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Page 1: Books  Methods of soil an 3 1011 2

Chapter 35

Organic Matter Characterization

ROGER S. SWIFT, CSIRO Division of Soils, Adelaide, Australia

INTRODUCTION

It is generally accepted that the term soil organic matter refers only to the non­living organic material in the soil, which makes up by far the major portion of the total organic components. The living organic components, which are part of the soil biota, comprise a minor portion of the total organic material, and they will not be considered in this chapter. The soil organic matter can be of plant, animal or microbial origin and may be relatively fresh or highly decomposed and trans­formed. It is to the characterization of this material to which this chapter will be devoted. For in-depth reviews on this topic, the reader should consult Hayes and Swift (1978), Stevenson (1994), Aiken et al. (1985), and Hayes et al. (1989).

In chemical terms, it is possible to identify in soil organic matter, compo­nents belonging to the main classes of naturally occurring organic compounds found in plants and animals. Each of these compounds can be found in a wide variety of physical environments and physicochemical associations. In addition to these identifiable compounds, there are much larger amounts of organic mat­ter which are not amenable to current methods of chemical characterization. To bring some semblance of order to this diverse and complex system, it is neces­sary to establish and superimpose a set of classifications and definitions in order to establish a common framework for discussion and investigation.

Given the complexity of the soil system, any attempt to rigorously catego­rize soil organic components is likely to be, at best, imperfect. Quite clearly the most likely basis for classification lies in readily observable physical, chemical and/or biochemical differences between the various components. A useful delin­eation based on physical characteristics is that drawn between recognizable remains of plants (or animal) debris and the highly degraded and transformed materials which contain no recognizable plant, animal or microbial structures. Although this classification is essentially based on visual observation of physical differences, in essence, it purposes to differentiate between the results of bio­chemical transformations. As such it is unlikely to be wholly successful. For example, the same classes of organic compounds (e.g., carbohydrates, peptides and amino acids) can be found in both fractions.

Copyright © 1996 Soil Science Society of America and American Society of Agronomy, 677 S. Segoe Rd., Madison, WI 53711, USA. Methods of Soil Analysis. Part 3. Chemical Methods-SSSA Book Series no. 5.

1011

Published 1996

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In an attempt to overcome this problem emphasis can be placed directly on the chemical characteristics themselves as the basis of classification. Using this approach, the nonliving organic matter can be divided into humic and nonhumic substances (Hayes & Swift, 1978). In this scheme, all of the recognizable plant debris plus all of the identifiable classes of organic compounds (such as carbo­hydrates and peptides) in their natural or transformed state are categorized as nonhumic substances. The remaining amorphous, highly transformed, darkly coloured material which cannot be identified as belonging to an established group of organic compound are classified as humic substances.

As well as the classifications referred to above, it would be possible to devise others based, for instance, on the distinction between free, discrete organ­ic particles in the soil as opposed to bound organic matter which is adsorbed by clays, oxides and other mineral surfaces in the soil. Once again, the actual ar­rangements in the soil are not quite so simple, as it is difficult to distinguish absolutely between free and bound organic matter. For example, organic matter which is apparently bound may be sorbed to the surface of a mineral particle, or may be involved in other processes, such as occlusion, clustering or aggregate formation of mineral particles. Once organic components are involved in sorption or occlusion, the physical and chemical conditions in which they exist may pro­tect them from other soil processes, such as degradation and/or complexation, possibly for long periods of time (Theng et aI., 1992).

As can be deduced from the above discussion, the order which we may appear to superimpose on the system by the use of definitions and classification systems, although helpful, is, nonetheless, illusory. Soil organic matter is, and will, remain a complex mixture of organic compounds in a wide variety of physicochemical environments and subject to a wide raIlge of associations with minerals, metal cations, and anthropogenic organic compounds. It is this system which we must study either in situ in all its complexity, or we may simplify the material to be studied by undertaking extraction, purification and fractionation procedures.

Soil organic matter is an important pool of C in the global C budget and is a major component of the more active fractions involved in the earth's C cycle in terms of both total amount and annual flux. The levels of soil organic matter are sensitive to changes in temperature, rainfall and atmospheric CO2 concentrations and will be both an important indicator of, and possible contributor to, global climate change.

It has been commonly estimated that soil organic matter contains approxi­mately 1500 x 1015 g C (Schlesinger, 1984). Although this figure is based on a large number of measurements, it is a very difficult quantity to estimate accu­rately compared with, for instance, the amount of C as CO2 in the atmosphere or dissolved in the surface waters of the oceans. This is due to the spatial variabili­ty of soils and the paucity of data from some regions of the world. Most estimates for soil organic C range from 1100 x 1015 to 3000 X 1015 g C (Schlesinger, 1984), but all estimates indicate that there is more C in the soil than there is in the atmos­phere or in living biomass.

Just as important as the total amount of C is the flux of CO2 from the soil organic matter which is estimated at 75 x 1015 g C yr-1 (Schlesinger, 1984). This

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ORGANIC MATIER CHARACTERIZATION 1013

is roughly equivalent to twice the amount of plant material entering the soil each year. There are, of course, much larger reservoirs of C on the earth than soil or­ganic matter (e.g., carbonate and fossil fuel deposits or CO2 dissolved in the oceans) but most of these are involved to a lesser extent in the annual fluxes pass­ing through the C cycle.

As indicated above, the physically and chemically heterogeneous mixture of materials which make up soil organic matter varies substantially in terms of both their amount and resistance to biological decomposition. In most mineral soils under equilibrium conditions the highly decomposed and biochemically transformed materials are the dominant component, whereas the recognizable plants and other biological fragments represent only a few percentage points of the total organic matter. Of the transformed materials, the humic substances gen­erally constitute two-thirds to three-quarters. Most of the remainder is made up of much lesser amounts of carbohydrate and proteinaceous materials. Lesser amounts of a wide range of other compounds can also be identified, including lipophilic compounds such as fats and waxes.

On entering the soil, some of the more labile plant materials are decom­posed very rapidly, sometimes within days or weeks, whereas the more resistant compounds may survive for several months or even years. Similarly, the trans­formed materials will consist of substances with widely varying rates of decom­position. One of the characteristics of the decomposition process is that many of the compounds which are produced are more resistant to decomposition than their precursors. This is one of the essential features of the humification process. Part of this stability is due to the chemical changes which result from the bio­logical transformations which occur during decomposition and part is due to the association of organic macromolecules with mineral surfaces and with one another.

The overall or mean turnover time of organic matter in mineral soils is fre­quently several tens to a few hundred years but, due to the factors outlined above, this is made up of turnover times of particular components from as low as hours/days through to several thousands of years or longer for the more intract­able, charcoal-like materials. Although peats and a number of other soils contain large amounts of organic matter, most mineral soils contain just a few percent­age points of organic matter (usually 1-5% C) in the surface layers. Neverthe­less, this relatively small amount of organic matter has a profound effect on a wide range of soil properties, such as: cation exchange capacity, soil structure, the retention and cycling of nutrients, the binding of heavy metals and organic pesticides, etc. Given the importance of these effects, it is not surprising that the nature and properties of soil organic matter have attracted the interest of soil sci­entists for many years. Although considerable progress has been made in the understanding of many of the properties of these materials, it is, perhaps, disap­pointing that more progress has not been made in other aspects, such as the elu­cidation of structure. As we shall see below, there are good reasons for this which are inherent in the properties of the molecules which make up soil organic mat­ter in general, and humic substances in particular.

The purpose of this chapter is to provide an outline of the experimental methodology currently being applied to the study of organic matter in soils.

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Those methods which are commonly used, or able to be implemented with min­imal cost, are reported in detail. Other methods which require expensive or spe­cialized equipment are mentioned, but the reader is referred to more detailed ref­erence texts for further information.

Some of the chemical methods are somewhat dated, but have not changed in recent years. However, the instrumental methods generally have changed sig­nificantly since the last edition. The methods outlined and/or referred to in this chapter are meant to be used as a guide to the methods which have been report­ed in the literature. Scientific research is a dynamic study and the researcher should be flexible with ideas and methods, and be able to adjust these when nec­essary.

EXTRACTION OF SOIL ORGANIC MATTER

Although a number of techniques are now available which allow organic matter to be studied in situ in its unaltered, natural state, most of these techniques still require the organic matter to be removed or extracted from the soil. Among other things, extraction results in separating the organic from the mineral soil components, removing other inorganic interferences, increasing the organic mat­ter concentration and rendering the organic matter soluble. The application of many chemical procedures requires that the material being studied should be in solution.

One criticism of extraction procedures is that, as a consequence of the extraction process, the organic matter constituents are modified to a greater or lesser extent. It is, of course, inevitable that associations with mineral surfaces and other organic constituents will be disrupted, as this is a necessary part of the extraction process. Of more concern is the possibility of chemical alteration of the organic matter itself, resulting in artifact formation caused by the extraction process. It is important that such chemical modification should be minimized or avoided, if at all possible. This is more likely to be achieved through the use of mild extraction reagents [e.g., neutral pyrophosphate (Na4PZ07)], as opposed to strong reagents [e.g., sodium hydroxide (NaOH)]. However, this usually results in a substantial decline in the overall yield of extracted organic matter.

Having decided to use an extraction procedure, it is as well to optimize its usefulness and efficiency by incorporating other steps, such as purification and fractionation into the overall procedure. Although the intention of an extraction procedure may be to isolate a particular component, the reality is that a complex mixture is usually obtained and, by the use of appropriate procedures, this can be turned to advantage. Thus, it is possible to obtain samples of humic substances and soil polysaccharides from a single extraction and to further fractionate them both. Indeed, it is highly desirable that such procedures are employed, in order to properly purify the particular component being studied.

Extraction and Purification of Humic Substances

The procedures used to extract, purify and fractionate humic substances exploit the basic chemical and physical properties of these materials (Hayes, 1985). If we are to understand how the various procedures operate and whether

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they can be improved or refmed, then it is necessary to have a reasonable under­standing of the composition, properties and structure of humic substances.

Humic substances are a closely related family of naturally occurring macromolecules, with a chemically ill-defined structure. For a detailed review of the composition and structure of humic substances the reader is referred to the book Humic Substances II: In Search of Structure (Hayes et aI., 1989), only sali­ent points will be noted here.

In terms of the chemical composition of humic substances, the following statements can now be made with some degree of certainty. The presence of sub­stituted aromatic rings in humic substances has now been independently con­firmed by a range of chemical and instrumental techniques. The substituents on the aromatic rings are predominantly carboxyl groups, hydroxyl groups, carbo­nyl groups, and aliphatic units. Multiple substitution on the aromatic rings is common. Significant amounts of aliphatic C are also present with chain lengths of 1 to 20 C atoms but with smaller chain lengths predominating. The aliphatic units may be present as side chains or as units within the main polymer chain attached to aromatic units at both ends. The functional groups, which are distrib­uted throughout the length of the molecule, are certainly attached to aromatic groups and they may also be attached to the aliphatic moieties.

The various substituted aromatic and aliphatic units described above are assembled together in a somewhat random or, at least, a largely disordered array, and joined mainly by strong C-C bonds and perhaps ether linkages as well. Such assemblies make up the molecular backbone of soil humic substances. Carbohy­drates and peptides can be attached to this backbone in lesser but significant amounts.

In terms of physical properties, humic substances are variable-charge materials in which the carboxyl and, to a lesser extent, phenolic hydroxyl func­tional groups dissociate progressively with increasing pH. The resultant negative charge gives rise to cation-exchange sites and to numerous inter- and intramole­cular charge interactions. Soil humic substances are extremely polydisperse and have molecular weights ranging from a few thousand to well over a million dal­tons (Swift, 1989). Together with the wide range of molecular weights there will, of course, be concomitant variation in molecular size but this will also be depen­dent on the molecular shape or conformation. Of the various possible models for macromolecular conformation it has been suggested (Swift, 1989) that the ran­dom coil structure is most appropriate for humic substances. Unlike other possi­ble conformations this model accommodates the disordered chemical structure, the solubility and solvation properties and the charge characteristics of these materials.

In summary, the molecules of humic substances consist of hydrophilic and hydrophobic groupings, charged sites and counter ions, the identity and propor­tion of which vary from molecule to molecule. In addition, there is a substantial molecular weight range to superimpose on the variation in chemical composi­tion. The limitations imposed by these properties on the application and success of extraction and fractionation procedures need to be recognized at the outset.

Soil humic substances are largely insoluble and may remain in the soil system for long periods of time before they are degraded by relatively slow

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chemical and biological oxidative decomposition processes. Humic substances in soil are rendered insoluble by: their own inherent chemical properties, the composition of balancing cations, interactions with other organic molecules, and interaction with mineral surfaces. In order to extract the humic substances, it is necessary to displace the insolubilizing cations and to disrupt the various intra­and intermolecular associations and surface interactions.

Generally, the cations responsible for the insolubility of humic substances are a combination of di- and trivalent ions (e.g., Ca2+, Mg2+, Fe3+, and Al3+) as well as H+. All of these ions are strongly held on the organic exchange sites so that there is only a small amount of dissociation and, as a consequence, very lit­tle intramolecular charge repulsion. In addition, the multivalent cations simulta­neously satisfy charged sites at different parts of the molecule, or between mol­ecules, thereby forming cationic bridges. Both of these effects cause the organic molecule to adopt a condensed conformation from which solvent (usually water) is largely excluded. The net effect of these processes is to cause the molecules to be insoluble.

Humic substances can form associations with other humic molecules or with different organic macromolecules such as lignin or partially decomposed plant materials. These associations may be via cationic bridges, polar interac­tions, hydrogen bonding or van der Waals forces. Whatever their nature these interactions will tend to confer insolubility on the humic macromolecule.

Similar bonding mechanisms are also possible with mineral surfaces. In this case, however, some of the associations may be stronger. For instance, humic substances may bond directly with Fe or Al atoms at a clay or sesquioxide sur­face. The net result of the many interactions is strong attachment to a surface and resultant insolubility. In addition to the effects listed above, the inherent chemi­cal properties of humic substances such as the relative amounts of polar and non­polar groups and the molecular weight will greatly affect their solubility proper­ties.

The selection of both the extractant and the method used for the extraction process should be made with an understanding of the chemical and physical characteristics of the fraction(s) of humic substances required to be separated from the soil matrix. Whitehead and Tinsley (1964) have proposed four criteria for solvents for humic substances. In their view, an effective extractant should have:

1. A high polarity and a high dielectric constant to assist the dispersion of the charged molecules.

2. A small molecular size to penetrate into the humic structures. 3. The ability to disrupt the existing hydrogen bonds and to provide alter­

native groups to form humic-solvent hydrogen bonds. 4. The ability to remove and immobilize metallic cations.

Stevenson (1994) has listed four criteria for the effective extraction method:

1. The method leads to the isolation of unaltered materials. 2. The extracted humic substances are free of inorganic contaminants such

as clay and polyvalent cations.

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ORGANIC MAlTER CHARACTERIZATION 1017

3. Extraction is complete, thereby ensuring representation of fractions from the entire molecular weight range.

4. The method is universally applicable to all soils.

Although the four requirements for an extractant are achievable, the four requirements for an extraction method represent more an ideal than a set of cri­teria that are achievable.

Extraction with Aqueous Solutions

In terms of the desirable criteria for an extractant listed above, water is a very good solvent. It is polar, has a high dielectric constant, is able to form hydrogen bonds and thereby disrupt other hydrogen bonds, and is small and pen­etrative. However, an accompanying solute or additional step is required to dis­place and immobilize the insolubilizing, multivalent metal ions by use of a reagent which is capable of forming soluble complexes with these cations or of removing them as precipitates. In this regard, solutions of pyrophosphate and EDTA {N,W-1,2-ethanediylbis[N-(carboxymethyl)glycine]} have become pop­ular extractants because of their ability to form stable, soluble complexes with metal ions at near neutral pH values. Alternatively, the multivalent cations can be removed by prior washing with dilute acid which gives a H+ -saturated soil. How­ever, the acidic conditions are not conducive to extraction of humic substances and the pH needs to be increased before proceeding further with the extraction process.

