically inactive [38] although they contain multiple chiral centres
and are generally considered to be branched and to act as networks that
cross-link cell wall components. No satisfactory exact definition is available
[11,38]. Broad and simple definitions leave out important aspects of lignins
and reflecting upon their obvious complexity, it is doubtful there will ever be
one. Several molecular models of lignins have been proposed [39-42] giving an
idea of the complexity involved (see Figure 3 for a softwood lignin model).
These models are meant to consolidate the available experimental data and
produce a qualitative representation of the native polymer, i.e., the
unmodified polymer as it exists in situ. New models have updated previous
ones as new features of lignins have been revealed. Nevertheless, no thorough
3-dimensional structural determination of any lignin has been achieved. The
closest to such a determination has been achieved by a Raman spectroscopy
study of spruce secondary cell wall. The study revealed how the aromatic
rings in the lignin are positioned relatively parallel to each other and in the
plane of the cell wall [43].
It is not possible to prepare a lignin sample that represents the whole
lignin in its original state with any of the currently available methods. There
are two reasons: lignin is heterogeneous within the structure itself and it is
strongly associated with cell wall polysaccharides in different ways [16]
making its complete isolation very difficult. Most structure determination
studies have been made on isolated lignin samples, which traditionally
involve such treatments that induce chemical changes [44-46]. Major
advances in lignin analysis have been achieved by the development of
acidolytic degradative methods, which selectively cleave certain bond-types in
lignins. These methods, together with different chromatographic and mass
spectroscopic tools, provide a good deal of structural information [47]. There
are also so-called nondegradative methods, which basically involve common
spectroscopic techniques like IR and UV. Among these, probably the most
powerful method for structural characterization of lignins is solution-state
nuclear magnetic resonance spectroscopy (NMR) [47-49]. However there are
three generic analytical limitations despite the latest developments [11]:
ƒ Only fractions of lignin can be characterized. There is no method to
completely depolymerize lignin and make it all available for analysis.
ƒ Lignin isolation is plagued by contamination by other cell wall
components, mostly carbohydrates.
ƒ Rough total lignin content determinations are uncertain.
5
Figure 3 Sof twood lignin model proposed by Brunow, et al 1998 [39] Reprinted with
permission. Copyright (1998) American Chemical Society.
Since there is no isolation procedure available today to produce
quantitatively and chemically unmodified lignin, its true molecular weight is
unknown. Instead, molecular weight determinations are made on isolated
lignins using a variety of methods such as Size exclusion chromatography
(SEC) among others. Reports of isolated lignin molecular weights range in
general from 10 2 to 10 6 g mol - 1 [46]. A molecular weight of 10 6 g mol - 1
corresponds roughly to a degree of polymerization (DP) of 5000. Molecular
weight averages of approximately 20 000 g mol - 1 (DP of ~100) have been
6
reported for spruce milled wood lignin [50]. For technical lignins, such as
lignosulfonates, the molecular weights range from 100 to 10 000 g mol - 1 [51]
(DP from ~1 to ~50). Anyhow, since the isolation procedures most likely affect
the size of the polymers, any molecular weight determination of such samples
will represent rather low values of the actual polymers in nature.
Chemical Heterogeneity
There are three monomers, called monolignols, from which most lignins
are built: coniferyl alcohol, sinapyl alcohol and p-coumaryl alcohol. In the
polymer, they are found as guaiacyl, syringyl and p-hydroxyphenyl residues
respectively (Figure 4).
Figure 4 The common monolignols - and carbon number
nomenclature - and their respective residues as f ound on the
lignin polymer.
The evident variety of lignins between different plant species has led to the
development of different classification systems. A general way, at first
adopted by most researchers, was to follow plant taxonomy. According to this
simple classification, gymnosperm lignin exhibits mostly guaiacyl residues,
angiosperms, a mix of guaiacyl and syringyl residues and grasses, a mix of phydroxyphenyl, guaiacyl and syringyl residues. However this classification
leaves out herbaceous angiosperm lignins, vascular cryptogam lignins (e.g.,
ferns) and overlooks exceptions such as compression wood of gymnosperms
and the “exceptional” conifers which display lignin of a guaiacyl-syringyl type
[52]. Thus a classification that is more consistent has been proposed [53],
based on the lignins’ chemical characteristics rather than on the taxonomy of
the plants from which the lignin comes. The basis for this classification is the
predominance of each of the three lignin residues, and has been developed
from two main lignin groups into 4 main groups [45]: Type-G lignins, Type GS lignins, Type H-G-S lignins and Type H-G lignins (G, S and H stand for
guaiacyl, syringyl and p-hydroxyphenyl respectively). The complexity of this
classification system for lignins represents one aspect of their substantial
chemical diversity.
7
Not only do lignins vary between plant species, they also vary within the
same plant. Differences in monomer composition, from fibres compared to
vessels, have been observed in a wide variety of hardwoods [54], and
differences in lignins between cell types have also been observed in spruce
[55]. Furthermore, physical and chemical differences have been noted within
the cell wall, between lignins from the middle lamella as well as from the
secondary wall [56,57].
Lignins also exhibit an important heterogeneity of substructures with six
major inter-unit structures along with a number minor variations and
additional features. The inter-unit bonds are generally very stable carboncarbon and ether bonds (Figure 5 and Table 1). An additional variation aspect
in inter-unit bond distribution is observed depending on the type of lignin,
e.g., syringyl lignin displays no inter-unit bonds involving the 5-carbon.
Apart from this, there are considerable variations in bond-type distribution
within the plant cell wall. Lignins originating from the middle lamella of some
gymnosperms contain more “condensed” lignin (i.e., lignin poor in β-O-4′
bonds) than those coming from the secondary cell walls [58,59]. A similar
tendency has been noted in poplar [60]. Monomer distribution in the cell wall,
which is probably associated with bond-type distribution, has been reported
to follow a general pattern in both gymnosperms and angiosperms [36].
According to this, the cell corner and middle lamella contain mostly phydroxyphenyl and/or guaiacyl residues while the secondary cell wall
contains mostly guaiacyl and/or syringyl residues. However, as far as
softwoods are concerned, there is some disagreement in this matter [61].
Figure 5 The most common bonds in lignins. Some evidence exists
that β -1′ and 5-5′ are actually derived from trimeric f orms in lignins:
spirodienone [62,63] and dibenzodioxocin [64] respectively as
revie wed by Ralph et al. 2004 [65]. The most common form of β -O-4′
structures in lignins is the benzyl alcohol (R’=OH). Non-cyclic benzyl
aryl ether (R′=Ar) are much less common. β -O-4′ may also display
lignin carbohydrate bonds (R′=carbohydrate)
8
Table 1 Frequency of some lignin substructures in Spruce and Birch.
Sources: Adler 1977 [66], Zhang et al 2006 [62] and Zhang and Gellerstedt
2007 [67]. One C9 unit corresponds to a phenylpropanoid unit in the
polymer.
Intermonomeric bondtype
Frequency per 100 C 9 unit
in Spruce
Frequency per 100 C 9
unit in Birch
β-O-4′
35-45
60
β-5′
9-12
6
β-β′
3-4
3
β-1′/spirodienone*
2 / 1-2
1 / 2-3
5-5′/dibenzodioxocin*
9-11 / 4
~5/ ~0
4-O-5′
~4
~7
*See Figure 5 f or f urther details about these structures
Lignin Biosynthesis
Monolignol Biosynthesis
The biosynthesis of lignin monolignols originates in the phenylpropanoid
metabolic pathway, which is now relatively well established [8,68]. As
previously mentioned, this pathway leads to a whole group of important plant
phenolics but only a short description of the monolignol pathway will be
presented here. The biosynthesis of the building units of lignin is thus a part
of the secondary metabolism of plants, i.e., a dispensable metabolism for
survival. Nonetheless, the plant spends important amounts of biochemical
resources on lignification, making it a very energetically expensive process
[69]. Also, substantial amounts of carbon are invested, as lignin is richer in
carbon than polysaccharides [70]. All this is dedicated to a non-reversible
process.
The general pathway starts with the amino acid phenylalanine. The
specific pathway leading to the monolignols involves at least eight different
enzymes and cofactors, biochemical energy (ATP) and reductive power
(NADPH). Here, the pathway is summarized in three steps (Figure 6) - not
necessarily in this order:
1. Removal of the amino group from phenylalanine and creation of a
double bond on the side-chain.
2. A series of hydroxylations and methoxylations of specific aromatic
carbons.
3. Reductions of the side-chain carbonyl group, from (a) carboxylic
acid to aldehyde and (b) from aldehyde to alcohol.
It is worth noticing that lignin-like dehydrogenation polymers cannot only
be formed with the alcohol monomers (the monolignols) but also with the
different precursors like the acid or aldehyde [71,72]. However, plants
apparently do “prefer” to build their lignin with the alcohol monomer
although they could save energy by using any of the phenolic precursors
instead.
9
Figure 6 Summarized phenylpropanoid path way leading to monolignols. In terms of
energy, step 2 requires one NADPH for each hydroxylation and step 3 requires one
ATP and two NADPH to reduce each monomer to alcohol. Thus f rom phenylalanine to
conif eryl alcohol one ATP and f our NADPH are required, which is equivalent to a
to tal of 13 ATP molecules [73]. For a more detailed description, see appended article
IV.
“Unexpected” Plasticity of Lignin
There is a considerable amount of observations describing alternative
phenolics apparently being naturally incorporated to the lignin polymer
(Figure 7a). Kenaf lignins contain over 50% γ-acetylated monomers [74] while
grass lignins have approximately 10% p-coumaric acid [75]. The presence of
γ-p-hydroxybenzoate substituted monomers in various plant lignins has been
observed repeatedly [76-79]. Ferulates have been shown to be incorporated
into lignin and to cross-link lignin to cell wall polysaccharides in grass [80].
In addition, other puzzling features in lignins are the reduced lignin
structures in end-groups and dimeric structures (Figure 7b), which have been
observed in softwood lignins [81-83].
Figure 7 a. Examples of phenolic monomers incorporated into lignins either naturally
or in mutants/transgenics. b. Reduced lignin structures of undetermined origin.
Also, the study of plant transgenics and mutants has revealed even more
chemical diversity of lignins. The genetic changes in all these specimens
affect the monolignol biosynthetic pathway. Up- or down-regulations of
specific enzymes, due to the genetic changes, lead to over- or under-
10
production of certain phenolics of this pathway. The plant’s lignin is thereby
affected: the relative amounts of the different monolignols are changed
[84,85] and in other cases, other phenolic species than the main monolignols
are found in the lignin [86-88], resulting in remarkably different lignins
deposited in the cell walls of these plants. The viability or growth is
considerably affected in certain cases but interestingly, a lignin is
nevertheless synthesized independently of the availability of the main three
monolignols. Thus lignin possesses a rather unique ability among
biopolymers of monomeric “plasticity” or flexibility, i.e., a capacity of
accommodating monomers of moderately varied chemical structures.
