Origins of humus variation. Effects of leaching and seasonal flooding on aromaticity

Org. Geochem. Vol. 17, No. 1, pp. 85-91, 1991 Printed in Great Britain. All rights reserved

0146-6380/91 $3.00+ 0.00 Copyright © 1991 PergamonPress plc

Origins of humus variation. Effects of leaching and seasonal flooding on aromaticity RIWANDI SIHOMBING1, W. DAVID JOHNSONl, MICHAELA. WILSON1'2., MERIKEJOHNSON3, ANTHONY M. VASSALLO2 and DAVID ALDERDICE1 ~Department of Physical Chemistry, University of New South Wales, P.O. Box l, Kensington, NSW 2033, Australia 2CSIRO Division of Coal Technology, P.O. Box 136, North Ryde, NSW 2113, Australia 3New South Wales Department of Minerals and Energy, Lidcombe, NSW, Australia (Received 1 February 1990; accepted 22 June 1990)

Abstract--Humic and fulvic acids from a range of depositional environments in the Myall Lakes area of New South Wales, Australia have been examined by high resolution solid state nuclear magnetic resonance spectroscopy. For the first time definitive evidence is presented which shows that the local oxidizing/ reducing environment greatly alters the gross structural group content of the soil organic matter. In addition, swamp water located between two soils in the area has been studied. The swamp water is shown to contain tannins and be terrestrially derived. However there appears to be a contribution from microorganisms to the organic matter in the swamp sediments. Key words--humus variation, leaching, aromaticity, Myall Lakes

INTRODUCTION

occur where the water table crops out in the dune swales. The soil is a weakly podzolized uniform coarse grain sand. The materials examined during the study for their humic characteristics, included leaf litter, surface soil, sub-soil, swamp water and ground water. The samples were analysed by 13C cross polarization/ magic angle spinning (CP/MAS) spectroscopy, infrared spectroscopy, gel permeation chromatography and wet chemical analysis. Full details of the infrared and wet chemical methods can be found elsewhere (Sihombing, 1990). The objective of this study was to look for changes in the structure of humic organic matter by CP/MAS spectroscopy and relate this to the depositional environment.

Notwithstanding variations due to inadequate quantitation or differences in extraction procedures, it is apparent that nuclear magnetic resonance (NMR) spectra of humic and fulvic extracts differ considerably from each other (Wilson, 1987). In general the spectra show that fulvic acid fractions contain more carboxyl carbon and more carbohydrate. It is also significant that the fraction of carbon which is aromatic in humic extracts can vary from I0 to 74% (Hatcher et al., 1980a,b, 1981, 1989; Skjemstad et al., 1983; Schnitzer and Calderoni, 1985; Ghosal and Chian, 1985), yet there is no clear evidence as to why such variability can occur. While these facts are interesting it is important to understand the biogeochemical processes which have led to these differences (Hatcher and Spiker, 1988). In this work we studied humic materials obtained from the Myall Lakes region on the central coast of New South Wales, Australia, a region which represents a microcosm of depositional environments. The coastal system consists of Carboniferous bedrock embayments which have been filled with Quaternary sand deposits to a depth of 100-200 ft below sea level (Thorn, 1965). Within the embayments, sand barriers enclose lakes where freshwater swamp and estuarine sediments have been deposited. A dual system of Pleistocene and Holocene beach ridges trends parallel to the shoreline. The sandbeds form an unconfined aquifer, and swamps and wet heaths

EXPERIMENTAL

Sample sites

A detailed map of the area under study is shown in Fig. 1. Figure 2 shows the locations of sample sites A, B, C, E, F, G, H. Samples come from the Neranie swamp area on the eastern shore of Myall Lake (Fig. 1). These samples consist of fallen leaves (A), surface soil (B) and a hole dug into the water table (C) (Fig. 2) and are at the eastern soil boundary of the swamp. Samples E (swamp water), F (swamp sediment) and G (surface soil) come from a swamp which is sometimes dry near the shore of Myall Lake. At H swamp water continuously flows from the main swamp into Myall Lake.