Having removed the polyvalent cations from the negatively charged organ­ic exchange sites, it is important to replace them with a counter-ion that dissoci­ates very readily, and Na+ and K+ are the ions most commonly used for extrac­tion. The resultant high degree of dissociation causes intramolecular charge repulsion and leads to molecular expansion together with ingress of water which effectively solvates polar groups and disrupts intermolecular bonds. The net result is that the humic substances become soluble and are extracted.

One of the oldest and most frequently used methods of extracting humic substances from soils uses sodium hydroxide as the extractant. In this procedure many of the polyvalent cations (if not previously displaced by acid pretreatment) are removed through the formation of insoluble hydroxides at high pH values and are replaced by sodium. The high pH of sodium hydroxide solutions also causes many organic functional groups to ionize resulting in a higher charge den­sity on the molecules. The importance of this effect can be judged by the fact that sodium hydroxide extracts far larger amounts of humic material at pH values of 12 and above than the sodium salts of complexing agents used at near neutral pH values (Table 35-1). The problems of artifact formation by oxidation of the humic substances which can occur at high pH values can be substantially reduced by performing the extractions under a Nz atmosphere.

Extracts obtained by using sodium hydroxide have higher average molec­ular weight and lower functional group content than those extracted from the same soil by sodium pyrophosphate (Na4PZ07) at neutral pH values. Observa­tions such as these clearly indicate that, at any given pH value, the solubility lim­itations of humic substances are predominantly determined by charge density and molecular weight considerations.

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Table 35-1. Yields and compositions of humic acid (HA) and fulvic acid (FA)t fractions extracted with different solvents from H+ soil (adapted from Hayes et aI., 1975).

Elemental composition

Extractant Fraction Yield C H N S Ash

%

OMF* HA 16 53.4 4.5 2.6 1.7 1.8 OMF* FA 2.0 49.9 4.0 3.1 1.5 4.7 Sulpholane HA 10.0 54.0 4.8 3.2 2.4 0.8 Sulpholane FA 12.0 51.8 4.3 3.2 1.7 3.3 OMSO* HA 17.0 53.5 4.1 3.2 1.9 2.7 OMSO* FA 6.0 51.5 4.1 2.1 1.2 6.5 Pyridine HA 34.0 54.7 5.0 4.3 NO§ 2.2 Pyridine FA 2.0 45.3 5.1 5.8 NO 3.8 EO~ HA 49.0 54.7 5.7 6.2 NO 3.7 EO~ FA 14.0 48.2 5.4 10.5 NO 5.8 O.5MNaOH HA 58.0 52.0 5.9 2.8 NO 2.0 O.5MNaOH FA 2.0 43.9 5.9 4.2 NO 2.5 1MEOTA HA 12.5 50.8 4.0 NO NO 2.6 1MEOTA FA 3.8 45.7 4.0 NO NO 5.6

t Excluding material lost during dialysis. * OMSO - dimethylsulfoxide, OMF = dimethylformamide, EOA = 1,2-diaminoethane § NO = not determined.

International Humic Substance Society Method. A number of methods for the extraction of humic substances from soil using sodium hydroxide solution have been published. These methods are generally successful and yield compara­ble results. The following is a method which has been developed by the Interna­tional Humic Substance Society (IHSS) as an acceptable method for the extrac­tion of humic substances from soils.

It has been clearly stated by IHSS that this is not meant to be a recom­mended or approved method, but a method that has been found to be satisfactory for most soil types and one which can be performed in most laboratories. It pro­duces relatively high yields and can be used as a standard method for compar­isons between and within laboratories. An important component of this method is the use of an adsorbent resin in the purification process. This can be replaced by dialysis if the resin is unavailable.

Materials

1. Hydrochloric acid (HCI), 1 M, 6 M 2. Sodium hydroxide, 1 M, 0.1 M 3. Potassium hydroxide (KOH), 0.1 M 4. Potassium chloride (KCI) 5. Hydrofluoric acid (HF) concentrated, 0.3 M 6. XAD-8 resin (Rohm & Haas Co., Philadelphia, PA) 7. Visking dialysis tubing (Visking Co., Chicago, IL) [MWCO (molecular

weight cut-oft)] 10 000 dalton Method

Remove roots and sieve the dried soil sample to pass a 2.0-mm sieve. Equi­librate the sample to a pH value between 1 to 2 with 1 M HCI at room tempera-

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ture. Adjust the solution volume with 0.1 M HCI to provide a final concentration that has a ratio of 10 mL liquid/1 g dry sample. Shake the suspension for 1 hand then separate the supernatant from the residue by decantation after allowing the solution to settle or by low speed centrifugation. Save the supernatant (FA Extract 1) for the isolation of fulvic acid using XAD-S.

Neutralize the soil residue with 1 M NaOH to pH = 7.0 then add 0.1 M NaOH under an atmosphere of N2 to give a final extractant to soil ratio of 10:1. Extract the suspension under N2 with intermittent shaking for a minimum of 4 h. Allow the alkaline suspension to settle overnight and collect the supernatant by means of decantation or centrifugation. Acidify the supernatant with 6 M HCI with constant stirring to pH = 1.0 and then allow the suspension to stand for 12 to 16 h. Centrifuge to separate the humic acid (precipitate) and fulvic acid (super­natant - FA Extract 2) fractions.

Redissolve the humic acid fraction by adding a minimum volume of 0.1 M KOH under N2. Add solid KCI to attain a concentration of 0.3 M [K+] and then centrifuge at high speed to remove the suspended solids. Reprecipitate the humic acid by adding 6 M HCI with constant stirring to pH = 1.0 and allow the suspen­sion to stand again for 12 to 16 h. Centrifuge and discard the supernatant. Sus­pend the humic acid precipitate in 0.1 M HClIO.3 M HF solution in a plastic con­tainer and shake overnight at room temperature. Centrifuge and repeat the HCI/HF treatment, if necessary, until the ash content is below 1 %. Transfer the precipitate to a Visking dialysis tube by slurrying with water and dialyze against distilled water until the dialysis water gives a negative Cl- test with silver nitrate AgN03• Freeze dry the humic acid.

Pass the supernatant designated "FA Extract I" through a column of XAD­S (0.15 mL of resin per gram of initial sample dry weight at a flow rate of 15 bed volumes per h). Discard the effluent, rinse the XAD-S column containing sorbed fulvic acid with 0.65 column volumes of distilled H20. Back elute the XAD-S column with 1 column volume of 0.1 M NaOH, followed by 2 to 3 column vol­umes of distilled H20. Immediately acidify the solution with 6 M Hel to pH = 1.0. Add concentrated HF to a final concentration of 0.3 M HE The solution vol­ume should be sufficient to maintain the fulvic acid in solution.

Pass the supernatant designated "FA Extract 2" through a column of XAD-8 (1.0 mL of resin per gram of initial sample dry weight). Repeat the back elution and acidification as for "FA Extract I" above. Combine the final eluates from each of the fulvic acid extracts and pass this solution through XAD-8 resin in a glass column (column volume should be one-fifth of sample volume). Rinse with 0.65 column volumes of distilled H20. Back elute with 1 column volume of 0.1 M NaOH followed by two column volumes of distilled H20. Pass the eluate through W-saturated cation exchange resin [Bio-Rad AG-MP-5 (Bio-Rad, Rich­mond, CA) using three times the mole of Na ions in solution]. Freeze dry the elu­ate to recover the H+ -saturated fulvic acid.

Comments

XAD-S is a nonionic, macroporous (pore size 25 Ilm), methyl methacrylate ester resin (see "Fractionation of Humic Substances Adsorption"). Because it is sometimes difficult to obtain it may be necessary to use an alternative resin such

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as Polyclar, which is a cross-linked poly(vinylpyrrolidone) (PVP) (Watanabe & Kuwatsuka 1991; De Nobili et aI., 1990a) or other equivalent resin.

Extensive purification procedures of the resins are required before use. These methods and methods used to store the resin are detailed by Thurman & Malcolm (1981).

If it is not possible to purify the fulvic acid using resin treatments, exhaus­tive dialysis against distilled H20 is an alternative but less satisfactory method of purification. If there is a significant concentration of polyvalent cations such as Al3+ present, these may form insoluble metal-humate complexes as the solution is neutralized. Therefore, the dialysis should be carried out against dilute HCI ini­tially until the concentration of any polyvalent cations has been significantly reduced, before finally dialyzing against distilled H20. Technically, a fraction obtained in this way should be referred to as a fulvic fraction, rather than fulvic acid, as it is likely to contain significant amounts of unbound soil polysaccharide.

Sequential Extraction. By extracting the humic substances from the soil at only one pH value as in the above method, variations in the physiochemical properties of the humic substances which reflect the environment in which they exist in the soil, may not be apparent. Sequential extraction of humic substances at a number of different pH may be a more sensitive method for differentially extracting the humic substances and for determining relative distribution in the soil and their degree of interaction with soil colloidal particles. This can be done by changing the pH alone or changing the nature of the extractant anion (Posner, 1966; Skjemstad, 1992). The sequence outlined by Posner (1966), 0.1 M pyro­phosphate (PH = 7), cold 0.5 M NaOH followed by hot 0.5 M NaOH, was found to produce fractions which were distinctly different with respect to their molecu­lar size, functional group content and infrared spectral characteristics (Posner et aI., 1968; Swift & Posner, 1971; Cameron et aI., 1972a). The method outlined below is based on that of Posner (1966) and involves three different extractants at two pH values. This methodology could be refined and developed to vary and/or increase the number of extractants or pH values using similar procedures as outlined by Posner (1966).

Materials

1. Sodium dihydrogen pyrophosphate/tetrapotassium pyrophosphate (Na2H2P207~P207)' 1:1, 0.1 M, pH = 7.0.

2. Hydrochloric acid, 0.1 M, 5 M. 3. Sodium hydroxide, 0.5 M, 5 M. 4. Visking dialysis tubing.

Method

Prepare the soil by crushing and air drying before passing it through a 2.0-rom sieve. Pretreat the soil with 0.1 M HCI (with stirring) for two consecutive periods of 24 h before commencing the extraction process.

Combine this soil sample with the first extractant (pyrophosphate), in the soil to solution ratio of 1 g/5 mL. Allow the suspension to stand for 48 h at the desired temperature (20 or 60°C) under N2. Separate the soil and extractant (Frac-

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ORGANIC MATI'ER CHARACTERIZATION 1021

tion 1) by centrifugation at 1200 g for 30 min. Wash the soil residue with distilled water several times. Repeat the above extraction for each extractant (e.g., 0.5 M NaOH) changing the temperature as required.

The humic acid components of these fractions can be obtained as follows. Acidify the fraction to pH = 1 with HCI and allow it to stand for 24 h before cen­trifugation. Adjust the solution volume to 600 mL with distilled water and raise the solution to pH = 7 using a suitable alkali (e.g., 5 M NaOH). Centrifuge the solution at 6000 g at 2 to 5°C for 30 min. Repeat this precipitation and dissolu­tion procedure a further four times, increasing the centrifugation times by 30 min each time, to remove the clay and humins from the solution. Retain all the super­natant solutions for fulvic acid fractionation.

Finally, precipitate the humic acid at pH = 1, allow the solution to stand for 24 h, centrifuge and then exhaustively dialyze the precipitated humic acid against distilled water in Visking dialysis tubing until chloride free. Freeze dry the slur­ry to recover the humic acid.

Separate the fulvic acid fraction from the combined supernatants using XAD-8, PVP or dialysis tubing, as outlined in the previous method. Freeze dry the fulvic acid.

Extraction with Non-Aqueous Solvents

Non-aqueous solvents have been used by a number of workers to extract humic substances from soils. The most successful solvents tried so far being pyri­dine and dipolar aprotic solvents such as dimethylsulfoxide (DMSO, C2H60S) and dimethylformamide (DMF, C3H7NO) (Hayes, 1985; Hayes et aI., 1975; Pic­colo, 1988; Piccolo & Mirabella, 1987; Ma'shum et aI., 1988).

These dipolar aprotic solvents have been found to work most effectively for humic substances where the ionization of the humic molecules has been sup­pressed so that they will behave essentially as if they are uncharged molecules. This is achieved by first washing the sample in dilute acid to remove the metal cations and then maintaining the solution pH at moderately acidic levels during the extraction with the aprotic solvent. Under these conditions the DMSO is able to efficiently solvate the humic molecules, extracting humic substances with sim­ilar yields and molecular weight and cation-exchange capacity as those obtained when extracting in aqueous solutions at neutral pH. By using an amphiphilic sol­vent consisting of a mixture of ammonium (NUt)/isopropanol Ma'shum et al. (1988) were able to isolate the hydrophobic organic matter from a soil.

A method of supercritical gas extraction of organic matter using an organ­ic solvent has been applied by Schnitzer et al. (1986) to extract specific organic components from a soil matrix. The method has been further extended to sequen­tially extract organic matter from soils using supercritical fluid extraction, togeth­er with a range of organic solvents (Schnitzer, 1990).

Extraction and Purification of Soil Polysaccharides

As the interest in understanding the role of soil organic matter has increased the focus has frequently been on the humic substances because they account for a large percentage of the soil organic C. The soil polysaccharides, which are com-

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posed of a wide range of monosaccharides in both simple and complex molecu­lar structures, form a smaller but nonetheless important part of the soil organic matter.

Polysaccharides in the soil may occur as an original fragment of once liv­ing organic matter or may have been formed as a result of microbial activity as part of the decomposition process. Polysaccharides exist in association with inor­ganic or organic colloidal components, either through sorption (van der Waals forces, H-bonding) or through chemisorption (e.g., phenolic glycoside linkages). Polysaccharides have a number of effects on soil properties such as the cation exchange capacity (uronic acid group), C metabolism, biological activity and the complexing of metals. But more importantly, it has been found that polysaccha­rides are involved with stability of aggregates in soils.

The percentage of the total soil polysaccharide content involved in aggre­gation of soil particles is probably quite low as it requires the interaction of the polysaccharide with multiple soil components. This interaction will depend on the type of functional groups on the polysaccharide and the conformational struc­ture of the sugar groups, as well as the nature of the other soil components (Williams et aI., 1967). The correlation between total polysaccharide and degree of aggregation has been found to be poor (Oades, 1967). However, many of the polysaccharides are not active (e.g., partly decomposed cellulose) and so do not contribute to aggregation of the soil particles. More recent work (Haynes & Swift, 1990) has shown that there are particularly active fractions of polysaccha­ride involved in the aggregation process. It has been noted (Greenland et aI., 1962) that the aggregating effects of soil polysaccharides are more noticeable in soils with low organic matter content.

As indicated above, one of the main reasons for studying polysaccharides in soils is associated with their ability to bind and stabilize soil particles. Ironi­cally, the polysaccharides involved in this process are probably those most firm­ly held in the soil system and, therefore, the most difficult to remove without degrading them. Any extraction method should, ideally, be able to extract all of the polysaccharides from the soil matrix and, if this is not possible, then be able to extract a representative fraction of the polysaccharides. It is possible to esti­mate the efficiency of the extraction procedure by comparing the amount of prod­uct from the extraction with that which is obtained using hydrolysis procedures to determine the total carbohydrate content of the soil (Cheshire, 1979). Howev­er, such a comparison does not clearly indicate whether preferential extraction of some polysaccharides is occurring in the extraction procedure, and hence whether the extracted fraction is representative of that in the original soil sample.

Extraction of polysaccharides from soil utilizes similar methods to those applied to extraction of humic substances, and often is part of the procedure of their isolation. None of the methods used are ideal and to some extent the most suitable method will depend on the nature and origin of the soil. The most effi­cient methods for general application are those using 0.1 or 0.5 M sodium hydroxide. Large amounts of other fractions of organic matter are simultaneous­ly extracted from the soil using this method and the efficiency of separating the polysaccharides from the bulk of the extract (especially the humic acid fraction) is doubtful. It has been found that yields are significantly improved by pretreat-

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ORGANIC MATIER CHARACTERIZATION 1023

ment of the soil with dilute HCI or HF (Swincer et aI., 1968), or sulfuric acid (H2S04) (Barker et aI., 1967) to acidify the soil organic components. Methylation (Cheshire et al. 1983) of the soil as a pretreatment is another alternative although this method may be unsatisfactory as a preparative method for some analyses.

