Soil Biology & Biochemistry 34 (2002) 139±162
www.elsevier.com/locate/soilbio
Review
The macromolecular organic composition of plant and microbial residues
as inputs to soil organic matter
Ingrid KoÈgel-Knabner*
È kologie, Wissenschaftszentrum Weihenstephan fuÈr ErnaÈhrung, Landnutzung und Umwelt,
Lehrstuhl fuÈr Bodenkunde, Department fuÈr O
Technische UniversitaÈt MuÈnchen, D-85350 Freising-Weihenstephan, Germany
Received 18 September 2000; received in revised form 27 June 2001; accepted 29 July 2001
Abstract
Plant litter and the microbial biomass are the major parent materials for soil organic matter (SOM) formation. Plant litter is composed of
complex mixtures of organic components, mainly polysaccharides and lignin, but also aliphatic biopolymers and tannins. The composition
and relative abundance of these components vary widely among plant species and tissue type. Whereas some components, such as lignin, are
exclusively found in plant residues, speci®c products are formed by microorganisms, e.g. amino sugars. A wide variety of chemical methods
is available for characterizing the chemical composition of these materials, especially the chemolytic methods, which determine individual
degradation products and solid-state 13C NMR spectroscopy, that gives an overview of the total organic chemical composition of the litter
material. With the development of these techniques, an increasing number of studies are being carried out to investigate the changes during
decay and the formation of humic substances. An overview is given on the amount of litter input, the proportion of various plant parts and
their distribution (below-ground/above-ground), as well as the relative proportion of the different plant tissues. Major emphasis is on the
organic chemical composition of the parent material for SOM formation and thus this paper provides information that will help to identify the
changes occurring during biodegradation of plant litter in soils. q 2002 Elsevier Science Ltd. All rights reserved.
Keywords: Litter; Polysaccharides; Lignin; Lipids; Biopolymers; Nuclear magnetic resonance; Plant residues; Microbial residues; Soil organic matter
1. Introduction
Plant litter materials provide the primary resources for
organic matter formation in soil. The amount of plant litter,
its composition and its properties are essential controlling
factors for the formation of soil organic matter (SOM) and
humi®cation processes in terrestrial ecosystems (Swift et
al., 1979; Scholes et al., 1997). The microbial biomass
also represents a signi®cant compartment of the terrestrial
biomass and microbial residues in soil are an important
parent material for humus formation (Haider, 1992). For
modeling the C transformations in soils, it is essential to
know the composition of input materials. The predictors
for plant litter decomposition dynamics include data on
the contents of cellulose, holocellulose (cellulose and hemicelluloses), lignin and tannins. Within a particular climatic
region, litter chemistry measurements, especially cellulose±
lignin±N relationships, are predictors for litter degradation
rates (Palm and Rowland, 1997; Moorhead et al., 1999;
* Tel.: 149-8161-713677; fax: 149-8161-714466.
E-mail address: [email protected] (I. KoÈgel-Knabner).
Berg, 2000). However, relationships often are not very
clear (Aerts, 1997). This is possibly be related to the inadequate methods used to determine the organic chemical
composition of plant litter. Often the data are obtained on
the basis of chemical degradative methods that can at most
be considered as proximate values. This has become especially evident for those plant components that provide
analytical problems, such as lignin or tannins. It is recognized that the conventional Klason lignin method is not
entirely suitable for lignin determination in non-woody
tissues (Zech et al., 1987; KoÈgel et al., 1988; Preston et
al., 1997). Nonetheless, model calculations and estimates
for litter degradation are still based on these data (Palm
and Rowland, 1997).
The properties and composition of plant residues can be
examined from various aspects. Oades (1988) demonstrated
that a number of factors are decisive for the formation of
humus in soils. In this review, humi®cation is considered as
the prolonged stabilization of organic substances against
biodegradation. The following factors have been found as
particularly important factors for controlling the humi®cation processes in soils (Oades, 1988):
0038-0717/02/$ - see front matter q 2002 Elsevier Science Ltd. All rights reserved.
PII: S 0038- 071 7( 01) 00158-4
140
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
² the amount of litter input,
² the proportion of various plant parts and their distribution
(below- or above-ground),
² the relative proportion of the different plant tissues, and
² their chemical composition.
Two groups of materials will be treated with respect to
their importance for SOM formation; namely on the one
hand, the plant residues, called primary resources and on
the other hand the secondary resources, i.e. microbial residues and exudates. Soil animals, particularly invertebrates,
play an essential role in controlling litter decomposition in
soils (Wolters, 2000). Quantitatively, the relative amount of
animal residues in the C-turnover and therefore also as
parent material for humi®cation in soils is small. Thus
they are not included in this review. Major information on
litter and SOM composition today comes from molecularlevel chemical analyses of speci®c plant or microbial
components (chemolytic techniques, analytical pyrolysis)
in combination with 13C NMR spectroscopy (Baldock et
al., 1997; Preston et al., 1997; KoÈgel-Knabner, 2000).
Thus, this review attempts to summarize the knowledge
on litter input to soils, molecular-level composition and
13
C NMR spectroscopic examples from different plant and
microbial residues and thus to provide a basis for studies on
the changes occurring during residue decay and humus
formation. Special emphasis is put on the organic chemical
composition of the parent material for SOM formation.
2. Amount, proportion and distribution of plant residues
in soil
2.1. Above-ground input
Forest litter consists mainly of foliage or coniferous
needles. Branches, bark and fruits, in comparison, represent
only 21% in cool-temperate climates (Jensen, 1974) and
20±40% in coniferous forests (Millar, 1974) of the total
above-ground litterfall. The contribution of herbaceous
vegetation to total litterfall amounts to less than 5% in
forests of the temperate zones. Meentemeyer et al. (1982)
estimated that the proportions of foliage in total aboveground litterfall in coniferous forests was to be 200±
600 g d.m. m 22y 21. Similar orders of magnitude apply
also for the above-ground litter input in deciduous forest.
Litterfall in coniferous forests (e.g. in spruce stands) is not
bound to a de®ned season. In general, the average amount of
total above-ground litter input in forests increases with
decreasing latitude and increasing productivity from the
boreal coniferous forests (100±400 g d.m. m 22y 21) to the
tropics (600±1200 g d.m. m 22y 21) (Waring and Schlesinger,
1985).
In natural forests, woody debris is not removed and thus
comprises an important component of the total organic
matter input (Harmon et al., 1986; Preston et al., 1998). In
contrast, in highly managed forests most of the woody
debris and the logs are removed and the litter input is shifted
in composition from woody to non-woody materials.
Less information is available on the organic matter input
for arable and grassland ecosystems. Input varies depending
on amount and type of crop residues and fertilizer applications (Table 1). Typical values for farm yard manure input
in different European long-term agroecosystem experiments
range between 100 and 360 g C m 22y 21 (KoÈrschens et al.,
1998). Values are much higher if the crop residues returned
to the soil and the below-ground C are also estimated.
2.2. Below-ground input
A considerable proportion of the organic material
becomes incorporated into the soil as below-ground input,
i.e. as root litter and rhizodeposition. On a global average,
approximately 30, 50, and 75% of the total root biomass
occur in the top 10, 20, and 40 cm of soil (Jackson et al.,
1996). Maximum rooting depth depends on the plant
species, but may be much deeper than is commonly estimated (Richter and Markewitz, 1995; Canadell et al., 1996).
The data summarized by Canadell et al. (1996) give a global
average of maximum rooting depth of 4.6 m. The data also
show that input of root litter into soils can occur to great
depths and places C into deeper soil horizons (Nepstad et al.,
1994; Trumbore et al., 1995). Tundra, boreal forests, and
temperate grasslands have 80±90% of their roots in the top
30 cm, whereas deserts and temperate coniferous forests
have only 50% of their roots in the upper 30 cm (Jackson
et al., 1996).
Root-to-shoot ratios compiled by different authors are
highly variable (Table 2). High values are found for tundra,
grasslands and cold deserts (4±7). Low values (0.1±0.5)
were obtained for forest ecosystems and croplands. Generally, grassland and steppe soils receive a higher proportion
of total carbon input as root litter in comparison to forest
ecosystems under similar climatic conditions. In forest soils,
the contribution of root litter to the input of organic matter
in the forest ¯oor in cool-temperate climates varies between
20 and 50%, depending on the tree species and the life form
(evergreen or deciduous) (Vogt et al., 1986). According to
Vogt et al. (1983) the major proportion of root input into
forest soils is localized in the forest ¯oor and the A-horizon.