*To whom correspondence should be addressed. 85

86

RIWANDISIHOMBINGet al.

SMITHS

LAKE

Bi.

(

Horse, Point'el

) \

Neranie

\ \

\

@

Bay NSW University Research Station I BUNGWAHL

EFG i

AB

~H

C

Fig. 1. Map showing area of study, Myall Lakes, NSW, Australia. (From Atkinson et al., 1981.) Preparation o f humic and fulvic acids

Humic substances were extracted from plant litter (A), from soils and sediments (B, F and G) and from swamp water (C, E and H), as follows: Washed air-dried plant litter was leached with distilled water and the leachate studied for up to 50 days by gel permeation chromatography (Sihombing, 1990). No change in molecular mass distributions were observed after 11 days. Samples from 11 days of extraction were separated as outlined below. Humic substances were extracted from soils and sediments with aqueous solutions of 0.1 M Na+P207/0.1 M NaOH by passing the basic solution through a bed of soil or sediment (500-1000 g) in a glass column. About 3-5 litres of basic solution was required for extraction over 24h. The extract was centrifuged

(500rpm for 10min) and the supernatant liquid filtered through sintered glass (porosity 4). Swamp water and plant litter extract were filtered, then rotary evaporated under vacuum at 40--45°C to about 10% of the original volume, then filtered through sintered glass (porosity 4). Humic and fulvic acids were separated and purified from these concentrated solutions of humic substances by acidifying with 1 M HCI to pH 1. Humic acid precipitated out overnight and collected by decantation and centrifugation. This precipitate was redissolved in ammonia and reprecipitated with HCI solution several times (5-7) until the mother liquors were colourless. The final precipitate of humic acids was dialysed against triply distilled water using spectraphor membrane (MW>2000) and then freeze-dried.

CROSS SECTION OF SAMPLING A R E A S

B

"

~ 'i L

.

Y. k e " ~. •,~ ~, - . _-- , : E ~ Swamp F

H

M

all

___

Fig. 2. Diagrammatic representation of sampling points. (A) Fallen leaves; (B) surface soil; (C) swamp water at the top part of the Neranie Swamp; (E) bottom end of Neranie Swamp adjacent to the lake; (F) sediment underneath swamp; (G) surface soil; (H) outlet swamp.

Origins of humus variation T a b l e I. Yields o f humic ( H A ) a nd fulvic acids ( F A ) Site

HA

FA

A B C E F G H

1.86° 1.65 a 5.3 b 27.3 b 6.19 ~ 2.93 ° 12.4b

2.37 a 0.17 a 17.9b 125.9b 1.62* 0.64* 65.3 b

q f i e l d as % mass, a ir dried. bYield as mg/I.

The supernatant and mother liquors were neutralized with ammonia, and stored for purification of fulvic acids using XAD-7 resin (Gregor and Powell, private communication, 1986). The crude fulvic acid solution, acidified to pH 1.7 and filtered, was adsorbed onto a column of XAD-7, washed of salts with HC1 (pH 1.7), and with water. The adsorbed fulvic acid was desorbed with 0.01 M KOH and the eluate passed over a cation exchange resin to obtain free fulvic acid in solution. This solution was filtered and freeze dried. Yields of humic and fulvic acids are listed in Table 1. Nuclear magnetic resonance spectroscopy