Other methods of extraction include using extractants such as hot water, which tends to have lower yields and is thought to be degradative, and dilute min­eral acids, which give lower yields but significantly reduce the amount of humic substance being extracted, so simplifying the purification of the polysaccharide. Organic reagents such as DMSO give a high percentage of polysaccharides in the product. The polysaccharide can be separated from the humic substances, by sorbing the latter on XAD-8 resin, but separating the polysaccharides from the DMSO is difficult. Complexing agents such as EDTA give low yields and hence a greater probability of unrepresentative fractions.

Resins such as XAD-8 or Polyclar AT [cross-linked poly(vinylpyrroli­done)] (Sanderson & Perera, 1966; Swincer et aI., 1968; Drijber & Lowe, 1990) may be used as a means of removing humic substances from the soil extract to obtain the polysaccharide fraction. With an appropriate eluant, the humic sub­stances are adsorbed on the resin and the polysaccharide is eluted through the col­umn. Purification of this polysaccharide fraction may then be achieved by dialy­sis to remove salts, solvents and other low molecular weight material. Adsorption of the coloured compounds onto charcoal is another method of purification, although it may be difficult to recover the polysaccharides from the charcoal.

Gel chromatography can be used to fractionate polysaccharides on the basis of molecular size. Ion exchange chromatography using diethylaminoethyl-cellu­lose {(C6H70)..{OHhx.a[OCH2CH2N(CH3CH2)]a} (DEAE-cellulose) can then be used to fractionate on the basis of charge density differences. In gel chromatog­raphy care must be taken to select a gel that isolates the polysaccharides as a group without losing a specific part of the fraction; such as the smaller molecules that bypass with the largest molecules in the case of using Sephadex G-lOO (Pharacia, Uppsala, Sweden). The gel chromatography method given below is based on that of Swincer et al. (1968).

Materials

1. Hydrochloric acid, 1 M. 2. Sodium hydroxide, 0.5 M. 3. Dowex 50 (W) (Dow Co., Midland, MI). 4. Polyclar AT resin. 5. Sephadex G-25.

Method

Pretreat the soil for 16 h with 1 M HCI at 20°C, then extract the soil twice for 16 h with 0.5 M NaOH at 20°C. Pass the extracts upwards through a column containing Dowex 50 (H+) to remove the humic acids and then concentrate the eluates in vacuo at 45°C. Pass the eluates through Polyclar AT to remove the residual coloured materials. Remove the salts and low molecular weight materi­al by separation in a column of Sephadex G-25.

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Comments

The cation exchange resin in the H+ form was used instead of precipitating the humic acid in acid solution, as it overcomes the problem of coprecipitation of the polysaccharides. New resins, such as XAD-4 (polystyrene divinylbenzene) can be used in place of Polyclar AT.

FRACTIONATION OF SOIL ORGANIC MATTER

Fractionation of Humic Substances

Because of the complexity of structure and interactions of soil humic sub­stances, the physical and chemical properties of these natural organic mixtures are difficult to define precisely. In order to simplify the study of humic sub­stances, a variety of techniques have been developed to fractionate samples into distinctive and hopefully less complex parts. Fractionating a sample of humic substances does not result in pure homogeneous compounds but rather fractions in which one or more of the physical or chemical properties has a narrower range of values than the original sample.

The particular method chosen for the fractionation process will depend on the chemical and/or physical characteristics being studied. Commonly used frac­tionation procedures are based on characteristics such as differences in solubili­ty, molecular size and electrostatic charge of the molecules within the system (Swift, 1985). Some of the techniques can be classed as preparative as they result in fraction samples of sufficient size to be used in further study (e.g., those based on solubility and separation according to molecular size). Other fractionation techniques, such as some of those based on charge characteristics (e.g., electro­focusing) are more appropriate for fingerprinting. Such techniques are able to characterize a sample for purposes of comparison with other samples chemicals, but do not easily produce sufficient sample for further study.

Fractionation Based on Solubility

The solubility of humic substances is not only dependent on the pH of the solution, but is also dependent on the type and concentration of the cations and other solutes present, and on the nature of the solvent system. Theoretically, all of these properties could be used to fractionate a humic substance sample.

Use of pH. Precipitation using changes in pH is the basis of the humic acidlfulvic acid fractionation (Fig. 35-1). The procedure can be refined by using smaller changes in pH to obtain more narrowly defined fractions (Flaig et aI., 1975). Unfortunately, it is difficult to obtain fractions with limited overlap due to significant coprecipitation which occurs in the experimental procedure. Howev­er, because of the variety of structural and functional groups within a sample of humic substance, and of the variety of interactions between these groups, frac­tionation of humic substances on this basis is unlikely to yield subfractions with substantially different chemical characteristics.

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ORGANIC MATI'ER CHARACTERIZATION

FRACTIONATION OF SOIL HUMIC SUBSTANCES

soluble in acid soluble in alkali

fractionation on the

basis of jOIUbility

insoluble in acid soluble in alkali

e.g. recognizable plant debris; plus polysaccharides, proteins. lignins, etc. in their natural or transformed states.

insoluble in acid insoluble in alkali

~ .... I[f----- Decreasing molecular weight ----­

..... f------ Decreasing carbon content ----­

...... f------ Increasing oxygen content ----­

.... I[f----- Increasing acidity and CEC -----

I( Decreasing nitrogen content

..... !----- Decreasing resemblance to lignin -----

1025

Fig. 35-1. Diagram showing the categorisation of soil organic matter into humic and nonhumic sub­stances, the fractionation of the humic substances and the property variations within these fractions.

A variation in using the range in solubility as a means of obtaining fractions is to sequentially extract a soil sample using a range of solutions of increasing pH, and ability to solubilize the humic substances present in the sample. This type of extraction is discussed in "Sequential Extraction."

Salting Out. Increasing the salt concentration of a humic substance solu­tion decreases the intramolecular charge repulsion and causes the polyelectrolyte macromolecules to shrink and to exclude the solvent. Simultaneously, increasing the salt concentration causes a decrease in the extension of the diffuse double layer of charge, thereby decreasing intermolecular charge repulsion allowing molecules to approach each other more closely, and hence to coagulate. It is pos­sible to fractionate humic substances based on this behaviour (Theng et aI., 1968) as outlined below.

Materials

1. Sodium hydroxide, 1.0 M. 2. Ammonium sulfate [(NH4)2S04], solid. 3. Sulfuric acid (H2S04), 1.0 M.

Method

Prepare 150 mL of 0.33% K+ -humate solution. Add 1.0 g of solid (NH4)2S04, readjust the pH to seven using 1 M NaOH and shake for 5 to 6 h. Sep­arate the precipitate by centrifugation and shake the solution overnight without

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adding more salt. Centrifuge the suspension and combine the precipitates and retain.

Add successive increments of 1.0 g of (NH4)zS04 and repeat the procedure until the amount of precipitate obtained after centrifugation is negligible. For each fraction obtained from the salting out procedure, dissolve the precipitate in distilled water or dilute alkali, adjust the pH to one using 1 M H2S04 and then remove the excess salt by exhaustive dialysis. Freeze dry the product and weigh.

Commen~ther Precipitation Methods

The interaction between heavy metal ions and humic substances in solution can affect the solubility of the humic substances. Using the insolubility of the metal-humic substances complex is one way of isolating that fraction of humic substance which interacts with the metal concerned. MacCarthy and O'Cinneide (1974a) used this method to study complexing of humic substances with both Cu and Co under both acidic and alkaline conditions. Metal-humate interactions are of interest agriculturally as they may influence the availability of trace metals in soiVplant systems. The ability of humic substances to interact with heavy metals is also of great interest environmentally with respect to the transport of heavy metals through soiVwater systems and, hence, the fate of toxic metals in soils and ground waters.

Fractionation on the basis of metal-humate solubility, and that based on salting out, are not used regularly as they are rather tedious and the fractions are ill-dermed due to problems of coprecipitation.

Use of Organic Solvents. Humic substances have relatively low solubility in many conventional organic solvents which can be used as a method of frac­tionation (Hayes, 1985). Historically, ethanol was used to extract the hymatome­lanic acid fraction from precipitated humic acid (Stevenson, 1994), but it can also be used to fractionally precipitate alkaline solutions of humic acid (Kyuma, 1964; Kumada & Kawamura, 1968). The method below is based on that of Kumada and Kawamura (1968). Other water-miscible organic solvents, such as acetone [(CH3)zCO) and methanol (CH30H), may be used in a similar way to obtain frac­tions of humic substances.

Materials

1. Sodium hydroxide, 0.1 M, 0.2M. 2. Ethanol (C2HsOH), absolute. 3. Hydrochloric acid, 1:3. 4. Sulfuric acid, concentrated.

Method

Dissolve 150 mg of humic acid in 20 mL of 0.2 M NaOH. Add absolute ethanol (~HsOH) to give a concentration of 10% (v/v), allow to stand overnight and then centrifuge the suspension at 7000 rpm for 20 min. Dissolve the precipi­tate in 0.2 M NaOH, add ethanol to the same concentration as above and recover the precipitate by centrifugation.

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ORGANIC MAITER CHARACTERIZATION 1027

Add ethanol to the combined supernatant to give an ethanol concentration of 20% and repeat the above procedure. Continue the process by increasing the alcohol concentration in steps of 10% until the fmal concentration is 80% ethanol and eight fractionated precipitates have been obtained.

Dissolve each precipitate in 0.1 M NaOH, precipitate by adding HCI (1:3) and then wash the precipitate with ether (C4HlOO) over a glass-filter. Dry the pre­cipitates in a desiccator containing concentrated H2S04 for 2 d and weigh the products.

Reduce the concentration of alcohol in the 80% ethanol soluble humic frac­tion by distillation in a water bath, precipitate, wash and dry this fraction as described above for the other fractions.

Comments-Use of Other Organic Solvents

Generally, extracting fractions using water-immiscible solvents gives very low yields. However, Rice and MacCarthy (1989) have achieved some success using a different approach with the polar water-immiscible solvent, methyl isobutyl ketone [CH3COCH2CH(CH3)2] (MIBK). They initially separated the aqueous fulvic acid fraction from the nonaqueous huminlhumic acid fraction at low pH using HCI to acidify the solution. Following this the pH of the nonaque­ous phase was made alkaline using NaOH allowing the separation of the humic acid in the aqueous phase from the humin associated with the MIBK nonaqueous phase. Rice and MacCarthy (1989) were then able to fractionate the humin resi­due into an acid soluble form and aqueous and nonaqueous phase products. The study of the humin fraction of the soil is a neglected but important aspect of soil research, and this technique provides an opportunity to study the humin fraction.

Adsorption. The desorption behaviour of humic substances sorbed to a gel (Swincer et al., 1968), resin (Yonebayashi & Hattori, 1990), charcoal (Forsyth, 1947) or alumina (Dragunov & Murzakov, 1970), can be used as a means of frac­tionation. The purification of the fulvic acid fraction using resins such as XAD-8 or PVP during the extraction of humic substances ("Extraction with Aqueous Solutions") is essentially an adsorption/desorption fractionation procedure.

Materials

1. Sodium hydroxide, 0.1 M, 10 M. 2. Ethanol, 50% (v/v). 3. Universal buffer, 0.02 M. 4. Sulfuric acid,S M. 5. XAD-8 resin. 6. Amberlite IR-120 resin (Rohm & Haas Co., Philadelphia, PA).

Method

Dissolve the humic acid sample in 0.1 M NaOH and treat with Amberlite IR-120 resin to convert it to the H+ -exchanged form. Dissolve 5 mg of the H+ -sat­urated humic acid in 2 mL of aqueous solution and load onto a column packed with XAD-8 resin. Elute the humic acid in a stepwise fashion with universal buffer solutions adjusted to pH = 7 and pH = 11, distilled water and 50% ethanol.

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Determine the elution profile by first adjusting the pH of the effluent to pH = 12 using 10 M NaOH and then measuring the optical density at 400 nm. Precipitate the humic acid contained in each effluent using sulfuric acid and then dissolve it in 0.1 M NaOH. If the humic acid fraction does not precipitate on acidification, absorb it onto a small column of XAD-8 at pH = 3 and elute with NaOH solution. Dialyze each of the eluates against distilled water and freeze dry.

Comments It is possible to refine this method by using a greater range of buffers to sep­

arate the humic acid into a larger number of fractions. Yonebayashi and Hattori (1990) extended the method by using both a pH gradient and a water-ethanol gra­dient to allow for the ability of obtaining a greater number of fractions if required. MacCarthy et al. (1979) used a similar method to fractionate a sample of humic substance. Instead of using buffers a pH gradient solution was generated by elut­ing simultaneously from two pumps, one containing 0.1 M H3P04 and the other 0.1 M NaOH. MacCarthy et al. (1979) separated out only two fractions, but it would be possible to adapt the method to obtain more fractions, depending on the characteristics of the humic substance and the nature of the pH gradient used.

Fractionation Based on Molecular Size

The extreme range in molecular weights associated with humic substances extracted from soils should theoretically allow the separation of the sample into many specific fractions. In reality the complex intermolecular associations make it difficult to obtain fractions with insignificant overlap in molecular content. Nevertheless, fractionation of humic substances on the basis of molecular weight is a powerful and attractive procedure. Gel permeation chromatography and ultra­filtration are techniques which have been used successfully with proteins and car­bohydrates for fractionations based on molecular size. Reasonable success has been achieved in using these rapid and reliable techniques to purify and fraction­ate humic substances.

Gel Permeation Chromatography. The gels used for molecular size chromatography have structures consisting of a system of pores, the sizes of which are determined by the degree of cross-linking in the polymer. When a solu­tion of humic substance is applied to the top of a gel column and eluted with sol­vent, large molecules which are unable to enter the pores in the beads will bypass the beads and will be eluted first from the column. Smaller molecules which are able to enter the pores will have their passage through the column retarded, the extent of which will depend on the actual size and shape of the molecule. The net result is that the solute molecules are eluted from the column in order of decreas­ing molecular size, and, for a given family of macromolecules, in order of decreasing molecular weight also.

The range of molecular sizes over which the gel is able to differentiate the molecules will depend on the type of gel chosen, but gels are available which are able to produce fractions in the range of several thousands to millions of daltons. Gels are selected on the basis of the exclusion limit of the gel and the molecular weight range over which the gel is suitable. Given the extreme polydispersity of humic substances, it is generally necessary to use a range of gels to achieve sat-

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ORGANIC MATTER CHARACTERIZATION 1029

isfactory fractionation. For discussion on the calibration of gels, see "Gel Chro­matography."

It is important to select gels that are inert to the solute molecules so that there are no chemical or physical interactions between gel and solute. Otherwise the resulting separation cannot be entirely attributed to molecular size and/or weight differences. Swift and Posner (1971) discussed these problems fully and show that they can be largely overcome by careful selection of the gel matrix and by the use of appropriate buffer solutions. By successively using gels of various exclusion limits and reapplying the excluded or included portion of a given gel to another with a higher or lower exclusion limit it is possible to successfully sepa­rate out fractions (Schnitzer & Skinner 1968, p. 41-SS). The method outlined below is that used by Swift et ai. (1992).

Materials

1. Sodium tetraborate, (Na2B407) 1 % w/v, pH = 8.S. 2. Sephadex G-7S, G-1S0. 3. Sephrose 6B (Pharmacia, Uppsala, Sweden).

Method

Dissolve the sample of humic acid in a borate buffer (1 % w/v, pH = 8.S) and run SO mL through Sephadex G-1S0 in a preparative (S-cm diam.) column to fractionate the sample with respect to molecular weight. Collect the eluate as 10-mL fractions and read the absorbance of these at 400 nm.

Repeat the above procedure four times and divide the eluate into four parts corresponding to four regions of the elution curve, namely the excluded peak (A) and three segments of the broad peak of lower molecular weights (E, C, D) (Fig. 3S-2). Dialyze and freeze dry each of the four combined eluates.

Redissolve the four fractions using borate buffer and then fractionate them using Sepharose 6B, Sephadex G-1S0 or G-7S gels for fractions high to low mol­ecular weight, respectively.