On the other hand, Raich and Nadelhoffer (1989) estimated
that the relation of the carbon allocation above-ground/
below-ground, the root-to-shoot-relation, is at approximately 2.5 in forests with a litter production of 200±
500 g m 22y 21. Generally, all these data are highly uncertain
as the accumulation of the root necromass impedes the
determination of the annual root litter input (Vogt et al.,
1998). High amounts of partially decomposed root residues
were found in different forest soils (Beudert et al., 1989;
Preston, 1992). Very limited information is available on
the amount of root necromass, the annual input of root litter
with respect to different biomes or plant types as well as on
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
141
Table 1
Typical values for organic matter input in some long-term agroecosystem experiments (compiled from KoÈrschens et al., 1998; Gerzabek et al., 1997; Bronson
et al., 1998; Buyanovsky and Wagner, 1997; Vanotti et al., 1997; Campbell and Zentner, 1997)
Site
Type of input
Input of C kg m 22
Bad LauchstaÈdt, Germany
Rothamsted, England
Prague, Chechia
Thyrow, Germany
Halle, Germany
Ultuna, Sweden
Farm yard manure
Farm yard manure
Farm yard manure
Farm yard manure
Farm yard manure
Green manure, animal
manure, or peat
0.13±0.16
0.35±0.36
0.10±0.12
0.15±0.16
0.12
0.18±0.19
IRRI, Phillipines
Aquatic photosynthetic
biomass, rice root biomass,
root exudates and ®ne root
turnover
0.40
0.48
N fertilizer
Bhairahawa, Nepal
Rice roots, rhizodeposition
0.30
NPK fertilizer
Sanborn Field, USA
Wheat tops
Wheat roots
Soybean tops
Soybean roots
Corn tops
Corn roots
Wheat, roots and stubble only
Wheat residues 1 manure
Corn residues
Corn residues 1 manure
0.22
0.15
0.22
0.13
0.58
0.34
0.06
0.15 1 0.33
0.09
0.18 1 0.33
No treatment
Manure
No treatment
Manure
Summer grain crop-winter
cover crop
Above-ground
Below-ground
Above-ground
Below-ground
0.58
0.29
0.71
0.22
Conventional tillage
0.21±0.75
0.03±0.57
Wheat-fallow, fertilized
Continuous wheat, fertilized
Horseshoe Bend, USA
Swift Current, Canada
Straw and roots
the turnover of roots (Scholes et al., 1997). Gill and Jackson
(2000) compiled available data on root turnover in different
ecosystems. The slowest average turnover was observed for
entire tree root systems (10% annually), followed by shrubTable 2
Root-to-shoot ratio (primary production) as an indicator for above- and
below-ground contribution of plant litter in different vegetation types
(Data from Oades, 1988; Raich and Nadelhoffer, 1989; Jackson et al., 1996)
Type of vegetation
Root-to-shoot ratio
Desert grassland
Steppe/prairie
Temperate grassland
Montane grassland
Short grass steppe
Tropical grassland
Forests, in average
Temperate forest
Boreal coniferous forest
Tropical deciduous forest
Tropical evergreen forest
Mediterranean forest
Tundra
0.3±6
6
3.7
6
13
0.5±2; 0.7
2
2.5; 0.20
4.0; 0.32
0.34
0.19
0.25
6.6
No fertilizer
No tillage
land total roots. Annual turnover was 53% for grassland ®ne
roots, 55% for wetland ®ne roots and 56% for forest ®ne
roots.
Rhizodeposition, i.e. all organic carbon released by living
roots, accounts for a substantial input of organic matter in
soils. The number of data for soils is limited, mainly
because most of the data obtained from sterile soil experiments or nutrient solutions cannot be applied to soils. Most
of the exudates are rapidly consumed by soil microorganisms. With the use of different labeling techniques, it has
been possible to quantify the amount of organic matter
translocated into the soil below-ground (Kuzyakov and
Domanski, 2000). Table 3 gives an estimate of the belowground C input under wheat and pasture. The higher belowground allocation under pasture is due to the longer
vegetation period of the pasture. From the limited and
non-complete studies available no generalization is possible
for the amount of rhizodeposition under forest (Kuzyakov
and Domanski, 2000). Total input of organic matter in
rice cropping systems in the Philippines and Nepal, consisting of aquatic photosynthetic biomass, rice root biomass,
root exudates and ®ne root turnover ranged between
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
142
Table 3
Rough estimation of total C input in the soil and root-derived CO2 ef¯ux from a soil under wheat with 0.6 kg m 22 grain yield (it is assumed that total aboveground plant mass is two times higher than the grain yield) and in a pasture of about 0.6 kg m 22 dry matter production (from Kuzyakov and Domanski, 2000)
% Of total assimilated
% Of below-ground
Wheat
Wheat
Pasture
kg C m 22a
Pasture
Wheat
Pasture
0.24
0.24
0.16
0.04
0.12
Shoot
Shoot CO2 b
Roots
Soil 1 MO c
Root CO2 d
50
25
13
3
9
30
30
20
5
15
52
12
36
50
13
38
0.48
0.24
0.12
0.03
0.09
Below-ground
25
40
100
100
0.24
0.32
100
100
1.0
0.8
Total assimilated C
a
b
c
d
C content in dry mass of shoots and roots is assumed to be 40%.
Shoot respiration.
C remains in soil and microorganisms.
Root-derived Cs; the sum of root respiration and rhizomicrobial respiration of rhizodeposits.
0.30 and 0.48 g C m 22y 21 (Bronson et al., 1998). Mean
below-ground input of C in a long-term experiment with
cereals, rape crops and fodder beet was between 30 and
50 g C m 22y 21 (Gerzabek et al., 1997).
3. Tissue types of plant residues
Essentially, two different types of plant tissue reach the
soil for decomposition: parenchymatic tissue and woody
tissue. Parenchymatic cells are found in the living green
tissue of leaves and in the cortex (bark) of young twigs
and ®ne roots. They are composed of cellulose walls, the
protoplast, rich in protein, and the vacuola. Woody tissues
form the woody part (xylem) and the supporting tissue
(sclerenchym) of stems, leaf epidermis, leaf ribs and
barks. The different layers of the woody cell wall (middle
lamella, primary wall, secondary wall, tertiary wall) can be
differentiated in their structure as well as in their chemical
composition (Fig. 1). The middle lamella, which acts as the
binding substance between the cells, consists of pectin and
in woody tissues also of lignin. Primary wall, secondary
wall, and tertiary wall consist of cellulose, polyoses (hemicelluloses) and lignin. The middle layer (primary wall and
middle lamella) has the highest lignin concentration (40±
60%). The largest part of the lignin (ca. 75±80%) is derived
from the secondary wall, whose lignin concentration only
amounts to 20±30% (Fengel and Wegener, 1984). A wartlayer, whose chemical composition is unknown, also exists
in the cell wall of coniferous and some angiosperm trees
(SjoÈstroÈm, 1993). The composition of wood and the relative
abundance of the individual components vary widely among
tree and cell types (Hedges, 1990).
4. Plant compound classes
Plant tissues can be divided into various compound
classes, including storage materials that are intracellular,
and structural components that occur in membranes, extracellular or as cell wall constituents. The storage materials
of plants are easily degradable and thus are important
carbon and energy sources for microorganisms. The major
organic compounds of plant litter are polysaccharides and
lignin. According to Millar (1974), spruce needles are
composed of 20% cellulose and lignin, 12% polyoses, 1±
5% protein and 1±6% ash. Leaf litter contains 8±14% ash,
10±19% hemicelluloses, 10±22% cellulose, 5±8% lignin
and 2±15% raw protein (Williams and Gray, 1974). These
data can only be seen as approximate values because, as
Fig. 1. Structure and chemical composition of the woody plant cell wall (from Fengel and Wegener, 1984); W, wart layer; P, primary wall; ML, middle
lamella; S, secondary wall.
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Table 4
Decomposition of different organic substrates in Green®eld sandy loam
(from Martin and Haider, 1986)
Substrate
Glycine
Glucose
Starch
Casein
Cellulose
Benzoic acid
Lima bean (Phaseolus lunatus)
straw
Cysteine
Caffeic acid
Wheat straw
Sudangrass (Sorghum
sudanense)
Walnut wood
Cow manure
Almond shells
Douglas ®r wood (Pseudotsuga
menzicsii)
Ponderosa pine needles (Pinus
ponderosa)
Catechol
Peat moss
Incense cedar wood
Arthrobacter sp. cells
Earthworms (Lumbricus
rubellus)
Penicillium cells
Azotobacter chroococcum cells
Azotobacter polysaccharide
Aspergillus glaucus cells
a
Mineralization after weeks a
1
4
12
20
28
74
73
48
58
27
68
36
83
82
69
72
52
78
57
89
89
81
84
77
82
75
90
90
84
85
79
83
78
91
90
86
85
84
84
79
54
40
26
34
69
60
45
54
75
67
59
64
77
68
63
65
78
68
67
66
7
18
12
2
28
33
24
5
45
43
37
15
52
48
39
29
53
50
41
34
12
23
28
30
32
11
,1
,1
18
3
,1
22
8
1
24
14
2
26
17
3
60
59
79
72
85
80
86
82
87
84
56
61
3
26
72
69
27
41
76
74
61
50
78
76
65
52
79
78
68
54
Percentage added C, evolved as CO2; incubation at 22 ^ 2.2 8C.
indicated above, the conventional methods for analysis of
the plant litter components may often be not speci®c for any
compound class (Swift et al., 1979; Ryan et al., 1990;
Preston et al., 1997). Data from different analyses for arable
crop residues showed a high variability for lignin and cellulose contents (Rahn et al., 1999). Only 50±60% of the total
organic matter of plant litter is accounted for by chemical
degradative techniques (KoÈgel et al., 1988).
