AN ABSTRACT OF THE THESIS OF
Evelyne L. Ndiaye for the degree of Master of Science in Soil Science presented on
December 15, 1998. Title: Winter Cover Cropping Effects on Integrative Biological
Indicators of Soil Quality
Redacted for Privacy
Abstract approved:
Richard P. Dick
Responses of biological indicators of soil quality to winter cover cropping were
measured on soil samples collected from 6 commercial growers' fields and two
experiment research stations in the Willamette Valley of Oregon. The research stations
were the North Willamette Research and Extension Center (Aurora, OR), and the
Oregon State University Vegetable Farm (Corvallis, OR). The research stations and
five on-farm sites compared winter cover crops or winter fallow in rotation with a
summer vegetable crop. In one on-farm site, minimum tillage or conventional till
following winter cover crops was compared. The objectives of this study were to:
1) monitor changes in soil biological properties under field managed with cover crops;
2) test potential of buried cotton strip as indicator of soil biological activity and as a soil
quality index; and 3) assess the degree of correlation between tensile strength and
cotton strip weight loss. The major findings were: 1) microbial biomass carbon and
B-glucosidase activity were the most sensitive to cover crop management; 2) cotton strip
decomposition was correlated to soil biological properties but was not very sensitive to
management changes; and 3) that measuring weight loss was nearly as effective as
tensile strength in assessing cotton strip decomposition in soils.
©Copyright by Evelyne L. Ndiaye
December 15, 1998
All Rights Reserved
Winter Cover Cropping Effects on Integrative Biological Indicators
of Soil Quality
by
Evelyne L. Ndiaye
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Master of Science
Presented December 15, 1998
Commencement June 1999
Master of Science thesis of Evelyne L. Ndiaye presented on December 15, 1998
APPROVED:
Redacted for Privacy
Major Professor, representing Soil Science
Redacted for Privacy
Dead of the Department of Crop and Soil Science
Redacted for Privacy
Dean /Graduate S
of
I understand that my thesis will become part of the permanent collection of Oregon State
University libraries. My signature below authorizes release of my thesis to any reader
upon request.
Redacted for Privacy
lE
yne Lydie
diaye, Author
ACKNOWLEDGEMENTS
I would like to express my gratitude to all those who contributed directly or
indirectly to my thesis project. It is the product of many hands and minds without
which it would not be possible.
I thank my major professor and leader of the Soil Quality Project, Dr. Richard
Dick, for his support, guidance, help in the field and inputs in data interpretation.
I also thank Dr. Lynda Ciuffiti, Dr. Dave Myrold and Dr. Jack Stang for serving
on my committee and for their contributions to the final product.
I thank Dr. Hsiou-Lien Chen for the use of the Instron tensiometer.
I express my sincere appreciation to ISRA (Institut Senegalais de Recherche
Agricole) and to the NRBAR (Senegal Natural Resources-Based Agricultural Research)
Project for sponsoring me these three years.
Thanks go also to the USDA (United States Department of Agriculture) and to
the SARE (Sustainable Agricultural Research and Education) Program for funding the
Soil Quality Project, and to all the commercial growers involved in this study. It was
quite an experience to work on their fields considering the Willamette Valley weather.
Thanks to the graduate students and to the staff of the Soil Science division,
particularly Jayne Smith, Pam Wegner and Tracy Mitzel.
Special thanks go to Joan Sandeno for her help in the lab, on the field and all the
treats at the Silverton bakery; and to Bob Christ and Beto Sattell for all their help with
my data.
Thanks go to my officemates Gilbert Buller, Hudson Minshew, Mary Schutter,
for all the jokes that kept my spirit up; and to my dear friends in Corvallis and in
Senegal for their help, love and attention during these years.
Finally, I want to express my deepest gratitude to my father Eugene Ndiaye, my
brothers and sisters for their love and support; and to my uncle Maurice Ndiaye and his
family for their love and prayers. God Bless you all!