Refine the fractions by taking central cuts of the resulting four individual peaks (shaded section, Fig. 3S-2) and dialyze these against distilled water exhaustively before freeze drying.

Comments The volume of the excluded fraction (void volume) can be determined by

using the nonadsorbing chemical, Blue Dextran 2000 (Pharmacia, Uppsala, Swe­den). To determine the total effective column volume of Sephadex gels use N-2,4-dinitrophenyl aspartic acid (a yellow compound), sucrose (C12H220 11), or glucose (C6H120 6) as the low molecular weight marker. Except for gels to which it is reversibly adsorbed (e.g., Sephadex gels) the sodium salt of 2,6-dichlorophenol indo-phenol (a blue dye) can also be used to determine the total effective volume of the column.

Typical elution patterns obtained by proper application of gel permeation chromatography (e.g., Dubach et aI., 1964; Swift & Posner, 1971) should be con­sulted. The elution pattern shown in Fig. 3S-2, using borate or tris buffer is an example of the type of curve that should be obtained.

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Primary fractionation (Sephadex G-150)

A B c

:, SecondarY,!ractionations".

, , , Sepharose 66 : Sephadex G-150: Sephadex G-150 : Sephadex G-75

Vt Vo Vt Va

Elution volume

Fig. 35-2. Composite diagram showing initial elution pattern (absorbance at 400 nm vs. volume elut­ed) for whole humic acid (upper section of diagram) and patterns for the subsequent runs of the sep­arate fractions, FI, F2, F3 and F4 (lower section of diagram). The shaded areas of the secondary elution patterns represent the "central cuts" taken to obtain the final secondary fractions.

Ultrafiltration. In recent years a range of polymer-based membrane filters have been developed with pore sizes ranging from several micrometers to a few nanometers_ The nominal molecular weight cut-off values of these range from 700 to 1 000 000 daltons and, theoretically, these membrane filters should be able to be used to fractionate the polydisperse humic substances according to molec­ular weight. However, it has been observed that charge-charge interactions between the solute and the membrane surface may interfere with the filtration process and humic acids are highly charged. This fact together with the difficul­ties encountered during the manufacture of the filters and the unresolved doubts about the relationship between molecular size and molecular weight for humic substances, create some problems concerning the overall usefulness of the results when studying humic substances.

Ultrafiltration is commonly used as a purification procedure in which the membrane is used to separate the wanted fraction from the unwanted fraction. It is also used in conjunction with other methods of fractionation, such as gel per­meation and ultracentrifugation to increase the flexibility of the fractionation pro­cedure (Cameron et aI., 1972a). The method below has been used to study the complexing capacity of humic substances in natural waters (Smith, 1976). It is possible to alter the method by changing the membrane types to alter the molec­ular size range of the fractions, and by incorporating a greater number of mem­branes to increase the number of fractions. The membrane selection will have to

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ORGANIC MATIER CHARACTERIZATION 1031

include some with higher molecular weight cut -off values for use with soil humic substances. It is advisable to begin the ultrafiltration using the membrane of high­est cut-off value and work downwards in size in order to limit clogging of the filters.

Materials

1. Ultrafiltration membranes, XM-l00 (Amicon, Beverly, MA), XM-50 (Amicon, Beverly, MA), PM-lO (Amicon, Beverly, MA), YM-2 (Ami­con, Beverly, MA), with molecular weight cut-offs 100 x 103, 50 X 103,

10 X 103, 1 X 103 daltons respectively. 2. Ultrafiltration cell.

Method

Flush the ultrafiltration cell and membrane with double distilled water until the absorbance of the filtrate solution records zero magnitude [using ultraviolet (UV)-visible spectroanalysis]. Introduce a sample of known volume (e.g., 400 mL) into the cell and pressurize the system with N2. Collect the ultrafiltrate and retain in a preweighed flask. When the sample has been concentrated to approx­imately 20 mL, depressurize the system and pour the concentrate into a pre­weighed flask. Determine the volume of this fraction by gravimetry assuming the specific gravity of 1.0.

Sequentially fractionate the ultrafiltrate from the above step using mem­branes with the next lower molecular weight cut-off until that having the small­est weight cut-off has been used. Determine the volume of the ultrafiltrate.

The above fractions could be further refined by repeating the ultrafiltration using the membrane associated with their retention. Dialyze the fractions against distilled water and freeze dry. Compute the mass per volume of the initial ultra­filtrate to determine the relative fraction of the original mass in the sample.

Fractionation Based on Charge Characteristics

The charge characteristics of humic substances in solution, originate pre­dominately from the presence of carboxylic acid, phenolic and enolic functional groups. The amount of charge relates to the degree of dissociation of these func­tional groups which is a function of the pH of the solution and the identity of the counterions in the surrounding medium. By taking advantage of the differences in charge density within a sample of humic substances it is possible to fraction­ate them according to their charge characteristics.

Electrophoresis. Electrophoresis is the movement of charged species in a solution in response to an applied electrical potential. Traditionally, elec­trophoresis was carried out in free solution so that movement of the species was able to be related to both charge and size of a molecule or ion. By carrying out the process in a gel medium the movement of the charged species can be retard­ed according to their size and the size of the pores in the gel. In this way the frac­tionation of humic substances can be related to both the charge and molecular size of the species. Experimental conditions can be modified by altering the buffer or the characteristics of the gel, to optimize the fractionation obtained.

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Gel electrophoresis can be used as both a preparative fractionation tech­nique, and as a means of characterizing humic substances. A preparative elec­trophoretic method (De Nobili et al., 1990a) which produces fractionated humic material able to be used in other analyses has been outlined below.

Materials

1. Potassium dihydrogen phosphate buffer (KH2P04) 0.02 M pH = 7.0. 2. Glycerol (C3Hg03). 3. Polyacrylamide gel. 4. Sodium pyrophosphate, 0.1 M. 5. Poly(vinylpyrrolidone).

Method

Prepare the gel rods by casting three different overlayed gels of decreasing acrylamide concentration (9, 7, and 5%). Prepare a sample containing 15 mglmL organic C, mix it with glycerol, and then filter the solution through a 0.2-llm membrane filter before application to avoid blocking the pores at the entrance to the gel. Perform the electrophoresis in 0.02 M phosphate buffer (PH = 7.0) using 15 rnA per rod (care must be taken to ensure that the gel in the rods does not become overheated.

Stop the run when the front of the migrating band reaches the bottom of the 9% gel segment. Immediately after the run take 2-cm long slices of the front frac­tion, the center fraction and the tail fraction of the migrating band of humic sub­stances. Discard the intermediate sections. If doing the fractionation in duplicate, mince the corresponding slices and extract them three times with 0.1 MNa4PZ07. Concentrate the fractions on PVP columns. These fractions are suitable to be used in the following electrofocusing analysis or can be dialyzed and freeze dried and prepared for some other analyses.

Comments

The equipment used by De Nobili et al. (1990a) consisted of 13 x 115 mm polyacrylamide gel rods and a Bio-Rad 155 Electrophoresis cell, powered with an LKB 2117 power supply (LKB, Bromma, Sweden) and cooled with water cir­culating at 4°C. Although presented as a preparative method, it should be noted that the amounts of material obtained by this procedures are very small.

The technique of electrophoresis has been refined by incorporating a pH gradient into the gel system. The charged species migrate until they reach the position corresponding to their isoelectric point, a pH at which they cease to be charged, in a process called electrofocusing. In this method, the gel has relative­ly large pores so that molecular size is not involved in the fractionation.

In electrofocusingt as described by De Nobili (1988), a pH gradient is set up by the use of a mixture of compounds which are good buffers and able to act as ampholytes in the separation medium. The range in pI (isoelectric point) val­ues of the ampholytes in the gel must be between the pH of the electrolytes in the anode and cathode compartments. The most acidic ampholyte (lowest pI) moves to occupy a position closest to the anode and correspondingly the most basic am­pholyte occupies the space closest to the cathode (highest pI), with intermediate

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ORGANIC MATTER CHARACTERIZATION 1033

ampholytes moving to occupy spaces arranged in order of their decreasing acid­ity in between these two.

Electrofocusing is a potentially useful technique for characterizing humic substances. However, a number of operational aspects indicate that any conclu­sions formed from using this technique should only be tentative (Duxbury, 1989). Duxbury believes that the process is not electrofocusing but is electrophoresis in a pH gradient. Some of the reasons for this are that humic molecules are not gen­erally amphoteric and once near their isoelectric point tend to lose their charge and may not even remain in solution. Those that diffuse past their pI are no longer charged and so are not focused back to that point, i.e., true electrofocusing does not occur.

Other problems which potentially may be associated with the process are the interactions of humic molecules with the ampholines and the aggregation ten­dencies of humic substances (Duxbury, 1989).

Fractionation of Polysaccharides

The polysaccharide fraction of soil organic matter will be made up of a mixture of components of varying molecular weight, charge density and mono­saccharide composition. Because of the similarity in some of the physical and chemical characteristics between polysaccharides and humic substances many of the inethods used for fractionation of humic substances can be applied to the frac­tionation of polysaccharides, such as fractional precipitation (Parsons & Tinsley, 1961), electrophoresis (Mortensen & Schwendinger, 1963; Barker et aI., 1965), density fractionation (Strickland & Sollins, 1987), ion-exchange chromatography (Finch et aI., 1966; Thomas et aI., 1967) and gel filtration (Barker et aI., 1965, 1967; Swincer et aI., 1968). The last two methods are those most commonly used.

lon-Exchange Chromatography

The method below is based on that of Barker et ai. (1967). Initially, a pH gradient elution is carried out to determine the elution characteristics of the par­ticular polysaccharide sample. From this information, it should be possible to dif­ferentiate logical fractions within the sample and the salt concentration associat­ed with each of these fractions. The elution is then repeated using a fresh sample of polysaccharide, and solutions of specific salt concentrations in a buffer to sep­arate the fractions. In this way the sample is fractionated on the basis of increas­ing charge density.

Materials

1. Sodium chloride, 2 M 2. Phosphate buffer (0.0371 M KH2P04, 0.0043 M Na2HP04), pH = 6 3. DEAE-cellulose

Method

Prepare a column (3.4 by 45 cm) of DEAE-cellulose and carry out a gradi­ent elution with 0 to 2 M NaCl in phosphate buffer. Monitor the elution profile by collecting lO-mL fractions and analyzing these for sugars (Dubois et aI., 1956),

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and absorbance at 400 nm. Determine the number of fractions to be eluted and the NaCI concentration at which their elution is maximized. For each fraction selected, prepare 600-mL portions of buffer containing the maximum NaCl con­centration associated with that fraction. This preliminary run can be carried out using a smaller amount of polysaccharide than that used in the main preparative experiment that follows.

Equilibrate the column with the phosphate buffer. Dissolve 200 mg of poly­saccharide in 50 mL of phosphate buffer, apply it to the column and elute it suc­cessively with the 600-mL portions of buffer prepared as above, using increasing concentrations of NaCl. Collect lO-mL fractions of the eluate and analyze as above. Combine the polysaccharide-containing fractions for each concentration of the salt, dialyze against distilled water and freeze dry.

Gel Filtration

This method is similar to the previous method, except that the separation is on the basis of size, rather than charge density.

Materials

1. Sodium chloride (NaCl), 1 M. 2. Sephadex G series gels. 3. Biogel P series gels.

Method

Pretreat the gel and equilibrate the column using 1 M NaCI to eliminate ion­ic and adsorption effects. Load the column (54 by 3.1 cm) with 3 mL of solution containing 200 to 300 Ilg polysaccharide/mL. Elute the column with 1 M NaCl, collect 5-mL fractions and analyze these using absorption measurements. Deter­mine the amount of polysaccharide (Dubois et aI., 1956) as a function of the elution volume and separate and combine fractions as outlined in "Gel Perme­ation Chromatography." Suitable gels for polysaccharide analysis are the Sepha­dex G series (Pharmacia, Uppsala, Sweden) and Biogel P series (Calbiochem, Los Angeles).

Density Fractionation of Soil Organic Matter

When soil organic matter becomes intimately associated with soil mineral particles, the density of the resulting combination is greater than that of the organ­ic matter alone. By fractionating the soil on the basis of the density of the com­ponent particles it is possible to separate the organic matter into fractions of dif­ferent physical characteristics and almost certainly different chemical behavior, based on their degree of decomposition and association with mineral particles. Density fractionation avoids the need for solvent extraction and decreases the possibility of artifact formation. It also is believed that this type of fractionation may partially separate the soil organic matter according to age, with the youngest fraction being the lightest.

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ORGANIC MA'ITER CHARACTERIZATION 1035

A number of methods have been developed to achieve this type of frac­tionation. These involve the separation of the soil particles on the basis of their density in water (Turchenek & Oades 1979), and in organic liquids or salt solu­tions of high specific gravity (Greenland & Ford 1964; Sollins et aI., 1984; Turchenek & Oades, 1979; Dalal & Mayer, 1986, Roth et aI., 1992; Baldock et aI., 1990). By combining particle size fractionation (e.g., sieving, sedimentation, and continuous flow centrifugation) with density fractionation, it is possible to separate the organic matter in the soil into multiple fractions (Turchenek & Oades, 1979; Oades et aI., 1987; Ducaroir et aI., 1990).

Initially, the sample is ultrasonified to break up all the aggregates and to thoroughly disperse all the colloidal particles. The sample is then sieved, and/or separated on the basis of settling time in water, before fractionating these resul­tant fractions on the basis of their density in a high specific gravity solution. The method below is based on that of Baldock et ai. (1990).

Materials

1. Poly tungstate solution (Na3 W04.9W03.H20).

Method

Add approximately 20 g of soil to 50 mL of deionized water in a 150-mL beaker and sonicate at constant temperature and at a medium output for 5 min. Pass the dispersed sample through a 53-J..lm sieve and then separate the sieved fraction into >2-J..lm and ~2-J..lm fractions using gravitational separation in deion­ized water. Dialyze the ~2-J..lm fraction against deionized water and freeze dry.

Combine the >53-J..lm and the >2-J..lm fractions and separate these into a light and heavy fraction by centrifugation in a sodium poly tungstate solution of density 2.0 g cm-3. Separate the light fraction from the supernatant by filtration using a screen with 5-llm mesh.

Resuspend the sediment in the poly tungstate solution, centrifuge and remove and filter the supernatant as before. Repeat this process until the super­natant is clear, then wash the combined light fractions and dry at 45°C. Wash the heavy fraction five times with deionized water and then dry it at 45°C.

This method gives three fractions; clay fraction (~2-J..lm diam. particles), light fraction (>5-J..lm diam. particles with a density of ~2.0 g cm-3) and heavy fraction (>2-J..lm diam. particles with a density >2.0 g cm-3). The method could be extended to achieve a greater number of fractions by using different density solutions (e.g., 1.6 g cm-3).

Comment

The earlier methods involving the use of dense organic liquids (Greenland & Ford, 1964) are not generally used nowadays, due to the health, safety and environmental problems associated with the use of halogenated hydrocarbons. Freeze drying, which is unlikely to alter the composition of the fractions, may be preferable to heat drying.

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CHARACTERIZATION OF HUMIC SUBSTANCES

Many of the chemical, physical, and spectroscopic methods already suc­cessfully used in general organic chemistry, and in the study of naturally occur­ring macromolecules have been applied to the study of soil humic substances to try to determine the composition and general structure of the component macro­molecules. However, because of the heterogeneous and polydisperse nature of humic substances, and the complexity of the inter- and intramolecular reactions, it is often difficult to interpret the results of these studies.

Because of the diverse nature of humic substances and the lack of any obvi­ous means of referencing data, it is difficult to quantify the characteristics of these samples. A set of standard and reference samples have been prepared by the IHSS to assist in the comparison of humic substances within and between labo­ratories, and these are available for purchase from IHSS.

Characterization by Chemical Methods

The three general characteristics of a chemical compound are the elemen­tal composition, the arrangement of these elements in the chemical structure, and the types and locations of the functional groups in the structure.