Oxidative Polymerization
Karl Freudenberg and co-workers are internationally recognized for their
extensive work on lignin polymerization using model polymers made from the
three common monolignols (Figure 4) [89]. The in vitro polymers formed from
the oxidation of these monolignols by an oxidizing enzyme were called
“dehydrogenation polymers” (DHP). The composition, the reaction products
and the spectral properties of DHPs from coniferyl alcohol displayed a great
resemblance with spruce lignin to the extent that they were originally
described as identical [90]. Later on, however, DHPs were repeatedly shown to
have noteworthy differences compared to lignin [91-94]. Although DHPs
produce largely the same substructures as lignin, these are not found in the
same proportions. DHPs are also difficult to produce in large molecular
weights. Moreover, DHPs produced in different conditions differ as well,
particularly by varying the rate of addition of the monolignols. Interestingly,
Freudenberg himself noticed this difference and coined two general types of
DHPs syntheses: Zulaufverfahren for one-time addition versus Zutropfverfahren
for gradual addition. As a result, the validity of DHPs as lignin models has
been discussed and questioned [95] but all the same they have been used and
studied in connection with lignin biosynthesis because of two main reasons:
DHPs produce all the major substructures of lignin and they can be produced
without the typical impurities of isolated lignins, such as carbohydrates. As
the differences were increasingly recognized numerous attempts have been
made to make up for them by synthesizing DHPs in a great variety of
conditions. These conditions represent different chemical and physical factors
in the cell wall during lignin polymerization, which could potentially affect
the structure of lignin. Apart from studying pH effects [96-98], the monolignol
rate of addition [99,100], different oxidation agents [101-103] and other
environmental conditions have also been tested. DHP syntheses in the
presence of carbohydrates [23,104,105], trying to mimic the cell wall
environment also proved to affect the structure of the polymer in different
ways. Although no DHP has shown identical properties with isolated lignin, it
has become evident that a DHP’s structural outcome is sensitive to its
polymerization conditions [106] and thus its properties can be indirectly
controlled. Which cell wall conditions actually influence lignin polymerization
and to what degree they do so, is poorly understood.
From a theoretical point of view, a model for lignin polymerization had
already been proposed by Erdtman in the 1930’s [107] in the light of a
dehydrogenation experiment performed as early as 1908 by two French
chemists [108]. This model described lignin as the product of the
dehydrogenation of propenyl phenol derivatives. Freudenberg and co-workers
11
provided solid evidence for this model with their extensive experimental work
on dehydrogenation models. The great value of Freudenberg’s work lies in the
structural elucidation of more than forty products of DHP formation, from
monomers to hexamers [89]. It became clear that the dehydrogenative
polymerization process follows certain rules and the knowledge of these rules
provided a basis for explaining the majority of the structures found in lignin.
Today, this theoretical model is essentially the same and it can be
summarized as follows:
Lignin polymerization is the result of radical coupling of resonance-stabilized
monolignol radicals and phenolic radicals on the lignin polymer, followed by
nucleophilic additions to the quinone methide intermediates (Figure 8).
Figure 8 β -O-4′ coupling of monolignol, conif eryl alcohol (CA), to guaiacyl polymer (PG)
end-group. The f inal step constitutes a nucleophilic addition to the α -carbon of the
quinone methide intermediate.
This model was supplemented by Sarkanen [95] with the idea of Bulk
polymerization vs. End-wise polymerization. A bulk polymer is the result of
predominant dimerization and subsequent coupling of dimer or oligomer
radicals to each other, whereas an end-wise polymer is the product of
monomer radicals’ coupling to a growing polymer chain. A bulk polymer will
have more of a branched character whereas the end-wise polymer is more
linear. According to Sarkanen [95], this then accounts for the differences
observed in DHPs structures depending on the monolignol rate of addition as
in Zulaufverfahren and Zutropfverfahren. The formation of certain bonds is
particularly favoured depending on the type of polymerization. As a result, an
end-wise polymer displays large amounts of β-O-4′ bonds while a bulk
polymer contains large amounts of β-5′ and β-β′. The bulk polymer will also
display a greater amount of α―β unsaturated side chains and phenolic endgroups. Since natural lignins contain β-O-4′ structures mostly, they are
considered to be the product of end-wise polymerization [65,66,95].
Challenges
Polymerization Control
The model offered by Erdtman and later supported by Freudenberg and
others implies no strict control over the coupling step which ultimately
defines much of the lignin structure. Thus according to this model, lignin is
not expected to display any particular sequence of monolignols and of the
bond-types created at each coupling event. As a result, lignin polymerization
has subsequently been described as a random or near-random coupling of
12
monolignols [9,109], giving the impression that any monolignol might couple
in any of the possible ways. Pure statistical randomness is not chemically
possible in phenolic radical coupling [65] and Erdtman already supposed
there would be a certain degree of order in lignins [110]. By looking at
monolignol and bond distributions in lignins of different plant-types (Table 1),
one would expect some kind of influence or “control” over the polymerization.
The inability of DHPs to completely reproduce lignins in vitro, especially the
large amounts of β-O-4′, makes clear the lack of understanding of the
regulating mechanisms at work in the plant cell. Several suggestions have
been made to explain how different factors might influence bond and
monomeric distribution, representing possible ways of indirect regulation in
the plant cell wall:
ƒ
The rate of monolignol addition to the reaction site (alternatively the
rate of radical generation) in the cell wall is very slow and very
difficult if at all possible to reproduce in vitro. It has been shown
that different addition rates will result in substantial differences in
bond distributions [66,95,100,111].
ƒ
Precursor/monolignol
availability
will
define
the
monomer
composition of lignin. The most obvious examples are mutants and
transgenics which apparently incorporate available phenolic
compounds as lignin monomers when lacking the common
monolignols. The different monolignol availability will also influence
the bonding pattern since depending on the monolignol structure
some bonds are generally not formed (e.g., no 5-carbon bonds in
syringyl lignin).
ƒ
The reaction environment, the heterogeneous polysaccharide matrix
of the cell wall plays an important role in lignin polymerization and
will ultimately influence bond formation [23,105,112,113]. As
mentioned previously local pH and hydrophobicity vary within the
cell wall.
ƒ
Purely chemical factors determine the outcome of the different
coupling reactions in lignin polymerization: the oxidation potentials
of the different phenolic species and their radical reactivities
[110,114]. If lignin is mostly the product of the addition of
monolignols to a growing lignin polymer, then cross-coupling (i.e., a
monolignol radical coupling to a phenolic end-group radical) must
be a main coupling event in lignin biopolymerization [65]. The
regioselectivity of such reaction is not well understood but some
experimental evidence suggests that the chemical effect, depending
on the chemistry of the different phenolic species, over bondformation can be quite strong. An interesting work by Syrjänen and
Brunow showed how β-O-4′ bonds are strongly favoured when crosscoupling coniferyl alcohol to a phenolic end-group model [111].
An alternative to the purely chemically “controlled” model has been
proposed by several researchers [115-117]. This model suggests that the
polymerization step in lignin biosynthesis is actually under strict control, as
is the general case for other biopolymers. Consequently this control dictates
both the sequence of the monomers and the bond-types formed. Such in vivo
control system could then explain why in vitro syntheses of lignin models
such as DHPs, have not been able to reproduce lignin correctly, particularly
the amount of β-O-4′ structures. According to this model a primary
lignooligomer is formed into arrays of dirigent sites in the cell wall, which
completely control the radical-radical coupling of monolignol radicals. The
13
nature of the dirigent sites has been proposed to be cell wall proteins, such
as hydroxyproline-rich gluco-proteins, which are present in the plant cell
walls. Thereafter, the first lignin chain, product of the arrays of dirigent sites,
works itself as a polymeric template which is replicated into a new lignin
chain, copy of the first. The second chain will then serve as a template for a
third and so on. Thus lignins are suggested to possess similar biochemical
properties to enzymes’ catalytic capabilities. In addition, the replication
mechanism of this template model produces a mirror image stereoisomerically
speaking. This mirror effect is suggested to explain the observed optical
inactivity of lignins. The proposal of this new model implying strict control, is
controversial since it stands in contrast to the purely chemically controlled
model and has generated much debate [38,65,115,117-122]. Although this
new model is in need of more experimental evidence to gain wider acceptance
[11,119], it has found its way into a text-book and is presented as the model
for lignin polymerization awaiting “[…]full clarification at the biochemical
level.” [123]
Another puzzle in lignification is the polymerization initiation sites. It is
believed that in the xylem, lignification starts at the cell corners and
proceeds towards the centre of the cell as previously described. Remarkably,
in other plant tissues, there are reports of just the opposite happening, i.e.,
lignin deposition starting in the innermost cellulosic layer [124-126]. How
does the common xylem cell prevent polymerization from happening in the
inner part of cell wall as monolignols are transported into the cell wall’s outer
layers? Is
lignification specifically “directed” to
start
at
specific
sites/molecular structures in the cell wall matrix? Several suggestions in the
literature attempt to answer these questions. Enzymes/proteins as well as
cell wall polysaccharides are possible candidates as initiation sites [17].
Cross-linking in the cell wall by ferulates has led some to suggest them as
starting points for lignification [127,128]. Ferulate dimers have been shown
to cross-link polysaccharides by ester bonds and so a polysaccharide ester
ferulate group in its phenolic state could be a starting point for
polymerization. Such groups have been shown to couple monolignols through
dehydrogenative coupling in vitro and in vivo [129,130]. The strict control
model of lignin polymerization mentioned above proposes hydroxyproline-rich
proteins as initiation sites. Interactions between lignin and these cell wall
proteins and others (proline- and glycine-rich proteins) are believed to exist
but neither the nature of the linkages nor the proteins’ functions have been
established [11].
Dehydrogenation
The oxidation step, creating a phenoxy radical on either a phenolic endgroup of the polymer or on a monomer, has been a key subject of many
studies. Generally, it is believed that hydrogen peroxide (H 2 O 2 ) is the main
oxidizing agent along with cell wall peroxidases (class III) as catalysts
[131,132]. It is difficult to assign specific functions to different isozymes
because of peroxidases’ wide substrate specificity but there are indications of
monolignol-specific peroxidases [133-135]. Moreover, there is significant
evidence for the involvement of other oxidases, such as laccase in
lignification [136-138].