Samples were thoroughly re-freeze dried before examination by NMR. High resolution solid state spectra were obtained at 22.5 MHz on a Bruker CXP 100 instrument. The samples were spun at 3.4 kHz at the magic angle (54.7°). A 90 ° proton pulse of 5ps was used and a recycle time of 1 s. Signal intensity was obtained by cross polarizing proton polarization to carbon for 1 ms. The resultant free induction decay was collected in 1 K of data points, zero filled to 4 K and Fourier transformed using a line broadening factor of 50 Hz. Data was collected for 1-24 h depending on the quality of the spectrum obtained as determined by signal to noise ratios. Although the quantitative nature of the cross polarization method on geochemical samples is the subject of considerable debate (Snape et al., 1989; Earl et al., 1987; Wilson, 1987; Wershaw and Mikita, 1987) the best compromise for work in which time limitations exclude extensive relaxation studies is the choice of a 1-2 ms contact time between protons and carbons. A contact time of 1 ms was used in the experiments reported here. Table 2. Integration areas of ~3C N M R

87

The ~3C N M R spectra were analysed by integrating the various areas assigned to each functional group. As integration limits the natural valleys between signals were used as integration cut-off points. No absolute accuracy is claimed in these measurements which are listed in Table 2. However, they are useful for comparative purposes. In general several structural types could be delineated; ketone and aldehyde carbon resonating at or around 202 ppm, carboxyl carbon including carboxylic acid and amide carbon at or around 172ppm (these are probably hydrolysed and exist as free acid after extraction), oxygenated aromatic carbon at or around 150 ppm, protonated and carbon substituted aromatic carbon at 130ppm, and anchiomeric dioxygenated carbon of carbohydrate at 103 ppm or thereabouts. We shall offer evidence for a contribution from tannins to this resonance. Alcoholic carbons of carbohydrates were also observed at or about 70-75 ppm. Sometimes a well defined resonance from methoxyl carbon in syringyl and guaiacyl units of lignin was observed at 55 ppm. Aliphatic resonances were observed centred around 44, 30 and 21 ppm. The presence of oxygenated structures such as carbohydate and methoxyl carbon was confirmed by C--O absorptions in the infrared at 1140-1040cm -l. In the case of sample F the carbohydrate content was also measured independently (Usman, unpublished results) using classical wet chemical techniques (Lowe, 1978). RESULTS AND DISCUSSION

Figure 2 is a useful diagrammatic representation of the positions in the lake system from which samples were taken. Figure 3 shows the N M R spectra of the leaf litter fulvic and humic acids from site A. Several things are obvious from the spectra. First, the spectra differ considerably from those of whole leaves (Wilson et al., 1983; Wilson et al., 1987). There is far less alcoholic carbon of carbohydrate (,,, 73 ppm) and much more carboxyl carbon (172-173ppm). It is significant that the resonance from aryl oxy-carbon at 147-150 ppm is not larger than the methoxyl resonance at 56 ppm. Thus the iignin aromatic rings are not demethoxylated. Most of the carbohydrate present in the sample should be as non-crystalline polysaccharides or spectra from humic and fulvic acids

H u m i c acid chemical shift (ppm)

Fulvic acid chemical shift (ppm)

Sample site

199

172

A B

4.6 --

13.8 22.2

150-147 7.1 --

150-101 33.7 49.5

72-70

56-52

50-0

199

172

18.4 --

10.7 --

18.9 28.3

2.9 --

18.4 27.0

C

6.7

15.9

I0.0

38.8

E

6.2

13.2

7.8

38.4

F G H

-5,5 4.2

8.9 13.8 15.1

6.8 8.3 10.9

26.3 41.4 44.5

8.1 12.4 10.9

150-147 6.8 --

8.2

9.4

27.7

6.0

23.7

10.9

14.0

23.7

--

33.7

-8.3 16,0

56.8 24.1 13.4

-2.4 1.6

21.0 15.5 19.4

150-101

72-70

56-52

50-0

26.4 25.4

16.8 8.7

8.0 --

27.5 39.7

--

23.4

20.7

2.2

30.2

--

25.7

13.4

--

27.3

-8.7 --

23.4 24.3 28.7

12.2 10.7 15.7

-6.3 --

43.5 43.2 34.7

88

RIWANDI SIHOMBING et al.