Procedures to determine the type and relative abundance of the elements within an organic compound are well defined and measurable experimentally. Such procedures are not dealt with in this chapter. Because of their relatively greater activity, it is usually far easier to determine the presence of the various functional groups in large organic compounds than it is to determine the arrange­ments of the elements within the structures. However, once the elemental com­position and the functional group characteristics are determined, and an estima­tion of the molecular weight has been made (see "Molecular Size and Shape"), it should be possible to make an assessment of the type of structures of which humic substances are composed.

Using general methods of organic analysis, it has been established that the major functional groups in humic substances are the carboxylic acids (and other groups containing the C=O functionality), phenols and alcohols, with Nand S located in the minor functional groups. To some extent, the accuracy of these analyses are open to question, but so long as they are reproducible they allow comparisons to be made of humic substances from different sources.

Acidic Functional Group Concentrations

The acidic behaviour of humic substances is such that they are considered to be a mixture of stronger (mostly carboxylic acids) and weaker organic acids (mostly phenolic acids). The pKa of these two types of acidic groups are around pH = 4.5 and pH = 10 respectively (Christensen et aI., 1976; Martell & Smith, 1977). Identical functional groups in a humic substance may not have identical pKa values (Perdue, 1985). The actual pKa of an acid is not only dependent on the nature of the acidic functional group, but also is dependent on the chemical envi­ronment in which it exists, as well as the nature of the compound to which it is bound. There would be considerable overlap in the range of pKa values of the

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ORGANIC MATIER CHARACTERIZATION 1037

acidic groups in a solution of humic substances, and the overall range of all of these values could span the pH range from 0 to 13 (Perdue, 1985). On the basis of pKa measurements, it would not be possible to distinguish unequivocally car­boxylic acids from other types of acids in humic substances (Perdue, 1985).

A number of different types of methods for the analysis of acids have been used to determine the acidic functional groups of humic acids (Stevensen, 1994; Perdue, 1978; Schnitzer & Khan, 1972). These include direct and indirect titra­tions, as well as nonaqueous and thermometric titrations.

Total Acidity. The total acidity of a sample should include all the acidic hydrogens present. Thus determination of total acidity would require the use of a basic reagent at high pH so that even the weakest acids present would be account­ed for. This type of method has several sources of error. First, the volume of base required to reach equilibrium at high pH values is relatively large compared with the amount required for reaction with the humic acid. Second, it is necessary to carry out the titration rapidly to limit the possibility of errors arising from any base-catalysed side reactions (Perdue, 1985). The results of the analysis are also dependent on the conditions in solution (e.g., properties of the pH electrode, com­position of the background electrolyte, etc.). Because of the foregoing consider­ations, the determination of the total acidity of a humic substance is a good ap­proximation only.

Direct TItrations. There is a tendency for the pH of the solution to decrease with time under alkaline conditions, and this effect can be considerably reduced by carrying out the titration in the absence of O2, This drift in pH is thought to be due to the production of acids from side reactions associated with the presence of the alkali (Swift & Posner, 1972). The titration can be carried either in the for­ward direction (i.e., against a dilute base) or in the reverse direction (i.e., against a dilute acid). As humic substances tend to be insoluble in acid solution, it may be difficult to achieve dissolution of the solid at moderately acidic pH values. For this reason the reverse titration, which involves dissolving a known amount \ ,f humic acid in KOH and titrating with dilute acid, is more rapid but the error asso­ciated with titrating in the high pH region (see above) is carried through the whole titration.

The method below is for the forward titration (Swift & Posner, 1972) but can easily be adapted to the reverse titration.

Materials

1. W -Dowex 50W resin. 2. Potassium chloride (KCI), 0.01 M. 3. Potassium hydroxide, 0.1 M.

Method

Prepare a solution of humic substance for titration by treating it with an excess of H+ -Dowex 50W resin. Prepare a solution containing 5 mg of the humic substances in 20 mL of 0.01 M KCI and titrate it against standardized 0.1 M KOH to pH = 7.0 (arbitrary end-point). Add the KOH stepwise (e.g., 50 steps) as rapid­ly as possible but still allowing for the solution to reach equilibrium before each

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pH reading. Repeat the procedure using a blank solution. Construct a plot of pH vs. titre volume for each solution to determine the acidity of the humic acid as a function of pH. All solutions should be CO2 free and the titration should be car­ried out under N2 at constant temperature.

Total acidity (mmolc g-l) = (V sample - VblanJ X MbaselWsample(g)

where V blank and Vsample are the titre volumes (mL) associated with the blank and sample titrations, respectively, and Wsample (g) is the mass of the humic substance used in the experiment. (Note, mmolc designated millimole of charge.)

Comment

During the titration, the solution may not reach equilibrium in a suitably short period of time. In this event, choose a set period of time between successive additions of alkali or a set time over which the pH is relatively constant, as the time at which the pH reading is taken.

Indirect titrations. The humic substance is allowed to equilibrate in an excess of Ba(OH)2' before the unused alkali is titrated with a standard acid. The total acidity of the humic substance is determined by calculating the difference in titre volume with respect to pH between an analysis including the humic sub­stance and a blank analysis under the same conditions. This method has been reg­ularly used as a means of determining the total acidity of a humic substance and as it involves a solution of pH > 13, it is considered that the result would be a rea­sonable estimation of the total acidity of the sample.

Traditionally, the method used has not specified the type of filter paper to remove the barium salts, and it has been found (Davis, 1982) that their occurrence in the titration solution would give an underestimation of the total acidity value.

The method below is based on that of Schnitzer and Khan (1972) with the modifications as suggested by Perdue (1985). Materials

1. Barium hydroxide (Ba(OH)2), 0.1 M. 2. Hydrochloric acid, 0.5 M.

Method

Place between 50 to 100 mg of humic substance in the W-form in a 125-mL ground-glass stoppered Erlenmeyer flask. Add 20 mL of 0.1 M Ba(OH)2 to the flask containing the humic substance and the same amount to a similar flask in which there is no humic substance, to act as a blank. Displace the air in each of the flasks with N2, stopper, and then shake for 24 h at room temperature. Fil­ter the solutions through 0.45-Jlm membrane, then wash the residues thoroughly with CO2-free distilled water. Using standardized 0.5 M HCI and with the aid of a pH meter, titrate the filtrate plus washings of both the sample and blank solu­tions to pH = 8.4. Determine the total acidity as described above for direct titra­tions, using the following equation.

Total acidity (mmolc g-l) = (Vblank - Vsample) X MaciJWsample(g)

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ORGANIC MATTER CHARACTERIZATION 1039

Stronger Organic Acids (Carboxyl Groups). As there are no distinctive equivalence points in the titration curves of humic substances, the determination of the pH at which a specified group of acids is neutralized is somewhat arbitrary. Because weak acids by definition must have a pKa above pH = 7.0 this value has sometimes been used as means of differentiating the acidity associated with weak and strong organic acids (Gamble, 1972; Burch et aI., 1978). Another more defin­itive method of delineation is to construct a first derivative curve of titre volume vs. pH to determine the maximum change in pH with change in titre volume. The volume below this point is then taken to be that associated with carboxyl groups and that above due to the phenolic groups and other weak acids.

Direct Titrations. The method detailed below is that of Oliver et al. (1983) and is carried out on the microscale. It is possible to scale all the apparatus up­ward in order to carry out the procedure using normal-sized measures and glass­ware. The results of this analysis give an operational estimation of the acidity associated with strong organic acids and hence carboxylic acids (Perdue, 1985). As for total acidity titrations, this method can be adapted to be carried out in the reverse direction.

Materials

1. Sodium hydroxide, 0.1 M.

Method

Suspend 10 mg of humic acid (H± form) in 2 mL of distilled water and titrate the solution to pH = 7.0 with 0.1 M NaOH in 50-ilL increments. Carry out a similar titration in the absence of humic substances.

COOH acidity (mmolc g-l) - (Vsample - Vblank) X MbasJWsample(g)

Indirect Titrations. The most common method used to determine the acidi­ty associated with the stronger organic acids is the calcium acetate method in which the amount of acetic acid generated from the reaction of calcium acetate with humic substance is determined by titration with standard sodium hydroxide solution.

There are a number of operational problems associated with this method. Because of poor buffering in the solution, the equilibrium pH of the solution is determined by the amount of humic acid added. Complexation of Ca2+ with the humic acid increases the release of protons to solution increasing the apparent acidity (Perdue et aI., 1980). As in the barium hydroxide method above, it is also important that the filtration step removes all solids from solution before titration. This separation can be achieved by using fine grade filtration (Davis, 1982) or ultrafiltration (Perdue et aI., 1980). The method below is based on that of Wright and Schnitzer (1959) with the alteration to filtration as suggested by Perdue (1985).

Materials

1. Calcium acetate [Ca(OAc)z], 0.5 M. 2. Sodium hydroxide, O. 1 M.

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Method

Place between 50 to 100 mg of humic substance in a 125-mL ground-glass stoppered Erlenmeyer flask. Add 10 mL of 0.5 M Ca(OAc)2 and 40 mL of COz­free distilled water to the flask containing the humic substance and to a similar flask in which there is no humic substance, to act as a blank. Shake the flasks for 24 h at room temperature, filter the solutions through 0.45-l1m membrane, and then wash the residues thoroughly with CO2-free distilled water. With the aid of a pH meter, titrate the filtrate plus washings of the sample and blank solutions to pH = 9.8 using standardized 0.1 M NaOH.

COOH acidity (mmole g-l) = (Vsample - Vb1ank) X MbasJWsample(g)

Comments

De Nobili et al. (1990b) has suggested an alternative method for the deter­mination of the carboxylic acid content of humic substance based on their solu­bility in the presence of cetyltrimethylammonium (CfA +). The CfA + is very effi­cient at quantitatively precipitating the humic substances in solutions, thus mini­mizing some of the problems in the calcium acetate method outlined by Perdue (1985). The method has several other advantages including flexibility of pH for the analysis and no apparent interference from the phenolic groups. However, the procedure has not been widely tested and it is not possible to recommend it as a general method at this stage.

Weaker Organic Acids (phenolic groups). The difference between the total acidity of a humic substance and the acidity associated with the stronger organic acids (carboxylic acids) is usually attributed to the phenolic groups, even though other acids such as weak carboxyl groups, alcoholic groups and enols are also probably involved. Before this assumption is accepted the structural impli­cations of the values should be investigated for consistency (see "General Com­ments" below). (Note, mmolf designated millimole of functional groups.)

Phenolic-OH groups (mmolf g-l)

= total acidity (mmole g-l) - COOH acidity (mmole g-l)

Comment

It is also possible to determine the acidity characteristics of the humic sub­stance using nonaqueous .titration methods (Stevenson, 1994) which tends to enhance the strengths of the weak acids. A method using DMSO is detailed in Yonebayashi and Hattori (1985) but other solvents which can also be used include pyridine (CsHsN) and dimethyl formamide. These methods warrant more inves­tigation than they have so far received.

Hydroxyl (OH) Groups

Total Hydroxyl (OH) Groups. The humic substances are acetylated with acetic anhydride in pyridine. Acetyl groups are then hydrolyzed to acetic acid (C2H40 2), which is distilled and titrated with standard base (Brooks et aI., 1958; Schnitzer & Skinner, 1965).

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ORGANIC MATTER CHARACTERIZATION 1041

Materials

1. Pyridine, 95% purity. 2. Acetic anhydride [(CH3CO)20] 97% purity. 3. Sodium hydroxide, 3 M, 0.1 M. 4. Sulfuric acid, 3 M. 5. Phenolphthalein, 0.5% in 95% ethyl alcohol.

Method

Using 5 mL of equal parts of pyridine and acetic anhydride reflux 50 to 100 mg of humic substance for 2 to 3 h under an atmosphere of N2. After cooling the mixture, pour it into distilled water and collect the precipitate by filtration. Wash the precipitate thoroughly with distilled water and dry under a vacuum in the presence ofP20 s. Reflux this acetylated sample (50 mg) with 25 mL of 3 M aque­ous NaOH solution for 2 h under N2. Cool the mixture, add 25 mL of 3 M H2S04

and 25 mL of distilled water and distill through a splash head. Collect 25 mL of distillate and titrate with standardized 0.1 M NaOH using

phenolphthalein as the indicator. Add 25 mL portions of distilled water to the dis­tillation mixture and continue the distillation. Repeat the collection of distillate, titration and addition of 25-mL portions of the distillation mixture until the sam­ple and reagent blanks titrate equally. Calculate the acetyl and hydroxyl contents as follows:

Acetyl (mmolf g-l) = (Vsample - Vblank) X MbasJWsample(g) Hydroxyl (mmolf g-l) = acetyl content/(1 - 0.042 x acetyl content)

Comments

The factor of 0.042 above is a consequence of the difference in molecular weight of 42 between the ROH and CH3COOR. The carboxylic and phenolic OH groups can be determined using a methylation procedure using methyl iodide as outlined by Schnitzer and Skinner (1965)

Phenolic (OH) Groups. The phenolic content of a humic substance is assumed to be equivalent to the concentration of weaker organic acids in the sam­ple [see "Weaker Organic Acids (Phenolic Groups)"], i.e.,

Phenolic OH (mmolf g-l)

= total acidity (mmolc g-l) - COOH acidity (mmolc g-l)

Alcoholic (OH) Groups. The alcoholic hydroxyl content of a humic sub­stance is assumed to be the difference between the total hydroxyl content and the phenolic content.

Alcoholic OH (mmolf g-l)

= total hydroxyl (mmolf g-l) - phenolic hydroxyl (mmolf g-l)

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Carbonyl (>C=O) Groups

Total Carbonyl (>C=O) Groups. The humic substances are allowed to react with an excess of hydroxylamine hydrochloride in methanoV2-propanoi. The unreacted hydroxylamine hydrochloride is titrated with standard HCI04 solution (Fritz et aI., 1959)

Materials

1. Dimethylaminoethanol [(CH3)2NCH2CH20H] 99% purity, 0.25 M (22.5 g in 2-propanol).

2. Hydroxylamine hydrochloride (NH20H . HCI), 0.4 M (27.8 g in 300 mL absolute methanol and dilute to 1 L with 2-propanol).

3. Perchloric acid (HCI04), 0.2 M.

Method

Place 50 mg of humic substance in a 50-mL ground-glass stoppered Erlen­meyer flask. Add 5 mL of 0.25 M 2-diethylaminoethanol solution and 6.3 mL of 0.4 M hydroxylamine hydrochloride solution to the flask containing the humic substance and to a similar flask in which there is no humic substance, to act as a blank. Heat the flasks on a steam bath for 15 min. Cool the solutions and back titrate potentiometrically the excess of hydroxylamine hydrochloride with stan­dard perchloric acid solution. Determine the end-point by plotting milliunits vs. milliliters of acid.

Quinone Groups. In the natural environmental humic substances undergo redox-type interactions with a number of soil components including iron (Waite & Morel, 1984). This type of interaction can be used to determine the concentra­tion of quinoid groups present in a sample. In the method outlined below the humic substances are reduced in alkaline triethanolamine (TEA) solution by Fe2+.

The excess reductant is back-titrated amperometrically with standard dichromate solution (Glebko et aI., 1970)

0= (d=0 + 2Fe2+ -triethanolamine ~ HO-<Q>-OH + 2Fe3+

Materials

1. Sodium hydroxide, 2 M. 2. Triethanolamine (TEA) 97% purity, 2 M. 3. Ferrous ammonium sulfate hexahydrate [FeS04(NH4hS04.6H20], 0.05

M. 4. Potassium dichromate (K2Cr207), 0.004 M.

Method Dissolve 20 mg of humic substance in a solution of 45 mL distilled water,

25 mL of 2 M NaOH and 25 mL of 2 M TEA in a 200-ml tall form titration cell,

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ORGANIC MATIER CHARACTERIZATION 1043

fit the lid and flush with N2 continually during the analysis. Stir the solution mag­netically for 30 min before adding 5 mL of 0.05 M ferrous ammonium sulfate hexahydrate solution and leave for 30 min. Using standardized 0.004 M potassi­um dichromate solution, back titrate the excess reductant in solution at a constant potential of -80mV, determined using a platinum foil (2.0 by 0.5 cm)/platinum wire electrode system connected to a polarograph. Carry out a blank titration under the same conditions.

