Chemical characterization of humic acid extracted from a Philippines agricultural soil

Material Information

Chemical characterization of humic acid extracted from a Philippines agricultural soil
Nefcy, Ann Mary
Publication Date:
Physical Description:
xi, 103 leaves : illustrations ; 29 cm

Thesis/Dissertation Information

Master's ( Master of science)
Degree Grantor:
University of Colorado Denver
Degree Divisions:
Department of Chemistry, CU Denver
Degree Disciplines:


Subjects / Keywords:
Soils -- Humic acid content -- Philippines ( lcsh )
Soils -- Humic acid content ( fast )
Philippines ( fast )
bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )


Includes bibliographical references (leaves 92-103).
General Note:
Submitted in partial fulfillment of the requirements for the degree, Master of Science, Department of Chemistry
Statement of Responsibility:
by Ann Mary Nefcy.

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Source Institution:
University of Colorado Denver
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Auraria Library
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All applicable rights reserved by the source institution and holding location.
Resource Identifier:
23281926 ( OCLC )
LD1190.L46 1990m .N43 ( lcc )

Full Text
Ann Mary Nefcy
B.S., University' of Michigan, 1971
A thesis submitted to the
Faculty of the Graduate School of the
University of Colorado in partial fulfillment
of the requirements for the degree of
Master of Science
Department of Chemistry
1 990

This thesis for the Master of Science
degree by
Ann Mary Nefcy
has been approved for the
Department of
Chemi s t ry
7 JyecIO

Nefcy, Ann Mary (M.S., Chemistry)
Chemical Characterization of Humic Acid Extracted
from a Philippines Agricultural Soil
Thesis directed by Graduate Faculty Advisor
Robert L. Wershaw
Humic substances comprise a general class of
biogenic, refractory, yellow-black organic substances
that are ubiquitous, occurring in all terrestrial and
aquatic environments. The formation of humic
substances remains a matter of debate. One proposed
model describes humic substances as being composed of
the partially degraded molecular components of living
organisms, which are held together in ordered,
membrane-1ike or micelle-like, aggregated structures
weak interactions, such as hydrogen bonding, pi
bonding, van der Waals interactions and hydrophobic
forces. This concept is explored while the author
chemically characterizes the carbohydrate components
humic acid extracted from a Philippines agricultural
soil on which corn was grown. The humic acid was
extracted using methods developed by the Internationa
Humic Substances Society. The humic acid was then
fractionated by gel filtration separation. The

carbohydrates, in both the whole humic acid and the
first fraction, were then analyzed by a modified
Hakomori procedure. The sugars were methylated,
hydrolyzed, reduced, and acetylated. The products were
analyzed by gas chromatography/mass spectrometry. The
results indicted the presence of partially degraded
plant fragments: dextrans, xylose, arabinose, rhamnose,
fucose, glucose, galactose, and mannose. The position
of the glycoside linkages suggest the presence of an
aggregated structure.
The form and content of this abstract are approved.
I recommend its publication
Si gned

Figures ..........................................vii
Acknowledgements..................................x i
1 . I NTRODUCT ION.....................................1
Purpose of the Study...........................2
Scope of the Study.............................2
2. REVIEW OF THE LITERATURE.........................k
Explanation of Terminology.....................5
Humic Substances...........................12
Degradation of Plant Cell Walls.............1 1*
Cellulose: Aerobic Degradation.............15
Cellulose: Anaerobic Degradation...........22
Hemice11u1ose: Aerobic Degradation.........25
Hemice11u1ose: Anaerobic Degradation......29
Lignin: Aerobic Degradation................30
Lignin: Anaerobic Degradation..............33
Hydrolysis of Humic Substances................35
3. EXPERIMENTAL......................................1*0

Source and Isolation of Humic Acids..........1*0
Gel Filtration...............................1*2

Colorimetric Analysis.........................^6
Infrared Analysis.............................^7
G 1 ucosy 1 1 i nkage Analysis..............A7
Hydrolysis and Reduction......................50
Alditol Acetate Derivatives...................51
Gas Chromatography Mass Spectrometry........51
k. RESULTS AND DISCUSSION...........................5^
Colorimetric Analysis.........................5^
Infrared Analysis..:..........................59
Gas Chromatography-Mass Spectrometry..........63
Fraction 1, Sephadex G-50, Supernatant..65
Fraction 1, Sepharose CL-6B..............72
Un f r a c t i ona t ed Humic Acid.......83
5 CONCLUS ION...,.................................91

Fi gure
2.1 The structure of cellulose..........................6
2.2 Cellulose: crystalline and amorphous regions.......8
2.3 Sugars found in the hydrolysates of hemi-
2.4 The building blocks of lignin polymers.............11
2.5 A structural model of softwood lignin..............13
2.6 Schematic representation of synergistic action
of cellulolytic enzymes........................18
2.7 Some lignin degradation products from woods
decayed by Phanerochaete chrysosporiurn........31
3.1 Schematics for humic acid fractionated on
Sephadex G-50..................................43
3.2 Schematics for humic acid fractionated on
Sepharose CL-6B................................44
3.3 Reaction schematics for g 1 ycoside-1inkage
4.1 Infrared spectra of A. First fraction of humic
acid (as the salt), Sephadex G-50
B. Unfractionated humic acid (as the salt),
C. Unfractionated humic acid precipitated,
following methylation and hydrolysis of 6....60
4.2a Total ion chromatogram (TIC), derivatized
supernatant from humic acid first fraction,
Sephadex G-50...................................66
4.2b Mass spectrum at retention time 7.271
minutes as determined by TIC shown in
Figure 4.2a............................................69
4.2c Mass spectrum at retention time 8.252
minutes as determined by TIC shown in
Figure 4.2a............................................70
v i i

4.2d Mass spectrum at retention time 9-014
minutes as determined by TIC shown in
Figure 4.2a............................................71
4.3 Possible structures for dextran..................73
4.4a (Top) Total ion chromatogram (TIC),
derivatized humic acid first fraction,
Sepharose CL-6B
(Bottom) Extracted ion current profile
for mass 1 1 8 (EICP)...........................75
4.4b Mass spectrum at retention time 5.935
minutes as determined by EICP shown in
Figure 4.4a............................................76
4.4c Mass spectrum at retention time 6.551
minutes as determined by EICP shown in
Figure 4.4a............................................77
4.4d Mass spectrum at retention time 7.019
minutes as determined by EICP shown in
Figure 4.4a............................................78
4.4e Mass spectrum at retention time 7.594
minutes as determined by EICP shown in
Figure 4.4a.....................................79
4.4f Mass spectrum at retention time 8.381
minutes as determined by EICP shown in
Figure 4.4a.....................................80
4.4g Mass spectrum at retention time 9.393
minutes as determined by TIC shown in
Figure 4.4a............................................8 1
4.4h Mass spectrum at retention time 9.553
minutes as determined by TIC shown in
Figure 4.4a............................................82
4.5a (Top) Extracted ion current profile for
mass 118 (EICP)
(Bottom) Reconstructed ion chromatogram
(RIC) derivatized unfractionated humic acid....84
4.5b Mass spectrum of scan #1073 as determined
by EICP shown in Figure 4.5a....................85
v i i i

4.5c Mass spectrum of scan #1156 as determined
by EICP shown in Figure 4.5 a...................86
4.5d Mass spectrum of scan #1216 as determined
by EICP shown in Figure 4.5a....................87
4.5e Mass spectrum of scan #1290 as determined
by RIC shown in Figure 4.5a.....................88
i x

2.1 E n do- 1 4 b e ta g 1 ucana ses...........19
2.2 Exo- 1 be t a g 1 uca na se s..............20
2.3 Cellobiose Hydrolases............................21
2.b Some Known Anaerobic Cellulolytic Micro-
organ i sms..................................23
3.1 Outline of Extraction Procedure..................41
A.1 Anthone Analysis of 2.5 ml. Samples Collected
off a Sepharose CL-6B Column.................55
b.Z Summary of Anthrone Analysis, Both Unfraction-
ated and Sephadex G- 50 Fractionation.................56

I am grateful to Jane Wallace, my supervisor at
the United Statfes Geological Survey, who encouraged me
to go back to school and move into a research project.
I wish to thank Dr. Robert Wershaw, currently my
supervisor at the United States Geological .Survey, for
transferring me into his project and for his continued
support throughout this research. Dr. Wershaw has
taken time out of his busy schedule to serve as my
thesis advisor, and also to patiently help me shape the
skills I need to become a research scientist. I am
indebted to Dr. Douglas Dyckes, Chair of the Chemistry
Department at the University of Colorado at Denver and
UCD Professor John Lanning for assuming the
responsibilities associated with thesis committee
membe r s.
I would especially like to thank Dr. Michael
McNeil, Department of Microbiology, Colorado State
University, who assisted with the GC/MS analysis and
was kind enough to patiently explain concepts to me.
i would also like to thank Colleen Rostad, U.S.
Geological Survey, who assisted with GC/MS analysis.
My special gratitude goes to Raymond McNamee. He
has shown me that even the handicapped and disabled can
achieve their goals.
x i

The formation of humic substances remains a matter
of debate. One concept is that the humic
macromolecules of nature arose from transformations of
polymeric and macromo1 ecu 1 ar components of plant,
animal, and microbial cells, such as polysaccharides,
mucopolysaccharides, proteins or peptides, lipids and
long-chain hydrocarbons, and lignins. Another concept
is that humic substances were synthesized through
interactions between component molecules released from
these, or between alteration products of such component
molecules, or between metabolic products from
biological activity in soil and water environments (l).
Wershaw (2) has proposed a model in which the humic
substances are composed of the partially degraded
molecular components of living organisms, which are
held together in ordered, membrane-1ike or micelle-
like, aggregated structures by weak interactions, such
as hydrogen bonding, pi bonding, van der Waals
interactions, and hydrophobic forces.
Humic substances are ubiquitous, and are found
wherever organic matter is being decomposed, or has
been transformed as in the case of sediments. Humic

substances are present wherever there is organic matter
in soils, sediments, and waters (T).
Puroose of the Study
There is extreme po1ydiversity in the composition
and structure of humic substances. This study explores
humic acid extracted from soil on which corn was grown
under controlled conditions. The study attempts to
focus on evidence that humic acids are composed of
partially degraded plant fragments.
Scope of the Study
Although humic substances represent a large
portion of the organic matter in soils, sediments, and
water, the compounds cannot be separated into pure
substances, and scientists must work with mixtures.
The problem is the extreme' heterogeneity of humic
substances. This study attempts to build on primary
and secondary structural work, but to approach the
issue from theories of tertiary and quaternary
structure. An understanding of the membrane-1ike
structure and of the weak interactions that hold the
aggregated structure together, will allow the proper
approach to the underlying structural components.