(a)

A i

I

Jo

I

)

Chemical shift, 6 (ppm)

Fig. 3. 13C CP/MAS NMR spectra of (a) fulvic and (b) humic acid from Sample A. hydrolysed forms of cellulose. Hence it might be expected that the resonance from anchiomeric carbon (e.g. the dioxygenated carbon of the pyranose ring in glucose) should be l/Sth the size of the resonance from alcoholic carbon. It is clear that the resonance at 103-106ppm assigned to this material is much larger than this. Elsewhere (Wilson and Hatcher, 1988) we have shown that the dipolar dephasing technique in which the N M R spectrometer decoupler is turned off for a short period to remove CH 1 and CH2 carbon resonances from spectra can be used to identify tannins since they are one of the few

(a)

Chemical ,h,ft, ~ (ppm)

Fig. 5. 13C CP/MAS NMR spectra of (a) fulvic and (b) humic acid from Sample C. structures which have non-protonated carbons that resonate in this region. Dipolar dephasing showed that the 103-106ppm resonance was indeed nonprotonated. Another interesting feature of the spectra is the increased presence of carbohydate in the humic acid rather than the fulvic acid. The reverse is more commonly observed for soil organic material. As expected, the carboxyl content of the fulvic acid is higher than that of the humic acid. The ~3C CP/MAS spectra of the samples from the soil at site B are shown in Fig. 4. Surprisingly the carbohydrate content as demonstrated by the magnitude of the resonances (72 ppm) of the humic acid is insignificant and that for the fulvic acid is lower than expected. The humic acid spectrum is highly aromatic and represents the residual components of soils after extensive oxidation, possibly residual lignin components in which the aromatic rings are linked

(a)

(b)

o ~

ur)

=oo,

,oo

N

0

o

Cheml(:al shift. 6 (ppm)

Fig. 4. '3C CP/MAS NMR spectra of (a) fulvic and (b) humic acid from Sample B.

~ / ' , v ¥, " |

200 J

100 I

0 ,

Chemloel shift. ~ (ppm)

Fig. 6. '~C CP/MAS NMR spectra of (a) fulvic acid and (b) humic acid from Sample E.

Origins of humus variation via biphenyl units (Hatcher et al., 1989). Because the aliphatic component of the humic acid is so small most of the carboxylic acid carbons must be attached to aromatic rings. Spectra of aquatic organics collected from the water hole at site C are shown in Fig. 5, There is a surprising resemblance between the spectrum of the fulvic acid and that of the fulvic acid from the leaf litter [Fig. 3(a)] which strongly suggests that the leaf litter has an important input into the swamp organic matter at this point. In particular this represents evidence for incorporation of tannins into the swamp. Indeed samples taken from mid-swamp (E) have very similar N M R spectra (Fig. 6) to those obtained from the sample from the water hole at C. Thus the spectra from mid-swamp water support this conclusion. Vance and David (1989) have shown that mineral soils preferentially adsorb fulvic acids. It appears that this is not the case here because the sandy podzols contain less sites capable of bonding the carboxyl groups which are present in great concentration in the fulvic acid. Sedimentary organic matter isolated at site F however is chemically quite different and has a highly aliphatic structure. In particular, polymethylene structures are present as demonstrated by resonances at 30 ppm in the spectra (Fig. 7). The influence of the surrounding litter detritus is also clear since the 103 ppm resonance is much larger than that of the 72 ppm carbohydrate resonance [e.g. Fig. 7(b)] and there are distinct resonances in spectra for aryl-oxy structures. Methoxyl carbon is absent, so there must be a concentration of phenols in the sediment. The spectra of humic and fulvic acids from soil at site G (Fig. 8) are quite different from those from soil at site B. Carbohydrates are present and there is significantly more aliphatic carbon. In addition, there is clear evidence for the presence of lignin in the form

89

0

N

g (b) v~

200

100

L

I

~

Chemk~al shift, 6 (ppm)

Fig. 8. I~C CP/MAS NMR spectra of(a) fulvic acid and (b) humic acid from Sample G.

of methoxyl and aryl-oxy carbon. Spectra of water from the outlet, site H, shown in Fig. 9, show that some demethoxylation occurs when the organic matter is solubilized. Whereas the organic material is largely humic acid at site G it is largely fulvic acid at site H. Spectra of humic acid at site G and fulvic acid at site H show only small resonances from ary-oxy carbon which suggests that phenols are not present in high concentration.