Quinoid C=O (mmolf g-l) = (Vblank - Vsample) X 6 x M(K2Cr207)/Wsample(g)

Ketonic C=O Groups

Ketonic C=O groups (mmolf g-l)

= total C=O groups (mmolf g-l) - quinone groups (mmolf g-l)

General Comments

Typical values of functional group content of soil humic substances from various origins are shown in Table 35-2. Because of the complexity of humic substances, it is not easy to assess the degree of accuracy of analytical results. However, it is possible to determine the consistency of the data with respect to the logical consequences of the relationships between each set of values. For a detailed discussion of these uses the reader is referred to Perdue (1985).

With the determination of the elemental composition and the number aver­age molecular weight, theoretical values can be determined for the amount of

Table 35-2. Distribution of oxygen-containing functional groups in humic and fulvic acids isolated from soils of widely different climatic zones (in cmollkg)t (Stevenson, 1994).

Climatic zone

Cool, Cool, temperate temperate

Functional group Arctic acid soils neutral soils Subtropical Tropical Range Average

Humic acids Total acidity 560 57~90 62()"'{)60 63~770 62~750 56~90 670 COOH 320 15~570 390-450 42~520 380-450 15~570 360 AcidicOH 240 32~570 21~250 21~250 22~300 21~570 390 Weaky acidic + 490 27~350 24~320 290 2~160 20-490 260

alcoholic OH Quinone C=O 230

{1~180 {45~560 {8~150 {3~140 {1~560 {290 KetonicC=O 170 OCH3 40 40 30 3~50 6~0 3~0 60

Fulvic acids Total acidity 1100 89~1420 64~1230 82~1030 64~1420 1030 COOH 880 61~50 52~960 72~1120 52~1120 820 AcidicOH 220 28~570 12~270 3~250 3~570 300 Weakly acidic + 380 340-460 69~950 26~520 26~950 610

alcoholic OH Quinone C=O 200

{17~31O {12~260 3~150 {120-420 {270

KetonicC=O 200 16~270

OCH3 60 30-40 8~90 9~120 3~120 80

t From Schnitzer, 1977.

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unsaturation in the system and hence upper limits of the main functional groups such as carboxylic acids, phenols alcohols and enols.

Characterization by Spectroscopic Methods

The use of spectroscopic methods of analysis is important in the character­ization of humic substances. The individual techniques may yield only small fragments of information but the importance of this is magnified when combined with information from other spectroscopic techniques and/or other methods of characterization.

The aim of this section is to outline briefly the types of spectroscopic tech­niques generally used in the study of humic substances, and how they have assist­ed in increasing the knowledge of these substances. For more detailed informa­tion about the techniques the reader should consult a recent text concerned prin­cipally with their use. For a comprehensive review of the applications of the methods to studies on humic substances the text Humic Substances II: In Search of Structure (Hayes et aI., 1989) is recommended.

Ultraviolet-Visible Spectroscopy

The absorption of electromagnetic radiation in the UV (200-400 nm) and visible region (400-800 nm) is associated with the electronic transitions of the bonding electrons. The absorption of UV -visible radiation by organic compounds is due to the presence of specific segments or functional groups (chromophores) which contain unbonded electrons (e.g., carbonyl groups, S, N or 0 atoms, and conjugated C-C multiple bonds). The electronic transition within a molecular orbital is termed local excitation and the electronic transition involving the trans­fer of an electron from one chromophore to another (e.g., from an aromatic ring to an OH group) is termed electron transfer.

Generally, measurement of the absorbance of a substance is carried out by dissolving it in a solvent and determining the difference in absorbance of that solution from that of a solution containing only the solvent, either in sequence (in single beam instruments) or directly (double beam instruments). The absorbance (A) is related to the concentration of the sample in solution (c) according to the Beer-Lambert law where

A = €/c

where I is the path length of the cell and € is the absorptivity

Applications of Humic Substances

The absorbance spectrum of a compound is a characteristic which can be used in its identification. However, because the peaks of absorbance spectra are relatively broad, it is difficult to identify a particular compound in a mixture of even simple molecules and certainly not possible in a sample of humic substance. The absorption spectra of humic substances are generally featureless, consisting of a relatively smooth curve, with increasing absorption with decreasing wave­length. The curve represents the summation of the absorbances of the component chromophores. Even though it is not known what proportion of molecules con-

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ORGANIC MATIER CHARACTERIZATION 1045

tribute to the spectra, the smoothness indicates that there are a very large number of different chromophores in the sample.

Despite the apparent lack of detailed information in the spectra, different samples and fractions of humic substances do show slight variations which can be measured in a number of ways to allow comparisons to be made. For exam­ple, the EJE6 value (the ratio of the absorbance at 465 nm and 665 nm, of a dilute aqueous solution of a substance) is commonly used to characterize humic sub­stances (Konova, 1966; Chen et aI., 1977; Stevenson, 1994) with the ratio for humic acids being generally less than five and that for fulvic acid more than five. Because of the known differences between humic acids and fulvic acids the ratio would appear to be a measure of the degree of humification of a sample of humic substance.

The magnitude of the absorbance at a given wavelength varies slightly with pH (MacCarthy & O'Cinneide, 1974b; Baes & Bloom, 1990) which is probably due to the ionisation of carboxylic and phenolic functional groups. This change in absorbance with pH is dependent on the wavelength so that the EJE6 ratios of humic acid and fulvic acid also vary with pH (Chen et aI., 1977; Ghosh & Schnitzer, 1979). In alkaline solution the ratio decreases with increasing pH but below pH = 5.5 the EJE6 decreases with decreasing pH (Chen et aI., 1977; Ghosh & Schnitzer, 1979).

Both absorbance and the EJE6 ratio may be affected by variations in the salt concentration of the solution. It has therefore been suggested that the EJE6

ratio be determined in 0.05 M NaHC03 (pH = 8) (Chen et aI., 1977). A more detailed study of the dependence of the absorbance on pH can be

made by referencing the absorbance reading against a spectrum of the sample at a different pH (Tsutsuki & Kuwatsuka, 1979). These difference spectra are char­acterized by stronger peaks but the origin and cause of these peaks is not yet understood with any certainty.

The UV -visible spectra of humic substances offer very little assistance in the identification of their structure. One peculiarity is the observed absorbance at wavelengths above 500 nm when it has been found that compounds do not nor­mally absorb light at these wavelengths. It has been suggested that this is due to complex unsaturated structures (Tsutsuki & Kuwatsuka, 1979) or to electron donor-acceptor complexes (Lindqvist, 1972, 1973).

Infrared Spectroscopy

Infrared spectroscopy is the study of the molecular vibrations of bonded atoms. The frequency of the absorption is characteristic of the atoms in the bond and the type of motion associated with the vibration. The observed frequency can be used to distinguish the component atoms as well as the bonding characteristics of those atoms. Hydrogen bonding may also be apparent as it causes greater sep­aration of the bond between H and the other atom in the covalent bond, thereby decreasing the frequency of the absorption and broadening the bands.

Detailed interpretation of the spectra to determine the structure is possible in simple molecules, but it is generally not so in complex molecules or mixtures of molecules. However, useful information can be gained by comparing the spec­tra of different samples and noting changes in the spectra after chemically alter-

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1046 SWIFT

ing a sample. Using a variety of approaches it is possible to gain information on the functional groups such as aromatic, aliphatic and quinone groups associated with the sample.

Infrared spectra are usually determined over the frequency range 4000 to 400 cm-l . Absorption bands in the frequency region 4000 to 1250 cm-l are rela­tively unaffected by the remainder of the molecule and this region is called the "characteristic group frequency region." Absorption bands below 1250 em-I are affected strongly by the molecular structure and so this region is called the "fin­gerprint region."

The spectral analysis is usually carried out on a solid sample, but as the technique depends on the absorption of radiation, the sample is dispersed in a medium, which is transparent to infrared radiation. The pressed pellet method of sample preparation requires a mixture of about 1 mg of humic m~terial per 100 mg of an alkali halide (KBr) both dried and preground very finely. The mixture is pressed (~7500 kg em-2) into a small disc about 100 mm in diameter and 1 to 2 mm thick (Stevenson, 1994). Potassium bromide is transparent over the con­ventional sample range 4000-400 cm-l and so does not interfere with the spectral analysis.

Alternatively the fmely ground and dry sample can be mixed with a low vapor pressure, medium molecular weight hydrocarbon [commonly "Nujol" (Harry Holland & Sons, Burr Ridge, IL)]. This substance is less satisfactory as it adsorbs radiation at 2900, 1460 and 1375 cm-l and so imposes its spectrum over that of the sample being analyzed, and is not very suitable for the analysis of soil humic substances.

A common problem is the incorporation of water into the sample matrix, especially when using KBr discs. The presence of water causes bands at 3300 to 3000 cm-l and 1720 to 1500 cm-l regions. This can be reduced by heating and evacuating the die prior to pressing or using a nonhygroscopic pelleting matrix (Stevenson, 1994). The problems associated with water in the sample can be avoided by using cast films or solutions in a cell (Bloom & Leenheer, 1989).

Applications to Humic Substances

The infrared (IR) spectra of humic substances are relatively simple with only a few broad bands and no well-defmed, sharp peaks, typical of the spectra of single compounds. As for UV -visible spectra, these undefined broad bands are evidence that the functional groups in the sample of humic substances exist in a wide variety of chemical environments. There are often similarities in the gener­al appearance of spectra of different samples of humic substances. This does not necessarily mean that the samples are composed of molecules with similar struc­ture, but does indicate that the overall functional group content is similar.

Derivatization techniques are frequently used to study the attribution of observed spectral bands to a particular functional group. An excellent outline of the bands associated with the main functional groups and the type of derivatiza­tion procedures which may assist in the assignment of the bands for a particular sample, is given in Bloom and Leenheer (1989).

In situations where the drying-out process may affect the chemical equilib­rium in the sample being studied, it may be necessary to analyze the sample in

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ORGANIC MATIER CHARACTERIZATION 1047

4000 3000 2000 1000

Wavenumber (cm-1)

Fig. 35-3. Diffuse reflectance Fourier-transformed infrared spectra of four International Humic Sub-stances Society humic acids (Neimeyer et aI., 1992). Note: the Kubelka-Munk units arise from the application of the Kubelka-Munk transformation (Kortum, 1969) of the reflectance spectra and result in spectra similar in appearance to absorbance spectra obtained from transmission measure­ments (Baes & Bloom, 1989).

Table 35-3. Fourier-transformed infrared (FTIR) bands of peat humic and fulvic acids (adapted from Baes & Bloom, 1989.

Band

3330-3380 3030 2930 2840 2600 1720 1610 1525 1450 1350 1270 1225 1170 1070 830

775

Assignment

OH stretch of phenolic OH (contribution from aliphatic OH, H20 and possibly NH) Aromatic CH stretch Asymmetric CH stretch of -CH2-

Symmetric CH stretch of -CHz-OH stretch of H-bonded -COOH -C=O stretch of -COOH Aromatic C=C stretch and/or asymmetric -COO- stretch Aromatic C=C stretch -CH deformation of -CH3 and -CH bending of -CHz Symmetric -COO' stretch and/or -CH bending of aliphatics -C-OH stretch of phenolic OH -C-O stretch and OH deformation of -COOH -C-OH stretch of aliphatic OH C-C stretch of aliphatic groups Aromatic CH out of plane bending Aromatic CH out of plane bending

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1048 SWIFr

the aqueous state. This is difficult in infrared spectroscopy as the absorption bands of water are large and centered around the major area of interest, 3400 to 1640 cm- I , and so interfere with the analysis. This may be overcome by using D20 instead ofHzO to shift the position of the bands (MacCarthy & Mark, 1975). More recently, Fourier Transform Infrared Spectroscopy (FTIR) and an adapta­tion to this method using diffuse reflectance rather than transmitted light (DRIFT), have been applied to the study of humic substances (Baes & Bloom, 1989; Niemeyer et ai., 1992; Ristori et ai., 1992). The Fourier transform tech­nique eliminates problems associated with the presence of water and improves the resolution. The diffuse reflectance adaptation permits the analysis of opaque samples so that whole soil samples can be studied (Ristori et ai., 1992; Nguyen et ai., 1991). Reflectance methods can be used with dispersive instruments, and KBr pellets can be used with FTIR. The combination of FTIR with diffuse reflectance, however, seems to offer the best method for analysis of humic sub­stances.

The small amount of sample required and the simplicity of the procedure make IR spectroscopy one of the most commonly used methods of analysis for humic substances. The information gained from the spectra is in many cases not definitive as the assignment of bands to particular groups is only tentative in com­plex mixtures of compounds. The results should be confirmed with other meth­ods of analysis.

Examples of typical FTIR spectra of humic acids are shown in Fig. 35-3. The assignment of structure generally given to the absorption bands in humic substances is well documented in Stevenson (1994) and a summary of the assign­ments is shown in Table 35-3.

Nuclear Magnetic Resonance

When certain nuclei with a particular type of spin are placed in a magnetic field, they align themselves such that some have their spin magnetic moment vec­tors parallel to the field vector and are at slightly lower energy to others that are antiparallel in the field.

If an oscillating magnetic field is superimposed on the steady magnetic field with a perpendicular magnetic vector, then for a particular steady magnetic field strength, absorption of radiation will occur at certain frequencies of oscilla­tion, allowing transitions between different energy spin states. This resonance condition can be met by varying the magnitude of either the steady state field or the oscillating field. In Fourier transform nuclear magnetic resonance (NMR) spectroscopy the sample is subjected to a pulse of radio frequency radiation com­prising all frequencies.

In a molecular environment, the atoms surrounding a nucleus partially shield it from the magnetic field so that the frequency of the oscillating field re­quired to achieve resonance is changed. As this shielding is a function of the chemical environment associated with the nuclei the resonance frequencies of the atoms in the compound are able to give information about the chemical structure of the compound. Common nuclei used for the studies with NMR are 1 Hand 13e.

The NMR frequency of a given nucleus (v sample) is measured relative to that of a standard compound {typically tetramethylsilane [(CH3)4Si] (TMS)) (v

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ORGANIC MATTER CHARACTERIZATION 1049

reference). The frequency for resonance is then given as the difference, or, chem­ical shift (0) between these two frequencies, expressed in parts per million (ppm).

o = (Vsample - Vreference) X 106/vreference

Common solvents used in liquid state 13C-NMR are NaOH and NaOO and these appear to display no absorptions. The OMSO-d6 provides a sharper spec­trum but the 30 to 40 ppm region is obscured unless the solvent is depleted in 13e. In IH-NMR the aqueous H is H20, 0 20 and aqueous NaOH obscures the 3 to 5 ppm region.

There are a number of problems associated with liquid state NMR such as the amount of sample required (100-200 mg), insolubility of some compounds in suitable solvents, interference from water in IH-NMR and the long analysis times required (2-12 h are common for IH-NMR and up to 1 wk for 13C NMR). In some situations the dissolution process interferes with the analysis. The develop­ment of solid state NMR has avoided these problems as well as achieving a high­er signal to noise ratio, generally giving greater sensitivity in the spectra. How­ever, the technical problems in getting good spectra are much greater.

Applications to Humic Substances

Nuclear magnetic resonance spectroscopy is generally used to compare the difference in concentration of the main functional groups between samples of humic substances. Both IH-NMR and 13C-NMR are able to be used for the char­acterization of humic substances, although the relatively low abundance of 13C (about 1.1%) in the latter technique places considerable demands on instrumen­tation and methodology. Nevertheless, these two methods provide powerful tools for studying humic substances (Wilson et aI., 1983; Wilson & Goh, 1983).

The most common type of NMR now used in the study of humic substances is called solid state CPMAS-NMR (cross-polarization, magic angle spinning­nuclear magnetic resonance) (Wilson et aI., 1987; Krosshaven et aI., 1990). An example of CPMAS 13C-NMR spectra of a soil humic substance is shown in Fig.

Chemical shift, S(ppm)

Fig. 35-4. Typical CPMAS 13C-NMR spectrum of a soil humic substance.

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1050

Table 35-4. Major proton resonance of humic materialst (Wershaw, 1985).