My approach is an effort to retain most of the
original g1ucosy1 l inkages in the carbohydrate fraction
of the humic acid, which was separated from the organic
substances in the soil. Gel filtration chromatography
was found to be a useful technique for concentration of
the carbohydrate fraction. Columns were chosen that
have minimal adsorption effects for the carbohydrates,
and the separation is primarily based on size.
Methylation analysis is the oldest, but still by
far the most widely used procedure for linkage analysis
in carbohydrate polymers (3). Methylation analysis can
be used on both fractionated humic acids and total
(unfract i onated) humic acids. Such procedures allow
the examination of possible structural changes during
fractionation. Gas chromatography/mass spectrometry
was used to identify the volatile derivatives of sugars
and of methylated and partially methylated sugars. By
identifying the substantial fragments, I was able to
work back from the fragments to a better understanding
of the humic acid complex.

Humic acids consist of a number of different
oligomers and simple compounds that arise from the
partial degradation of plant remains. In order to
better understand some of these degradation products
this review will cover the main components, of plants
from a chemist's viewpoint, degradation of these
components, and analytical difficulties encountered
in the hydrolysis of the humic acids formed from the
plant remains.
The degradation of plant cell walls is an
extremely complex subject and only a brief overview
is covered here. The main emphasis is on the
synergism of the reactions, and on the transfer
reactions of glycosyl groups (which can also be
simple hydrolysis in the case of water as an
acceptor). Both aerobic and anaerobic degradation i

Explanation of terminology
Cel 1u1ose
Cellulose is the chief component of wood and
plant fibers; cotton, for instance, is nearly pure
cellulose. It is insoluble in water and tasteless.
These properties, in part at least, are due to its
extremely high molecular weight.
Cellulose has the formula (C H 0.) .
6 10 5 n
Complete acid hydrolysis yields D-(+)-g1ucose as
the only monosaccharide. Hydrolysis of completely
methylated cellulose gives a high yield of 2,3,6-tri-
0-methy1 -D-g1ucose Therefore, cellulose is made up
of chains of 0-glucose units, each unit joined by a
glycoside beta-linkage to the C-^ of the next {k).
Figure 2.1 shows the structure of cellulose. Each
unit has three hydroxyl groups; one primary, in the 6
position, and two secondary, In the 2 and 3 positions.
Native cellulose is a huge molecule containing more
than 10,000 glucose units with a molecular weight
above 1.5 million. Cellulose molecules are arranged
in parallel polymer chains which are firmly bound
together by a large number of hydrogen bonds to form

Figure 2.1.
The structure of cellulose.

elementary fibrils. In these fibrils, areas of
complete order (crystalline regions) alternate with
areas of less order (amorphous regions). The fibrils
are arranged to form microfibrils and fibres (5).
Figure 2.2 shows cellulose fibrils.
Hemice11u1oses and Pectins
Hemice 1 1u1ose is a collective term for various
complex populations of polysaccharide molecules from
higher land plants. No single, much less short or
simple, definition would satisfactorily embrace all
of the materials called hemice11u1oses since the name
was coined 90 years ago. The following is a composite
of definitions satisfying many of the past and present
usages. Hemicelluloses are polysaccharides from, or
in, higher land plants and are abundant in cells that
have undergone lignification. In the cell walls they
form an aqueous gel in which bundles of cellulose
molecules (microfibri1s) are embedded in regular or
irregular orientations. The hemicelluloses are
dissolved by alkali; cellulose is not. They are much
more readily hydrolyzed by acid than cellulose and
then, depending on the plant, give L-arabinose, D-
xylose, D-mannose, some D- and less L-, galactoses, D-
glucose, L-rhamnose, D-glucuronic acid, ^-0-methy1-D-

The Structure of Wood Cell Wall*
Very long cellulose chain molecules built of up to 15,000 glucose units
combine to form strands with a largely crystalline structure. Sur-
rounded by shorter carbohydrate chain molecules (hemicelluloses) and
lignin (an amorphous high-molecular-weight polymer of aromatic com-
pounds), cellulose strands form fibrils. In the growing cell, fibrils are
laid down onto the middle lamella, which separates the newly formed
cells, in discernible layers (p, primary wall; S(, St, S3, outer, inner,
and terminal secondary walls, respectively). This principle is the same
for all cells, but the thickness of the individual cell wall layers may
differ among cell species. Within and between fibrils, the void spaces
and capillaries in the cell walls can be filled with water or air. During
heartwood formation, they can partly be filled with secondary extrac-
Figure 1.1. Cellulose: crystalline and amorphous
Reprinted with permission from Hoffmann, Per, and
Jones, M. A., 1989, Structure and degradation process
for waterlogged archaeological wood, in Rowell, R. M.
and Barbour, R. J., eds., Archaeological wood:
Properties, chemistry and preservation: Advances in
Chemistry Series 225, Washington, O.C., American
Chemical Society, p.36. Copyright 1989 American
Chemical Soc i'e t y .

glucuronic acid, D-ga 1 acturonic acid, and other sugars
(6). Figure 2.3 shows sugars found in hydrolysates of
This long definition is not comprehensive. It
implicitly includes some materials (the pectic
substances) other definitions would exclude and
excludes materials ( e g cereal gums) chemically
similar to hemice11u1ose but present in the non-
lignified cell walls of endosperms.
Lignin is a complex three-dimensional polymer
produced in vivo by an enzyme-initia ted dehydrogenative
polymerization of three pheny1propane monomers, p-
hydroxycinnamyI, coniferyl, and sinapyl alcohols
(Figure 2. *0 The monomeric pheny1propane units in
lignin, unlike those of other naturally occurring
polymers, are linked to each other not by a single
intermonomeric linkage but by several different carbon-
to-carbon and ether linkages, most of which are not
readily hydrolyzable. Thus, it is not surprising that,
in contrast to cellulose and other polysaccharides in
plants, lignin is resistant to degradation by most
microorganisms. Indeed, a main function of lignin
pro.bably is to protect plant cell walls from microbial


Figure 2.3. Sugars found in hydrolysates of
hem iceiluloses.
Reprinted with permission from Wilkie, K.C.B., 1983,
Hemice1 1u 1 ose: Chemtech, v. 13, no. 5, p. 3 0 8 .
Copyright 1983 American Chemical Society.

Figure 2 A .
The building blocks of
lignin po1yners.
Reprinted with permission from Hedges, J. I., 1989, The
chemistry of archaeological wood, in Rowell, R. M., and
Barbour, R. J., eds., Archaeological wood: Properties,
chemistry and preservation: Advances in Chemistry
Series 225, Washington, D.C., American Chemical
Society, p. 1 15- Copyright 1989 American Chemical

degradation, in addition to serving as a binding agent
and reinforcing material for cell walls (7,8). Only
certain fungi and probably certain strains of bacteria
are capable of decomposing lignin efficiently in nature
(9). Figure 2.5 shows a structural model of softwood
1 i g n i n .
Humic Substances
Humic substances comprise a general catagory of
naturally occurring, biogenic, heterogeneous, organic
substances that can generally be characterized as being
yellow to black in color, of high molecular weight, and
There are three.major fractions of humic
substances, and these fractions, once isolated from the
environment, are operationally defined in terms of
their solubilities.
Humin: That fraction of humic substances that is
not soluble in water at any pH value.
Humic Acid: That fraction of humic substances
that is not soluble in water under acid
conditions (below pH 2), but becomes soluble at
greater pH.
Fulvic Acid: That fraction of humic substances
that is soluble under all pH conditions (10).

Figure 2.5. A structural model of softwood lignin.
Reprinted with permission from Hlguchi, Takayoshi,
1985, Biosynthesis of lignin, in Higuchi, Takayochi,
ed., Biosynthesis and biodegradation of wood
components: New York, Academic Press, p. 143.
Copyright 1985 Academic Press. The experimental data
from which this diagram was drawn were taken from
Adler, E arid Harton, J., 1 959, Zur kenntnis der
carbony1gruppen im lignin. I: Acta Chemica
Scandinavica, v. 13, no. 1, p. 75~96.

Degradation of Plant Cell Walls
Humic substances are ubiquitous, and are found
wherever organic matter is being decomposed, or has
been transposed as in the case of sediments. Hence
they are present wherever there is organic matter in
soils, sediments, and waters (11).
Almost half of the global carbon fixed annually
via photosynthesis is incorporated into the
lignocellulosic cell walls of arborescent plants
(12) Lignin biodegradation is central to the earth's
carbon cycle because lignin is second only to cellulose
in abundance and, perhaps more significantly, because
lignin physically protects most of the world's
cellulose and hemice11u1oses from enzymatic hydrolysis
(13) . Living organisms produce a wide variety of
polymeric carbohydrates, of which cellulose is the most
abundant. The main deterioration of cellulose is
caused by microorganisms, and the microbiological
degradation of lignocellulosic materials is one of the
most important processes in nature (1*0.