TABLE OF CONTENTS
Page
Chapter 1. LITERATURE REVIEW
1
Introduction
2
Soil quality
3
Cover crops
7
Soil Biological properties
8
Literature Cited
Chapter 2. WINTER COVER CROP EFFECTS ON MICROBIAL
BIOMASS, ENZYME ACTIVITIES, AND COTTON STRIP
DECOMPOSITION
13
19
Abstract
20
Introduction
20
Materials and Methods
24
Results
32
Discussion
46
Literature Cited
51
TABLE OF CONTENTS (Continued)
Page
Chapter 3. TILLAGE EFFECT ON SOIL BIOLOGICAL PROPERTIES
56
Abstract
57
Introduction
57
Materials and Methods
58
Results and Discussion
60
Literature Cited
67
Chapter 4. COTTON STRIP ASSAY AS A SOIL BIOLOGICAL
INDICATOR: A LABORATORY EXPERIMENT
69
Abstract
70
Introduction
70
Materials and Methods
72
Results
75
Discussion
83
Literature cited
86
Chapter 5. SUMMARY
89
BIBLIOGRAPHY
92
APPENDICES
102
LIST OF FIGURES
Figure
2.1
2.2
2.3
2.4
2.5
2.6
2.7
2.8
2.9
2.10
3.1
3.2
Page
Winter cover crop effects on MBc at on-farm research sites
(* indicates significant treatment effect at P < 0.05).
33
Winter cover crop effects on MBC on experiment research sites
(* indicates significant treatment effect at P < 0.05).
35
Winter cover crop effects on soil respiration at on-farm research sites
(* indicates significant treatment effect at P < 0.05).
37
Winter cover crop effects on soil respiration on experiment research
sites (* indicates significant treatment effect at P < 0.05).
38
Winter cover crop effects on arylsulfatase and 13- glucosidase activities at
on-farm research sites (* indicates significant treatment effect at
P < 0.05).
40
Winter cover crop effects on arylsulfatase activity on experiment
research sites (* indicates significant treatment effect at P < 0.05).
41
Winter cover crop effects on B-glucosidase activity on experiment
research sites (* indicates significant treatment effect at P < 0.05).
42
Winter cover crop effects on cotton strip decomposition at on-farm
research sites. Within each day, treatments bars followed by the same
letter are not significantly different (P < 0.05).
43
Winter cover crop effects on cotton strip decomposition at NWREC.
Within each day, treatments bars with the same letter are not
significantly different (P < 0.05).
44
Winter cover crop effects on cotton strip decomposition at Vegetable
Farm. Within each day, treatments bars with the same letter are not
significantly different (P< 0.05)
45
Fig 3.1. Tillage effect on MBc and soil respiration at on-farm research
site (* indicates significant treatment effect at P < 0.05).
61
Tillage effect on mineralizable N at on-farm research sites (* indicates
significant treatment effect at P < 0.05).
63
LIST OF FIGURES (Continued)
3.3
3.4
4.1
4.2
4.3
4.4
4.5
4.6
Page
Tillage effect on arylsulfatase and B-glucosidase activity at on-farm
research site (* indicates significant treatment effect at P < 0.05).
64
Tillage effect on cotton strip decomposition at on farm research site.
Within each day, treatment bars with the same letter are not
significantly different (P < 0.05).
66
Effect of temperature, soil moisture and N rate on cotton strip
decomposition in a Chehalis silt loam soil. Bars followed by the same
letter within a soil moisture, temperature and sampling time were not
significantly different at P < 0.05.
76
Effect of temperature, soil moisture and N rate on MBc in a Chehalis
silt loam soil. Bars followed by the same letter within a soil moisture,
temperature and sampling time were not significantly different at
P < 0.05.
77
Effect of temperature, soil moisture and N rate B-glucosidase activity in
a Chehalis silt loam soil. Bars followed by the same letter within a soil
moisture, temperature and sampling time were not significantly different
at P < 0.05.
79
Effect of temperature, soil moisture and N rate on cotton strip
decomposition in Willamette silt loam soil. Bars followed by the same
letter within a soil moisture, temperature and sampling time were not
significantly different at P < 0.05.
80
Effect of temperature, soil moisture and N rate on MBc in a Willamette
silt loam soil. Bars followed by the same letter within a soil moisture,
temperature and sampling time were not significantly different P < 0.05.
81
Effect of temperature, soil moisture and N rate B-glucosidase activity in
a Willamette silt loam soil. Bars followed by the same letter within a
soil moisture, temperature and sampling time were not significantly
different at P < 0.05.
82
LIST OF TABLES
Table
2.1
Page
Winter cover crop and crop rotation on-farm research sites and
experiment research stations.
27
Chemical characteristics of soils from on-farm and experiment
research sites at canopy closure in 1996.
29
Treatment effects on mineralizable N at on-farm and experiment
research sites
36
Matrix of Pearson correlation coefficients for 11 of the measured soil
biochemical parameters.
47
3.1
Physical chemical properties of soil in 1996.
59
3.2
Pearson coefficients (r) for association between biological parameters
and Total C.
67
Analysis of variance of soil environment effects on cotton strip
decomposition on a Chehalis silt loam soil.