5. Intracellular and storage materials
5.1. Proteins
Proteins represent the most abundant group of substances
in plant cells. They consist of polypeptides, long chains of
various amino acids. Proteins serve manifold purposes, e.g.
as enzymes, transport proteins, regulators, storage substances or as structure proteins. They are usually composed
of the 20 most frequent amino acids, which can be subdivided into basic, neutral or acidic amino acids. Further
143
rarely occurring amino acids in plants and microorganisms
have been described. The proteins from plant and microbial
tissues can be decomposed by a multitude of microorganisms and are considered to be less stable plant components
with high turnover rates (Table 4). Nonetheless peptide type
material is found in soils and has been shown to be stabilized in soils over longer periods of time.
The determination of proteins in litter and soils consists
of an acidic hydrolysis (usually with 6 M HCl) followed
either by a chromatographic separation of the individual
amino acids (usually by HPLC, or gas chromatography) or
by the photometric determination of the total concentration
of a-amino groups in the hydrolysate (Stevenson and
Cheng, 1970; KoÈgel-Knabner, 1995).
5.2. Starch, fructans
Starch is an important storage polysaccharide in vascular
plants, but is also present in some algae and bacteria. Starch
consists of two different polymers of glucose, amylose and
amylopectin, where amylose composes, on average, 25% of
the starch (Fig. 2). Amylose consists of long chains of a-dglucose which are connected by (1-4)-glycosidic bonds,
producing a helical tertiary structure. Amylopectin is
composed of similar glucose chains, but is distinguished
from amylose by branching with (1-6)-glycosidic bond
side chains. This branching takes place after approximately
24±30 glucose units. Starch is easily degraded by aerobic as
well as anaerobic microorganisms.
We often ®nd fructan as a further storage polysaccharide
in grasses. This is a water-soluble polymer of fructose with
a-d-glucose as an end group. Besides their storage function,
they are essential for osmoregulation and freezing point
depression in plant cells. The two most important groups
are inulin, composed of b-(2-1)-linked fructose, and levans
or phleins, with b-(2-6)-linkage of the fructose units. The
hydrolytic enzymes necessary to decompose fructans are
widespread among bacteria (De Leeuw and Largeau, 1993).
5.3. Chlorophyll and other pigments
Chlorophyll consists of four pyrrole rings which, together
with a ®fth ring, build a porphyrin structure. A long phytol
chain is bound to the porphyrin structure (Hendry, 1988).
Chlorophyll is present in all photosynthetically active cells.
During leaf senescence, chlorophyll is decomposed, whereas the carotenoids (yellow pigments) accumulate and new
red anthocyanins are synthesized, giving the autumnal
coloration of foliage (Matile, 2000). The porphyrin ring is
cleaved via different intermediate stages to colorless
products (Hendry, 1988; HoÈrtensteiner, 1999). Supposedly
these colorless decomposition products as well as the carotenoids and anthocyanins are decomposed in the soil; however, their fate and relevance in soils remain largely
unknown. The brown color of dead leaves is due to the
oxidation and subsequent polymerization of secondary
144
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 2. Structure of starch components: amylose and amylopectin.
phenols, occurring when the subcellular compartmentation
collapses after cell death (Matile, 2000).
6. Plant cell wall components
6.1. Polysaccharides
6.1.1. Cellulose
Cellulose is the most abundant biopolymer, as it
comprises the major structural component of the cell walls
of lower and higher plants. We ®nd high cellulose contents
in stalks and stems and in other woody parts of plants.
Cellulose is also a component of the cell walls of algae
and fungi, whereas it is only seldomly found in bacteria
(Peberdy, 1990; De Leeuw and Largeau, 1993).
Cellulose is a linear polymer glucan and is composed
of glucose units (.10 000) which are linked by b-(1-4)glycosidic bonds (Fig. 3a). The regular arrangement of the
hydroxyl groups along the cellulose chain leads to the
formation of H-bridges (Fig. 3b) and therefore to a ®brillar
structure with crystalline properties (Fig. 3c). Approximately 15% of the cellulose molecule has an amorphous
structure. The cellulose ®brils build a basic structure,
which is closely associated with hemicelluloses and in the
woody cell wall with lignin.
In soils under aerobic conditions, cellulose decomposes
slowly (Martin and Haider, 1986). We ®nd cellulosedecomposing organisms, above all, among the fungi. Also,
many eubacteria are able to decompose cellulose. Thus
cellulose is found only in traces in mineral soils. Particular
groups of bacteria can also decompose cellulose slowly to
low molecular acids under anaerobic conditions. However,
a relative enrichment of cellulose usually occurs under
anaerobic conditions, as in the formation of peat (De
Leeuw and Largeau, 1993).
The wet-chemical determination of cellulose is
accomplished after a two-stage hydrolysis procedure,
consisting of a ®rst treatment with concentrated H2SO4
(room temperature) and a second hydrolysis step with
dilute H2SO4 at elevated temperature to release the
glucose monomers (KoÈgel-Knabner, 1995). These steps
are necessary to break up the stable, crystalline cellulose
structure.
6.1.2. Non-cellulosic polysaccharides
The non-cellulosic polysaccharides of the plant cell walls
are often summarized as hemicelluloses or polyoses. Noncellulosic polysaccharides differ from cellulose in their
composition of sugar units (mainly pentoses, hexoses, hexuronic acids and desoxyhexoses), side chains and branching
(Fig. 4). Hemicelluloses are a group of polysaccharides of
different composition, which consist of cellulose-like sugar
units, bound together with glycosidic linkages, but are more
or less strongly branched and have a lower degree of polymerization than cellulose. Content and composition of
hemicelluloses are different in deciduous and coniferous
wood and litter (Table 5). Deciduous wood contains 3/4
pentoses and 1/4 hexoses, whereas the relation in coniferous
trees is reversed.
Xylans are a widespread hemicellulose group, consisting of (1-4)-glycosidic units of b-d-xylose. Additionally,
they contain, among other substances, a-l-arabinose and
4-O-methyl-d-glucuronic acid linked in the C2 or C3
position of the xylose. They comprise 5±30% of the polysaccharides in woody tissues. Mannans are composed of a
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
145
Fig. 3. Basic unit and supramolecular structure of cellulose; (a) basic unit with b-(1-4)-glycosidic bond, (b) intramolecular hydrogen bonds in native
crystalline cellulose between O-3-H and O-5 0 , and between O-2-H and O-6 0 (from Klemm et al., 1998), (c) fringed ®bril model of cellulose supramolecular
structure (from Klemm et al., 1998).
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
146
duous trees. Coniferous plants possess glucomannans
with galactose side chains.
Galactans are water-soluble, highly branched polysaccharides composed of (1-3/6)-glycosidic-bound b-dgalactose. The side chains consist, among other substances,
of l-arabinose and l-rhamnose which are linked via a (1-6)glycosidic bond. Pectins are complex, strongly branched
polysaccharides, consisting mainly of galactose, arabinose
and hexuronic acids. Pectins form the binding substance
between the cells, especially in herbaceous plants and fruits;
additionally, they occur in the primary wall. Pectin only
composes a small proportion of the plant material in
woody tissues and in grasses (approximately 1% in woody
material). Fig. 5 shows an exemplary structure of a plant
hemicellulose unit. Hemicelluloses and pectin are decomposed by many aerobic and anaerobic bacteria and fungi.
Their decomposition rate is higher than that of cellulose
(Swift et al., 1979).
Similar heterogeneous non-cellulosic polysaccharides are
found not only in plants, but also in bacteria, fungi and
algae. In contrast to crystalline cellulose, most hemicelluloses are soluble in alkaline solutions (SjoÈstroÈm, 1993), so
that they will at least be partly extracted during a traditional
humic substances extraction (KoÈgel-Knabner et al., 1989).