Cellulose: Aerobic Degradation
The group of hydrolytic and other enzymes which
degrade cellulose are known collectively as the
cellulases. The term "cellulase" usually refers to a
mixture of enzymes rather than to a single enzyme.
The function of the cellulases may be classified into
three distinct areas. Firstly, they can be produced
by plants as morphogenic agents which weaken the
ce11u1ose-rich cell walls in preparation for growth,
differentiation or abscissions such as dropping of
leaves, flowers or seeds, or in the ripening process.
Secondly, they can be produced by some plant pathogens
as invasive agents which facilitate the penetration of
the plant. Thirdly, they can function as digestive
agents which permit the cellulose itself to be used as
a carbon source and, in so doing, may make other non-
cel lulosic plant tissue accessible to degradative
enzymes (15).
The primary reaction in the enzymatic degradation
of cellulose is the same as in degradation by acids,
viz. hydrolysis. This was recognized at least as long
ago as 1912 (16). The enzymes which catalyze the
hydrolysis of cellulose fall into one of two groups
according to the sites of cleavage in the

polysaccharide chain, viz. at random sites (endo-
enzymes) producing a range of fragment sizes, or at a
specific site adjacent to the non-reducing end
producing a unique low molecular weight fragment (exo-
enzymes) (17). It has long been established that the
rate of hydrolysis of crystalline cellulose by a
combination of endo- and exo-enzymes is much greater
than the sum of the individual actions of the enzymes.
However, as yet, the molecular interactions involved in
this synergistic action are not clearly understood and
there is little consensus on the matter (18).
The present state of knowledge of the degradation
of cellulose is based mainly on studies of cellulolytic
fungi. Trichoderma has three major enzymes in the
cellulase complex, 1,4,beta-D glucan cellobiohydrolase
( ce 1 1obiohydro1 ase) 1,l,beta-D glucan ^-glucano-
hydrolase (endog 1ucanase) and beta-g1ucosidase
(ce 1 1obiase) Cellobiohydrolase hydrolyzes the beta 1-
h glucosidic bond removing cellobiose units from the
non-reducing end of the chain. It is capable of
hydrolyzing amorphous and micro-crysta 1 1 ine cellulose.
Endog 1 ucanase randomly hydrolyzes the cellulose chain
to produce glucose and cellobiose. It is capable of
hydrolyzing amorphous cellulose but shows very little
activity against crystalline cellulose.. Beta-

glucosidase hydrolyzes celloblose to glucose and is not
capable of hydrolyzing cellulose. The sequence of
events is shown in Figure 2.6. Endog1ucanase is
considered to be the enzyme which initiates attack by
randomly cleaving the internal beta 1-* linkages.
Ce11 obiohydro1ase then sequentially removes cel 1obiose
from the non-reducing end of the chain (19). A
characteristic property of the exog1ucanases is that
the beta-configuration is inverted on hydrolysis (20).
The combined action of the endog1ucanase and cellobio-
hydrolase yields cellobiose and oligosaccharides which
are attacked by the beta-g1ucosidase to produce
This is definitely an oversimplification of the
process. Precise details of the initiation reaction
for cellulose degradation is still uncertain. There
is little consensus regarding the biochemical details
of the reactions. This could partially be attributed
to use of heterogeneous, poorly-defined substrates
that are not reproducible in the various laboratories
(18) But the problem is also one of complexity.
Some endo- and exo-enzymes which have been identified
are listed in Tables 2.1 and 2.2. Table 2.3 lists
cellobiose hydrolases. Many, but not all, endo-
glucanases display transferase activity towards

A/nor amt

- qiucaiiJaio

Figure 2.6. Schematic representation of synergistic
action of cellulolytic enzymes.
Reprinted with permission from Harrison, L. A., 1987,
Microbial degradation of cellulose polymers used in
cosmetics and toiletries: London, Chapman 6 Hall, Int.
J. Cosmet. Sci., v. 9, p. 78. Diagram drawn from the
theories of Wood, T. M., and McCrae, S. I., 1979,
Synergism between enzymes involved in the
solubilisation of native cellulose: Advances in
Chemistry Series 181 Washington, D.C, American
Chemical Society, p. 181-209.

Table 2.1. Endo-1,4-beta-giucanases.
Aspergillus niger 26,000 A.
Cellulo bacillus thermocellum 90,000 B.
Fusarium solani 37,000 c.
Pea 20.000
(Pisum sativum) 70.000 D.
Penicillium notatum 35.000 E
Ruminococcus albus 30,000-1.500.000 F.
Sporotrichum pulverulerttum 32.000 36,700 28.300
37,500 G.
Stereum sanguinolentum 20.500 H.
auranticus 34.000 I.
Trichoderma koningii 13.000 48.000 31.000
48,000 J.
Trichoderma viride 12.500
50.000 K.
A. Hurst, P.L., Neil sen, J., Sullivan, P.A., and
Shepherd, M.G., 1977 Blochem. J., v.165, p.3 3 .
B. Ng , T.K., and Zeikus, J.G., 1981, Appl. Environ.
Microbiol., v.kl, p. 2 3 1
C. Wood, T.M., 1969, Biochem.J., v.115, p.^57.
0. Wong, Y.-S., Fincher, G.B., and Maclachlan, G.A.,
1977, J.Biol. Chem. v. 252 p.1*02.
E. Aimin', K.E., and Eriksson, K.-E., 1968, Arch.
Blochem. Biophys., v.12^, p.129.
F. Wood, T.M., Wilson, C.A., and Stuart, C.S., 1982,
Biochem. J., v.205, p.129.
G. Almin, K.E., Eriksson, K.-E., and Pettersson, B.
1975, Eur. J. Biochem., v.52, p. 207.
H. Bjorndal, H., and Eriksson, K.-E., 1968, Arch.
Biochem. .Biophys., v. 12A, p.14 9 .
1. Tong, C.C., Cole, A.L., and Shepherd, M.G., 1980,
Biochem. J., v. 191 p.83.
J. Wood, T.M., and McCrae, S.I., 1978, Biochem. J.,
v 1 71 p 6 1 .
K. Berghem, L.E.R., Pettersson, L.G., and Axio-Freder
icksson, 1976, Eur. J. Biochem, v.6l, p.621.

Table 2.2. Exo-1.4-beta-glucanases.
Cellovibrio gilvus
Fusarium solani
Irpex lacteus
Penicillium funiculosum
Sporotrichum pulverulentum
Trichoderma koningii
Trichoderma reseii
Trichoderma viride
A Storvick, w 0 . Cole, F.E., and King, K.W., 1963,
Biochemistry, v.2, p.1106.
B. Wood, T.m., and McCrae, S.I., 1977, Carbohyd. Res.,
v.57. p.117.
C. Kanda. T., Nakakubo, S., Wakayabashi, K., and
Nisi zawa, K, 1978, Adv. Chem. Ser. v 181 p.211.
0. Wood, T.M., and McCrae, S.I., 1982, Carbohydr. Res.,
v.110, p. 291.
E. Eriksson, K.-E., and Pettersson, B., 1975, Eur. J.
Biochem., v 5 1 p 2 1 3 -
F. Halliwell, G., and Griffin, M., 1973, Biochem. J.,
v.135. p-587.
G. Berghem, L.E.R., Pettersson, L.G., and Axio-Freder-
icksson, V B 1976, Eur. J. Biochem., v.6l, p 6 Z1.
H. Berghem, L.E.R., and Pettersson, L.G., 1973, Eur. J.
Biochem., v 3 7 p.21.

Table 2.3. Cellobiose Hydrolases
Bacillus theobromae
Clostridium thermocellum
Sporotrichum pulverulentum
Stereum sanguinolentum
Trichoderma rescii
Trichoderma viride
A. Umezurike, G.M., 1971, Biochim. Biophys. Acta,
v 22 7 p. 9 .
B. Ait. Creuzet, N., and Cattaneo, J., 1979,
Biochem. Biophys. Res. Commun., v.90, p.537.
C. Deshpande, v., Eriksson, K.-E., and Pettersson, B.,
1978, Eur. J. Biochem., v.90, p.191.
0. Bucht, B., and Eriksson, K.-E., 1969, Arch. Biochem.
Biophys., v.129, p.416.
E. Inglin, M., Feinberg, B.A., and Loewenberg, J.R.,
1980, Biochem. J., v.l85, p.515 -
F. Berghem, I.E.R., and Pettersson, L.G., 1974, Eur. J.
Biochem., v.46, p.295.
G. Ladisch, M.R., Gong C.S.,and Tsao, G.T., 1977, Dev.
Ind. Microbiol., v. 18, p.157-
H. Li, L.H., Flora, R.M., and King, K.W., 1965, Arch.
Biochem. Biophys., v.111, p.439.

ce11odextrins. A general feature of glycoside
hydrolyses is their ability to catalyze synthetic
reactions leading to glycosides, oligosaccharides or
polysaccharides. All such reactions are formally
transg 1 ycosv1 ations (15).
An additional variable is the stereochemical
consideration. Enzymes should exhibit a high degree
of stereospecificity. Studies have shown that the
attack by cellulase (complete system) appears to be
confined to the surfaces of the elementary fibril
that contain the accessible 2,6 and 2,3,6 hydroxyl
groups. (21, 22).
Cellulose: Anaerobic Degradation
The understanding of anaerobic degradation of
cellulose is only just beginning. The introduction
of specific isotope labelling for cellulose components
has expanded knowledge of these processes. Federle
and Vestal (23) prepared ( 14C-ce 1 1 u 1 ose)- 1ignoce11u1 ose
from white pine by feeding live plant cuttings a C-
labelled cellulose precursor. After 50 days of
incubation under anoxic conditions, a range of 16 to
23% of ( 14C-ce 1 1u1ose) white pine was mineralized to
C02 and 'CH4. Some known anaerobic cellulolytic
microorganisms are shown in Table 2.A.