For the polymer to grow, the oxidation of phenolic end-groups on the
polymer must be at least as effective as the oxidation of monomers. To
14
imagine cell wall enzymes being involved in the oxidation of diffusing
monomers in the cell wall matrix might not be too difficult, but to picture
them oxidizing the phenolic residues on the polymer as effectively as the
monomers raises several questions. The lignifying cell wall is increasingly
hydrophobic, turning it into a hostile environment for most enzymes. The
oxidation of phenolic residues on the lignin polymer, which is expected to be
more or less immobile as it is attached to a solid phase, will be dependent on
the diffusion rate of the enzyme in an already crowded cell wall matrix.
Additionally, enzymatic oxidation of a phenolic residue on the polymer will
need to overcome a higher oxidation potential, compared to a monolignol
[114], and steric hindrance by the polymer itself. Recently, however, a cell
wall peroxidase was discovered in poplar callus, which effectively oxidizes
polymeric lignin and plays a significant role in secondary cell wall
lignification [139,140].
Other possibilities for the oxidation step have been envisaged, possibly
solving these challenges. Phenolic radicals might act as oxidative carriers or
shuttles that, once oxidized by an enzyme, can diffuse more freely than
enzymes onto the growing polymer and oxidize phenolic residues
[98,122,141].
Low molecular weight oxidants such as superoxide radical
●¯
anion, O 2 , [142] and manganese, Mn 3 + [103], have been proposed to carry
out the same function.
Lignin among Other Polycyclic Biopolymers
There are other phenolic biopolymers that, although still poorly
characterized, exhibit noteworthy similarities with lignins, i.e., suberin,
polymeric tannins and melanin. In the literature, melanin has been compared
to lignin but only superficially [143-145]. A short description of melanin
function and biosynthesis is presented here to account for a number of
similarities which might be of interest in elucidating these complex
biopolymers.
Melanin(s) is a group of polycyclic biopolymers that are produced by a
wide variety of organisms: bacteria, fungi, animals, protozoans and plants.
They can be described as “generally black biological macromolecules
composed of various types of phenolic or indolic monomers…” [146].
Interestingly, as with lignin, most structure determinations have been done
on harshly isolated samples and so the structure of intact melanin is little
understood
[146].
In
comparison,
lignin
research
and
structural
characterization has come a long way. There are essentially three types of
melanin [147]: eumelanin (nitrogen containing melanin derived from tyrosine
and phenylalanine), phaeomelanin (like eumelanin but generally also
containing sulphur), allomelanin (nitrogen-free melanin derived from
naphthalene derivatives and catechol among others). A rough generalization
would classify eumelanin and phaeomelanin as animal melanin and
allomelanin as plant, fungi and bacterial melanin [144].
General properties usually attributed to melanin are: absorbance of a wide
spectrum of light (UV and visible light); to participate in one- and twoelectron redox reactions and to display cation chelating properties [148].
More specifically there are several interesting benefits which melanins convey
15
to the organisms that synthesize them. Under anaerobic conditions bacteria
may use them as electron acceptors or carriers, making possible energy
producing processes, similar to oxidative phosphorylation [147]. Melanin
protects some fungi against UV-light [149]. They provide a virulence factor for
certain pathogenic bacteria and fungi by acting as radical scavengers and
thus defending them against their host’s own defence mechanisms [150,151].
Also in fungi, production of melanin is associated with cell wall modification,
thus providing structural strength and protection it against extreme
environments [146]. In insect cuticle formation, melanin is believed to provide
mechanical strength [152] and act as a structural component in wound
healing in arthropods and plants [153,154]. Apart from this, melanin in
animals are usually pigments that provide camouflage and mimicry properties
or ornamentation for sexual display [143]. In humans, melanin has been
reported to offer protection to skin cells from damage caused by UV-light
[155].
Melanin biosynthesis bears some interesting similarities with lignin
biosynthesis. As with lignin, melanin synthesis is part of the secondary
metabolism of the organism and thus non-essential for survival. Also, the
final polymer is a product of oxidative polymerization of phenolic (and indolic)
compounds or precursors [146,148]. In Figure 9, some of the known melanin
precursors are presented to be compared with lignin monolignols and
alternative phenolic monomers in Figures 4 and 7. Animal melanin precursors
are mainly 3,4-dihrydroxy-phenylalanine (DOPA) and cysteinyl DOPA while
fungi melanin precursors are diverse but mostly 1,8-dihydroxynaphtalene
(DHN). Other known precursors are homogentisic acid in a few bacteria [156]
and Glutaminyl-4-dihydroxybenzene (GDHB) in certain fungi [157].
In fungi, the polymerization process is believed to be achieved by a
phenoloxidase. Tyrosinase, peroxidase and laccase are considered possible
candidates [146,158]. Very little is known about the coupling step and about
what the DHN polymerization products are or what types of bonds are formed.
In the case of animal melanin it is known that DOPA polymerizes to melanin
through oxidation by tyrosinase. The oxidation leads to formation of quinones
which react with diphenolic precursors to form oligomers. In vitro
experiments have shown that in the presence of tyrosinase and a pH of at
least 6.8 DOPA forms melanin and at pH 8 or above DOPA polymerizes without
the oxidase [159]. An intriguing fact is that melanosomes, the cell organelles
where animal melanin is synthesized, have an acidic pH. A low pH is an
unfavourable condition for polymerization, which requires deprotonation. It
has thus been suggested that this is a way for the cell to control melanin
polymerization [160]. An important difference between animal and fungi
melanin is that in animals melanin is synthesized and kept inside the cell (in
melanosomes) while fungi melanin occurs in the cell walls or extracellularly
[146,161].
In insect cuticle, diphenolic precursors such as DOPA have been isolated
and generally believed to build melanin contributing to the sclerotization
process, which involves the darkening and hardening of cuticles
[148,152,162]. However, only the diphenolic monomers N-acetyl-dopamine
(NADA) and N-beta-alanyldopamine (NBAD) (Figure 8) are believed to play an
important role, at least in the hardening process [163]. Although it is not
known to which extent any of these present diphenolic precursors actually
16
polymerize in the cuticle, there is evidence for an uncharacterized
polyphenolic polymeric fraction [164,165]. As for NADA and NBAD there is
evidence supporting their participation in cross-linking and polymerizing
proteins of the cuticle [163]. Through oxidation by phenoloxidases these
precursors form quinones and quinone methides which react with
nucleophilic groups in the cuticle from e.g., proteins [145,163]. Interestingly,
dimers of these precursors (Figure 9) have been found in insect cuticles [166].
Figure 9 Melanin precursors f rom different organisms and proposed
coupling mechanisms. a. Animal melanin precursor (DOPA: 3,4-dihrydroxyphenylalanine) oxidation and polymerization. b. Fungi melanin precursors
(GDBH:Glutaminyl-4-dihydroxybenzene and DHN:1,8-dihydroxynaphtalene)
c. Bacterial melanin precursor (homogentisic acid) d. Sclerotization agen t
in insects (NADA: N-acetyl-dopamine )
Maybe the most relevant and consistent difference between lignin and
melanin precursors is the diphenolic nature of melanin precursors. The
polymerization process is also chemically different: the oxidation of melanin
precursors is believed to lead to quinones, not stabilized radicals as in lignin,
which then polymerize. Despite these differences, from a biosynthetic point of
view, lignin and melanin monomers display the same chemical potential.
17
Upon activation by an oxidative agent they are able to spontaneously
polymerize without any catalytic assistance, as opposed to other biopolymers.
The monomers of nucleic acids, proteins, polysaccharides among others, do
not polymerize by themselves after activation; instead they all depend on a
carefully orchestrated enzymatic process for the coupling step [167]. Although
it has not been determined whether melanins are polymerized in nature under
a strict control system, the possibility exists that they are not. At least, in the
case of lignin biopolymerization there is substantial evidence supporting the
idea that lignin is deposited in the plant cell wall without such control. This
does not mean, of course, that there is no careful control over the
polymerization step, but as an alternative, the control mechanisms might be
indirect.
In summary, the similarities between lignin and melanin include:
ƒ
A role in hardening of extra-cellular layers or cell wall
ƒ
Cross-linking extracellular components
ƒ
A role in defence against pathogenic attacks or wound healing
ƒ
Being
the
product
of
polymerization
of
phenolic
compounds
through oxidative coupling
ƒ
Being recalcitrant due to the stability of the inter-unit bonds
ƒ
The coupling/polymerization step is probably purely chemically
controlled.
It seems that in nature, common functions such as mechanical strength,
cross-linking polymers and pathogenic protection are, in a wide variety of
organisms, achieved by synthesizing polycyclic biopolymers such as lignins
and melanins through oxidative polymerization of phenolic/diphenolic
monomers.
18
Methods
Detailed and individual descriptions of materials and methods can be
found in each paper appended to this thesis. A general description of the
methods used in this work is presented here.
Coniferyl Alcohol Synthesis
Coniferyl alcohol (Figure 4) was repeatedly synthesized for the majority of
DHP experiments and was consistently used throughout all the studies. It
was obtained from the reduction of commercially available coniferyl aldehyde
with sodium borohydride, as described elsewhere [168].
Dehydrogenation Polymers – DHP
The in vitro synthesis of dehydrogenation polymers in different conditions
and the analyses of them using common organic chemistry analytic methods
is the principal basis of all the experimental work presented in this thesis.
Karl Freudenberg and co-workers, who originally introduced model
dehydrogenation experiments and with them elucidated the formation and
structure of many lignin substructures, already stated two general ways of
synthesizing the polymers that would affect their final properties:
Zulaufverfahren and Zutropfverfahren: a one-time addition versus a drop-wise
(or gradual) addition of monomers, respectively. Although, no standardized
method exists for them, these methods (or principles for synthesis) have been
used by many groups and in each case adapted or modified depending on the
desired effect: the one-time addition giving more bulk, branched polymers
and the drop-wise giving more linear polymers. Since lignin is recognized as a
generally linear polymer and one objective of this work was to synthesize
polymers as similar to lignin as possible, the Zutropfverfahren or drop-wise
principle was chosen in all DHP experiments. It is certain that DHPs, as
lignin models, do not fully represent lignins and their polymerization.
Nevertheless, the fact that they can form most of the bonds that exist in
natural lignins and that the contamination problems of lignin isolation are
avoided means they are useful experimental modelling systems to elucidate
the formation and structure of lignins.
The general procedure for DHP synthesis can be described as follows.