6.2. Lignin
Fig. 4. Basic structures of major sugar monomers in plant hemicelluloses
(from Fengel and Wegener, 1984); bold: example in ®gure (e.g. xylose),
structural isomers are indicated (e.g. ribose, arabinose).
chain of (1-4)-glycosidic-linked b-d-mannose, which are
partly supplemented with side chains of a-d-galactose
(bound by (1-6)-glycosidic bonds). Glucomannans with a
glucose±mannose-ratio of 1:2 are mainly found in deci-
Lignin is a high molecular, three-dimensional macromolecule consisting of phenyl propane units. Lignin ®lls
out the cell walls, which consist predominantly of linear
polysaccharidic membranes, providing structural rigidity.
Lignin is an important element of the cell walls of vascular
plants, ferns and club mosses. In comparison, mosses, algae
and microorganisms do not contain lignin (Higuchi, 1990).
Together with hemicellulose, lignin is found in the primary
wall, the secondary wall, and in the middle lamella of the
voids of the cellulose-micro®brils. It serves as a connection
between the cells and reinforces the cell walls of the xylem
tissue. Furthermore, it protects the woody cell wall against
microbial attack. After the polysaccharides, lignin is the
most abundant biopolymer in nature and a large contributor
to the residues of the terrestrial biomass.
The primary building units of lignin (monolignols) are the
cinnamyl alcohols coniferyl alcohol, sinapyl alcohol and pcoumaryl alcohol, shown in Fig. 6 using the conventional
Table 5
Amount and composition of the most important hemicelluloses in deciduous and coniferous wood (from Fengel and Wegener, 1984)
Polyoses
Deciduous wood
Coniferous wood
Content (%)
Units
Content (%)
Units
Xylans
25±30
5±10
Mannans
Galactans
3±5
0.5±2
Xylose, 4-O-methylglucuronic
acid
Mannose, glucose
Galactose, arabinose, rhammose
Xylose, 4-Omethylglucuronic acid
Mannose, glucose, galactose, acetyl groups
Galactose, arabinose
20±25
0.5±3
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
147
Fig. 5. Example of a representative hemicellulose unit: Arabino-4-O-methyl-glucurono-xylan.
terminology of the carbon atoms (deviating from the IUPACterminology). The monomers react through the so-called
dehydrogenative polymerization to a three-dimensional
macromolecule, which contains a multitude of C±C and
ether-linked compounds (Fig. 7). The arylglycerol-barylether (b-O-4) linkage dominates by far, followed by
biphenyl (5-5) and phenylcoumaran (b-5) linkages. Table 6
provides an overview of the most frequent types of bonds and
their structure in gymnosperm and angiosperm lignin. Most
of the linkages in lignin molecules are not hydrolyzable.
Lignin in gymnosperms, angiosperms and grasses is classi®ed based on differences in monolignol composition. The
lignin of gymnosperms is composed almost exclusively of
guaiacyl propane monomers, which are derived from coniferyl alcohol. Angiosperm lignin contains approximately
equal proportions of guaiacyl propane units and syringyl
propane units, derived from sinapyl alcohol. Lignin of grasses
is composed of about equal proportions of guaiacyl propane,
syringyl propane and p-hydroxyphenyl propane units. Additionally around 5±10% p-coumaric acid and ferulic acid,
which are predominantly esteri®ed to the terminal hydroxyl
groups of the propyl side chains, is found in lignin. The
proportions of coniferyl, sinapyl and p-coumaryl alcohol
amount to 94:1:5 in spruce lignin, 56:40:4 in beech lignin
(Fengel and Wegener, 1984) and 1:1:1 in grass lignin. Fig. 8
shows the model of spruce lignin as described by Adler
(1977), which contains all essential structural elements.
Nimz (1974) was the ®rst to develop a structural model for
angiosperm lignin using European beech as an example. In
these models, the ultrastructure of lignin is considered to be
heterogeneous and formed by random polymerization. In
contrast, Faulon and Hatcher (1994) proposed a three-dimensional lignin model based on a helical structure.
Part of the cellulose or hemicelluloses is bound to lignin
in the so-called ligno-cellulose- or lignin±polysaccharidecomplex (Fengel and Wegener, 1984; SjoÈstroÈm, 1993). The
structure of this complex is far from clear and no detailed
structural model can be given. It is supposed to be held
together by hydrogen bonds and covalent (ester or ether)
linkages, as exempli®ed in Fig. 9.
Lignin is comparably resistant against microbial decomposition; only a limited group of fungi (white-rot fungi) is
able to completely decompose lignin to CO2. Other fungi
(soft rot and brown rot fungi), in fact, induce structural
changes in lignins, but they are not able to induce a
complete mineralization. In soils, lignin degradation is
most probably mediated by consortia of decomposer microorganisms (Haider, 1992). As it is an oxidative decomposition process, lignin is not decomposed under anaerobic
conditions (Kirk and Farrel, 1987). During biodegradation,
lignin undergoes a gradual oxidative transformation process
that introduces carboxyl groups in the molecule, so that the
Fig. 6. Structure of lignin precursors: (I) coumaryl alcohol, (II) coniferyl alcohol, (III) sinapyl alcohol.
148
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 7. Structure of the major bonds in lignin.
transformed molecule is extractable by NaOH and is thus
found in the humic acid fraction (KoÈgel-Knabner et al.,
1988; Shevchenko and Bailey, 1996).
The analysis of lignin in plant litter and soils is dif®cult
because of the heterogeneous composition of monomers and
the different types of bonds. In proximate compound group
analyses, lignin is determined gravimetrically as a hydro-
lysis residue after the extraction of lipids and hydrolysis
of polysaccharides. This is the well-known determination
of Klason-lignin in wood chemistry. In plant litter and
soils, attention must be paid to the fact the residue contains,
besides lignin, further aliphatic plant components, e.g. cutin
and suberin (Zech et al., 1987; Preston et al., 1997). Thus
this method, although often used in litter bag and other
Table 6
Major bonding types in lignin of deciduous and coniferous wood (from SjoÈstroÈm, 1993)
Bonding type
b-O-4
a-O-4
b-5
5-5
4-O-5
b-1
b-b
% of total bonds
Arylglycerol-b-arylether
Non-cyclic benzyl-arylether
Phenylcoumaran type
Biphenyl
Diarylether
1,2-Diarylpropane
Resinol type
Deciduous wood
Coniferous wood
50
2±8
9±12
10±11
4
7
2
60
7
6
5
7
7
3
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
149
Fig. 8. Model of spruce lignin (from Adler, 1977).
studies, does not provide an appropriate value for lignin
content. The chemolytic analysis of lignin with CuO or
TMAH thermochemolysis has proven more suitable for
the assessment of the decomposition of lignin in soils
(Hedges and Ertel, 1982; KoÈgel, 1986; Hatcher et al.,
1995; Clifford et al., 1995).
6.3. Tannins and other polyphenols
Tannins are de®ned as polyphenols that occur in higher
plants. They precipitate proteins in aqueous solutions
and therefore act as tanning substances (Haslam, 1981).
Besides tannic substances, plants contain a multitude of
other secondary phenolic substances. Tannic substances
are distinguished in two groups, the condensed or nonhydrolyzable tannin (also termed proanthocyanidine) and
the hydrolyzable tannins (Haslam, 1981).
The basic structural unit of condensed tannin (¯avon-3-ol)
is presented in Fig. 10. The condensed tannins are polyphenols from polyhydroxy-¯avan-3-ol units, which are linked
mostly through C±C bonds between C-4 and C-8 and
sporadically between C-4 and C-6 and therefore, not
acid- or base-hydrolyzable. Normally, condensed tannins
consist of less than 10 ¯avan units, but up to 40 monomers have been found. Due to the presence of different
functional groups, an immense heterogeneity exists
within this compound class. Polymer proanthocyanidines
possess two phenolic OH-groups on the B-ring, whereas
prodelphinidines posses three OH-groups. The proanthocyanidines are bound to polysaccharides by glycosidic
bonds, e.g. on hemicellulose.
Hydrolyzable tannins have two basic units, namely sugar
(mostly d-glucose or similar polyoles) and phenolic acids.
They yield both units upon acid or alkaline hydrolysis. They
are a heterogeneous group of macromolecules, which can be
differentiated into gallotannin and ellagitannin. Gallotannins have a central sugar unit, which is esteri®ed with
several molecules of gallic acid (Fig. 11). Ellagic acid is the
basic phenolic unit of ellagitannins. Tannins are quantitatively important components of various plant parts. They
150
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 9. Most frequently suggested types of lignin±polysaccharide linkages (from Fengel and Wegener, 1984).
occur in various organs of higher plants, especially in
dicotyledones. Tannins in fungi, algae, moss and grasses
are of minor importance (Haslam, 1981). Condensed tannins
in high concentrations are found in the bark of trees (De
Fig. 10. Structural units of condensed tannins.
Leeuw and Largeau, 1993) and also in the shell material of
hazelnuts (Preston and Sayer, 1992).