Table 2.4. Some Known Anaerobic Cellulolytic Microorganisms.
ORGANISM References
Acetivibrio cellobiopanun A., B.
Anaerobic fungi
(Neocallimastix frontalis,
Sphaeromonas communis.
Piromonas communis) C., D., E.
Bacteroides succinogenes F.
Butyrivibrio fibrisolvens
(Diplodinum spp.) H., I., J.
Clostridium cellobioparum K.
Clostridium thermocellum L., M.
Eubacterium cellulosolvens N.
Rumincoccus albus 0., P.
Rumincoccus flavefaciens Q.
A .
C .
0 .
E .
F .
I .
J .
N .
0 .
P .
Khan, A.W. , 1980, FEMS Microbiol.Lett., v 9 p2.
Khan,A.W., Saddler,J.N.,Patel,G.B., Colvin,J.R., and
Martin, S.M., 1980, FEMS Microbio1.Le11. v.7, p.**7.
Akin, D.E., Gordon, G.L.R., and Hogan, J
App1.Environ.Microbio1., v.k6, p.738.
Bauchop, T., 1981, Agric. Envlr., v.6, p
1981, J. Gen. Microbiol., v
and Ooetsch, R.N., 195^, J
Orpin, C G. ,
Bryant, M P.
v.3 7, p.1176
Cheng K.-J.,
8a c t e r i o I v
Mauldin, J.K
1983 ,
3 39 .
123, p.287.
Oa i ry Sc i. ,
J.W. 1977, J.
and Cos terton,
129, p.1506.
, 1977, Insect Biochem., v.7,
Prins,R.A. ,and Prast.E.R. ,1973,Protozool ,v.20,p.1*71.
Vogels, G.O., Hoppe, W.F., and Stumm, C.K., 1980,
Appl. Environ. Microbiol., v.1*0, p. 608.
Hungate, R.E., 19 ^ ^, J. Bacteriol., v.18, p. 380.
Ng, T.K., Weimer, P.J., and Zeikus, J.G., 1977, Arch
Microbiol., v 114, p.1.
Weimer, P.J., and Zeikus, J.G., 1977, Appl. Environ,
Microbiol., v.33, p.289.
Bryant, M.P., Small, N., Bouma, C.,
1958, J. Bacteriol., v.76, p.15.
Miller, T.L., and Wolin, M.J., 1973
v. 118, p. 836.
Smith, W.R., Yu, 1., and Hungate, R
Bacteriol., v 11h, p 72 9.
A.K.Sijpesteijn, J.Gen.Microbiol.,1951, v 3,p.289.
and Chu H ,
J. Bacteriol.
. E. 1973, J-

Schink (2*0 has indicated that the lack of
suitable techniques for the cultivation of strictly
anaerobic bacteria before the 1970's caused consider-
able confusion about their metabolic capabilites. He
proposes that the following reactions, observed in
natural environments, can be catalyzed in.the absence
of oxygen: hydrogenations, dehydrogenations,
hydrations, dehydrations, hydrolyses, condensations,
photoreactions carboxy1 ations, decarboxylations, lyase
reactions and coenzyme B 12 dependent rearrangements of
a carbon skeleton.
Anaerobic degradation of cellulose does not
depend solely on microorganisms. Light promotes the
deterioration of cellulosic products, particularly
cotton fabrics. With highly purified cellulose,
there is no light absorption in the visible region.
However, the mercury emission wavelength at 253.7 nm
can induce photodegradation and produce free radicals
in the photoirradiated cellulose. Although it was
initially thought that the presence of oxygen
influenced the direct photolytic rate of degradation,
it is now clear that the degradation by 253.7 nm
radiation is independent of the presence of oxygen.
Although light of wavelengths greater than 310 nm
cannot induce degradation of cellulose directly,

certain dyes and related compounds, lignin or metal
ions, are capable of absorbing light in the near-
ultraviolet or visible part of the spectrum and in
their excited state can induce degradation of
cellulose (25).
Hemice11u1ose: Aerobic Degradation
The hetero- 1 ,4-1 inked xylans (or heteroxy1 ans)
constitute a well characterized group of poly-
saccharides which form the major components of the
hemiceI 1u1 osic fractions of terrestrial plants (26).
They constitute a group of complex polysaccharides
composed of a backbone chain of 1 ,4-beta 1 inked D-
xylose residues to which are attached various
appendages. These may be L-arabinose; D-glucuronic
acid; various short oligosaccharide chains consisting
of D-xylose, L-arabinose, galactose and D-glucuronic
acid; 0-acetyl groups; feruloyl and p-coumaroyl esters
linked via L-arabinose residues; and benzyl ether
groups as occur in lignin-carbohydrate complexes (27).
Enzymes attacking the hemice11u1oses are hydro-
lytic by nature and are referred to as hemice11u1o1ytic
enzymes or hemice1 1u1 ases (28). Hemice 1 Iu1 ases (glycan
hydrolases) specifically degrade those glycans that
make up the backbone chain of the hemice11u1oses.

Hemice11u1 ases are classified like the ceilulases,
i.e., enzymes which catalyze the hydrolysis of hemi-
cellulose fall into one of two groups according to the
site of cleavage of the polysaccharide chain. An exo
enzyme degrades the polysaccharide by successive remov-
al of terminal glucose or o1igosaccharide units and
proceeds in a stepwise manner, usually from the non-
reducing end of the polysaccharide chain. These include
the exog1ucosidases, which cleave mono-saccharide and
short side-chain oligosaccharide linkages, and are also
involved in hydrolyzing the low molecular weight end-
products (e.g., oligosaccharides) released during
depolymerization of the main backbone chain. Endo
enzymes attack polysaccharides in a random manner
causing multiple scission that is accompanied by a
marked decrease in the degree of polymerization (DP)
of the substrate. The polymer is progressively
degraded into shorter fragments until nondegradable
products (usually mono-and disaccharides) are formed.
These endo enzymes include the glycanases, or poly-
saccharide hydrolases, and these are responsible for
attack on the polymer backbone Itself. Hemice11u1 ases
of the endo type constitute the most common group of
the hemicellulases. They are classified according to
the nature of the polysaccharide chain they act upon.

In the case of the heteroxy1 ans, they are known as the
1,^-beta-D-xylanases, or xylanases.
The total hydrolysis of the heteroxylan chain
requires the endoxy 1 anases acting in synerg-ism with
the exoglycosidases, which include the beta-
xylosidases, alpha-arabinosidases, galactosidases,
and a 1pha-g1ucuronidases (27).
Hemice11u1 ases of microbial origin have been
reported to be produced both constitutive1y (produced
irrespective of growth substrate) and inductively
(produced when grown on hemice11u1ose or similar
inducers (29). Cellulases, by analogy, are inductive
enzymes and are produced only when the microorganism
is grown on cellulose or cellobiose (30).
Degradation schematics are complicated by a
general class of enzymes called transg1ycosy1 ases.
Transg1ycosy1 ases operate by transferring the glycosyl
group of the donor to a suitable acceptor. If the
enzyme fails to discriminate against water as an
acceptor, the obvious end-result of the transferring
reaction is simple hydrolysis (31). A yeast xylanase,
Cryptoeoccus nffiUut xylanase (an endo type),
manifested transfer reactions from hydrolysis products
during degradation (32), i.e., not only did this
xylanase hydrolyze the xylan chain, but it was also

capable of synthesizing oligosaccharides from the low
molecular weight hydrolysis products. Several fungal
xylanases have exhibited transg1ycosy1 ation activity
(33, 34). Vrsanska et al. (35), describe the
mechanism for the binding and hydrolysis of xylose
oligosaccharides by the fungal endo 1 ,4-beta-D-xy1 anses
from !Asptrrji[[us nigcr str. 15. They showed that the
xylose substrate tended to be utilized during
degradation for resynthesis into xylose oligo-
saccharides of higher degree of polymerization (DP).
This indicated that xylose could be utilized as a
glycosyl receptor in transfer reactions. Two xylanases
have been isolated from the bacterium 'Bacittus
Both xylanases degraded xylan, but produced no xylose
in the course of the hydrolysis. The production of
higher oligosaccharides indicated transxy1 osy1 ation
activity (36).
The pectic substances should be included as hemi-
cellulose as there is no adequate clearcut boundary
differentiating the two sets of materials (37). Pectins
are complex heteropolysaccharides, mainly composed of
rhamnose, with a backbone of a 1pha 1 ,4 1 inked
galacturonic acids associated with neutral sugars.
Because of the co-valent linkages between pectin and
hemiceI 1u1oses in natural plant tissue, pectin

degradation always implies to a certain extent
hemice11u1ose degradation as well (38).
Hemice11u1ose: Anaerobic Degradation
Brenner et al. (39) conducted long-term studies
in three predominantly anaerobic wetlands: a salt
marsh, an acidic freshwater marsh, and a mangrove
swamp. [ 14C-po1ysaccharide] 1ignoce11u1 oses were
prepared from indigenous plants (**0) and incubated
under strictly anaerobic conditions with anoxic
sediments collected from the habitat sites. The
samples were periodically analyzed for the release of
14C as 14C02 and 14CH4 over a period of 1*0 weeks. The
radiolabel led polysaccharide fraction of Spartina alterni-
flori the ubiquitous salt marsh cord grass, was the
most amenable to degradation, with almost 30% of the
original 14C recovered as labelled gases. Twenty-four
percent of the labeled polysaccharide in the needle-
rush Juncus roemerianus was mineralized to 14C0a and
14CH4 after 280 days, while a surprising 20% of CartJ^
wa£uriana rad i o 1 a be 1 1 ed (a freshwater macrophyte) was
mineralized in the highly acidic (pH 3.8) anoxic peat
sediments of Okefenokee Swamp. Those results indicate
that slow but significant turnover of native
hemice 1 1u1ose may take place in anoxic sediments.

Lignin: Aerobic Degradation
Ulmer et al. (1*1) and Leatham (1*2) have
experimental work that suggests that lignin does not
support growth or its own degradation, and that it is
degraded to many different low molecular weight
intermediates. There are at least three modes of
degradative reactions that are operative in the
decomposition of the lignin macromolecule by white
rot fungi ( * 3 kk) :
1. Oxidative cleavage of side chains between
alpha- and beta-carbons leading to the formation of
a romatic acids.
2. Cleavage of beta-aryl ether bonds and
modification of side chain structures.
3. Degradation of aromatic nuclei through
oxidative ring opening.
More than 75 different degradation products were
identified from spruce and birch woods that had been
partially decayed by Phanerochaete chrysoporium (1*5, kb, k7)t
Representative structures are shown in Figure 2.7.
Low molecular weight components from birch cover a
broad spectrum. About 100 compounds were identified,
including aromatic acids, aliphatic acids, phenols,

Figure 2.7. Some lignin degradation products from woods
decayed by Phanerochaete chrysosporiurn.
Source: Kirk, T. K., USDA Forest Service, 1987,
Enzymatic combustion: The degradation of lignin by
white-rot fungi: Lignin enzymatic and microbial
degradation, Colloq. INRA, Paris, v. 40, p. 52.

and even some hydrocarbons (44). Structural
identification of the low molecular weight compounds
is relatively easier than the characterization of the
polymeric biodegraded lignins. However, such analyses
includes several uncertainties:
1) whether an identified compound is a fungal
degradation product or a fungal metabolite,
2) whether it is a primary degradation product
or a secondary degradation product,
3) whether a major identified product is a true
major lignin degradation product or an accumulated
product that the fungus is not capable of
assimilating, and
4) whether there are major degradation products
not identified because of rapid assimilation by the
fungus after their formation (48).
Whether white rot fungi can further degrade all
or even most of the unusual degradation intermediates
from lignin, such as V-X in Figure 2.7, is not known.
It Is conceivable that the total weight of lignin
converted to these "uncommon" products is minuscule,
and that their further degradation is accomplished by
other microbes. Or perhaps they are simply repolymer-
ized into humus. It seems unlikely, however, that the
fungi stop with these products (49).