Phenolic monomers (normally coniferyl alcohol) were dissolved in an organic
solvent, miscible with water, and diluted in a larger volume of aqueous
buffered solution, generally 3-50 mmol/l H 2 KPO 4 with regulated pH, generally
near-neutral or slightly acidic in some specific cases. A reaction mixture was
prepared with an oxidant, generally hydrogen peroxide and a catalyst,
horseradish peroxidase (HRP), which were dissolved separately from the
monomers in a buffered solution of the same type. The monomers were
dropped into the reaction slowly, using a peristaltic pump at flow-rates
ranging between 6.5 and 2.5 ml/h depending on the experiment. After 24
19
alternatively 48h of reaction at room temperature and protected from light,
the product was precipitated (if needed) by evaporation of the organic solvent
initially used. The precipitate was washed with pure water by centrifugation.
This step was taken to get rid of enzymes, salts and unreacted hydrogen
peroxide. The product was then suspended in water and freeze-dried. The
dried product, generally a powder was weighed, and in certain cases, washed
in an appropriate organic solvent, e.g., ethyl acetate, to get rid of lowmolecular weight products, mostly dimers but also unreacted monomers.
Derivatization
Both the low-molecular weight and the polymeric
derivatized for the different chromatographic methods used.
fractions
were
Acetylation
To make DHP polymers more soluble in THF, which was the solvent
in the SEC system (described below), acetylation was required. One to
two mg of polymeric DHP were dissolved in 1-2 ml pyridine:acetic
anhydride (1:1) and left to react overnight. The reaction was stopped
with an excess of methanol, alternatively water:methanol (1:1) and
pyridine co-evaporated with toluene to dryness.
Silylation
For analysis with GC-FID and GC-MS, sylilation of low molecular
weight compounds such as dimers and monomers was carried out under
nitrogen for at least 1h. The product was dissolved in methylene
chloride, alternatively acetone, ideally at 1mg/ml concentration and N,
O-Bis(trimethylsilyl)acetamide (BSTFA) was added together with
pyridine (10%) as catalyst.
Gas Chromatography - Flame Ionization Detector, GC-FID
GC-FID was used to check purity of different synthetic models and to
quantify dimer amounts using tetracosane as internal standard and
calibrated with authentic models when available. The GC-FID instrument was
a Hewlett-Packard 6890 with a RTX 5 column (30m, 0.25 µm I.D., 0.25 µm
film thickness).
Gas Chromatography - Mass Spectroscopy, GC-MS
GC-MS was used to confirm chemical structures of synthetic models and
identify monomers and/or dimers in product mixes such as low molecular
weight fractions from DHPs. The instrument was from Thermo Finnigan Trace
GC-MS, 2000 series. The column used was RTX 5MS (30 m, 0.32 mm, 0.25
µm).
Size Exclusion Chromatography – SEC
Samples of acetylated high molecular weight DHP products were analyzed
by SEC to study their hydrodynamic volume and obtain an idea of the relative
20
molecular weight compared to elution times of monodisperse polystyrene
standards. The analyses were performed on a system of three Cross-linked PS
columns in series (Steerage, Waters HR4+HR2+HR0.5) using THF as effluent
at 0.8 mL min - 1 . Detection was made with UV-light detector at 280 nm.
Nuclear Magnetic Resonance - NMR
Samples of underivatized, usually only high molecular weight DHP
fractions were dissolved in deuterated solvents for NMR analyses. Both 1-D
(qualitative and quantitative 1 3 C) and 2-D (HSQC and HSQC-TOCSY) NMR
experiments were run to detect and quantify specific DHP substructures. All
NMR experiments were run on a Bruker Avance 400 MHz instrument.
21
Results and Discussion
Reduction during Oxidative Coupling (Paper I)
In native lignin there are some types of structures that cannot be
explained solely by oxidative coupling. Although the Erdtman/Freudenberg
model explains the large majority of lignin structures, a number of structures
that cannot be explained by this chemistry alone have long been noted in
lignins, including the reduced lignin structures [169,170]. The most common
structure is dihydroconiferyl alcohol (Figure 10), found as an end-group and
suggested to be incorporated as such by dehydrogenative coupling since the
plant cell may synthesize it and subsequently use it as a monomer [87].
However, there is a more puzzling structure, reduced β-β′s (Figure 10), found
in softwood lignins [38,65,170]. Since this is a dimeric reduced structure it is
even more intriguing to consider its incorporation as such into the polymer,
thus maybe originating from a specific lignan: secoisolaresinol. Another
possibility is that this structure is formed subsequent to polymerization, e.g.,
by the reduction of a resinol structure, in the same way the lignan is formed
from pinoresinol [171]. A third possibility is that it is formed during the
dehydrogenative polymerization process, e.g., by the reduction of a quinone
methide intermediate. Any of these possibilities convey immediate problems.
Lignans are optically active, meaning that if the lignan secoisolaresinol has
been incorporated then only one stereoisomer would be found in lignin, which
is not the case [38]. A post-lignification reduction seems even less likely
because a lignified cell wall is very compact and rules out the only probable
candidates able to achieve such a reduction: enzymes [172]. Although an
enzyme that generates the lignan secoisolariciresinol could be trapped in the
cell wall during lignification the product would still be optically active.
Therefore, the third possibility was considered the most likely. We reasoned
there must exist a reductive agent able to achieve a reduction in the overall
oxidative environment of lignin polymerization. This was the objective of our
investigation: is there a naturally occurring reductive agent able to carry out
the reduction of a quinone methide intermediate during dehydrogenative
polymerization of monolignols?
Quinone Methide Model Reduction
Four low molecular mass biological reducing agents were screened for
their capability to reduce a model compound (Figure 11a) for the β-ether
quinone methide intermediate in lignin biopolymerization. The reducing
agents were nicotinamide adenine dinucleotide (NADH), reduced glutathione,
both being common reducing agents produced in virtually all living cells
[167], ascorbic acid known as a redox buffer agent in plant cell walls [173]
and the citric acid intermediate α-ketoglutaric acid. The extracellular enzyme
cellobiose dehydrogenase (CDH) was included in the screen as an example of
a reducing enzyme [174]. Sodium borohydride was used as a positive control
reducing agent. The β-ether quinone methide model was only successfully
reduced by NADH or NaBH 4 . Gas chromatograms of the sodium borohydridetreated sample and the NADH-treated sample displayed an equivalent peak
with identical MS spectra, corresponding to the silylated reduced model
22
compound (Figure 11b). Three NMR experiments ( 1 H, 1 3 C-DEPT-135 and 2D
HSQC), run on the sodium borohydride-treated sample, confirmed the identity
of the product.
Figure
10 Reduced lignin structures
present in sof twood lignin [170].
Reduced Structures in DHPs
NADH was tested in synthetic dehydrogenation polymers (DHPs). The
interest was to determine how well oxidation and reduction could proceed in
the same reaction medium. The 2D NMR technique HSQC was employed to
reveal the DHP structures formed. Figure 12 shows the presence of β-aryl
ether A, phenylcoumaran B, resinol C, and α,β-diaryl ether A 2 structures
which are common in conventional DHPs [49,66]. Furthermore, reduced βether structures A R (Figure 12) of the same type as compound b (Figure 11)
were observed from DHPs made in the presence of NADH. Disappointingly
secoisolariciresinol structures C R (Figure 12) were not observed, nor were
dihydroconiferyl alcohol X 5 moieties found.
Figure 11 Reduction of a quinone methide intermediate f rom a β -O-4′ model compound.
First step achieved according to Brunow, G. et al. [175]. See appended article I for
details.
From these results it can be argued that the origin of reduced structures
like secoisolariciresinol C R is an uncatalyzed, protein-independent reaction of
a biologically significant reductant such as NADH with the quinone methide
intermediate, formed during lignin polymerization. It is not known whether
this is the actual compound responsible for such a reduction and the
inability of CDH to reduce the quinone methide here does not imply that there
are no other enzymes able to catalyze the reaction. The substantial amount of
reduced β-aryl ether units A R detected (~48% of the β-ethers) indicates that
23
NADH was able to reduce the β-aryl ether quinone methide in an oxidative
polymerization system. NADH was able to reduce a β-O-4′ quinone methide
structure but not the β–β′ quinone methide; at least not in detectable
amounts. The conditions and/or reductants needed for such a reduction
remain uncertain. It would seem the reduction of a β-aryl ether (such as β-O4′) quinone methide competes with a nucleophilic attack by an external
nucleophile (water or a phenol), whereas the β–β′- or β–5′-quinone methides
have intramolecular hydroxyl groups out-competing the attack on the αcarbon. Since we observed resinol (β-β′) and phenylcoumaran (β-5′) structures
in the DHPs, NADH was apparently not effective in competing against the fast
intramolecular post-coupling reactions under these conditions.
Until these results were published, reduced structures had only been
observed in end-groups X 5 and in β-β′ C R units in natural lignins
[87,169,170,176]. Recently, however, reduced β-O-4′ structures (A R ) have
been observed in lignin [177], analyzed by NMR spectroscopy with a special
cryo-probe that increases the analysis sensibility. While it is exciting to
confirm that these structures do occur naturally and are not only a
laboratory product, it is also important to note that they apparently occur in
lesser amounts than the reduced β-β′s. The question that immediately arises
is: what process favours the formation of the structure which is seemingly
more difficult to produce?
In this work we did not detect any reduced β-β′ structures C R , but
succeeded in producing reduced β-O-4′ structures A R . Although it cannot be
confirmed, this could be due to the low sensitivity of the NMR analyses but
more probably to the difference in the structure and the polymerization
environment between lignin and DHPs. It is a known fact that DHPs are
sensitive to polymerization conditions [106]. If the latter explanation is the
case, then there must be some kind of mechanism directing the reducing
effect towards β-β′-units in lignin polymerization. There are of course other
options: enzymatic catalysis, low molecular weight catalysts or unusual
conditions during the reduction. Or, are β-β′ quinomethides more long-lived
in the natural conditions of the biopolymerization of lignin?
Whatever the reduction agent consumed in such a reaction, it appears to
be a waste of energy if reduced structures are just the products of sidereactions. Spending one molecule of NADH corresponds to 3 of ATP [73]. What
could be the benefits of reducing the lignin? Such structures could be
expected to be more hydrophobic than their “normal” counterparts. Also, they
offer fewer possibilities for hydrogen bonding and for the formation of
covalent bonds to polysaccharides. This could potentially mean less LCC
formation through ester and ether bonds to the α-carbon in lignin monomers,
and this might in turn have an effect on the properties of the cell wall.
Furthermore, a higher rotational freedom and thus flexibility should be
expected of a reduced β-β′ compared to the cyclic resinol structure in the
normal β-β′. Although reduced lignin structures might involve a biochemical
cost they could be a way for the plant to control the properties of its lignin.