Tannic substances play an important role as antifeedants,
i.e. they are defense of the plant against chewing phytophaguous insects or animals (Haslam, 1981). But tannins
are also reported to have residual effects outside the plant
and are therefore considered to be an important controlling
factor for litter decomposition in soils (Swift et al., 1979; De
Leeuw and Largeau, 1993). The proanthocyanidines, which
often occur together with lignin, are an important structural
component of the woody cell wall. They possess strong
antimicrobial effects due to their interactions with proteins.
They were also found to accumulate after the death of the
plant cell.
Few reliable analyses on the decomposition of tannins
in soils are available. Their chemical composition is
pronouncedly complex, and it is dif®cult to measure the
quantities of tannins in plant residues. Thus, little is
known in detail of the metabolism and turnover fate of
tannins in dying plant residues (Harborne, 1997). Most
probably, hydrolyzable tannins are decomposed more
rapidly than condensed tannins. As a total measure, the
concentration of water- and alkalisoluble polyphenols are
often determined by means of color reactions (Box, 1983).
Yu and Dahlgren (2000) found that the Folin±Ciocalteu
method was most suitable for total phenols and the butanol±
HCl method for condensed tannins. With these techniques
the tannin content may be underestimated, as the extraction
of tannins may not be complete and the extractability of
tannins may change during biodegradation. Yu and Dahlgren
(2000) reported that the recovery of tannins from foliage
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
151
Fig. 11. Structural units of hydrolyzable tannins: (a) gallic acid, (b) ellagic
acid, (c) pentagalloyl glucose, Gall, gallic acid.
was limited by the formation of protein±tannin complexes.
Hernes and Hedges (2000) developed a technique for analysis of condensed tannin monomers after depolymerization
of the whole litter or soil material without previous extraction. Preston et al. (1997) emphasize that the conventional
Klason lignin procedure includes cutin as well as tannins in
the Klason lignin residue and suggest the use of 13C NMR
spectroscopy in combination with molecular level techniques to analyze tannins in plant residues. Yu and Dahlgren
(2000) also found higher tannin contents with 13C NMR
than with extraction and colorimetric analyses.
Colorimetric analysis indicates there is a rapid loss of
tannins during biodegradation (Scho®eld et al., 1998;
Lorenz et al., 2000), either due to biodegradation or leaching. However, tannins in these studies may have escaped the
analytical window by not being extractable any more due to
formation of non-extractable complexes or due to slight
structural modi®cations that change their reactivity with
the color reagent.
6.4. Lipids
Lipids are organic substances that are insoluble in water
but extractable with non-polar solvents, e.g. chloroform,
hexane, ether or benzene (Dinel et al., 1990). Fig. 12
shows the most important components of the soil lipids,
which are already found as components of plant lipids.
Fig. 12. Major components of lipids in plants and microorganisms (from
Dinel et al., 1990).
Lipids are a heterogeneous group of substances that occur
both in plants as well as in microorganisms. Table 7 presents
an overview of the composition and occurrence of various
lipid classes in plants and microorganisms. A detailed
survey of the structural composition of lipids in plants and
microorganisms is found in Harwood and Russell (1984).
The surface lipids of plants are comprised of different structural groups. They cover the surface of leaves and needles
with a thin layer as a component of the plant cuticle. Table 8
gives an overview of the occurrence of various lipid classes
in the leaves of deciduous trees. The lipids in soil originate
from plants as well as microorganisms, whereas soil animals
only play a minor role.
6.5. Cutin and suberin
Cutin and suberin are polyesters that occur in vascular
plants. Cutin composes the macromolecular frame of the
plant cuticle in which the low molecular waxes and fats
are embedded. Together they form the cuticle. The cuticle
covers the epidermis and protects the surface of plants
against desiccation by the atmosphere. In contrast, suberin
is a cell wall component of cork cells, which compose the
periderm layer of sur®cial as well as subterranean parts of
woody plants. Suberin is also found in the endodermis and
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
152
Table 7
Occurrence ( £ ) of various lipid classes in the surface lipids of plants and
microorganisms (from Dinel et al., 1990)
Lipid class
Plants
Algae
Fungi
Bacteria
Hydrocarbons
N-alkanes
Branched alkanes
Ole®nes
Cyclic alkanes
£
£
£
£
£
£
£
£
£
£
£
£
Ketones
Monoketones
b-diketones
£
£
£
Secondary alcohols
Alcandioles (diester)
Free fatty acids
£
£
£
£
Wax esters
Primary alcohol esters
Triesters
£
£
£
Primary alcohols
Aldehydes
Terpenoids
£
£
£
£
£
£
£
in bundle sheath cells of grasses. The content of suberin is
particularly high in bark and in plant roots.
The cutin polymer is composed of di- and trihydroxy and
epoxy fatty acids with a C16 and C18 chain length (Fig. 13).
In the C16 group, dihydroxypalmitinic acid dominates, and
in the C18 group, oleic acid and hydroxyoleic acid dominate.
These are mainly linked by ester bonds and some ether
bonds (Kolattukudy, 1981). The individual composition
of cutin polymers is dependent on the plant species, stage
of development, and environmental conditions. Suberin is
composed of aliphatic and aromatic components (Table 9).
In contrast to cutin it contains monomers with a higher
chain length of C20 ±C30, in particular 1-alcanols, fatty
acids, v-hydroxy fatty acids and especially a,v-dioic
acids with a C16 or C18 chain length. In addition, suberin
contains phenolic acids, especially hydroxycinnamic acids.
Whereas it was supposed for a long time that the aliphatic
and aromatic units are linked by ester bonds in one macromolecule, recent research indicates that there are distinct
aromatic and aliphatic domains (Bernards and Lewis,
1998). Most of the recent work on suberinized tissues has
used potato tubers as a model system. It is not clear if the
®ndings obtained from this work also apply to other plants.
Recent investigations show that the cuticle of some
plants, especially Agave americana, contains a nonhydrolyzable biopolymer which consists of polymethylene
chains in addition to the hydrolyzable polyester material.
The structure of this polymer is still in debate and probably
contains also functionalized benzene rings in addition to
the aliphatic components (McKinney et al., 1996). This
macromolecule, classi®ed as cutan, seems to be comparably
resistant to decomposition and thus may supposedly accumulate in soil if present in the starting material (Tegelaar et
Table 8
Lipid composition of the sur®cial wax layer of leaves of various deciduous
trees comprised from GuÈlz et al. (1989), Prasad and GuÈlz (1990), and
Prasad et al. (1990)
Lipid class
Maple (%)
Beech (%)
Oak (%)
Hydrocarbons
Wax esters
Benzylacyl esters
Triterpenolacetates
Aldehydes
Primary alcohols
Triterpenoids
Fatty acids
6.9
5.5
2.1
14.4
38.1
10.2
4.9
17.1
17.0
17.4
0.9
±
10.3
34.8
±
8.1
6.4
1.1
±
0.5
38.8
36.0
3.6
6.1
al., 1989). Briggs (1999) points out that occurrence of cutan
in plant cuticles other than Agave americana remains to be
investigated in detail. The cutan components were only
present in small proportions in some major plant litter
materials and the corresponding forest ¯oor samples
(KoÈgel-Knabner et al., 1992). However, recently they
were isolated from a forest soil (Augris et al., 1998).
With the conventional extraction of litter with mixtures of
organic solvents only the low molecular (extractable) lipids,
fats and waxes are obtained (Ziegler, 1989; Bridson, 1985).
The polyesters of cutin and suberin can be degraded for
analysis by various depolymerization reactions (Holloway,
1984). Up to now the saponi®cation with BF3/methanol and
the TMAH thermochemolysis have been applied to litter
and soils (KoÈgel-Knabner et al., 1989; del Rio and Hatcher,
1998).
Cutin and suberin, which should be relatively easily
decomposed because of their chemical structure, are both
detected in soils and sediments. The proportion of suberin
from root litter, as compared to cutin, increases with
increasing soil depth (Riederer et al., 1993; Nierop, 1998).
7. Speci®c components of fungi and bacteria
7.1. Fungi
As in the cell walls of plants, the cell walls of fungi
consist mainly of homo- and heteropolysaccharides (Rogers
et al., 1980; Wessels and Sietsma, 1981; Peberdy, 1990).
Cell walls of some fungi also contain relatively high proportions of proteins. Lipids and melanins are quantitatively
minor components of fungal cell walls. Table 10 gives an
overview of the macromolecular components of fungal cell
walls.