Anaerobic Degradation
Aromatic monomers are known to be released by
aerobic lignolytic organisms such as white rot fungi.
Their fate in natural systems has yet to be determined.
It has been postulated that if further biodegradation
does not occur in aerobic environments, such components
may eventually enter anaerobic environments, where they
would be subject to anaerobic transformations (50).
Although elucidation of detailed pathways is
still lacking, anaerobic metabolism of aromatic
monomers in the absence of molecular oxygen is now
known to occur during anaerobic photometabolism (51),
under nitrate-reducing conditions (52), in microbial
consortia where fermentation is often coupled with
methanogenesis (53, 5^, 55), and by pure cultures of
fermentative bacteria (38, 50, 56).
Accumulation of nutrients close to surfaces is
of general advantage for microbes adsorbed to surfaces
under conditions of nutrient limitation (57). for
anaerobes, this enhanced metabolic activity of an
adsorbed microbial population also has the advantage of
an efficient protection from oxygen. Oxygen can
diffuse basically only from one direction and will be

used up very efficiently by aerobic microbes. Precipi-
tates of metal sulfides which commonly cover nearly
every surface in an anoxic habitat serve as an
additional oxygen protectant and redox buffer. Ferrous
sulfide has been used successfully as a reductant in
the cultivation of fastidious anaerobes (58). Metals
sulfides may also act as ion exchangers which maintain
a well- equilibrated balance of dissolved trace
elements, toxic heavy metals, and free hydrogen
sulfide. Clay and other insoluble minerals may
function in a similar manner and have been found to
enhance the metabolic activities of some sulfate-
reducing bacteria (59).
Surfaces may also contribute significantly to
the establishment and maintenance of stable anaerobic
microbial communities which depend on or benefit from
any kind of interspecies metabolic transfer (60).
Processes like this are of far greater importance
among anaerobic than among aerobic bacteria. Many
anaerobic bacteria form clumps or filaments which
lend themselves to adsorption to surfaces.

Hydrolysis of Humic Substances
Acid hydrolysis has been commonly used to
analyze the carbohydrate fraction of humic substances.
Various types and strengths of acids have been used, as
well as varying times and temperatures. Of course, a
wide varietv of substances have been elucidated from
these varying techniques. The following is a review of
some methods used for acid hydrolysis of humic
substances, with emphasis on structural changes that
When using sulfuric acid for hydrolysis, it has
been shown that part of the carbohydrate may become
sulfonated, leading to erroneous results (61). It has
been suggested that in aqueous solutions, sulfuric
acid can cause the formation of additional radicals
from acid catalyzed photopolymerization of radicals
(62). The lability of the carboxyl group makes the
identification of residues in polysaccharides difficult
due to decarboxylation occurring during hydrolysis,
resulting in the incorrect interpretation of the
primary structure (63). In addition to decomposing
carbohydrate material, acid may also convert sugars
into anhydro derivatives that may have the same gas-
chromatographic properties as other components in a

mixture. Thus, it has been found that under conditions
used to hydrolyze wood and pulp polysaccharides, D-
glucose gives 62% of 1,6-anhydro-beta-D-g1ucopyranose,
which has the same retention time as a 1pha-D-xy1ose
(64) . Problems are encountered in acid hydrolysis of
polysaccharides, since the monosaccharides obtained
will normally exist as mutarotated equilibrium mixtures
(65) . Great care must be taken in the interpretation
of data from the hydrolysis of humic substances. Humic
acids are believed to contain a number of functional
groups, including carboxyls, phenolic OH, alcoholic
OH, and carbonyls (66). These functional groups may
also contribute to the re-arrangement of polysacchar-
ides during hydrolysis, leading to incorrect inter-
pretations of the structures.
Schnitzer and Chan (67) noted that hot acid
hydrolysis (6m HC1) removed practically all the pro-
teinaceous materials and carbohydrates from a melanin
and a soil humic acid. Almendros et al. (68), indicated
that oxidative degradation methods can degrade the
humic acid completely. Preston and Ripmeester (69) have
used NMR to study acid hydrolysis (6N HC1) on fulvic
and humic acids. Refluxing for 24 hours caused removal
of amino acids, proteins and carbohydrates, as well as
low molecular weight phenolics, leaving a residue with

a much higher aromatic content. They suggested that it
is possible that some of the increase in aromaticity is
caused by reactions occurring during the hydrolysis.
Martin et al. (70) have generated data from strong
acid hydrolysis (reflux in 5N sulfuric acid) of fungal
melanins. They indicate that either complex poly-
saccharides are not present, or they have been
destroyed during acid hydrolysis and acetylation
procedures. Steelink (71) has suggested that perhaps
milder methods of hydrolysis will have to be developed
Anhydrous hydrogen fluoride hydrolyzes
polysaccharides and 1ignoce 1 1u1 osic materials more
rapidly and at lower temperatures than other acids
which have been studied for this purpose. The hexosans
are, at ambient temperature in HF, in a concentration-
dependent equilibrium between a 1 pha-D-g1ucopyranosy1
fluoride and oligomeric reversion products. HF
evaporation will shift this equilibrium towards
oligomeric reversion products. For pentosans, the
equilibrium favors exclusively the oligomeric reversion
products. Alpha- 1,6-oligomers (hexosans), (l * 3) ,
and (1 ^ k)- linked oligomers (pentosans), and branched
oligomers are predominant. The lignin ester and ether
linkages are possibly not cleaved by the action of this
reagent, but extensive temperature-dependent and

reaction time- dependent autocondensation, or
condensation with carbohydrates, takes place (72).
Mild acid hydrolysis procedures (trif1uoroacetic acid)
have been found useful by Albersheim et al. (73, 7*0
in examining the polysaccharide structures of cell
walls of higher plants. The group extensively studied
the structural characterization of oligosaccharides
isolated from the pectic polysaccharide rhamnoga1act-
uronan II (RG-ll). RG-II is a structurally complex
polysaccharide (D-ga 1 actosy1 -uronic acid-rich)
polysaccharide that is present in the primary (growing)
cell-walls of higher plants, and is composed of
approximately 60 glycosyl residues. It was found that
this compound could be characterized by a series of
partial hydrolysis, enzymatic hydrolysis, and analysis
of the residues. An advantage to the use of trifluoro-
acetic acid is that it is volatile and this readily
The strength of the acid affects the type of
compounds released. When subjected to hydrolysis with
acid (2M TFA, for 1 hour at 121 degrees), RG-II yielded
at least ten different monosaccharides. When RG-II was
subjected to milder hydrolysis conditions (2M TFA, for
70 minutes at 60 degrees), a heptasaccharide was
isolated that was structurally characterized and found

to contain aceric acid. Aceric acid is the only acidic,
branched-chain, deoxy sugar that has been identified in
nature. Hydrolysis with still milder conditions (o.lM
TFA for 2^ hours at **0 degrees) released a disaccharide
that was structurally characterized and found to be
composed of L-rhamnose and 3-deoxy-D-manno-2-
octulosonic acid (KDO).

Source and Isolation of Humic Acids
An agricultural soil sample was collected from the
experimental farm of the University of Southern
Mindanao in the town of Kabacan (7 degrees 6 seconds
North latitude and 12A degrees ^9 seconds East
longtitude) in the northeastern part of Cotabato by
personnel from the University of the Philippines
System. The sample was obtained by taking vertical
sections from the surface to a depth of 6 inches.
Collections were made from each side of a 0.25ha plot
on which corn was grown.
Humic acids were extracted from the soil by the
method of the International Humic Substances Society
(IHSS) as shown in Table 3*1- The humic acids
were extracted at the Natural Sciences Research
Institute, University of the Philippines System by
Noel M. Miraflor. The sample collection and extraction
procedures were the only parts that I did not actively
participate in, with respect to this research.

Table 3.1.
Outline of Extraction Procedure.
1. 1-2 kg. of air dried soil was sieved through a
2-nim sieve.
2. Soil sample was extracted with a volume of 0.1 M
HC1 equal to ten times the weight of the sample.
The pH of the solution was then adjusted to
between 1 and 2 with 1 M HC 1 .
3. Soil-HCl mixture was shaken for 1 h and
suspension then allowed to settle.
k. Mixture was centrifuged, and the supernatant
separated from the sediment. The sediment was
neutralized with 1 M NaOH to pH 7, and a volume
of 0.1 M NaOH equal to ten times the weight of
the sample was added under nitrogen.
5. The mixture from step U was shaken for k h and
then allowed to settle overnight.
6. Supernatant was separated from the sediment by
centrifugation, and the sediment was discarded.
7. The pH of the supernatant was adjusted to 1.0
with 6 M HCI and allowed to stand for 12-16 h.
8. The sediment was separated from the supernatant
by centrifugation and then redissolved in 0.1 M
KOH under nitrogen. The Kx concentration was
adjusted to 0.3 M with KC1.
9- The solution was. centrifuged and any sediment
present was discarded.
10. The supernatant was acidified as in step 7 and
the sediment (humic acid) separated by
centri fugation.
11. The humic acid from step 10 was suspended in a
solution 0.1 M in HCI and 0.3 M in HF overnight
to remove mineral matter.
12. The mixture was centrifuged, the precipitated
humic acid transferred to a dialysis casing and
dialyzed against distilled water until a negative
chloride test was obtained with AgN03.
13. The humic acid was freeze-dried.
Source: Wershaw, R.L., Pinckney, D.J., Llaguno,
E.C., and Vicente-Becke11 V., 1990, NMR
characterization of humic acid fractions from different
Philippine soils and sediments: Analytica Chimica Acta,
v. 232 p. 35.