24
Figure 12 2-D NMR spectra (HSQC, correlating a carbon nuclei to its
bonded hydrogen’s nuclei) of the non-aromatic region of DHPs synthesized
in the presence of NADH. Signals f or X5 and CR are only the expected
signals of these structures according to Zhang et al. [170]
25
Effects of an Anti-oxidant during Lignin Polymerization (Paper II)
In our experiments with reduced lignin structures (Paper I), ascorbic acid
could not reduce the quinone methide model and was discarded as a reducing
agent able to generate the observed lignin reduced structures. Nevertheless,
the presence of ascorbic acid in the cell wall has been reported [178,179] and
it is suggested to play a role in scavenging hydrogen peroxide in the cell wall
as it does in vacuoles [180,181]. Ascorbic acid is proposed to reduce
hydrogen peroxide indirectly by reducing phenolic radicals which in turn have
been generated by cell wall peroxidases and hydrogen peroxide. If there are
reducing agents present during lignin polymerization, what other effects
might they have upon the structure of lignin? The phenolic monomer radical
concentration is an important factor affecting the structural outcome of the
lignin polymer [95,100,111]. As a result, the presence of an anti-oxidant,
such as ascorbic acid, during lignin polymerization should influence it. In the
present work, we investigated the effect of the anti-oxidant ascorbic acid
during the synthesis of coniferyl alcohol dehydrogenation polymers (DHPs).
DHPs in the Presence of Ascorbic Acid
Dehydrogenation polymers were synthesized in the presence and in the
absence of ascorbic acid at pH 5 by enzymatic oxidation of coniferyl alcohol
using horseradish peroxidase and H 2 O 2 . The amounts of three common lignin
inter-unit bond types (β-5′; β-β′ and β-O-4′) were quantified by 1 3 C-NMR of the
polymeric fraction and compared to a control synthesis, i.e., an analogous
DHP synthesis without ascorbic acid. A portion of the spectra and signal
assignments, according to Landucci [92], are presented in Figure 13 and Table
2. The quantification was done as follows. The amount of β-5′ structures was
calculated by taking the mean of the integration values of the two isolated
signals for the α- and γ-carbons of this substructure (Figure 3, signals 1 and 4
respectively). Then, this value was subtracted from the integrated value of the
overlapping signals of the β-carbons of β-5′ and β-β′ (signal 8) in order to
determine the amount of β-β′ structures. The value for β-β′ structures was
subtracted from the signal 2 - i.e., the overlapping signals from β-carbons
from the benzyl alcohol β-O-4′ and non-cyclic benzyl aryl ether β-O-4′ (α-O-4′)
(see structures in Figure 5) and α-carbons from β-β′ - to obtain the amount of
all β-O-4′ structures. The amount of the benzyl alcohol form of β-O-4′ alone
was obtained from the integrated value of the overlapping α-carbon and γcarbon signals from β-O-4′ and β-β′ respectively (signal 3), also by subtracting
the β-β′ value obtained earlier. The amount of total β-O-4′ structures was
generally no more than 3% higher than the benzyl alcohol β-O-4′ form alone
in each case. Thus the contribution to the total amount of β-O-4′ by the noncyclic benzyl aryl ether form was low, which is consistent with the
observations that low pH favours the formation of the benzyl alcohol form of
β-O-4′ [182]. Signal 6 (Figure 4) was also assessed to determine the total
amount β-O-4′ structures but some contamination is suspected to have
occurred since 20 to 50% higher quantification values were obtained from
this signal compared to the β-O-4′ values obtained from signal 2 and 3.
26
Figure 13
1 3 C-NMR
partial spectra of the high molecular
weight f raction of DHPs produced in the presence of 50%
ascorbic acid (relative amount) – 1,3,5-trimethoxy benzene.
Signals assignments in Table 2.
1 3 C NMR spectra in Figure 13. β -O-4′ represents the
benzyl alcohol f orm and β -O-4′( α -O-4′) is the non-cyclic benzyl aryl ether f orm (see
structures in Figure 5).
Table 2 Signal assignments f rom
Signal
δ
1
87
2
83-85
β-carbons from β-O-4′, β-O-4′(α-O-4′) and α-carbons from β-β′
3
71-72
α-carbon from β-O-4′ and γ-carbons from β-β′
4
63
γ-carbons from β-5′
5
62
γ-carbon from coniferyl alcohol end-group
6
60
γ-carbons from β-O-4′ and β-O-4′(α-O-4′)
7
56
Methoxyl carbons from DHPs
8
53-54
β-carbons from β-5′ and β-β′
IS-a
93
Unsubstituted aromatic carbon from internal standard
IS-b
55
Methoxyl carbon from internal standard
Assignment
α-carbons from β-5′
Ascorbic acid clearly increased the formation of β-O-4′ structures and
decreased the formation of β-β′, at a 25% amount, relative to the coniferyl
alcohol, compared to the control experiment (Figure 14). Additional DHPs were
produced with larger amounts of ascorbic acid (50 and 75%) to study any
potential concentration effects. The effect of ascorbic acid was enhanced with
its concentration up to 50% for β-O-4′ and slightly enhanced for β-β′
structures (Figure 14). However β-5′ structures were practically not affected.
Since ascorbic acid was added into the reaction mix at the same rate as the
other reactants it was expected to continuously slow the reaction down by
reducing the radical concentration. Consequently, the larger the amount of
ascorbic acid, the slower the polymerization would proceed, enhancing an
end-wise polymerization [95]. The observed increase in β-O-4′ structures is in
agreement with other reports where the reaction was slowed down by limiting
the monomer diffusion rate alone [100,111]. Limiting the monomer
concentration limits the monomer radical concentration which in turn limits
27
dimerization. Dimerization is a known problem in DHP syntheses because of
generally high monomer concentrations [65]. As previously described, the
regioselectivity in dimerization reactions is different compared to crosscoupling and for end-wise polymerization to occur, monomer-oligomer crosscouplings are necessary. The negative effect that ascorbic acid had on the
formation of β-β′ structures might be due to a favoured end-wise
polymerization assuming β-β′ structures are mainly a product of dimerization
[183]. Furthermore, the anti-oxidant had apparently no effect on the
formation of β-5′. This could be due to the fact that these structures can be
both formed through dimerization and end-wise coupling. Interestingly, at
75% relative ascorbic acid concentration, the effect was reversed for β-O-4′
and β-β′ structures and the formation of β-5′ structures was affected
negatively. In addition, the total DHP amount was monitored as well (Figure
14) since ascorbic acid was expected to consume H 2 O 2 , the reaction yield
would consequently be affected. Accordingly, the yield started to decrease at
50 % ascorbic acid and was seriously affected by 75%. The effects observed at
75% ascorbic acid might suggest that the polymerization was continuously
quenched until ascorbic acid was depleted. Then, the polymerization would be
allowed to take place, producing DHPs with a different substructure
composition because of the accumulation of coniferyl alcohol.
Figure 14 Substructure quantif ication by 1 3 C NMR of DHPs
synthesized at pH 5 in the presence of different amounts of
ascorbic acid. The control experiment is represented as 0%
ascorbic acid. Total amounts of β -O-4′ are presented here.
Although it is likely that ascorbic acid primarily slows down the
polymerization rate by reducing phenolic radicals it is not possible to rule out
any effects of the anti-oxidant with the enzyme, either as a substrate or as an
inhibitor, generating a similar structural outcome. However, horseradish
peroxidase apparently prefers phenolics as substrates rather than ascorbic
acid [184,185]. Neither is it possible to rule out any direct effect of ascorbic
acid upon the regioselectivity of the coupling events themselves. However, if
such a “catalytic” effect were the case, one would not expect an inverted
effect after a certain concentration, as observed here.
28
The molecular weight determinations by size exclusion chromatography
showed that the DHPs displayed similar sizes independently of the presence
of ascorbic acid and up to a 50% (Figure 15). The results imply that similarsized polymers were produced with differences in inter-unit substructure
amounts. At 75% ascorbic acid, the sizes of the DHPs appeared to have been
somewhat affected (Figure 15). Based on the NMR inter-unit substructure
quantification results, it is likely that the polymerization occurred under
different conditions, as previously described, thus affecting the sizes of the
polymers.
All DHPs synthesized displayed a common maximum size (Figure 15), as if
the polymerization conditions did not “allow” them to grow larger. Generating
large molecular weight DHPs by conventional ways is known to be a problem
[112]. The conditions here used for the polymerization involved a mixture of
dioxane:water (2:3) which allowed the reaction to stay in solution all along,
compared to purely aqueous systems where DHPs tend to precipitate very
early in the reaction. This system was chosen to make all polymeric species
available for oxidation and coupling as much as possible. However, the
solubility of this complex mixture of polymers is probably not complete after
all. It is likely that in our system, solubility is still an issue which defines the
maximum size of the polymers. Improving the solubility of DHPs is believed to
affect their molecular weight positively [104].
Figure 15
Size
exclusion
chromatograph
of
DHPs
synthesized at pH 5 in different ascorbic acid concentrations
(0, 25, 50 and 75% ascorbic acid relative to conif eryl
alcohol).
The most common inter-unit substructure in lignins is the β-O-4′. These
results provide further support to the idea that a low monomer radical
concentration - in this case provided by ascorbic acid - enhances the
formation of β-O-4′ bonds by favouring end-wise over bulk polymers. Since
ascorbic acid has been reported to be present in the plant cell wall, it seems
plausible that this component or similar ones may play a role in the
biopolymerization of lignin, as an additional component besides monolignols,
peroxidases and hydrogen peroxide. The presence and concentration of a
phenolic anti-oxidant may also be a way for the plant to control and regulate
the lignin structure, possibly explaining the differences in lignin structure
29
between, e.g., cell wall layers, in addition to pH, polysaccharide composition
and monolignol addition rate.
Table 3 Molecular weight values (in g mol - 1 relative to polystyrene
standards) of DHPs synthesized at pH5 in different ascorbic acid (Asc A)
concentrations.
Mw
Mn
P
MP
Control (0%)
2469
1751
1.4
1626
Asc A (25%)
2498
1753
1.4
1667
Asc A (50%)
2641
1813
1.5
1704
Asc A (75%)
2520
1781
1.4
1913
Mw = Average molecular weight. Mn = Weighed average molecular
weight. MP = Peak molecular weight P = Polydispersity.
Testing the Lignin Template Polymerization Model (Paper III)
In this study an experiment was specifically designed in order to test the
template hypothesis of lignin polymerization [117]. This model proposes strict
control through replication of a lignin template. For the sake of simplicity we
considered synthesizing dehydrogenation polymers in the presence of a lignin
model, an isolated coniferyl alcohol lignan called pinoresinol, which would
represent the common lignin substructure (β-β′), Based on this hypothesis
this lignin was expected to act as a template and replicate itself during in
vitro polymerization of coniferyl alcohol. The phenolic groups of the lignin
model were protected through methylation to prevent it from coupling during
polymerization (Figure 16).