The basic unit of the cell walls of fungi and also the
exoskeleton of insects is chitin. Chitin is composed of Nacetyl-d-glucosamin in b-(1-4)-glycosidic bonds. Fungi
also synthesize various matrix polysaccharides (glucans)
as cell wall components (Table 9) that differ in the type of
bonds between the glucose units. The structural polysaccharides, chitin and b-glucan, are high crystalline,
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
153
Fig. 13. Monomers and structure of cutin (a) and suberin (b) (from Kolattukudy, 1981).
non-water-soluble substances whereas the matrix polysaccharides are amorphous or only weakly crystalline and
mostly water-soluble.
Fungi, but also some bacteria synthesize various melanins
which occur as components of the cell walls, either incorporated in the structure of the cell wall or as its outermost
layer (Butler and Day, 1998). Melanin pigments contain
protein, carbohydrates, lipids and a polymeric core that
consists of various types of phenolic, indolic, quinone,
hydro-quinone and semi-quinone monomers. The intramolecular structure of the individual components is poorly
understood and there is no knowledge of the intact structure
of any fungal melanin (Bell and Wheeler, 1986; Butler and
Day, 1998). Melanins absorb visible light in the entire
wavelength spectrum and are therefore black- to browncolored. Due to their non-hydrolyzable structure they
protect the fungal cell wall against microbial decomposition
by hydrolytic enzymes (Butler and Day, 1998).
It is often assumed that melanins represent precursors of
humic substances in soil based on their humic acid-similar
attributions (Saiz-Jimenez, 1996). Unfortunately, very little
is known about their composition or decomposition in soils.
This is mainly due to the problems associated with the
extraction and analysis of melanins. Butler and Day
(1998) report that ligninase enzymes from white-rot fungi
are able to completely degrade fungal and bacterial
Table 9
Major components of cutin and suberin (from Kolattukudy, 1981)
Monomer
Cutin
Suberin
Dicarbonic acids
Substituted acids
Phenols
Long-chain fatty acids (C20±26)
Long-chain alcohols
Minor component
Major component
Scarce
Rare, minor component
Rare, minor component
Major component
Minor component
Major component
Frequent, substantial component
Frequent, substantial component
154
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Table 10
Macromolecular components of fungal cell walls (from Peberdy, 1990)
1. Skeletal components
Chitin
b-glucan
2. Matrix components
a-glucan
3. Further components
Chitosan
d-galactosamine polymer
Polyuronides
Melanins
Lipids
b-1,4-linked homopolymer of Nacetyl-d-glucosamine
b-1,3-linked homopolymer from
d-glucose with 1,3- and b-1,6glycosidic bonds
a-1,3-homopolymer of glucose,
a-1,3- and a-1,4-linked glucane
b-1,4-polymer of d-glucosamine
melanins. Large differences within the structural composition of fungal melanins, but also in comparison to humic
acids were observed by Knicker et al. (1995). Gomes et
al. (1996) investigated the melanins of various actinomycetes from Brazilian soils. As indicated by IR spectroscopy, the melanins had a higher aliphaticity than humic
acids from the same soils. There was also evidence for
high contents of proteinaceous materials and varying
amounts of polysaccharides.
7.2. Bacteria
Bacterial cell walls are composed of a peptidoglucan
(murein), which contains carbohydrate as well as amino
acid elements (Rogers et al., 1980; Koch, 1990). The
carbohydrate backbone of murein is composed of N-acetylglucosamine and N-acetylmuramic acid. Whereas glucosamine is also found in insects and fungi, muramic acid is
only found in bacteria. In addition to the 20 major
amino acids of proteins, bacterial cell walls also contain
a series of unusual amino acids, linked in a two-dimensional structure, which provides rigidity and elasticity to
the bacterial cell wall. Cell walls of Gram-positive
bacteria contain approximately 20±40 murein layers,
whereas the cell walls of Gram-negative bacteria are
composed of fewer, even possibly only one murein
layer. Therefore, murein amounts to approximately
50% of the dry weight of the Gram-positive but only
10% of the dry weight of the cell wall of Gram-negative
bacteria. Dextrans are extracellular polysaccharides of
bacteria. They are composed of a main chain of (1-6)-linked
a-d-glucose which is often branched in (1-3) and (1-4)
bonds.
Although the cell wall polysaccharides of microorganisms
are relatively easily decomposed, the basic units such as
glucosamine, galactosamine or muramic acid are found in
soil after hydrolysis (Stevenson, 1994) and they accumulate
during litter decomposition (KoÈgel and Bochter, 1985;
Coelho et al., 1997). Bacteria additionally produce a
multitude of structural components such as teichonic acid,
teichuronic acid, lipoteichonic acid and lipopolysaccharides
(De Leeuw and Largeau, 1993). Little is known about their
fate in soil.
During the last decade, a number of algae and bacteria
have been reported to contain substantial amounts of
insoluble, non-hydrolyzable aliphatic biomacromolecules,
termed algaenan and bacteran. They derive from condensation of complex lipids and are located in the cell wall
(Largeau and De Leeuw, 1995). They might be relatively
resistant to biodegradation and thus have high potential to
accumulate in soils (Augris et al., 1998). However, some of
these compounds may be artifacts produced in a melanoidinlike condensation reaction during the isolation procedure
(Allard et al., 1997). The importance of these insoluble
aliphatic high molecular components in soil microorganisms, as well as their biodegradation pathways and their
potential role as precursors for aliphatic components of
SOM, remain to be investigated.
8. Composition of various plant and microbial residues
In the following, the composition of various plant components and microorganisms will be demonstrated as it can
be deduced from solid-state- 13C NMR-spectra. Molecular
level information can be obtained from analytical pyrolysis
or (thermo)chemolysis. Examples for the information
obtained from these techniques are also given. This provides
an overview of the organic chemical composition of the
most important primary materials in soils and can serve as
a comparison to the structural information on SOM or SOM
fractions as investigated also by these techniques. For
detailed descriptions of these analytical techniques the
reader is referred to Knicker and Nanny (1997) and SaizJimenez (1994).
Nuclear magnetic resonance (NMR) spectroscopy is a
powerful experimental method for atomic and molecular
level structure elucidation. During such an NMR experiment the sample of interest is placed into an external static
magnetic ®eld that forces the nuclei spins to distribute themselves among different energy levels. The energy difference
(DE) between those levels is dependent upon the magnetic
properties and the strength of the surrounding magnetic ®eld
of the affected nuclei. Consequently, DE is different for
nuclei in different chemical and physical environments.
Spin transitions between those levels can be induced if an
additional electromagnetic ®eld with a frequency corresponding exactly to DE is applied. In this case, the induced
transitions can be detected as a resonance signal at a speci®c
resonance frequency in a spectrum. Note that different to
mass spectrometric techniques an NMR signal represents
only one certain kind of nuclei that is typical for a speci®c
chemical functionality. As mentioned above DE and consequently the resonance frequency is also dependent upon
the strength of the external magnetic ®eld which makes it
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
155
Table 11
Signal assignment in solid state 13C NMR-spectra of plant residues (from
Wilson, 1987; H. Knicker, unpublished PhD Thesis, Regensburg university,
1993)
Signal (ppm)
Assignment
Major compounds
containing these
groups
15
21
32
Terminal CH3-groups
CH3 in acetyl groups
Alkyl-CH2 in R±(CH2)±CH2 ±
CH3
Methoxyl C, C in amino groups
C5 in xylan, Cy in
phenylpropane units
Glucan-C2, -C3, -C5: C3 in
xylan
Ca and Cb, in arylether
C4 in arabinosefuranoside
C1 in b(1-4)-glucan
C1 in glucopyranosid
C2, C6 in syringyl unit
C3, C5 in p-hydroxyphenyl unit
C6 in guaiacyl unit
C2, C6 in p-hydroxyphenyl unit,
Ca and Cb in Ar±CHyCH±
CHOH
C1 in guaiacyl, syringyl, pcoumaryl
C4 in syringyl unit
C3, C4 in guaiacyl, C3 in
syringyl
C3, C5 in syringyl
C4 in p-hydroxyphenyl
Aliphatic amide, carboxyl C
Non-speci®c
Lipids, hemicelluloses
Lipids
56
65
72
13
Fig. 14. Classi®cation of the signals in solid C NMR-spectra of a humic
acid from peat (modi®ed from Bortyatinski et al., 1996).
dif®cult to compare results obtained with two different
NMR spectrometers. In order to circumvent this problem,
the resonance frequency is given relative to a reference
as chemical shift in ppm in relation to a standard
(tetramethylsilan).
A spectrum can be divided in various chemical shift
regions (Fig. 14). Table 11 provides a tentative assignment
of the resonance lines to functional groups of the 13C NMRspectra in plant materials (Wilson et al., 1983a; H. Knicker,
unpublished PhD Thesis, Regensburg University, 1993;
Preston and Trofymov, 2000). The chemical-shift region
0±50 ppm (alkyl-C) covers unsubstituted C in paraf®nic
structures. Signals at 30 ppm show polymethylene C in
long-chain aliphatic structures (e.g. lipids, cutin), whereas
branched structures produce signals around 20±30 ppm.