Gel Filtration
The humic acid extracted from the agricultural
so i 1 samp 1e wa s fract i onated on ge
Thi s thesis will refer to chemical
t h i s humic acid and w i1 11 deal with
a c i d s and fract ions as:
1. Unfractionated humic acid
2. First fraction separated off Sephadex G-50
gel filtration media.
3. The supernatant made from the first fraction
separated off Sephadex G-50 gel filtration media
k. First fraction separated off Sepharose CL-6B
gel filtration media.
Figure 3.1 shows schematics for humic acid frac-
tionated on Sephadex G-50. This diagram is for a
column that was pumped from the bottom. A gravity feed
system was used initially, but was abandoned because
more reproducible separations were obtained with the
pumped system. Figure 3.2 shows schematics for humic
acid fractionated on Sepharose CL-6B. (Sephadex G-50
and Sepharose CL-6B are manufactured by Pharmacia LBK
Samples of the freeze-dried humic acid, weighing
approximately ,30 mg., were added to distilled water.
The humic acid was solublized, as the salt, by

(% of fraction)
24.7 7.
11.3 7.
6.4 7.
10.1 7.
first fraction
+ HC1
supernatant 7^777
direction of
floui of
elution solvent
Figure 3.1. Schematics for humic acid fractionated on
Sephadex G-50

first fraction
4 direction of
flow of
elution solvent
Figure 3.2. Schematics
Sepha rose CL-6B.
for humic acid fractionated on

adjusting the pH to 9 with NaOH. All calculations were
adjusted for the additional weight by assuming 6 meq
sodium per gram of humic acid.
Samples of the solublized humic acid were
repetitively added to the columns. The total elution
volume was arbitrarily separated into fractions based
on the appearance of dark bands. Four visible
fractions were identified for both gels. The fractions
were eluted with distilled water, then combined and
freeze-dried. The first fraction volume was
approximately **0 milliliters, based on measuring the
first dark brown band that appeared.
Fractionation was done on a Pharmacia XK-26
pressurized column packed with gel to a height of 225
mm, with an internal diameter of 26 mm. The flow rate
was 0.8 ml per minute. The void volume on this XK-26
column, packed with Sepharose CL-6B, was determined
using blue dextran, and found to be 37 milliliters.
To make the supernatant, the first combined
fraction eluted from the Sephadex G-50 was adjusted to
pH 1 with 1M HC1. A precipitate formed. The supernatant
was then withdrawn from the precipitate, and dialyzed
against distilled water until a negative chloride test
was obtained with AgN03. Membrane tubing, with a (MWCO)
mo 1 ecu 1 ar-weight-cut-off of 1 000, was used. The super-
natant was then freeze-dried.

Colorimetric Analysis
Two different colorimetric methods were used to
quantitatively determine the sugars in the humic
1. Anthrone method for hexoses (75): To a test
tube which contained 2.5 ml of less than 40 ppm
hexose in water, 5.0 ml of 0.21 anthrone in
concentrated sulfuric acid was carefully added. The
sampled was mixed thoroughly. The sample was heated
in a boiling-water bath for 5 minutes, and the
absorbance was read at 620 nm. Glucose was used to
calibrate a standard curve.
2. Orcinol method for pentoses and uronic acids
(76): To a test tube which contained 2.5 ml of less
than 20 ppm of pentose and/or uronic acid, 0.34 ml.
of 6% orcinol in 95% ethanol was added. Then 5 ml of
0. 1 !£ FeCI3'6H70 in concentrated HC1 was added and
the sample was mixed thoroughly. The sample was then
heated in a boiling water bath for twenty minutes,
and allowed to cool. The absorbance was read at 665
nm. D-(+)-Ga1 acturonic acid monohydrate was used to
calibrate a standard curve.

The absorbances were read on a Cary 118 UV-VIS
Infrared Analysis
Potassium bromide pellets were made during several
steps of the analytical process. These pellets were
scanned on a Perkin-Elmer 580 Infrared Spectro-
G1ucosy1 1inkage Analysis
Methylation: This method has been adapted from
the work of Albersheim et al. (77). The methylation
procedure is essentially a modified Hakomori procedure
(78) .
Sodium dimethy1su1finy1 anion was prepared as
follows: Sodium hydride (Z.k g 50% NaH in mineral oil)
was put into a 50 mi three necked flask with a
magnetic stir bar. The flask was fitted with a
thermometer and an entrance and exit port for
nitrogen gas. Nitrogen gas was flushed through the
flask during the entire procedure. Mixed hexanes (25
ml) were added, and the suspension was stirred. The
sodium hydride was then allowed to settle. Being
careful not to disturb the sodium hydride, the

hexanes, which contained the mineral oil, were
transferred into a beaker containing absolute ethanol
(which quenched small amounts of sodium hydride that
were left in suspension, and thus reduced the
possibility of explosion). This washing procedure was
repeated a total of three times. With nitrogen gas
rapidly flushing over it, the washed sodium hydride
(free from mineral oil) was magnetically stirred
until a dry powder was obtained. An aliquot of
10 ml dimethyl sulfoxide (DMSO) was added. The
suspension was heated to 55 degrees and maintained at
that temperature until no further bubbling from the
reaction was observed.
A freeze-dried humic acid sample of
approximately 2 mg was placed in a Pyrex test tube
with a screw-on Teflon cap. An aliquot of 1.00 ml
DMSO was added to the test tube and the contents were
kept under nitrogen throughout this procedure. The
mixture was stirred magnetically until the humic acid
was dissolved. An aliquot of 0.100 ml sodium di-
me thy1su1finy1 anion solution (DMSyl-base) was added
and the mixture was stirred for two hours at room
temperature. At least three more additions of DMSyl-
base were added until tripheny1 methane turned red in
the presence of an aliquot of this solution. The red

color indicated a positive test for excess DMSyl-
base. Then an equimolar amount of methyl iodide was
slowly added (0.024 ml for each aliquot of DMSyl-
base). The mixture was maintained at 20-25 degrees
by cooling in water during the addition of methyl
iodide. The mixture was stirred for one hour. At one
hour intervals, DMSyl-base and methyl iodide were
alternately added a total of two more times. The last
addition of methyl iodide was a four-fold excess
(0.096 ml), and the reaction mixture was stirred
overni g h t.
The per-O-methy 1 ated polysaccharides were
isolated by adding 1 ml of water to the solution and
dialyzing overnight against 1 liter of water, using
membrane tubing with a molecular weight cut off
(MWCO) of 1000. The material that diffused out of the
tubing was discarded in hazardous waste and the
polysaccharides were again dialyzed against water
overnight. The material inside the dialysis tubing
was again placed in a Pyrex test tube. The test tube
was placed in a freeze-dried flask. The contents were
then 1yopho1yzed .

Hydrolysis and Reduction
To the test tube containing the freeze-dried
sample, 0.50 ml trif1uoroacetic acid was added. The
test tube was sealed with a screw-on Teflon cap and
heated for one hour at 121 degrees Centigrade. The tube
was allowed to cool to room temperature and the
trif1uoroacetic acid was evaporated with nitrogen. To
the test tube, 0.50 ml isopropanol was added. The
sample was mixed and the isopropanol was evaporated at
room temperture.
The resulting partia I 1y 0-methy1 ated aldoses
were reduced to the corresponding partially 0-
methylated alditols by dissolving them in 0.4A ml of
95% ethanol and adding aqueous NaBD4 (0.4^ ml of 10
mg/ml in 1 M NH40H). The test tube was sealed and
kept for one hour at room temperature. To convert
the excess borodeuteride into borate, 0.10 ml acetic
acid was added to the tube. The sample was cleaned
up by adding acetic acid-metha no 1 (0.^0 ml of 1:9
v/v), mixing and evaporating with nitrogen. This
procedure was repeated three more times. The sample
was then evaporated twice from 0.40 ml methanol.

Alditol Acetate Derivatives
Alditol acetates were prepared by adding 0.10 ml
of acetic anhydride to the test tube containing
compounds from the previous step. The test tube was
sealed and heated for three hours at 121 degrees
Centigrade. The tube was allowed to cool to room
temperature and 1.00 ml water was added. Solid sodium
bicarbonate was added, a small amount at a time (25 mg)
until effervescence ceased. Additional water was added
to dissolve all the sodium bicarbonate. Approximately
1 ml dichloromethane was then added to the test tube,
and the contents mixed thoroughly. The dichloromethane
phase was then removed to a fresh tube and carefully
evaporated. Alditol acetates containing any number of
0-methyl groups, or none at all, can be separated on
the same chromatographic column.
Gas Chromatography Mass spectrometry
Two different instruments have been used in this
1. Hewlett Packard 5890A Gas Chromatograph with
a DB-23 column (30 meters, 0.25 mm I.D.) coupled to a
Hewlett Packard 5970 Mass selective detector. The

sample was dissolved in 50 microliters of acetone. The
injection was made using the split mode (split ratio
10:1). Helium was the carrier gas. The initial oven
temperature was 80 degrees Centigrade for two minutes,
increased to 170 degrees Centigrade at thirty degrees
per minute, then to 240 degrees Centigrade at four
degrees per minute, and held for five minutes at 240
degrees. The electron impact source was operated, at 70
electron volts.
2. Finnigan TS Q.-46 GC/MS/MS/DS using a Restek
Rtx5 column (30 meters, 0.25 mm I.D.). The sample
was dissolved in 100 microliters ethyl acetate. The
injection was made using the split mode. Helium
was the carrier gas.. The initial oven temperature
was 50 degrees Centigrade, increased to a temperature
of 300 degrees Centigrade at a rate of ten degrees per
minute, and then held at 300 degrees Centigrade for 12
minutes. The electron impact source was operated at 70
electron volts.
A summary diagram showing the reaction schematics
for the g1ycoside-1inkage analysis is in Figure 3.1.

| NoBD4/ nh4oh
COCH, (CHiC0)j0
CH,0 C ----
Figure 3.3- Reaction schematics for g1ycoside-1inkage

Colorimetric Analysis
The procedure for glycosy1 1inkage analysis should
ideally be applied to humic acid samples that have high
levels of hexoses. The anthrone method has been found
useful in quantitating hexoses from plant cell walls.
Anthrone analyses of the Philippines humic acid sample
is shown in Tables L.l and k.2. The humic acid sample
was extracted from a plot on which corn was grown and
found to contain a significant concentration of
The aggregation of humic acid creates difficulties
in estimating the molecular weight range for gel
filtration chromatography. Two different molecular
weight range gels were used in order to cover a wide
range and minimize problems of potential adsorption
effects. The Sepharose CL-6B gel has an approximate
molecular weight range of 10,000 to 1,000,000 for
dextrans (79). The analysis shown in Table ^ 1
indicates that the hexoses are dispersed across the
entire elution volume, indicating a broad range of

Table 1*. 1 Anthrone Analysis of 2.5 ml. Samples
Collected off a Sepharose CL-6B Column.
Sample § ppm hexoses Sample tf ppm hexoses
1 2.00 27 51.61
2 2.26 28 51.61
3 1 ** 3 29 53.59
1* 1 5** 30 55.51
5 1 .30 31 53.50
6 1 1 k 32 56.88
7 1.51 33 50.66
8 1.21 3** 1*4. 1 5
9 1 .29 35 1*1 .11*
1 0 1 .93 36 32.29
1 1 1 .61* 37 35 .**
1 2 0.86 38 26.21
1 3 1.22 39 22.85
l * 2.65 1*0 19.1**
l 5 1 .59 1*1 18.52
1 6 5-32 hi 19.00
l 7 33.56 *3 21 .98
18 1*8. 11* kk 25.22
19 51 .82 *5 31.52
20 52.28 1*6 1*6.28
21 **9.3 2 *7 1*6.28
22 *7.75 1*8 18.57
23 **5.21 *9 6.53
2k **3-03 50 7.70
25 **5 **0 51 1 .67
26 53-98 52 1.12
The first dark brown band begins eluting at Sample
Number 16, which is consistant with the calculated
void volume of 37 milliliters.