Figure 16 a. Pinoresinol and b. Pinoresinol dimethyl ether.
DHPs in the Presence of a β-β ′ Model Structure
Dehydrogenative polymers (DHP) were prepared from coniferyl alcohol in
the presence of pinoresinol methyl ether and a control polymerization was
performed in the absence of this dimer. During the reaction no precipitation
of the formed DHPs was observed due to the dioxane/water environment
used. The final material was fractionated into two parts for separate analyses:
a low molecular weight part, containing dimers produced from polymerization
and the methylated pinoresinol model; and a polymeric part. The low
30
molecular weight fraction was analyzed
The polymeric material was studied
substructure amounts were determined
as described in Paper II except that only
by GC-FID for dimer quantification.
by quantitative 1 3 C NMR and the
by peak integration in the same way
peaks 1, 3, 4 and 8 were quantified.
NMR quantification results indicated no increase in the relative amount of
β-β′ structures in the template experiment compared to our control (Table 4).
The amounts of two other common substructures (β-O-4′ and β-5′) were
determined as well in order to see if a template effect would quantitatively
influence their formation, either positively or negatively. It was concluded
that there was no specific structural influence of β-β′ model dimer as a
template for replication on the polymerization of coniferyl alcohol. Rather the
same pattern of bond distribution in the template experiment was reproduced
in the control. Of the three inter-unit structures studied here, β-5′ was the
predominant one followed by β-O-4′ and β-β′ last. This is in line with two
known observations: in vitro dimerization of coniferyl alcohol produces these
three structures with β-5′ as the major product [186], and dimerization is an
over-represented reaction in DHPs [65,95]. Interestingly, the polymerization
system used here provided relatively large amounts of β-O-4′ structures
(benzyl alcohol form), compared to purely aqueous systems with enzymic
oxidants at near-neutral pH [106,187]. The formation of β-O-4′ bonds is
otherwise known to be favoured at lower pH [187,188]. It is not possible to
determine if dimerization itself has been affected by the solvent polarity,
possibly favouring the formation of β-O-4′ [189,190] or if the solubility effects
of this water:dioxane system influences the final bond distribution.
Dimerization and cross-coupling reactions – as in coupling between a
monomer and a phenolic dimer/oligomer – do not display the same
regioselectivity. β-O-4′ becomes the predominant bond-type in the crosscoupling of a phenolic end-group model and coniferyl alcohol [111]. The
longer phenolic oligomeric species stay in solution, competing with monomers
in coupling other monomers, the more they will affect the final bond
distributions.
Table 4
1 3 C NMR quantif ication of
three specif ic
substructures in DHPs produced in the presence and
the absence of pinoresinol dimethyl ether.
Template DHP
Control DHP
(μmol g )
(μmol g )
β-β'
460
480
β-5'
960
940
β-O-4'
810
820
Substructure
-1
-1
Two dimers were quantified from GC-FID analyses: the added model dimer,
pinoresinol dimethyl ether, and β-β′ dimer products of polymerization (Table
5). The dimer quantification indicated no increase in the relative amount of ββ′ dimers and thus no replication effect. Other dimers, such as β-O-4′ and β5′, could not be properly quantified because of the lack of pure substances
needed for calibration, but relative amounts are presented. The identity of the
three dimers was determined by GC-MS of both silylated and acetylated
samples (see appended article III for details).
31
Table 5 GC-FID quantif ication of specif ic dimers f rom the low molecular weight
fractions. Proper quantif ication of β -5′ and β -O-4′ dimers could not be obtained; the
area ratios (dimer:internal standard per mg sample) are presented here.
Dimer
Template DHP
dimers
-1
(μmol g )
Control dimers
-1
(μmol g )
Template DHP
dimers
(Area ratios)
Control dimers
(Area ratios)
β-β′
4.7 ±0.5
β-5′
-
-
0.76 ±0.03
1.00 ±0.1
β-O-4′
-
-
0.15 ±0.003
0.19 ±0.01
94 ±11
17 ±0.8
12 ±1.,4
Pinoresinol
dimethyl
ether
137
±6
4.4
±0.2
0.07 ±0.01
0.07
±0.003
One might argue that the lignin model used, a dimer, is too short to fulfil
a template function as described by Chen and Sarkanen [117]. Hypothetically,
other bond-types and higher molecular weight lignin might have a stronger
catalytic or template replication effect. However, since monolignol radicals
can indeed couple and polymerize in the absence of any template, and the
mechanism suggested by these scientists implies a strict control over the
primary structure, i.e., bond-type and monolignol type, the template effect of
the lignin on monolignol polymerization ought to be strong. Therefore, some
replication effect, if not complete, should be expected from our template
experiment and we believe that if the template mechanism proposed is valid,
then any kind of lignin structure ought to have a template effect on the
polymerization of monolignols, even at the dimer level.
The proposal of the proteinaceous dirigent array/template polymerization
model, implying complete synthetic control over lignin polymerization, stems
from scientists who question the “traditional” theory, which only confers
certain chemical “control”. Although the traditional model does not
comprehensively explain all known lignin features, the experimental evidence
supporting this newer model is very limited. It consists mainly of two studies:
the effect on the molecular weight of synthetic lignins (DHP), when
macromolecular
kraft
lignin
components
are
present
during
DHP
polymerization – the kraft lignin being suggested to serve as template [191];
and the determination of the monomer sequence of a lignin hexamer fragment
from Eucalyptus globulus, claimed to be a repeating fragment and used as
evidence that lignin has a well defined primary structure [192]. A recent
analysis of the latter [119] indicates however, that small amounts of this
structure can be expected also by uncatalyzed radical polymerization, i.e., the
Erdtman/Freudenberg-model. Apart from this, there are also studies
localizing so-called dirigent arrays of proteins in the cell wall [115]. However,
no actual “catalytic” involvement of such proteinaceous elements with lignin
polymerization has been shown. From a more philosophical point of view,
there is a potential problem in one of the premises, repeatedly used as an
argument by the proponents of this new model [116,120,123,193-195]. The
authors assume that lignin cannot be an exception compared to how
biopolymers are usually synthesized in nature as expressed by the following
statement: “[…]lignin formation is under full biochemical control, in harmony
with the formation of all other known biopolymers.” [195]. However, no
reasons are given to explain why lignin could not be an exception. It is argued
that lignin polymerization could not be left to chance [116]; but no other
alternatives than complete control, are envisaged. No convincing reasons are
given to establish why an indirect or partial control cannot be enough.
Furthermore, few lignin chemists, if any, would argue today that lignin
polymerization is a completely random and uncontrolled process. On the
32
contrary, all the available reports on the different pH, hydrophobicity,
monolignol radical concentration effects on the polymerization of DHPs are
attempts at determining which factors influence and might “control” lignin
polymerization in situ.
Part of the critique aimed at the traditional model of lignin polymerization
is actually directed against the limitations of lignin analysis [121]. The
critique is justified to some extent because most knowledge of lignin
structure is based only on partial samples. It is assumed that the small
samples are representative of the rest of the polymer. If lignin analysis has
never been representative then no structure characterization should be used
as conclusive evidence for (or against) the conventional theory of lignin
polymerization. But then again, this criticism implies that most of the
knowledge on lignin structure, developed through hundreds of studies, is
seriously mistaken.
A probably relevant criticism lies in the choice of oxidase when modelling
with dehydrogenation polymers [193]. Originally, oxidases from mushrooms,
which do not synthesize lignin in vivo, were used by Freudenberg [196] and
later also peroxidases and laccases [90]. In the last decades, horseradish
peroxidase has probably been the most used enzyme in dehydrogenation
polymer experiments. Although peroxidases have a wide substrate specificity
and they certainly produce dehydrogenation polymers from monolignols, they
do not necessarily exhibit the same activity with different substrates
[98,140]. The structure of the lignin polymer should be expected to vary
depending on the activity of the peroxidase used [197], especially when
considering that lignin polymerization depends on an effective oxidation of
monomers as well as polymer end-groups. One of the reasons for the general
failure of DHPs in reproducing lignin structure more accurately might very
well be the choice of oxidase.
On the Role of the Monolignol γ-carbon (Paper IV)
The biosynthesis of the monolignols consumes important amounts of
energy. The normal monolignols used for lignin biosynthesis carry alcohol
groups at the γ-carbon. In order to produce this alcohol the plant needs to
reduce it from carboxylic acid to aldehyde and from aldehyde to alcohol
(Figure 6). The γ-carbon is not actually involved in the coupling of other
monolignols through radical polymerization so it is logical to ask why the
plant needs to reduce this group. In addition, ferulic acid and
coniferaldehyde
(Figure 17)
can
form
lignin-like
polymers
through
dehydrogenation [71,72,198]. Is there a reason for the plant to invest energy
in producing the alcohol for lignin? What effects would a lignin based on the
γ-aldehyde/carboxylic acid have on the cell wall? The aim of this study was to
chemically and physically characterize DHPs synthesized from three different
monomers that are produced by plants: coniferyl alcohol, coniferyl aldehyde
and ferulic acid. In addition, the biological significance of the γ-carbon
reduction is also discussed.
33
Figure 17 Monomers used in different dehydrogenation synthesis of
lignin-like polymers.
Characterization of DHPs from Different Phenolic Monomers
Dehydrogenation polymers (DHP) from coniferyl alcohol, coniferyl aldehyde
and ferulic acid (Figure 17) were synthesized by oxidation by horseradish
peroxidase and H 2 O 2 . The weight yields of the DHPs were 35, 47 and 58%
respectively, after extraction of low molecular weight compounds. In the
conditions used here, coniferyl alcohol had a propensity to precipitate at the
beginning of the reaction which could explain the relatively low DHP yield.
Interestingly, the acid-based DHPs did not precipitate during the reaction in
contrast to the other two DHPs.
Size exclusion chromatography analysis (SEC) (Figure 18) revealed that the
three DHPs contained dispersed polymeric material. The weight average
molecular weight (Mw) and the number average molecular weight (Mn) (Table
5) of the alcohol and the acid DHPs were similar, but the molecular weight at
the peak (MP) of the latter was almost twice the MP of the other DHPs
indicating that ferulic acid polymerization might have produced some
important populations of larger polymers compared to the alcohol and
aldehyde. However, caution should be exercised when comparing sizes of
polymers containing different functional groups, as determined by SEC.