The terminal methyl group is found at approximately
14 ppm. Amino acid C also contributes to signal intensity
between 17 and 50 ppm. Aliphatic alcohol and ether structures (O-alkyl-C), such as those found in carbohydrates,
have resonances between 50 and 110 ppm. The C2, C3
and C5 in hexoses (cellulose and hemicelluloses), but also
C2 and C3 in pentoses (in hemicelluloses) contribute to the
dominating doublet signal at 72 ppm in plant residues. The
shoulder at approximately 64 ppm is caused by hexose-C6
or pentose-C5 and the shoulder at 85/89 ppm by the corresponding C4 (crystalline/non-crystalline). A clearly distinguished signal at 105 ppm is characteristic for the anomeric
C1 of polysaccharides in a glycosidic bond (Wilson et al.,
1983b). The signals of aromatic carbon with a chemical shift
at 110, 130, and 150 ppm are predominantly classi®ed as Catoms of lignins and namely the protonated C (110 ppm), Csubstituted aromatic C and phenolic C (150 ppm). The
splitting of the phenolic group signal in lignin at 148 and
153 ppm is frequently found in lignin from angiosperms
(Gil and Pascoal Neto, 1999). In this case, the signal for
84
93
105
115
124
128
134
136
148
153
160
172
Lignin, proteins
Polysaccharides
Polysaccharides
Lignin
Polysaccharides
Polysaccharides
Polysaccharides
Lignin
Lignin
Lignin
Lignin
Lignin
Lignin
Lignin
Lignin
Lignin
Acids, amides
the C3 and C4 of the guaiacyl unit (148 ppm) superimposes
the signal of the C3 and C5 carbon of the syringyl unit at
153 ppm. Tannins also show signals in this chemical-shift
area and thus partly coincide with the phenolic group signals
of lignin. However, a signal at 145 ppm is highly indicative
for a contribution of tannins (Preston et al., 1997). The
resonances of the ring carbon atoms C2 and C6 of the
syringyl units occur at 105 ppm (Wilson, 1987; Hatcher,
1987). Several structure elements of lignins occur in the
O-alkyl chemical-shift region. The signal of the methoxyl
group (also from hemicelluloses) is found at 56 ppm. The
lignin side chain shows signals throughout the chemicalshift region of 60±90 ppm, depending on the type of linkage
and the side chain substitution. The signal with a maximum
at 175 ppm is mainly attributed to carboxyl groups, also
esteri®ed, but also to amide-C, especially from proteins.
Fig. 15 shows the solid-state 13C NMR-spectra of various
plant parts from beech. The spectrum of beech wood has
signals, which are clearly attributable to polysaccharides
(cellulose and hemicelluloses) as well as lignin (75,
105 ppm; 130, 150 ppm). The acetyl groups of hemicelluloses are identi®ed by signals at 175 ppm for the COOHgroup and at 22 ppm for the acetyl group. The lignin peak at
56 ppm is caused by the carbon of methoxyl groups. The
156
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 15. Solid-state 13C NMR-spectra of various plant parts of beech (Fagus
silvatica L.): (a) leaf litter, (b) wood, (c) coarse roots .5 mm, (d) ®ne roots
,2 mm (from KoÈgel et al., 1988; unpublished c and d).
typical signals of wood are also found in beech litter
(Fig. 15a). Additionally, signals occur in the chemicalshift region of alkyl carbon with a maximum at 30 ppm.
They originate from extractable lipids and cutins of the
leaf surface. These compounds also give rise to larger
concentrations of COOH-groups. Tannins that also cause
signals in the chemical-shift region of aromatic carbon
and O-alkyl C, are not to be distinguished from lignin and
polysaccharides in conventional CPMAS 13C NMR-spectra.
A tannin resonance at 105 ppm can be differentiated from
the polysaccharide signal at the same chemical shift by
spectra obtained using the dipolar dephasing technique
(Wilson and Hatcher, 1988; KoÈgel et al., 1988; Preston
and Sayer, 1992). Polysaccharides and aromatic components dominate in the roots and in the bark of beech.
Fig. 16 shows the information obtained from pyrolysis of
different plant parts of Calluna vulgaris. Typical pyrolysis
products of polysaccharides are furans, reduced furans,
pyrans and anhydrosugars. Major aromatic signals in ¯owers, leaves and roots are phenols, methylphenols, catechol,
4-methylcatechol and ethylcatechol, indicative of polyphenols (Nierop et al., 2001). The pyrograms of the stem
wood are dominated by guaiacols and syringols from lignin.
Both the ¯owers and leaves as well as the roots show a much
lower contribution of syringols, compared to the wood.
High amounts of alkanes are observed in the pyrolysate
of Calluna ¯owers and leaves, mainly C31 and C33, with
smaller contribution of C27 and C29. The stem wood is
composed of C16 and C18 fatty acids, 2-methylketones
Fig. 16. Pyrolysis-GC-traces obtained from Curie-point pyrolysis of Calluna vulgaris; (a) ¯owers and leaves, (b) stem wood, (c) roots; from Nierop et al.
(2001). Signal assignment and source of pyrolysis product: 3 ˆ acetic acid (Ps), 4 ˆ 2-butenal (Ps), 6 ˆ 2,5-dimethylfuran (Ps), 8 ˆ toluene (Pp, Ps),
9 ˆ (2H)-furan-3-one (Ps), 10 ˆ 3-furaldehyde (Ps), 12 ˆ 2-furaldehyde (Ps), 14 ˆ vynilbenzene (styrene) (Pp), 16 ˆ 2,3-dihydro-5-methylfuran-2-one
(Ps) 17 ˆ 5-methyl-2-furaldehyde (Ps), 18 ˆ 4-hydroxy-5,6-dihydro-(2H)-pyran-2-one (Ps), 19 ˆ phenol (Pp, Lg, Pr), 23 ˆ 2-methylphenol (Pp, Lg, Pr),
25 ˆ 3/4-methylphenol (Pp, Lg, Pr), 26 ˆ guaiacol (Lg), 27 ˆ Levoglucosenone (Ps), 31 ˆ ethylphenols (Pp, Lg), 33 ˆ 4-methylguaiacol (Lg), 38 ˆ
dihydroxybenzne (catechol) (Pp, Lg), 39 ˆ 4-vinylphenol (Pp, Lg), 43 ˆ 1,4-dideoxy-d-glycero-hex-1-enpyranos-3-ulose (Ps), 44 ˆ 4-methylcatechol
(Pp), 45 ˆ 4-vinalguaiacol (Lg), 46 ˆ syringol (Lg), 50 ˆ 4-formylguaiacol (vanillin) (Lg), 54 ˆ 4-methylsyringol (Lg), 55 ˆ trans 4-(2-propenyl)guaiacol
(Lg), 56 ˆ 4-acetylguaiacol (Lg), 59 ˆ 4-ethylsyringol (Lg), 60 ˆ 4-vinylsyringol (Lg), 61 ˆ anhydroglucosan (levoglucosan) (Ps), 62 ˆ 4-(1-propenyl)syringol (Lg), 64 ˆ cis 4-(2-propenyl)syringol (Lg), 65 ˆ 4-formylsyringol (Lg), 66 ˆ trans 4-(2-propenyl)syringol (Lg), P n-alkene and n-alkane (pair) (Lp,
Abp), B methyl-alkene and -alkane (pair) (Lp, Abp), V fatty acid (Lp), X alcohol (Lp), n-alkane (Lp); Ps: polysaccharides, Pp: polyphenols, Pr: proteins,
Lg: lignin, Lp: lipids, Abp: aliphatic biopolymer.
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 17. Solid-state 13C NMR-spectra from (a) leaves of perennial ryegrass
(Lolium perenne) and (b) wheat straw (from H. Knicker, unpublished PhD
Thesis, Regensburg university, 1993); (c) roots from Lolium multi¯orum
(from Leinweber et al., 1993).
(odd-numbered C25 ±C31), C22 and C24 alcohols. These are
supposed to derive from suberin present in the outer bark of
Calluna stems. Small contributions of n-alkanes and n-alk1-enes are supposed to be derived from suberan (Nierop et
al., 2001). These results con®rm that the composition of the
components of different plant parts may be variable. It
should also be noted that speci®c plant compounds have
different susceptibility to pyrolysis, as the major component
of all Calluna parts are polysaccharides (Nierop et al.,
2001).
Fig. 17b shows the 13C NMR spectrum of wheat straw.