(a)

(b) (b)

A

~~~ Chemical ihlft, 8 (ppm)

Fig. 7.13C CP/MAS NMR spectra of (a) fulvic acid and (b) humic acid from Sample F.

OO l 100 Chemical shift, 6 (ppm)

Fig. 9. ]3C CP/MAS NMR spectra of(a) fulvic acid and (b) humic acid from Sample H.

90

RIWANDI S1HObtBINOet al.

O x i d i z i n g the reducing environments

The major problem in identifying why different recent sedimentary organic deposits have different types of organic matter has been to delineate the wide range of factors which cause soil organic matter variability. Geography and climate are obvious factors but so is the type of plant life, degree of microbial activity and also transport factors, i.e. whether organic matter is transferred from one system to another by water. The system studied here sheds light on a number of these important phenomena. For example, site B is a soil which is in a highly oxidizing environment while site G represents a soil which is constantly flooded and hence, relatively, is in a predominantly reducing environment. We note that the N M R spectra of the humic acids at site B [Fig. 4(b)] are those of a highly aromatic residue in which most other functional groups have been converted to carboxyl functionality. On the other hand, humic acids from site G (Fig. 8) are quite different. They retain many of their aliphatic structures including polymethylene (30 ppm) and methoxyl and aryl-oxy structures of lignin, and tannins. There is some degree of oxidation as identified by the presence of carboxyl (172 ppm) and ketone/aldehyde functionality (193 ppm) but this is much less pronounced than at site B. It is also noteworthy that the organic matter in the swamp water resembles the humic substances extracted from the litter, and this result is supported by the fact that the organic matter from the water hole at site "C" is also similar. There is great variability in the structure of aquatic humic substances (Wilson et al., 1981; Hatcher et al., 1985; Hatcher and Orem, 1985; Little and Jacobson, 1985; Gillam and Wilson, 1985; Buddrus et al., 1989; Thorn, 1987; Lundquist et al., 1985). This variability probably reflects differing amounts and types of terrestrial input. The results shown here clearly indicate that terrestrial input dominates the organic structure. The reducing environment of the sediment at the swamp bed preserves many types of organic matter and it is significant that the organic matter at site F is highly aliphatic, and more so than soil at site B or G. Nevertheless there is some evidence for oxidation. It is impossible to accommodate all the carboxyl groups in the fulvic acid as being bonded to aromatic structures in this type of organic matter as proposed for the aromatic structures in the humic acid at site B. Thus the carboxylic groups must be bonded to an aliphatic backbone. That is, the sediment represents an intermediate in which aliphatic carbon has been oxidized to carboxylic groups but not completely to carbon dioxide. Finally some comment should be made on the origin of the polymethylene component present in spectra of the sedimentary swamp material. There are two possible origins for this material. If it is material

akin to aliphatic polymers resistance to degradation by microorganisms and detected in cuticular material then the aliphatic polymer would have to be transported through the swamp water from the surrounding terrestrial environment. There is no evidence for this supposition from the spectra of the water organics. Moreover it is difficult to see how a non-polar entity such as polymethylene should be water soluble. It is more probable that this represents microbial detritus which is known to consist of long polymethylene chains (Wilson et al., 1988; Baldock et al., 1989, 1990a--c). CONCLUSIONS