Table k.2. Summary of Anthrone Analysis, both
Unfractionated and Sephadex G-50 Fractionation.
Humic acid unfractionated: average: 12.2% hexoses 12.2% 11.9* 12.5* 12.0% hexoses hexoses hexoses hexoses
Humic acid (Sephadex G-50) first fraction
supernatant: 69.0% hexoses
58.0% hexoses
average: bk.5% hexoses 62.0% hexoses
65.0% hexoses
67.2% hexoses
65.8% hexoses
Humic acid
fractions e luted off Sephadex G-50:
(ave rages)
1 . 2k .1% hexoses
2. 11.3* hexoses
3. 6.1*% hexoses
k . 10.1% hexoses
Void volume calculated with bl ue dext ran to be

molecular weights. The Sephadex G-50 gel has an
approximate molecular weight range of 500 to 10,000 for
dextrans (79). The analyses shown in Table k.2
Indicates that the majority of the hexoses are eluted
in the first fraction. This indicates that the hexoses
are coming off in the void volume of the column and
that the approximate molecular weight of these sugars
exceeds 10,000. Sephadex G-50 is noted by the
manufacturer to have two types of gel-solute
interaction, ionic and aromatic. My studies indicated
that an immovable colored fraction always remained on
the Sephadex G-50 columns. This does not lessen the
value of the two types of columns for separating the
carbohydrate fraction. _
Gel filtration chromatography is based on separ-
ation primarily by size. As a solute passes through a
chromatographic bed, its movement depends upon the
bulk flow of the mobile phase and upon the Brownian
motion of the solute molecules which causes their
diffusion both into and out of the stationary phase.
The separation in gel filtration depends on the
different abilities of the various sample molecules to
enter pores of the stationary phase. Very large
molecules which never enter the stationary phase, move
through the chromatographic bed fastest. Smaller

molecules, which can enter the gel pores, move more
slowly through the column, since they spend a
proportion of their time in the stationary phase.
Therefore, molecules are eluted in the order of
decreasing molecular size (79). This would also suggest
that the broad dispersal of hexoses on the Sepharose
CL-6B gel indicates that the hexoses are in the 10,000
to 1,000,000 molecular weight range.
The average of 6A.5% hexoses in the supernatant
(separated from the acidified first fraction eluted off
a Sephadex G-50 column) poses an interesting problem.
The hexoses were separated from the precipitate upon
acidification. However, the separation should have
occurred during the original humic acid extraction.
Since humic acids exist as membrane-1ike or micelle-
like aggregates of plant degradation products, a likely
explanation is that the aggregated structure prevented
the separation of the 1ignin-saccharidic complex. The
gel filtration broke up the aggregated structure. Humic
acid samples may interact with Sephadex G-50. It has
been recognized that this gel has aromatic/hydrophobic
interactions as well as ionic interactions (79). Once
the protective aggregate has been broken up, the

1ignin-saccharidic complex can be separated by pH
adjusting. This leaves the hexoses In the supernatant,
with the lignin and other associated plant fragments in
the precipitate.
An additional colorimetric analyses was performed
on the supernatant of the hydrolyzed first fraction
eluted off a Sephadex G-50 column. The orcinol method
is useful in quantitating pentoses and uronic acids.
The orcinol method showed only 0.33% pentoses/uron i c
acids in the supernatant (average of three samples).
Infrared Analysis
Figure 4.1 shows three I.R. spectra taken at
various steps of the experiment. Spectra were taken of
(A) the first fraction of humic acid (as the salt)
separated off Sephadex G-50, (B) The unfractionated
humic acid (as the salt) and (C) the unfractionated
humic acid precipitate, which resulted following
methylation and hydrolysis of (B). Humic acid will
elute in the salt form from the gel filtration
In most mineral soils, practically all of the
humic material occurs in association with clay,
probably as a c1 ay-meta 1-humus complex (80). An

Frequency, cm
Figure 4.1. Infrared spectra of A. first fraction
of humic acid (as the salt), Sephadex G-$0
B. unfractionated humic acid (as the salt)
C. unfractIona ted humic acid precipitated
following methylatlon and hydrolysis of B.

examination of (B) indicates that the humic acid was
indeed separated from the clay by the extraction
process. Clay structure would show non-hydrogen bonded
OH as sharp shoulders along the broad OH stretch
between 3600 and 3 700 cm"1 (81). There would also show
a doublet around 600 cm-1 and another band around 1100
cnfwhich arise from the Si-0 bonds in the clay. These
bands are not present in (B)f the unfractionated humic
acid (as the salt) (82).
Bands near 3 44 8 cm"1 and 1639 cm"1 due to
moisture, frequently appear in spectra obtained by the
pressed KBr disc technique (83). This can obscure the
intensity caused by hydrogen bonding between hydroxyl
groups present in lignins and carbohydrates, and also
between these OH groups and various carbonyl groups.
The CH2 antisymmetric and symmetric stretching modes
gives rise to bands at 2920 cm-1 and about 2860 cm-1 ,
respect i ve1y (84).
The carbonyl band of humic isolates occurs in
different regions depending on whether it is present as
the free acid or the humic salt. The twin bands (in A
and B) at approximately 1 600 cm-1 and 1400 cm"1 indicates
that this is the salt of a humic acid. The C=0
stretching vibration occurs at 1720 cm"1 in humic acids.
When titrated to pH 7, the peak at 1720 cm"1 largely

disappears, and is shifted to about 1 600 cm*1 (anti-
symmetric stretch) and the absorption intensifies at
about 1^00 cm"1 ( symmet r i c stretch). Unfortunately, the
carboxylate band overlaps aromatic carbon-carbon bands
at 1590 cm'1. The effect can easily be seen in (C),
where the ester has been formed. The C=0 stretch now
appears at about 17^0 cm"1, and the C-0 stretch appears
about 1220 cm'1 (83, 85, 86). There remains
stretching from C = 0 at. about 1 600 cm"1, since sugars
which have been methylated and hydrolyzed will have a
carbonyl group on the anomeric carbon. This is also the
group now causing the hydrogen bonding near 3^00 cm"1.
The presence of an intensive absorption band
between 1100 and 1000 cm"1 is typical of carbohydrates
in general (87), enabling this region to be used in
analyzing carbohydrate-containing biopolymers. This
band is produced by stretching vibrations of single
C-0 and C-C bonds with the participation of bending
vibrations of C-H and 0-H. This band is notably
present in A and B, however; when the carbohydrates
have been methylated and hydrolyzed (C), the band has
been significantly diminished.
Strong absorption bands in the 9 0 9 ~ 6 5 0 cm"1 region
generally indicate an aromatic structure. Aromatic
and heteroaromatic compounds display a strong out-of-

plane C-H bending and ring bending absorption bands
in this region that can frequently be correlated with
the substitution pattern (88). The bands shown in
Spectrum B at 705 cm"1 and 835 cm"1 are characteristic
of meta- and para-disubstituted aromatic rings (89).
Figure 2.k shows the building blocks of lignin polymers
as meta- and para-disubstituted aromatic rings. Figure
2.5 shows the rigid structure that holds these building
blocks. Upon methylation and hydrolysis of B, the
bands at 705 cm"1 and 835 cm"1 are split, and a strong
methylene band forms at 1^50 cm"1 as shown in Spectrum
C. Strong absorption bands indicative of aromatic
rings are absent in Spectrum A, which is the first
fraction separated from the aggregated lignin-
saccharidic complex.
Gas Chromatography Mass Spectrometry
Gas chromatographic methods for carbohydrate
analysis involve the formation of derivatives of
sufficient volatility and thermal stability. Alditol
acetate derivatives can be used, formed by acetylation
of the hydroxyl groups with acetic anhydride. The
methods used in this research allow the determination
of ring size and substitution pattern. First, free

hydroxyls are methylated, creating ethers. Next, the
g1ycoside-I inkages are broken and the rings are opened.
The resulting -C=0 group on the number one carbon is
reduced and labelled with sodium borodeuteride. Hydro-
xyl groups now present on the molecule are acetylated.
In hexoses, the number one carbon will then be labeled,
and acetylated. The number five carbon will be acetyl-
ated, since it is the carbon attached to the ring
oxygen. Other linkages can then be determined by the
positions of acetylation (refer to Figure 3.3 )
Etherification of polysaccharides is dependent on
a sufficient degree of ionization of hydroxyl groups to
achieve alkoxide formation with enhanced
nuc1eophi 1icity toward the alkylating agent, methyl
iodide. Effective reaction is also dependent on the
polysaccharide being soluble in dimethy1su1foxide, a
highly polar aprotic solvent (90). Dimethy1su1foxide
can dissolve ionic compounds, and solvate cations very
well. However, it will not solvate anions to any
appreciable extent. SN2 reactions, in general are
strongly favored by the use of polar aprotic solvents
(91 ) .
The formation of molecular aggregates by humic
acids through hydrogen bonding can be a formidable
problem in obtaining a sufficient deg'ree of ionization

of hydroxyl groups. Repetitive additions of DMSyl-
base, followed by methyl iodide were necessary to
overcome this difficulty. The humic acid was
also not completely soluble in dimethy1su1foxide. It
seemed to be partially suspended in the solvent. The
compounds were stirred overnight to achieve as much
soiubility as possible from this heterogeneous mixture.
Some methylations have been achieved with partially or
completely insoluble substrates, e.g., with mixtures of
p o 1 y s a c c h a r i.d e s in plant cell wall preparations (90).
However, caution needs to be taken in interpreting the
results of any compound other than a pure compound.
Fraction 1, Sephadex G-50, Supernatant
Three hydrolysis steps greatly improved the sugar
yield of the supernatant. Figure 4.2a shows the total
ion chromatogram (TIC), derivatized supernatant from
humic acid first fraction, Sephadex G-50. This sample
was methylated, hydrolyzed, reduced, and acetylated.
Positive identification was made based on retention
times, compared to a standard sugar mixture carried
through the same procedure. Mass spectrometry allows
the unambiguous identification of hetero-
polysaccharides containing only one sugar of each