Larger polymers might be expected of the ferulic acid polymerization because
it was the only reaction that did not precipitate and solubility is important for
the availability of the different oligomers for further oxidation and coupling.
Figure 18 Size Exclusion Chromatogram of acetylated DHP
materials.
34
Table 5 Molecular weight values (in g mol - 1 relative to polystyrene
standards) from SEC analyses of the three acetylated DHP materials.
Mw
Mn
P
MP
Acid DHP
4489
3395
1,3
5017
Aldehyde DHP
2709
1898
1,4
2222
Alcohol DHP
4464
3206
1,4
2725
Mw: Weight average molecular weight; Mn: Number average molecular
weight; P: polydispersity; MP: Peak molecular weight.
All three DHPs were analyzed by 1 3 C NMR and the spectra can be viewed in
Figure 19. Signal assignments were suggested (see appended article IV) based
on reported values for synthesized model components but also NMR studies of
isolated lignins [92,199-201]. The different substructures found are shown in
Figure 20 and the following observations were made from the comparison of
the three spectra:
ƒ
The different DHPs could generally form the same type of bonding
structures: β-O-4′, β-5′, β-β′ and 5-5′.
ƒ
Obvious structural differences were observed in the δ 60-90 ppm
region of the three spectra. The alcohol DHP displayed a number of
signals in this region corresponding to saturated α, β and γ
carbons while the acid and aldehyde DHPs practically lacked any
such signals. Saturated inter-unit substructures are formed when
coniferyl alcohol radical coupling occurs through at least one βcarbon, i.e., β-O-4′, β-5′, β-β′ and β-1′. Although some of these
bond-types occurred in the DHPs from ferulic acid and coniferyl
aldehyde the structures formed were predominantly unsaturated
and their signals appear in the 100-140 ppm region.
ƒ
There was a general lack of cyclic structures (e.g., β-5′ and β-β′) in
both the acid and aldehyde DHPs compared to the alcohol DHP.
The acid DHP did produce some signals, although weak, for cyclic
β-5′ and β-β′ structures which contain α and β single bond
structures.
ƒ
The acid and aldehyde DHPs lacked benzyl alcohol groups, i.e., αhydroxyl groups, and benzyl aryl-ether structures that are typical
of alcohol DHPs [66].
35
Figure 19 1 3 C NMR Spectra of a. Ferulic acid DHP b. Conif eraldehyde DHP and
c. Conif eryl alcohol DHP
Figure 20 Inter-unit substructures in the three diff erent DHP materials
36
AFM images of very smooth spin-coated DHP model surfaces are presented
in Figure 21. The contact angle of water was determined on the DHP model
surfaces as a relative measurement of the hydrophobicity of the different
polymeric materials. The results (Table 6) indicate that the coniferyl alcohol
DHP was less hydrophobic than the coniferaldehyde DHP which was less
hydrophobic than the ferulic acid DHP. All three DHPs indicated much higher
hydrophobicity than has been previously determined for kraft lignin [202],
but this was expected since the latter is a lignin derivative where both free
phenolic groups and carboxyl groups are present. There are reasons to believe
that the hydrophobicity of the aldehyde DHP was somewhat underestimated,
since this material was incompletely dissolved in pyridine (the solvent used
for making the surfaces). Therefore, the fraction of the DHP that was
dissolved by the pyridine and used for making the surfaces, most likely
represented a somewhat lower molecular weight DHP that was less
hydrophobic.
The DHPs were also characterized with a solvent panel (Table 7). Generally,
the aldehyde DHP displayed the lowest solubility; it was in some cases
troublesome to dissolve completely. The carboxylic acid and alcohol DHPs
showed rather similar patterns, although the ferulic acid DHP was more
soluble in the more polar solvents than the alcohol (Table 7).
Figure 21 AFM tapping mode height images (1 × 1 μm) and rms roughness
determinations of the diff erent spin-coated DHP model surf aces that were used in the
measurements of the contact angle of water. a) DHP from conif eryl alcohol, rms
roughness 0.349 nm. b) DHP f rom conif eryl aldehyde, rms roughness 0.411 nm. c) DHP
from f erulic acid, rms roughness 0.404 nm.
Table 6 Initial equilibrium contact angles of water on
DHP model surf aces.
Sample
Contact angle (º)
DHP from ferulic acid
63
DHP from coniferaldehyde
60
DHP from coniferyl alcohol
58
Kraft lignin (spruce)
46*
*f rom Norgren, et al. 2006 [202]
37
Table 7 Ocular inspection of the solubility of the different
unmodif ied DHP materials in organic and aqueous solvents
of increasing dielectric constants (except Dioxane:Water
and Acetone:Water, which are mixes of solvents at 1:1
ratio).
Solvent
Acid DHP
Aldehyde DHP
Alcohol DHP
Dioxane
no
no
partly
Ethyl acetate
no
no
no
THF
yes
no
no
CH2Cl2
no
no
no
Pyridine
yes
partly
yes
Acetone
partly
no
no
DMF
yes
partly
yes
DMSO
yes
yes
yes
Water
no
no
no
Dioxane: Water
no
no
yes
Acetone:Water
yes
No
yes
In summary, our investigation has demonstrated that also monolignols
with unreduced γ-carbons can form polymers with generally the same types of
inter-monolignol bonds as in natural lignin. Thus, there is no reason to
believe that reduction of γ-carbon is necessary to obtain a polymer. Based on
the contact angle measurements and the solubility panel, we reasoned that
coniferaldehyde generated the most hydrophobic polymeric material and the
coniferyl alcohol the less hydrophobic. Therefore, the reduction of the γcarbon could be a way to make the lignin slightly more hydrophilic. This is
surprising for two reasons:
ƒ
One of the biological roles of lignin is to make the cell wall
hydrophobic and waterproof, which is needed in the development of
water-conducting tissues in plants. However, a lignin structure that
is too hydrophobic may interact poorly with cellulose and
hemicelluloses and thereby not fulfill its biological functions
efficiently.
ƒ
The carboxylic acid in ferulic acid is a charged structure at
physiological pH as opposed to the primary alcohol of coniferyl
alcohol. It was therefore expected that the DHP made of ferulic acid
would be considerably more hydrophilic than the DHP made from
coniferyl alcohol, and not the opposite as in our contact angle
results.
The characterization by 1 3 C NMR demonstrated that in addition to the
expected differences in the γ-carbon functional group, there was one main
difference between the coniferyl alcohol DHP on one side and ferulic acid
/coniferyl aldehyde DHPs on the other. The latter DHPs displayed almost only
α―β unsaturated inter-unit substructures whereas the coniferyl alcohol DHP
displayed both unsaturated and saturated α―β bonds. In alcohol DHPs
unsaturated α―β structures commonly exist in end-groups whereas the
unsaturated structures are formed through β-coupling. In lignins, α―β
unsaturated structures are much less common than in DHPs. In addition,
benzyl alcohols, a frequent product of β-coupling in coniferyl alcohol DHPs
38
and especially in lignins (β-O-4′ see Figure 6), may contribute considerably to
the overall hydrophilicity of the polymer, offering an explanation for its lower
hydrophobicity in our results. However, the relative hydrophobicity of the
DHP made of ferulic acid was nonetheless surprising. Part of the explanation
might lie in a loss of charge through loss of carboxylic acid groups.
Apparently there is a tendency for decarboxylation of certain substructures
during the oxidative polymerization of ferulic acid [72,199,203]. If and how
much decarboxylation has occurred in our polymers is difficult to assert from
our NMR studies, since signals from such structures were difficult to
distinguish from the non-decarboxylated ones.
The structural differences between these DHPs are due to different
rearomatization pathways of the quinone methide intermediates which are
formed after oxidative coupling. A quinone methide intermediate is formed
when a coupling occurs through at least one β-carbon. In the case of coniferyl
alcohol, rearomatization always occurs through an intra- or intermolecular
nucleophilic attack on the α-carbon. Due to the carbonyl group at the γposition on both ferulic acid and coniferaldehyde, upon coupling through the
β-carbon, the β-proton is acidic enough to be released for the rearomatization
of the quinone methide intermediate. For ferulic acid, decarboxylation is an
additional way of rearomatizing a quinone methide. These additional
rearomatization pathways in the cases of the ferulic acid and coniferyl
aldehyde dehydrogenative polymerization apparently limit any intermolecular
nucleophilic attacks by water or a phenol compared to coniferyl alcohol
polymerization. Because of the carbonyl groups at the Cγ, the recreation of a
Cα―Cβ double bond during coupling is predominant for both ferulic acid and
coniferaldehyde polymerization. In Figure 22, the hypothetical mechanisms are
summarized for the creation of the most common intermonomeric bond in
lignin, the β-O-4′ bond for coniferyl alcohol, conifer aldehyde and ferulic acid
DHPs. Rearomatizing the quinone methide intermediate by an intramolecular
nucleophilic attack leads to a cyclic structure. Although the possibility of
forming cyclic structures, i.e., β-5′and β-β′ for ferulic acid and in the case of
β-5′ for coniferaldehyde has been shown elsewhere [72,200], only weak
signals from possible cyclic β-5′ and β-β′ structures were observed for the
ferulic acid DHPs here. Other reports of coniferyl aldehyde dehydrogenative
polymerization support the lack of cyclic β-5′ structure [71,204]. However, the
cyclic β-5′ might be formed first but as it is not thermodynamically stable, it
is subsequently transformed into the non-cyclic form [200]. In the case of the
acid, a suggestion has been made that cyclic and non-cyclic dimers of β-5′
and β-β′ structures could be both present and in some sort of equilibrium
[72,205].
How do these structural differences explain the large investment by plants
in reducing the γ-carbon? Apart from an increased hydrophobicity, a
coniferaldehyde or ferulic acid lignin might be expected to form less covalent
bonds with hemicelluloses and other wood cell wall polysaccharides. Such
covalent bonds, forming lignin carbohydrate complexes (LCC) [26,30,206],
may be created by nucleophilic attacks on the α-carbon of the quinone
methide intermediate of alcohols or carboxylic acids on the polysaccharide
[207,208], i.e., these hydroxyl groups replace water or a phenol in the
mechanism of Figure 22a. As the suggested mechanisms describe, and the
results of this study show, the possibilities for extramolecular nucleophilic
attacks appear to be limited and consequently the formation of this type of
LCC might be limited as well. It has recently been demonstrated that LCC
form entire networks in wood, where lignin crosslinks polysaccharides of
different kinds. These networks have been suggested to play an important
role for the mechanical properties of the wood [28].