(1) Two soils from the Myall Lakes area in close proximity have been shown to contain organic matter of widely different structure. Whereas the soil subjected to seasonal flooding has a ~3C N M R spectrum consisting of a wide range of resonances from lignin, carbohydrate, polymethylene and their mildly oxidized counterparts, the other soil is highly aromatic and shows strong evidence for oxidation. (2) The organic matter in the swamp between the two soils is largely terrestrially derived, although the sediments at the bottom of the swamp are highly aliphatic and contain large quantities of polymethylene probably derived from microorganisms. REFERENCES Atkinson G., Hutchings P., Johnson M., Johnson W. D. and Melville M. D. (1981) An ecological investigation of the Myall Lakes region. Aust. J. Ecol. 6, 299-327.

Baldock J. A., Oades J. M., Vassallo A. M. and Wilson M. A. (1989) Incorporation of uniformly labelled ~3C glucose carbon into the organic fraction of a soil. Carbon balance and CP/MAS measurements. Aust. J. Soil Res. 27, 725-746. Baldock J. A., Oades J. M., Vassallo A. M. and Wilson M. A. (1990a) Solid state CP/MAS ~3C NMR analysis of particle size and density fractions of a soil incubated with uniformly labelled t3C glucose. Aust. J. Soil Res. 28, 193-212. Baldock J. A., Oades J. M., Vassallo A. M. and Wilson M. A. (1990b) Solid state CP/MAS 13C NMR analysis of bacterial and fungal cultures isolated from a soil incubated with glucose. Aust. J. Soil Res. 28, 213-215. Baldock J. A., Oades J. M., Vassallo A. M. and Wilson M. A. (1990c) Significance of microbiological activity in soils as demonstrated by solid state 13C NMR. Environ. Sci. TechnoL 24, 527-530. Buddrus J., Burba P., Herzog H. and Lambert J. (1989) Quantitation of partial structures of aquatic humic substances by one and two dimensional solution ~3C nuclear magnetic resonance spectroscopy. Anal. Chem. 61, 628~53 I. Earl W. L., Wershaw R. L. and Thorn K. A. (1987) The use of variable temperature and magic angle sample spinning in studies of fulvic acids. J. Mag. Res. 74, 264-274. Ghosal M. and Chian E. K. (1985) An evaluation of aromatic fraction in humic substances. Soil Sci. Soc. Am. J. 49, 616~18. Gilliam A. H. and Wilson M. A. (1985) GC-MS and NMR characterization of dissolved marine humic substances. Org. Geochem. 8, 15-25.

Origins of humus variation Hatcher P. G. and Orem W. H. (1985) Structural inter° relationships among humic substances in marine and estuarine sediments as delineated by CP/MAS 13C NMR. In Organic Marine Geochemistry. Am. Chem. Soc. Ser. 305, 142-157. Hatcher P. G. and Spiker E. C. (1988) Selective degradation of plant biomoleeules. In Humic Substances and their Role in the Environment (Edited by Frimmel F. H. and Christman R. F.), pp. 59-74. Wiley, Chichester. Hatcher P. G., Maciel G. E. and Dennis L. W. (1981) Aliphatic structure of humic acids; a clue to their origin. Org. Geochem. 3, 43--48. Hatcher P. G., Rowan R. and Mattingly M. A. (1980) ~H and 13C NMR of marine humic acids. Org. Geochem. 2, 77-85. Hatcher P. G., Vander Hart D. L. and Earl W. L. (1980) Use of solid-state ~3C NMR in structural studies of humic acids and humin from Holocene sediments. Org. Geochem. 