Chemical shift, 1) (relative to TMS as 0)

ppm

13.0 10.0

Assignment

Carboxylic acid protons Hydroxyl protons Aromatic protons Lactone protons Methoxyl protons

SWIFr

6.0-7.5 4.0-5.5

3.7 2.6 1.3 0.9

Aliphatic protons attached C atom IX to a benzene ring:j: Aliphatic protons ~ to a benzene ring:j: Aliphatic protons y to a benzene ring:j:

t Spectra measured in deuterodimethylsulfoxide [(CD3hSOj. :j:--CaH;z-CpHrCrH2·

35-4. It is now possible to compare the NMR spectra of humic substances in whole soil samples with those of the associated extracted fractions. Using this type of approach, Krosshaven et al. (1992) concluded that conventional humus fractionation does not significantly change the content of the different functional groups in a sample. Assignments of chemical shifts for IH-NMR and CPMAS 13C NMR are shown in Tables 35-4 and 35-5, respectively.

Characterization of humic substances using NMR spectroscopy is a rela­tively new but widespread technique, both in the number of research groups involved and the type of research being undertaken. Nuclear magnetic resonance spectroscopy is proving to be important, both as a source of information relating to the structure of humic substances, as well as supporting information gained from other types of analyses. However, there is a need to standardize the proce­dures and techniques used, especially in solid state NMR, to take full advantage of this type of research with respect to humic substances.

Detailed information on the theoretical aspects of NMR spectroscopy, and the application of this theory to the studies of humic substances, is given in Hayes et al. (1989).

Electron Spin Resonance

When molecules containing unpaired electrons in a magnetic field are irra­diated with electromagnetic radiation, the electrons can be excited to a higher energy state and those which are already at the higher energy level can be induced to move to the lower energy state. For a system initially in a state of thermal equi­librium there will be slightly more electrons in the lower state giving a net ab­sorption of energy. The absorption of this energy is the basis of Electron Spin Resonance (ESR) spectroscopy which involves the study of molecules with unpaired electrons (free radicals).

The energy difference between the two spin states (aE) is

where h is Plank's constant, v is the frequency of the electromagnetic radiation, g is the spectroscopic splitting factor, ~ is the Bohr magneton (a constant) and Ho is the applied field.

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ORGANIC MATIER CHARACTERIZATION 1051

Table 35-5. Chemical shift assignments in the CPMAS DC NMR spectra of fulvic and humic acids (adapted from Malcolm, 1989).

Shift range

ppm

0-50 10-20 15-50 25-50 29-33 35-50 41-42 45-46 50-95 51-61 57-65 65-85 90-110

110-160 110-120 118-122 120-140 140-160 160-230 160-190 190-230

Possible assignments

Unsubstituted saturated aliphatic C atoms Terminal methyl groups Methylene groups in alkyl chains Methine groups in alkyl chains Methylene C (l,~, 0, E from terminal methyl group Methylene C atoms of branched alkyl chains (l-C in aliphatic acids R2 NCH3

Aliphatic C singly bonded to one 0 or N atom Aliphatic esters and ethers; methoxy, ethoxy Carbon in CH20H groups; C6 in polysaccharides Carbon in CH(OH) groups; ring C atoms of polysaccharides; ether-bonded aliphatic C Carbon singly bonded to two 0 atoms; C, anomeric in polysaccharides, acetal or ketal Aromatic and unsaturated C Protonated aromatic C, aryl H Aromatic C ortho to O-substituted aromatic C Unsubstituted and alkyl-substituted aromatic C. Aromatic C substituted by 0 and N; aromatic ether, phenol, aromatic amines Carbonyl, carboxyl, amide, ester C atoms Largely carboxyl C atoms Carbonyl C atoms

It is possible to determine the concentration of unpaired electrons (spins) by comparing the integrated absorption with that of a standard under the same conditions, although the accuracy of this method is believed to be quite low. Mea­surement of the field strength and the frequency of the electromagnetic radiation at resonance determines the g-value, the magnitude of which is associated with the molecular structure of the molecule. The hyperfine structure of the spectra can be used to determine the number and types of nuclei interacting with the free electron, and hence the structure of the molecule. The concentration of free radi­cals is determined by the width of the absorption line.

Application to Humic Substances

The ESR spectra of humic substances are generally only a single line which limits the amount of information that can be gained from the analysis. The hyper­fine structure in ESR spectra, typical of simple systems, is absent although a few instances have been reported with splitting (Atherton et aI., 1967; Senesi et aI., 1977). An example of an ESR spectrum of a humic substance is shown in Fig. 35-5 with some typical values of the measurable parameters for soil humic sub­stances in Table 35-6. These values are only meant to be indicative of those com­monly found as the magnitude is dependent on solution pH, metal ion content and solvent effects.

Results obtained from this type of research have led to the realization that the free radicals in humic substances are remarkably stable with respect to time and chemical attack. This stability has been attributed to the stabilization of an unpaired electron over an aromatic system (Theng & Posner, 1967) suggesting that humic substances contain free radicals of the semi-quinone type (Steelink,

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1052 SWIFT

A

B

-5 o +5

Fig. 35-5.ESR spectra of a podzol FA in NaOD-D20: (A) immediately after mixing fulvic acid with NaOH; (B) 48 h after mixing. g = 2.0040; line width = 5.0 x 10-4 T; frequency = 9.451 GHz; field center = 3342.0 (Steelink et aI., 1983).

1964). Basic solutions of humic substances contain exceptionally long-lived free radicals. It has also been found that the spin content generally increases marked­ly with increasing pH.

The concentration of free radicals in humic substances is relatively small (Schnitzer & Skinner, 1969), and as the concentration varies with the type of sol­vent used in the extraction procedure it may even be an artifact of extraction tech­nique used (Hayes et aI., 1975). Other techniques such as NMR and infrared (IR) are more useful than ESR in characterizing the functionality of soil humic sub­stances. However, ESR is a source of additional information and should be included in the general research procedure. If the technique was to become more reliable and informative the potential of its use would increase as it is concerned with one of the more reactive fractions of humic substances, the free radical con­tent.

Pyrolysis-Mass Spectroscopy

Pyrolysis (Py) is the degradation of a substance through the action of heat. This process is usually carried out in a vacuum, or in a rapid stream of inert gas to restrict the formation of secondary products. The absorption of thermal energy causes excitation of the bond vibrational modes resulting in the cleavage of the weaker bonds. The number and variety of the products formed by using this tech-

Table 35-6. Electron spin resonance parameters for various humic and fulvic acids (Steelink et aI., 1983).

Free-radical Spectroscopic Sample State concentration Line width splitting factor Reference

spinsg-1x 10-17

T (g value)

Soil humic Solid 5-10 4.8 x 10-4-5.2 x 10-4 2.0032-2.0047 Chen et al. (1978) acid

Soil fulvic Solid 1-2 4.5 x 10-4-7.5 x 10-4 2.0037-2.0050 Chen et al. (1978) acid

Soil humic Aqueous 2.1 2.5 x 10-4 2.0037 Varadachari et al. acid solution (1983)

Soil fulvic Aqueous 1.3 2.5 x 10-4 2.0038 Varadachari et al. acid solution (1983)

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ORGANIC MAlTER CHARACTERIZATION 1053

nique to study humic substances is very large. It is therefore essential to incorpo­rate mass spectrometry (Py-MS) or gas chromatography (Py-GC) into the system, to separate and identify the reaction products. A further, and highly desirable, improvement on the technique is to use a combination of both gas chromatogra­phy and mass spp.ctrometry (Py-GC-MS). This system allows the volatile reaction products to be separated prior to the analysis using the mass spectrometer. Alter­natively, soft-ionization techniques such as field ionization (FI) and field-desorp­tion combined with pyrolysis mass spectrometry, (Py-FIMS, Py-FDMS) extend the range in size of molecular ions which can be observed by Py-MS by includ­ing the small and larger molecular ions, respectively.

The data gained from each analysis can be used as ajingerprint of that sam­ple under the particular pyrolysis conditions used. This fingerprint can then be compared with the results of analysis on other samples of humic substances or on known chemical compounds. Because of the large amount of data collected using these analytical techniques, multivariate analysis with the aid of computers is used to collate the results (Howarth & Sinding-Larsen, 1983). The theoretical and technical aspects of these pyrolysis techniques and the methods of treatment of the accumulated data are described by Bracewell et ai. (1989).

Applications to Humic Substances

Pyrolysis techniques are used to study humic substances in an attempt to correlate samples of humic substances with their parent biopolymers (Meuzelaar et aI., 1982). Evidence from the studies using pyrolysis indicate that the humic macromolecules have polysaccharides, polypeptides and lignins incorporated into their structures (Greenland & Oades, 1975; Halma et aI., 1978; Bracewell et aI., 1980). Pyrolysis techniques have also been used to determine differences between the various fractions of humic substances (Saiz-Himinez et aI., 1978, 1979). The studies carried out so far indicate that there is very little variation in the pyrolysates of humic acids from different soils and different soil types, where­as the pyrolysates of a fulvic acid fraction are dependent on the origin of the soil and the method of extraction (Bracewell et aI., 1989).

One of the most important aspects of pyrolysis with respect to applications to humic substances is the ability of this technique to analyze whole soil samples. This permits a relatively rapid analysis of soil organic matter in general and hence, comparison of organic matter associated with soils from different origins (Baldock et aI., 1991) or as a result of different management practices (Schulten & Hempfling, 1992). Figure 35-6 shows Py-FI mass spectra [intensity of the ion vs. the mass of m/z (mass to charge ratio of the ion)] of the pyrolysates obtained from a soil as a result of different intensities of crop rotation and manure treat­ment. These two soil samples exhibited almost identical total concentrations of C and N. Schulten and Hempfling determined that there was little difference between the pyrolysates of the soil samples in the mass range m/z 50 to 200, but in the higher molecular weight products of the pyrolysis the intensities were greater for the soil that had less intense cropping and had been treated with manure.

Comparison of Py-FIMS spectra of whole soil samples with those of the extracted humic fractions permits the verification of consistency of results when

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1054 SWIFf

5.0 96 r--xl1f-(a)

4.0 58

3.0 82 110

1208 i 2.0

67

l1.0 ,!;' 0 Cf) 50 100 150 200 250 300 350 400 c: ~ 5.058-_96

(b) c: .£ 4.0 «i ~ 3.0 110

2.0

1.0 340

0 50 100 150 200 250 300 350 400

m/z-

Fig. 35-6. Averaged Py-FI mass spectra of soil samples from the same site and with the same Nand C content, but under different management systems and (a) with manure addition, (b) without manure addition.

characterizing the fractions (Hempfling & Schulten, 1991). It has been found that humic acid extracts from soils are enriched in polypeptides relative to the whole soil pyrolysate, and the fulvic acid extracts are deficient in polypeptides but enriched in polysaccharides or pseudo-polysaccharides (Haider et aI., 1977). The "humin" extract had a similar fingerprint to that of the humic acid indicating that the nonextracted material of a soil humic substance has similar chemical content to that of the extracted material. The effects of different methods of extraction of fractions of humic substances have also been observed using Py-FIMS by Haider & Schulten (1985).

Pyrolysis techniques also provide supporting data for the results obtained when using other types of chemical analysis and instrumental analysis. Zech et al. (1990) used chemical analysis combined with IR, 13C-NMR and Py-FIMS instrumental methods to characterize and compare the organic matter in soils from different soil horizons. Beyer et al. (1992) concluded that the Py-FIMS results confirmed and extended the results obtained from wet chemistry methods and NMR methods.

Pyrolysis techniques used to study whole soil samples allow the character­ization of the organic matter without going through the potentially destructive and time-consuming processes of extraction and purification of the humic sub­stances. It is for both of these reasons that this approach to organic matter char­acterization is receiving increasing interest and holds considerable promise.

Fluorescence Spectroscopy

The UV -visible radiation absorbed by a molecule may be dissipated as heat and/or electromagnetic radiation at a longer wavelength than the incident radia­tion. This emitted radiation is known as fluorescence and it is a characteristic of humic substances.

There are three types of spectra generally associated with this technique. Excitation spectra are obtained by scanning the incident radiation and determin-

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ORGANIC MAITER CHARACTERIZATION 1055

ing the intensity of radiation emitted at a fIxed wavelength. Emission spectra are obtained by fixing the incident radiation and determining the intensity of radia­tion emitted over a wavelength range. Synchronous excitation spectra are obtained by setting the difference between the emitted and absorbed radiation to be a constant (LlA) and determining the intensity of the emitted radiation for a range of wavelengths.

Application to Humic Substances

In general, only a small fraction of molecules are known to fluoresce, and so the proportion of molecules in a sample of humic substance contributing to the observed fluorescence is relatively small. Additionally, this observed fluores­cence is not the total fluorescence as humic substances absorb radiation over a wide range of wavelengths and some of the fluorescence radiation will be reab­sorbed by the sample.

Fluorescence spectroscopy has been used to characterize samples of humic substances but for a number of reasons it has not gained general acceptance (Sen­esi, 1990). Because of the reabsorbance of the fluoresced radiation the relative intensity of the observed radiation at a particular wavelength is a function of the concentration of humic substances in solution (Spark & Swift, 1994). When using UV -visible absorbance spectroscopy increasing the solution concentration of dilute solutions increases the absorbance in a regular manner. In fluorescence spectroscopy increasing the concentration of humic substances in dilute solution does initially increase the observed intensity of fluorescence, but eventually this reaches a maximum and then begins to decrease. The concentration at which the fluorescence reaches a maximum is dependent on the wavelength of excitation and emission as well as on the nature of the humic substance. As yet it has not been possible to correlate the detail in the observed fluorescence spectra of humic substances with known characteristics of these substances so the usefulness of this technique is limited.

Measurements of Physical Properties

The physical properties generally determined for humic substances are molecular size, shape and weight, and charge characteristics. The principles of these determinations are theoretically soundly based, but in practice the results are somewhat limited in value by the large variation in range of the properties in even the most highly fractionated samples.

Molecular Size and Shape

Molecular size and shape of humic molecules are two of the most elusive properties of humic substances. Considerable research on humic substances has been directed towards the determination of these properties, but as knowledge of humic substances has increased, so has the realization of the complexities asso­ciated with assigning values to the size and shape of humic molecules. It is prob­ably because of these problems, as well as the rapid development of spectro­scopic instrumentation, that this area of humic substance research has made little progress in recent years with respect to other areas.

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1056 SWIFT

The techniques generally used to study the molecular size and shape of humic substances are gel permeation chromatography and viscosimetry. A vari­ety of other techniques such as light scattering (see "Light Scattering"), Flow Field Flow Fractionation (FFFF) (Becket et aI., 1987), ultracentifugation (Posner & Creeth, 1972; Ritchie & Posner, 1982), and colligative data calculations (Reuter & Perdue, 1981) have also been used.

Gel Chromatography. The general principles relating to gel chromatogra­phy have previously been discussed with respect to the fractionation of humic substances (see "Ion-Exchange Chromatography"). For molecules of the same shape and structure it is possible to calibrate the gel column with respect to elu­tion volume vs. molecular weight using known compounds, and hence determine the molecular weight of an unknown compound. As elution time is dependent on the molecular shape and structure, the elution time for two molecules of the same molecular weight, one of which is spherical and the other a long chain molecule, may be quite different.

The elution volume for a given solute is dependent on the geometry of the column, the length of the gel bed and differences in the packing density, and so cannot be used to characterize a solute. The distribution coefficient (Kd) can be used to compare the migration rates of different solutes in different experiments.

where Vc is the elution volume, Vo is the void (excluded) volume Vi is the inner volume (i.e., the solvent volume inside the gel beads). Because of the difficulty in measuring the magnitude of Vi> the total volume of the column (VI) is used to determine an approximation of Kav according to the equation

Applications to Humic Substances

A variety of media are now used in this type of work including dextran (C6HlOOS)m polyacrylamide {[-CH2C(CONH2)O-]}, agarose gels and porous glass beads. The elution characteristics that can affect the elution time are inter­actions between the gel and solute, ionic strength and pH of the solution (Tsutsu­ki & Kuwatsuki, 1984), concentration of the sample and the type of buffer used (Swift & Posner, 1971). Swift and Posner (1971) found that two buffer systems tris (2-hydroxy-2-aminopropane-1,3-diol) and borate buffers at pH 9, are partic­ularly well suited for use with humic substances.