39
Figure 22 Mechanism suggestions f or the rearomatization of the β -O-4′ bond during
conif eryl alcohol, conif eraldehyde and f erulic acid oxidative coupling. Quinone methide
rearomatization in the case of (c) ferulic acid can occur as described f or (b) coniferyl
aldehyde too.
Thus, reducing the γ-carbon of the monolignols might convey at least one
very tangible benefit, which is improving the formation of ligninpolysaccharide covalent networks. Interestingly, a mutant loblolly pine was
discovered with a deficient expression of the enzyme cinnamyl alcohol
dehydrogenase [87] which affects the phenylpropanoid pathway (see appended
article IV for details). The viable plant exhibited decreased lignin content, but
also abnormally large amounts of aldehyde groups from coniferyl aldehyde
end-groups, among others. So, coniferyl aldehyde has the ability to copolymerize with normal lignin monomers to some degree. However, this does
not indicate that the reduction is unnecessary for obtaining a functional
lignin, since the lignin of this mutant still contains mostly alcohol monomers
which can more easily form α-carbon LCCs. Lignin has a unique ability
among biopolymers for “metabolic plasticity”, i.e., that non-conventional
monomers can be incorporated into the polymer [87,209]. However, this
should not be understood as if any potential monomer can form a highly
40
functional lignin. The standard monomers are very well designed by nature
for lignins to fulfill their biological functions, as discussed here for the
reduction of the γ-carbon.
41
Conclusions
The present work adds one more factor - reducing agents - to the growing
list of variables described in the literature as having influence over the
structure of lignin: carbohydrate composition of the cell wall matrix, pH,
oxidase activity and monolignol rate of addition. Biological reducing agents
were shown to be able to affect the structure of DHPs in two ways: in one
case, by producing reduced lignin structures during oxidative coupling and in
another case by altering the frequency of inter-unit bonds, especially by
favouring the formation of the most common lignin substructure, the β-O-4′.
In addition, one part of this work has focused on a different aspect which also
affects the structure of the polymer: the chemical structure of the monolignol
itself. The expensive reduction of the lignin monomers’ γ-carbon all the way to
an alcohol has considerable implications for the properties of the final
polymer in that it allows the formation of saturated inter-unit substructures
and possibly improves the formation of lignin carbohydrate complexes,
important for the cell wall properties as a whole. Some attention has been
given to the relatively newly suggested model of lignin polymerization
implying a strict control of the lignin primary structure. Although this
hypothesis is difficult to test, it is nevertheless interesting to consider
whether lignin has any catalytic capabilities (or template replication
capabilities). Our results here indicate no support for a replication effect as
such. Nevertheless, any effect of the lignin macromolecule itself (and other
cell
wall
macromolecular
components
for
that
matter)
upon
the
polymerization cannot be excluded, but remains to be clarified.
The data in this thesis suggests that it is very unlikely that plants use a
catalytic system, such as the one suggested by the proteinaceous dirigent
array/lignin template replication hypothesis, to control the structure of lignin
in great detail. Instead, it shows that plants can influence lignin
polymerization and structure by ensuring the presence of reducing agents
(probably of low molecular weight) during lignification. Furthermore, even
though lignins are reportedly flexible in the choice of monomers, investing in
the monomer γ-carbon reduction is indeed a way for the plant to generate a
lignin that fulfils its biological functions fully in the plant cell wall.
In several ways, our understanding of lignin polymerization chemistry is
only tentative. The control mechanisms at work defining lignin structure,
especially during polymerization, are today still unclear. Although there is a
suggestion for a strict catalytical system controlling it in the cell wall, there
is no convincing evidence supporting it. It is therefore all the more exciting to
elucidate an alternative system in nature of apparently rather subtle control:
one that is not enzymatically based. For that reason, establishing how and
what factors in the plant cell wall, chemical and physical, influence the final
structure of the lignin polymer is necessary; this might be useful, e.g., when
attempting to alter the structure of lignin in plants for the purpose of
improving pulping results.
Future research with DHPs as models of lignin polymerization would
benefit from some highly desired improvements. For example, the choice of
peroxidase should be considered more carefully in light of the findings that
have revealed differing activities of enzymes oxidizing monomers compared to
polymer end-groups. The kinetics of radical couplings between the different
phenolic species (monomer-monomer and monomer-oligomer) should be
42
determined to permit the better understanding of how different radical
concentrations favour the formation of either bulk or end-wise polymers.
Finally, the structure of DHPs is dependent on the solubility of the different
oligomeric species, but this is probably not the case in the cell wall where
growing lignin oligomers are more likely to be fixed to a “solid phase”, and
thus also less likely to react with each other. These oligomer-oligomer
reactions cannot be avoided in conventional “solution” DHPs. Thus, the
development of an effective system where oligomers are fixed to a solid phase
might improve end-wise polymerization and make the model polymers even
more similar to natural lignins.
43
Acknowledgements
I would like to express a few words of gratitude to the many persons who have
supported my work. Without them it would not have been possible.
First and foremost my two supervisors:
Professor Gunnar Henriksson, for always being available for discussions and
always encouraging, as well as being a constant idea-generator in this
project; and
Dr. Liming Zhang, for all your experimental guidance and ideas, and
especially for all the practical assistance with the NMR.
And also:
Professor Gösta Brunow, for our many rewarding discussions and your vast
knowledge of the relevant literature as well as for your cooperation in Paper I.
Your practical and theoretical knowledge in the field has been invaluable.
Professor John Ralph, for our many valuable discussions and your genuine
interest in our research project as well as for your cooperation in Paper I.
Dr. Magnus Norgren, for your valuable cooperation in Paper II.
Inga Persson, for taking care of all the formalities related to the project and
especially for going the extra mile in many things and making the whole
group feel more at home.
I would also like to thank the Swedish Research Council for Environment,
Agricultural Sciences and Spatial Planning (FORMAS), contract no 229-20041240, for financial support.
Many thanks to some special colleagues, Martin, Maria, Andrea, Lotta,
Viviana, Sverker, Stefan, Jiebing, David, Juanjo and Dimitri, for many
discussions and help, and for making my work more fun; and many thanks to
all colleagues from the Fibre and Pulp technology groups for contributing to a
great working atmosphere.
Last but not least, I want to thank my family which has doubled in size since
I started my Ph.D. studies. You are a daily encouragement and I cannot
imagine life without you. Many thanks also to my parents for all the support
and valuable suggestions while writing the thesis, and …to God for helping
me keep perspective on life.
44
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Glossary
Angiosperm: Vascular plant in which
the ovule is fertilized and develops
into a seed in an enclosed hollow
ovary.
ATP: Adenosine triphosphate. Energy
rich molecule and enzyme co-factor
involved
in
many
important
metabolic processes.
Club mosses: A small creeping vascular
plant which reproduce sexually
through spores.
Coumarins:
Heterocyclic
organic
molecules
with
characteristic
odours, involved in plant defence.
Ferns: The most diversified group of
vascular plants which display true
leaves
and
reproduce
through
spores.
Flavonoids:
Heterocyclic
organic
molecules functioning as pigments,
for example in flower petals.
Gymnosperm:
Vascular
plant
that
reproduces by means of an exposed
seed, or ovule.
Hardwood:
An
angiosperm
woody
plant, generally more dense and
with a more complex structure than
softwood.
Hemicellulose: Complex heteropolymer
of
different
sugar
monomer
composition found in the cell wall
of plants.
Horsetails:
Small
vascular
plant
growing a single hollow jointed
stem, which reproduce through
spores.
Kenaf : Herbaceous annual (or biennial)
plant with a woody base growing
several
meters
high
and
reproducing trough seeds.
Laccase: Copper containing enzymes
capable of carrying out oneelectron oxidation of phenols by
reducing
molecular
oxygen
to
water.
Librif orm f ibre: Angiosperm cell with a
mechanical support function.
Lignans:
Heterocyclic
oligomeric
molecules synthesized from the
monolignols and involved in plant
defence.
Lignosulf onates:
Water
soluble
polyelectrolyte
lignin-based
polymers recovered as a by-product
from
the
sulfite
pulp-making
process.
Lumen: The cell space left when
emptied from all organelles and
cytosolic
compounds.
Lumen
occurs in for example dead xylem
cells.
Melanosome: Organelle of animal cells
(usually called melanocytes) which
synthesize and contain melanin.
Monolignol: Specific name for the three
common
lignin
monomers:
pcoumaryl alcohol, coniferyl alcohol
and sinapyl alcohol.
NAD(P)H:
Nicotinamide
adenine
dinucleotide (phosphate) reduced
form. Electron rich molecules and
enzyme
co-factors
involved
in
important metabolic processes.
Nucleic acids: Biopolymers such as
DNA or RNA, built of nucleotides
usually
carrying
genetic
information.
Number average molecular Weight (Mn):
It is the common average weight of
any individual polymer in a mix of
varied-sized polymers.
Oxidative phosphorylation: The process
in which biochemical energy (ATP)
is formed through an electron
transfer mechanism involving the
reduction of molecular oxygen.
Parenchyma: Living plant cells with
thin cell walls dedicated to storage,
found in both xylem and phloem
tissues.
59
Pectins: Heterogenous polysaccharides
common in the cell walls of plants.
Phloem: Vascular plant tissue
carries organic nutrients.
that
Polydispersity: A way of describing the
distribution of molecular weights in
a polymer sample.
Protozoans:
Single
cell
eukaryotic
organisms generally very mobile.
Some
are
parasitic
like
the
amoebas.
Raman
spectroscopy:
Spectroscopic
technique used to study, e.g.,
vibrational energy in molecular
systems,
complementary
to
IR
spectroscopy.
Schlerenchyma: Plant cells with heavily
thickened
and
lignified
walls
dedicated to mechanical support.
Sclerotization:
The
process
of
hardening and darkening of insect
cuticles.
Sof twood: A Gymnosperm woody plant
with a generally less dense and
with a simpler structure than
hardwood.
Stillbenes:
Polyphenolic
compounds
involved
in
plant
defense,
particularly
in
hardwood
protection.
60
Styrylpyrones: Heterocyclic compounds
involved
in
plant
defence,
structurally close to stillbenes.
Suberin: Complex hydrophobic plant
substance in plants preventing
free water diffusion. It is found in
the roots and bark.
Tracheids: Xylem cells developed for
water conduction and structural
support.
TEM: Transmission electron
microscopy.
Vascular cryptogram: Collective term
for all vascular plants which do not
reproduce through seeds.
Vessel elements: Angiosperm xylem
cells
specifically
developed
for
water transportation
Weight average molecular weight (Mw):
It is an average molecular weight
that has been weighed against the
size
of
the
different
polymer
populations.
Xylem:
Vascular plants tissue that
carries
water
and
dissolved
minerals.