TC of AnnSol
Figure k.2a. Total Ion chromatogram (TIC), derlvatized
supernatant from humic acid first fraction,
Sephadex G-50.

class (e.g., one pentose and one hexose). However, If
the polysaccharide contains different sugars of the
same class (e.g., two or more different hexoses), an
unambiguous identification of the different methylated
sugars can only be made by comparing the GC retention
times to suitable standards (92). When dealing with
complex polysaccharide mixtures, standard compounds are
not always available and the analyst must be satisfied
with identification only by major classes of sugars.
The electron fragmentation patterns of the mass
spectra of partially 0 acety1 ated, partially 0-
methylated alditols are well known. Some of the rules
that can be used to determine the position of 0-methyl
and 0-acetyl groups are reviewed here (93)-
Rule 1. Primary fragments are formed by cleavage
of the alditol backbone.
Rule 2. The charge always resides on the fragment
with a methoxy-bearing carbon adjacent to the cleavage
po i n t.
Rule 3. Fragmentation between two adjacent
methoxy-bearing carbon atoms is favored over
fragmentation between a methoxy-bearing carbon and an
acetoxy-bearing carbon atom, which itself is highly
favored over fragmention between two acetoxy-bearing
ca rbon a toms.

Rule h Secondary fragment-ions are produced by
the loss of methanol or acetic acid. The loss of the
substituent on the carbon beta to the carbon bearing
the charge is strongly preferred.
Rule 5. When the partially O-acetylated, partially
O-methylated alditols are labelled at C-l with a
deuterium atom, the (nominal) mass-to-charge ratio
(m/z) of a fragment ion that contains C-1 is even,
whereas m/z of a fragment ion that does not contain C-1
is odd.
These fragmentation rules allow the arrangement of
the 0-acetyl and 0-methyl groups of the most common
partially O-acetylated, partially O-methylated alditols
to be readily determined. However, some electron-
impact mass spectra are not readily interpreted by
these rules. O-acetylated alditols can also lose a
fragment ion of MW 102, formed from two adjacent
acety1-groups, with a formula -Ac20.
Figures k.2 b, c, and d show the identification
of a dextran, formed from termina 1 1 ?nked, 6-linked and
3,6-linked glucose. Dextrans are bacterial poly-
saccharides of alpha- (16)D Glcp units differing only
in chain length and degree of branching which occurs
through a 1 p h a -(1 > 3) and a 1pha-(1k) branch points.
Many bacteria synthesize dextran from sucrose (9*0.


HOHc 205 '
.....I- -.
CH,0 C-
\ HORc
Figure A.2b. Hass spectrum at retention time 7.271
minutes as determined by TIC shown In Figure A.2a.

C-OCH, /
... |----<

c one
fltO c--
, ---I-------
/ CH.OC----
/ HOfle

'* ISO
Figure h. 2d. Mass spectrum at retention time 9.014
minutes as determined by TIC shown in Figure 4.2a.

Rapid methylation of dextrans has been
achieved through the use of the Hakomori procedure,
which utilizes dimethyl sufoxide as a solvent for the
dextran. Careful control of the reaction temperature
would, however, appear to be essential when a dextran
is methylated in this solvent, as hot methyl sulfoxide
has been reported to depolymerize native dextrans (95).
The bacteria that synthesize dextran belong to the
genera Lactobacillus, Leucohostoc, and Streptococcus
Dextran-1 i ke polysaccharides are also produced by
Acetobacter species, but by transfer of glucose
residues rom amyIodextrins. It is not known whether
the dextran structure is comb-like, laminated, or
ramified. Figure * 3 shows some possible structures for
dextrans. Another structural problem is whether the
side chains are regularly distributed. The methods for
studying this type of problem are not adequate.
However, the considerable variation in degree of
branching between different dextrans may indicate a
less regular distribution (96).
Fraction 1, Sepharose CL-6B
pentoses and hexoses linked in a variety of
are evident in the first fraction collected

Figure b.3
Possible structures
for dextran.
Re p r i nted
L i n dbe r g,
As p i na 1 1 ,
New York,
Ac a dem i c
with permission from Kenne, Lennart, and
Bengt, 1983, Bacterial polysaccharides, i
G. 0., ed., The polysaccharides, volume 2
Academic Press, p. 3**6. Copyright 1985

off a Sepharose CL-6B column. This is consistant with
the broad dispersal of hexoses from colorimetric
analysis, indicating a wide molecular weight range.
This sample was methylated, hydrolyzed, reduced and
acetylated once. The GC/MS data results are shown in
Figure k.k a b c,d,e,f,g, and h. Extracted ion current
profile (EICP) refers to the identification of only
those peaks in the TIC that contain an ion fragment of
a selected mass-to-charge ratio. Figure b.ka shows an
EICP for mass 118 and was used to determine the mass
spectrum of the compounds shown in Figures ^.^b through
4.4f. Figures and show the mass spectrum of
compounds determined using an EICP for mass 115. These
compounds are positively plant fragments (see Figure
2.3). Compounds elute in order of increasing molecular
weight. First, eluted are furanosides linked at the
one position, followed by pyranosides linked at the one
position. The next elution is furanosides linked at
more than one position', followed by pyranosides linked
at more than one position. The last group to elute
contains totally linked positions on pyranosides.
These compounds indicate that the compound was linked
in all positions so that none became methylated and all
positions became acetylated. These linkages can occur
because pyranoses exist in chair conformations to

Figure k.ka. (Top) Total ion chromatogram (TIC),
derlvatlzed humic acid first fraction, Sepharose CL-6B
(Bottom) Extracted Ion current profile for mass 118
(El CP) .


\ aaic

' HORc


Figure 4.4d. Hass spectrum at retention time 7*019
minutes as determined by EICP shown In Figure 4.4a.

C Oflc
Figure 4.4f. Hass spectrum at retention time 8.3 81
minutes as determined by EICP shown In Figure 4.4a.

Figure 4.4g. Hass spectrum at retention time 9-393
minutes as determined by TIC shown In Figure 4.4a.

151 ic,0. d-OAt **/
IQ0r\ \

362 .
C Me
| --
--C OAc
Figure 4.4h. Hass spectrum at retention time 9.553
minutes as determined by TIC shown In Figure 4.4a.

minimize torsional and van der Waals strain. It is not
unusual to find sugar esters linked in all positions,
as tannins, which are commonly found in a wide range of
plant tissues (97). The lack of methylation could also
be attributed to the aggregated complex that the sugars
exist in. It is possible that the aggregated structure
protects hydroxyls from methylation, and these become
acetylated following hydrolysis.
Unfractionated Humic Acid
The total unfractionated humic acid was also
examined to assure that there was no contamination from
the gel filtration procedures. The sample was carried
through two hydrolysis steps. Examination of the GC/MS
data from methylation, hydrolysis, reduction, and
acetolysis shows the presence of the termina 1 -1inked
glucose, 6-linked glucose and 3,6_linked glucose, as
found in the supernatant of fraction one. There is
also evidence of plant fragments, as was found in
Fraction 1, Sepharose CL-6B. GC/MS data are shown in
Figures 4.5 a,b,c,d, and e. Reconstructed ion
chromatogram (RIC) and total ion chromatogram (TIC)
is different terminology for essentially equivalent
methods of representing GC/MS output. Figure 4.5a
clearly indicates a large variety of sugars with a

SCANS 300 TO 1300
11/01/90 10110:00 CAL It 103IQ1 II
Figure k.$a. (Top) Extracted ion current profile for
mass 118 (EICP)
(Bottom) Reconstructed Ion chromatogram (RiC)
derlvatized unfractlonated humic acid.

tl/01/90 10i10:00 17sS3 CALI: 1031Q1 *t
M/2: 102
Figure 4. 5b. Mass spectrum of scan #1073 as determined
by EICP shown In Figure 4.5a.

11/01/90 10:18:00 19:16 CALI: 1031Q1 II RIC: 18944.
Figure 4.5c. Mass spectrum of scan #1156 as determined
by EICP shown in Figure 4.5a.

11/01/90 10:18(00 20116 CALI: 1031Q1 II
BASE M/Z: 110
RIC: 27640.

11/01'90 10:18:00 21:30 CALI: 1031Q1 11
Fnu^irpn I5B 2H 0T)
BASE M'Z: 115
RIC: 130560.
Figure *. 5e.
by RIC shown
Hass spectrum of scan #1290 as determined
In Figure 4.5a.

primary fragment mass-to-charge ratio of 118. This is
indicative of a first carbon deuterated and 0-
acetylated and a second carbon 0-methy1 a ted. The
termina 1 1 inked glucose, 6-1Inked glucose and 3,6-
linked glucose were selected from the EICP for mass
lift. Figure ^.5e shows data from secondary
fragment 115, which are indicative of total acetylated
hexoses, suggesting the presence of sugars in
aggregated structures or completely substituted.
Corn was grown on the plot from which these humic
acids were extracted. This enables a much better
definition of chemical decay processes leading to the
formation of humic structures in soil. In 1953,
Whistler, Conrad, and Hough found that corn cobs
contained a mixture of monosaccharides and
oligosaccharides (98). In 1956, Whistler and BeMiiler
extracted a hemice11u1ose from corn fiber, which they
called "corn fiber gum" and concluded was a highly
branched structure (99). With the advent of the
Hakomori procedure and advanced instrumental methods,
understanding of structural units in corn has improved.
Sugar analyses of the pectic polysaccharide fraction
from maize coleoptiles have indicated the presence of
rhamnose, fucose, arabinose, xylose, mannose,
galactose, and glucose (100, 101). The root-caps of