2, 87-92. Hatcher P. G., Breger I. A., Maciel G. C. and Szeverenyi N. M. (1985) In Humic Substances in Soil Sediment and Water Geochemistry, Isolation and Characterisation (Edited by Aiken G. R., McKnight D. M., Wershaw R. L. and MacCarthy P.), pp. 275-302. Wiley, New York. Hatcher P. G., Schnitzer M., Dennis L. W. and Maciel G. E. (1981) Aromatieity of humic substances in soils. Soil Sci. Soc. Am. J. 45, 1089-1094. Hatcher P. G., Schnitzer M., Vassallo A. M. and Wilson M. A. (1989) The chemical structure of highly aromatic humic acids in three volcanic ash soils as determined by dipolar dephasing. Geochim. Cosmochim. Acta 53, 125-130. Little B. and Jacobus J. (1985) A comparison of two techniques for the isolation of adsorbed dissolved organic material from seawater. Org. Geochem. 8, 27-33. Lowe L. E. (1978) Carbohydrates in soil. In Soil Organic Matter (Edited by Schnitzer M. and Khan S. U.). Elsevier, Amsterdam. Lundquist K., Paxeus N., Bardet M. and Robert D. R. (1985) Structural characterization of aquatic humic substances by NMR spectroscopy. Chem. Scripta 25, 373-375. Saiz-Jimenez C. and Leeuw J. W. de (1987) Chemical structure of a soil humic acid as revealed by analytical pyrolysis. J. Anal. Appl. PyroL 11, 367-376. Schnitzer M. and Calderoni G. (1985) Some chemical characteristics of paleosol humic acids. Chem. Geol. 1985, 53, 175-184.

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Sihombing R. (1990) Ph.D. thesis, University of New South Wales School of Chemistry, P.O. Box l, Kensington NSW 2033, Australia. Skjemstad J, O., Frost R. L. and Barton P. F. (1983) Structural units in humic acids from south-eastern Queensland soils as determined by 13C NMR spectroscopy. Aust. J. Soil Res. 21, 539-547. Snape C. E., Axelson D. E., Botto R. E., Delpuech J. J., Tekely P., Gerstein B. C., Pruski M., Maciel G. E. and Wilson M. A. (1989) Quantitative reliability of aromaticity and related measurements on coals by ~3C NMR. A debate. Fuel 68, 547-560, Thorn B. G. (1963) Late quarternary costal morphology of the Port Stephens-Myall Lakes area, NSW. J. Proc. R. Soc. N S W 98, 23-36. Thorn K. A. (1987) Structural characteristics of the IHSS Suwannee river fulvic and humic acids determined by solution state C-13 NMR spectroscopy. Sci. Total Environ. 62, 175-183. Vance G. F. and David M. B. (1989) Effect of acid treatment on dissolved carbon retention by spodic horizons. Soil Sci. Soc. Am. J. 53, 1242-1247. Wershaw R. L. and Mikita M. A. (1987) N.M.R. o f Humic Substances and Coal. Techniques, Problems and Solutions. Lewis, Chelsea. Wilson M. A. (1987) N M R Techniques and Applications in Geochemistry and Soil Chemistry. Pergamon Press, Oxford. Wilson M. A. and Hatcher P. G. (1988) Detection of tannins in modern and fossil barks and in plant residues by high-resolution solid-state t3C nuclear magnetic resonance. Org. Geochem, 12, 539-546. Wilson M. A., Barron P. F. and GiUam A. H. (1981) The structure of freshwater humic substances as revealed by 13C NMR spectroscopy. Geochim. Cosmochim. Acta 45, 1743-1750. Wilson M. A., Batts B. D. and Hatcher P. G. (1988) Molecular composition and mobility of torbanite precursors. Implications for the structure of coal. Energy Fuels 2, 668-672. Wilson M. A., Heng S., Goh K. M., Pugmire R. J. and Grant D. M. (1983) Studies of soil organic matter fractions using t3C cross polarization nuclear magnetic resonance spectroscopy with magic angle spinning. J. Soil Sci. 34, 83-97. Wilson M. A., Verheyen T. V., Vassallo A. M., Hill R. S. and Perry G. J. (1987) Selective loss of carbohydrates from plant remains during coalification. Org. Geochem. 11, 265-272.