Calibrations of the gels are usually made using polysaccharides or proteins of known molecular weights. It has been found (Cameron et al., 1972b) that there is a better correlation between humic substances with polysaccharides than with proteins. This may be because the more diffuse or open molecular configuration of polysaccharides is closer to that of the humic substances in solution, than are the condensed protein configurations.

Calibration of the column is carried out by determining the elution charac­teristics of humic substances of which the molecular weight has been estimated

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ORGANIC MATI'ER CHARACTERIZATION 1057

using some other technique, such as ultracentifugation (Cameron et aI., 1972b). Dawson et al. (1981) had some success calibrating the column using chemicals thought to be similar to those of humic substances.

Ultracentrifugation. Ultracentrifugation is the centrifugation of solutions at high speed (100 0()(}-600 000 g) under a vacuum to cause sedimentation of solute species. The velocity of sedimentation can be used to separate molecules and to determine molecular shape and size.

Sedimentation of a macromolecule requires its movement through solution which may be hindered by a number of factors including frictional resistance from the solvent, molecular enlargement due to solvation, and interactions with other sedimenting molecules and/or solute components. In addition to these fac­tors if the sedimenting molecule is charged its sedimentation velocity will be decreased due to the accumulation of small counter ions behind it. The sedimen­tation velocity may also be affected by the sedimentation characteristics of the buffer or background electrolyte.

A number of different techniques are used in ultracentrifugation. Sedimen­tation velocity involves the use of high rotor speed and requires the development of well-formed boundaries, which is not possible in polydisperse systems. Sedi­mentation equilibrium uses lower rotor speeds for long periods of time, until the macromolecules in solution are in equilibrium. The number average (Mn), weight-average (Mw) and z-average (Mz) molecular weights of the sample can be determined using this method for samples with a limited amount of poydispersi­ty without the need for any other measurements.

Both of the above techniques rely on the use of analytical centrifuges which have optical detection systems attached. When preparative centrifuges, which do not have associated optical systems, are used, it is necessary to carry out the sep­aration using the density gradient technique to overcome problems of convection and mechanical disturbance. The density gradient is formed in the centrifuge tube by using increasing concentrations of a solute such as sucrose, an inorganic salt or mixtures of H20 and D20.

Applications of Humic Substances

There are a number of problems associated with applying this technique to the study of humic substances. The color of humic substances restricts the trans­mission of light and hence limits the accuracy of the refractive index determina­tion. The uncertainty of the molecular structure and conformation of humic sub­stances makes it difficult to nominate suitable compounds to be used as internal standards. However, the method is still valuable in being able to determine rela­tive values of molecular weight between different fractions, or changes in the molecular weight of a fraction associated with certain conditions or treatments. It is necessary to use a relatively high concentration of background electrolyte concentration (0.1-0.2 M) when analyzing humic substances to limit interfer­ences associated with the highly charged nature of the molecules in these sam­ples. This concentration of electrolyte will also influence the configuration and degree of expansion of the humic substance in solution.

The three types of techniques, sedimentation velocity (Stevenson et aI., 1953; Cameron et aI., 1972b), equilibrium ultracentrifugation (Posner & Creeth,

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lOS8 SWIFf

1972) and density gradient methods (Ritchie & Posner, 1982) have been applied to samples of humic substances. In the latter two techniques, carried out using fractionated samples of humic substances, the large variation in the values of (Mn), (Mw) and (MJ are strong evidence for the polydispersity of humic sub­stances (Swift, 1989). Because of the highly polydisperse nature of humic sub­stances the density gradient technique is potentially the most applicable, particu­larly for preparative procedures.

Light Scattering. The scattering of light by particles in solution can be used to determine their molecular weight, a technique commonly used for the study of polymers. The molecular weight (M) of the particles in a monodisperse system at a density p can be determined by the using the relationship M = P V. The volume of the particle (V) is determined using the Raleigh equation where the ratio of the intensity of the scattered light [/(9)] to the intensity of the incident light (/0) (Hunter, 1989) is given by

1(9) = 91t2W(n2 - 1)2(1 + cosZS)

10 2,.z~..4(n2 + 2)

where N, and r are the number and radius of the of particles respectively, n is the refractive index of the light scattering particles relative to that of solvent, A. is the wavelength of the light and 9 is the angle between the incident beam and the scat­tered beam. It is assumed that the particles are uniform in size and spherical and that the particle size is small relative to the wavelength of the scattered light.

Applications to Humic Substances

Very little use has been made of this technique to study humic substances. Theoretically, light scattering techniques may be extended to study the changes in solution in response to change in pH (Wershaw, 1989) and to study the com­plexation of humic particles in the presence of heavy metal ions (MacCarthy & Mark, 1976; Ryan & Weber, 1982).

In practice problems arise from the absorption and fluorescence of light by humic substances which would significantly interfere with the analysis at wave­lengths below 500 nm (Wershaw, 1989). Reid et aI., (1991) used laser light at 633 nm to study the aggregation characteristics of humic molecules in solution. Using laser lights with the new technique of quasi-elastic light scattering is also a pos­sibility (Wershaw, 1989).

Viscosity. Information about the size and shape of humic and fulvic acids, as well as their weight, polyelectrolytic characteristics and interaction behavior with other macromolecules can be determined from measurements of viscosity. The equipment required for this type of analysis is relatively inexpensive and the experimental procedures are easier to conduct than other methods available for determining the characteristics of humic substances. However, it must be noted that the values obtained from the measurements tend to be relative values only. Both the theory behind this technique, as well as the experimental information, are well documented by Clapp et al. (1989).

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Application to Humic Substances

The viscosity of humic substances in solution has been found to depend on both the pH and the salt concentration of the solution (Chen & Schnitzer, 1976; Stevenson, 1994). The results obtained from studies using different concentra­tions of salts (Ghosh & Schnitzer, 1980) support the theory that humic macro­molecules in solution are polyelectrolytic in character with the degree of coiling dependent on the concentration of the salt. It has been found that for a given mol­ecular weight, humic substances from different sources exhibit varying degrees of expansion (Visser, 1985).

Besides being used simply as a method of characterizing a humic substance (Tomar et aI., 1992), viscosity measurements have also been used to study the interactions between contaminants and humic substances (Chen et aI., 1992) and between soil minerals and humic substances (Zhang et aI., 1991).

Other Techniques

A number of other techniques are used to characterize humic substances. Two of these are chemical degradation studies and electron microscopy. Neither of these techniques are discussed in detail, and the reader is referred to other sources of information.

Chemical Degradation

The macromolecular size, the diverse arrangement of functional groups and the multitude of interactions possible between these groups make the task of de­termining the structure of humic substances extremely difficult. The aim of chemical degradation studies of humic substances is usually to break down the large macromolecules into smaller, more recognizable units. From these units it may then be theoretically possible to determine the origin of the parent molecules and, hence, build up a picture of the structure of humic substances.

The process of rebuilding the molecular structure of humic substances from the component molecules is complex in itself, but this work is made much more difficult by the inability to control the degradative process to achieve a satisfac­tory set of simpler molecules. The ideal method should yield high amounts of degradative products of moderate molecular complexity (Stevenson, 1994). If the degradation producer is too mild, the total amount of each type of recognizable molecules is too small to be determined analytically. And, if the degradation pro­cedure is too extreme, the types of recognizable molecules produced are so sim­ple that little information is gained about the nature of the parent molecule. Some of those problems may be overcome by using a combination of different proce­dures and a range of severity (Stevenson, 1994; Hayes et aI., 1989).

The types of chemical procedures used in degradative studies include oxi­dation, reduction, hydrolysis and depolymerization.

The compounds produced when applying these procedures to humic sub­stances are numerous, including complex phenolic and benzenecarboxylic acids, hydroxycarboxylic acids, nitrophenols and polycyclic ring compounds (Steven­son, 1994). The text Humic Substances II. In Search of Structure (Hayes et aI., 1989) provides a detailed reference for this type of research.

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Electron Microscopy

Electron microscopy has been used to study the physical appearance of humic substances. This technique requires that the sample is dried which, undoubtedly, alters some of the physical characteristics of the humic substances; It is known that factors, such as pH, ionic strength, metal complexation and con­centration, affect the size and shape of the dried particles of humic substances (Chen & Schnitzer, 1989). The method of drying the sample also affects the resultant appearance of the particle.

It is because of these types of problems that the results from electron microscopy are thought to accentuate or distort the characteristics of humic sub­stances in the natural environment, but do provide an insight into the possible physical appearance of those substances. Recent advances in electron microscopy allow samples to be studied under moist conditions, and it is important that this technique be applied to the study of humic substances.

CHARACTERISTICS OF SOIL POLYSACCHARIDES

Many of the chemical and physical methods for characterizing humic sub­stances can similarly be used to characterize soil polysaccharides (Cheshire, 1979; Cheshire & Hayes, 1990). This section will concentrate on those methods peculiar to polysaccharides.

Determination of Monosaccharide Composition

Characterization of a sample of soil polysaccharides has similar problems to humic substances with respect to the complexity of the sample hindering most attempts at detailed analysis. However, because of the polymeric nature of this soil fraction it is theoretically possible to convert the larger macromolecules using hydrolysis procedures, into smaller and perhaps more recognizable units.

Hydrolysis of polysaccharides usually involves the reaction with hot acid to the sample to break the glycosidic bonds joining the different sugars. This method is generally used to carry out an estimation of total sugars. Because of the different natures of the monosaccharide units, the nature of the hydrolyzing acid and the analytical conditions needed to effect link breakages without destruction of the monosaccharide units depends on the type of sugar units present. Side­reactions such as acid-reversion [Acid-catalyzed disaccharide or oligosaccharide formation (Overend et aI., 1962)] are possible.

Another approach to the characterization of polysaccharides is the methyl­ation technique (Cheshire et aI., 1979, 1983). By methylating the polysaccharides prior to hydrolysis, it is theoretically possible to identify the monosaccharide linkages (unmethylated sites on the monosaccharide products) within the poly­saccharide macromolecule. Complete methylation is difficult even when using extracted samples which may be due to steric hindrances arising from the associ­ation of hydroxyl groups with nonsugar components (Cheshire & Hayes, 1990).

No ideal conditions exist whereby the hydrolysis conditions are optimal for all the general groups of monosaccharides. Therefore a variety of analysis are usually performed for each of the main groups of monosaccharides. These

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include hexosamines, hexoses, deoxyhexoses, pentoses and uronic acids. The conditions generally used are: hexosamines, HCI (4-8 M); hexoses, H2S04 (0.5 or 1 M); deoxyhexoses, HCI or H2S04 (0.1 M) (Swincer et aI., 1969). It is more difficult to achieve a satisfactory degree of recovery for the uronic acids and the pentoses.

Hydrolysis of Polysaccharides

The method below based on that of Oades et ai. (1970), uses 2.5 M H2S04 to extract one fraction of polysaccharides prior to a more vigorous extraction pro­cedure with 0.5 and 12 M H2S04,

Materials

1. Sulfuric acid, 12 M (72% w/w, sp. gr = 1.634), 2.5 M 2. Sodium hydroxide, concentrated.

Method

Combine 2 g of finely ground air dried soil with 25 mL of 2.5 M H2S04 in a 50-mL round bottom flask and reflux the suspension for 20 min. Filter the mix­ture through a sintered glass filter, wash the residue with distilled water and dry over P20S' Retain the filtrate and washings for analysis or to be combined with the hydrolysate from the next part of the method.

Soak the residue in 12 M H2S04 for 16 h and then dilute the suspension to 0.5 M H2S04, Stopper the flask with a capillary air leak and heat at 100°C for 5 h. Cool the mixture and filter through a sintered glass filter (porosity x 3). Com­bine the filtrate with that from the above procedure, neutralize this resulting solu­tion to pH = 7 using NaOH, make the volume up to 100 mL and then filter again (Whatman no. 1 paper). This solution is then ready for analysis of hydrolysates.

Analysis of Hydrolysates

The analysis of the sugar mixture released by the hydrolysis procedure is often carried out using an alkaline ferricyanide and anthrone method (Cheshire, 1979). An alternative to this involves reduction and acetylation of the sugars fol­lowed by analysis of the solution by gas-liquid chromatography (glc) or high per­formance liquid chromatography (hplc). For a simple monosaccharide mixture hplc is probably the most suitable technique. However, monosaccharide mixtures derived from samples of humic substances are relatively complex. It has been rec­ommended that glc is a more suitable method for quantitative and qualitative analysis of these samples (Chaplin & Kennedy, 1986).

The method below, based on that used by Oades et ai. (1970), is designed for analysis of the sugars using Gc.

Materials

1. Sodium borohydride (NaBH4) 2. Acetic anhydride (CH3CO)zO 3. Chloroform (CHCI3) 4. Methylene chloride (CH2Cl2)

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Method

Place a solution (water or methanol solvent) of the sugars (1-10 mg), including internal standards, in a 12.4 by 1.6-cm screw top test tube, add 1 mg of solid sodium borohydride and leave to stand overnight. Place this reduced sugar solution in a warm bath at 60°C and blow to dryness using a stream of air.

Add 1 to 2 mL of methanol containing 10% (v/v) glacial acetic acid to each tube in the 60°C water bath and blow it to dryness again using a jet of air. Repeat this procedure four times. Add 1 to 2 mL of acetic anhydride, then seal the tubes using screw caps fitted with Teflon inserts and heat at 130 to 135°C for 2 h. Dry the reaction mixture using the water bath and stream of air as before.

Extract the alditol acetates using chloroform and filter the solution under pressure through a small column (5 by 10 mm) of Silica Gel G (Bio-Rad, Rich­mond, CA) using chloroform as the eluant to remove the fine black solids present when analysing soil hydrolysates. Evaporate the chloroform filtrates to dryness then dissolve the alditol acetates in methylene chloride before injection into a gas chromatograph.

Comment

Two chemicals suitable for use as an internal standard for the above method are myoinositol (C6H120 6) and quebrachitol (C7H140 6) (Oades et aI., 1970). A suitable internal standard to be used for plant-derived carbohydrate analysis is pentaerythritol (CSH120 4) (Chaplin & Kennedy, 1986).

FUTURE DEVELOPMENTS

It must be obvious to the reader that there is still much to learn about the nature of soil organic matter. As each new technique is applied to this area of sci­entific research it becomes more apparent that, in the short term, the chances of a major breakthrough' are unlikely. The amount of information gained from each individual study or from the application of a single technique is usually small, but the significance of that information may then be magnified when combined with that from other studies or results. In this context, it is important that as many tech­niques as possible are used to study the same material and the information ob­tained integrated to maximize our knowledge of this material. The standard and reference samples prepared by the International Humic Substances Society will prove to be invaluable for this purpose.

It is anticipated that the type of analysis associated with soil organic matter research will tend towards the desirable situation in which the analytical proce­dures are capable of studying organic matter in situ in the soil matrix. This is like­ly to involve instrumentation of the complexity and costliness of the type of solid­state NMR, with other nondestructive instrumental techniques also playing a major role. In the long term the future directions of soil organic matter research will be governed by the requirements of our society. As the needs for increased agricultural production and decreased environmental pollution grow, so will the need for a better understanding of the chemical and physical characteristics of soil chemistry and, hence, soil organic matter.

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ACKNOWLEDGMENT

The author is very grateful to Dr. Kaye Spark for invaluable assistance in the preparation of this chapter.

REFERENCES

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Baldock, I.A., G.J. Currie, and J.M. Oades. 1991. Organic matter as seen by solid state 13C NMR and pyrolysis tandem mass spectrometry. p. 45-{i0. In W.S. Wilson (ed.) Advances in soil organic matter research: The impact on agriculture and the environment. Proc. Jt. Symp., Univ. Essex, 3-4 Sept. 1990. R. Soc. Chern., Cambridge, England.

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Yonebayashi, K., and T. Hattori. 1990. A new fractionation of soil humic acids by adsorption chro­matography. Geoderma 47:327-336.

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