The polysaccharides dominate and account for approximately 80% of the total signal intensity. The concentration
of aromatic carbon, essentially lignin, is distinctly lower
compared to forest litter. Also the proportion of alkyl-C
structures (30 ppm), whose signals originate from lipids,
157
Fig. 18. GC-trace of TMAH thermochemolysis products obtained from
Agave americana cuticles (from del Rio and Hatcher, 1998). 2 ˆ nonane1,9-dioic acid dimethyl ester, 7 ˆ tetradecanoic acid methyl ester, 12 ˆ
hexadecanoic acid methyl ester, 15 ˆ octadecanoic acid methyl ester,
16 ˆ 16-methoxy-hexadecanoic acid methyl ester, 19/20/21 ˆ 8,16-, 9,16and 10,16-dimethoxy-hexadecanoic acid methyl esters, 22 ˆ eicosanoic
acid methyl ester, 24 ˆ mixture of hydroxy-methoxy hexadecanoic acid
methyl ester, 26 ˆ 9,10,16-trimethoxyhexadecanoic acid methyl ester,
28 ˆ 10,18-dimethoxyoctadecanoic acid methyl ester, 31 ˆ tetracosanoic
acid methyl ester, 32 ˆ 9,10,12,18-tetramethoxyoctadecanoic acid methyl
ester, 34 ˆ hexacosanol methyl ether, 36 ˆ hexacosanoic acid methyl
ester, 38 ˆ octacosanol methyl ether, 40 ˆ octacosanoic acid methyl ester,
41 ˆ triacontanol methyl ether, 42 ˆ triacontanoic acid methyl ester, 43 ˆ
dotriacontanol methyl ether, 44 ˆ dotriacontanoic acid methyl ester.
cutins and peptides, is comparably lower as in deciduous
and coniferous litter. The signal at 22 ppm is again caused
by the acetyl groups in hemicelluloses. Similar results have
been established for straw of other species and for hay
(Table 12). The spectrum of Lolium perenne (Fig. 17a)
compared to wheat straw shows intensive signals at 30
and 175 ppm from alkyl-C and carboxyl and amide groups
originating from lipids and cutin, and mainly from proteins.
The roots of grasses are distinguished by high concentrations of polysaccharides and low contents of lignin and
suberin. This is con®rmed by the absence of signals at
30 ppm and low aromatic signal intensity (Fig. 17c).
With tetramethylammonium hydroxide (TMAH) thermochemolysis speci®c compounds have been identi®ed from
cuticles. Fig. 18 gives the results from Agave americana
cuticles as an example. Both C16 and C18 components are
Table 12
Composition of various plant residues in solid-state 13C NMR-spectra (from FruÈnd and LuÈdemann, 1989)
Plant material
Carboxyl C 210±160 ppm (%)
Aromatic C 160±110 ppm (%)
O-Alkyl C 110±45 ppm (%)
Alkyl C 45±0 ppm (%)
Wheat straw
Barley straw
Oat straw
Rye straw
Hay
Beed wood
Pine wood
Spruce wood
2.1
1.2
2.5
3.3
5.6
2.0
0.7
0.9
10.7
11.6
13.4
12.6
12.4
13.2
19.0
18.2
83.8
83.9
80.7
80.0
72.5
82.5
78.9
79.0
3.4
3.0
3.3
4.0
9.5
2.3
1.4
1.9
158
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
Fig. 20. Solid-state 13C NMR-spectra of mixed fungal cultures isolated
from biodegradation experiments of Lolium perenne in soil with varying
decomposition conditions; (a) water saturation, (b) 60% water holding
capacity; (c) Solid-state 13C NMR-spectrum of the algae Chlorella (all
from H. Knicker, unpublished PhD Thesis, Regensburg university, 1993).
Fig. 19. Solid-state 13C NMR-spectra of a mixed culture of fungi and
bacteria in a forest soil under Mediterranean climate (from Baldock et
al., 1990).
observed, with a predominance of C18 monomers, mainly
the methyl derivative of 9,10,18-trihydroxyoctadecanoic
acid. Major other components identi®ed are octacosanol
dihydroxyhexadecanoic acids, mainly the methyl derivatives of 8,16-, 9,16- and 10,16-dihydroxyhexadecanoic
acid. Very little information exists on the composition of
cutin/suberin and other aliphatic biopolymers from trees and
agricultural crops.
Up to the present, few analyses of the quantitative organic
chemical composition of microbial biomass in soils with the
use of 13C NMR spectroscopy have been made. The ®rst
results from Baldock et al. (1990) on Mediterranean-climate
forest soils demonstrate that the composition of bacterial
biomass differs distinctly from fungal biomass. Both groups
contain high proportions of alkyl-C structures and small
amounts of aromatic carbon, but fungi have more O-alkyl
C and less alkyl C compared to bacteria (Fig. 19). The
fungal culture from a biodegradation experiment has a
composition relatively similar to the fungi in soil (Fig.
20a and b) with large proportions of polysaccharides and
alkyl-C. A mixed culture of the Ah horizon of a deciduous
forest under temperate climate conditions also shows high
concentrations of alkyl-C (predominantly non-extractable
aliphatic biopolymer), proteins and polysaccharides (KoÈgelKnabner et al., 1992). The 13C NMR-spectra of an algae
(Chlorella) demonstrates that the composition of these
algae is similar to those of bacteria (Fig. 20c). Golchin
et al. (1996) concluded that microbial materials synthesized from glucose by soil microorganisms in a laboratory incubation experiment were mostly O-alkyl, alkyl
and carboxyl carbon. Also in this experiment, phenolic
and aromatic structures were only found in small
amounts. Knicker et al. (1995) investigated the composition of melanins from various fungal species (Fig. 21).
Whereas some melanins were high in aromatic carbons,
other melanins showed high proportions of carbohydrate-like O-alkyl carbon structures (60±110 ppm).
One melanin was dominated by signals in the region
for alkyl C between 0 and 45 ppm. They found a high
variability in carbon structures of melanins derived from
the different fungal species. Obviously, there is a high variability in carbon structures of melanins derived from the
different fungal species.
These are the ®rst results of the molecular composition of microbial biomass as a parent material for humus
formation. Hopefully, the characterization of the composition of the secondary resources by means of thermolytic,
chemolytic and 13C NMR-spectroscopy can be extended to
further soils and various environmental conditions in the
next years.
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
159
Fig. 21. Solid-state 13C- and 15N-NMR-spectra of melanins extracted with NaOH from pure cultures of various fungi (from Knicker et al., 1995).
9. Conclusions
Future work should consider the detailed chemical
composition of plant and microbial components when
investigating their fate during biodegradation in soils.
Only a limited data set is available, mainly for the more
common types of plant litter and crop residues that enter
soils. Major gaps exist in the knowledge on the organic
chemical composition of relevant species contributing to
the organic matter in soils under pasture, arable land and
forests. This concerns mainly the composition of the aliphatic components, especially the cuticle as well as other lipids.
But also the composition of the lignin±polysaccharide
complex is not known in detail. The variability of different
plant components, such as lipids and tannins is only known
for a limited number of plant species. This also applies to
microbial components, especially lipids and melanins,
whose structure and composition are not yet known in detail.
Amount and composition of below-ground C input have
been analyzed only for a very limited number of species.
Future work should consider that the composition of belowground C input may be substantially different to the aboveground input.
A number of techniques exist for a detailed and indepth analysis of speci®c compounds in plant litter and
microbial residues and could be used in biodegradation
studies. Nonetheless most of the investigations use only
the classical Klason lignin or van Soest analysis of proximate fractions, that are loaded with analytical problems and
do not analyze speci®c compound classes. Major advances
could be made if available chemolytic, thermolytic and
spectroscopic techniques would be used and developed
that allow a detailed analysis of individual plant or
microbial components. This is possible with the presently
available techniques as shown in the examples above.
Conventional chemolytic techniques or pyrolysis in
combination with gas chromatography allow to analyze
speci®c monomers from a number of compounds, such
as lignin, polysaccharides, lipids, etc. In combination
with mass spectrometry an unambiguous identi®cation
of these compounds is possible, even in complex mixtures. Emerging techniques of liquid-chromatography in
combination with mass spectrometry (LC±MS) provide
new analytical approaches for more polar compounds.
Additional information on the composition of organic
matter in heterogeneous macromolecular mixtures is
160
I. KoÈgel-Knabner / Soil Biology & Biochemistry 34 (2002) 139±162
obtained from non-destructive spectroscopic methods,
such as NMR spectroscopy. Although speci®c compounds
are hardly identi®ed, this technique can give good results
concerning the gross chemical composition of plant
residues. The combination of the spectroscopic techniques
with thermolytic and chemolytic methods may add substantially to the understanding of the changes occurring
during biodegradation of organic residues. This will allow
to elucidate the composition and degradation pathways of
a number of plant and microbial compounds that are still
vague.
Acknowledgements
I would like to thank Georg Guggenberger, John Waid,
Heike Knicker and an anonymous reviewer for valuable
comments.
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