75
Analysis of variance showing source of effects on cotton strip
decomposition on a Willamette silt loam soil.
78
2.2
2.3
2.4
4.1
4.2
LIST OF APPENDICES
Appendix
A
B
C
Page
Winter cover crop effects on MBA at each on farm-site
(* shows significance at P < 0.05).
103
Winter cover crop effects on B-glucosidase activity at each on-farm
site (* shows significance at P < 0.05).
104
Winter cover crop effects on arylsulfatase activity at each on- farm
site (* shows significance at P < 0.05).
105
I dedicate this thesis to:
My mother Marie Jeanne Gomis
My grandmother Anne Marie Sylla.
WINTER COVER CROPPING EFFECTS ON INTEGRATIVE BIOLOGICAL
INDICATORS OF SOIL QUALITY
CHAPTER 1
LITERATURE REVIEW
2
INTRODUCTION
Food production depends on many social, economic, and political factors that
affect farmers' motivation and ability to produce. Food supply also is affected by
physical and biological components such as availability of natural resources (especially
water), technologies and supply of production inputs (manure, lime, fertilizers,
irrigation water etc.) (Brady and Weil, 1996). Ongoing population growth has resulted
in increased demand for food and fiber, which has caused growers to farm more
intensively to maintain increased food production. Unfortunately, this rapid increase in
food production has declined these past few years due to economic and environmental
reasons. As society approaches the 21st century, policymakers and the scientific
community are becoming aware of the crucial need to develop more sustainable
agricultural systems. Commercial growers, who play a key role in food supply, are
testing various management practices to improve soil physical, chemical and biological
properties. However, these systems should be increasing or maintaining crop yields.
Adoption of management practices such as crop rotation, conservation tillage or cover
cropping are being applied to improve soil health by reducing soil erosion, compaction
and nutrient leaching. These practices are believed to contribute to a significant
increase in soil quality.
Farmers have intuitive abilities to sense changes in soil quality over time
because of their close association with their fields. However, it is necessary to
determine quantifiable early indicators of soil health and define management practices
that will improve soil quality and sustain productivity. In response to this, a
multidisciplinary project to investigate alternative agricultural systems in relation to
3
arthropods and soil quality dynamics was initiated in 1996. The project focused on the
intensively managed vegetable row crop system of western Oregon. The objectives of
this project were to:
1- Identify and explore sensitive early indicators of changes in soil quality
useful to agroecosystem analysis and farm management;
2- Identify and explore linkages between changes in soil management
[vegetation/soil management and associated carbon (C) inputs] with soil
community structure and processes, and above-ground arthropod dynamics;
3- Test and adopt strategies for conducting participatory research and education
programs.
The objectives of my part of the project were to:
a) Identify simple biological indicators of soil quality that are sensitive to changes in
soil management of cover crops;
b) Test potential of buried cotton strip as an indicator of soil biological activity and as
a soil quality index.
We hypothesize that conservation management, such as winter cover crop or reduced
tillage, would improve soil biological properties by increasing soil enzyme activity and
microbial biomass, and in the long term improve soil quality. It would be desirable to
have indicators of soil quality that respond to management within a short period of time.
SOIL QUALITY
Soil might be defined differently according to disciplinary and philosophical
considerations. A geologist might define soil as the eroded surface of rocks, whereas an
engineer might define soil in terms of soil physical characteristics such as compressibil­
4
ity or permeability to water. The soil scientist or agronomist might define soil as the
unconsolidated cover of the earth, made up of mineral, organic material, water, and air,
and capable of supporting plant growth. The quality of a soil is its ability to perform
various functions which includes: (i) provide a medium for plant growth, (ii) regulate
and partition water flow in environment and (iii) serve as an environmental buffer in the
formation, attenuation and degradation of environmentally hazardous compounds
(Larson and Pierce, 1991). Because soil is a non-renewable natural resource, it is crucial
to increase or at least maintain its quality for future generations.
Publication of the book Soil and Water Quality: An Agenda for Agriculture
(National Research Council, 1993) by the National Academy of Science brought
national attention to the concept and importance of soil quality in natural resource
management. Doran and Parkin (1994) define soil quality as "the capacity of the soil to
function within ecosystem boundaries, maintain environmental quality, and promote
plant, animal, and human health". The quality of the soil is therefore a fundamental
factor to sustain biological productivity on the earth.
Determination of the ability of soil to perform functions is based on measuring
key soil physical, chemical and biological properties. A single indicator or suites of
indicators are chosen according to the particular soil function of interest. For example,
agronomic indicators for crop growth would likely be different than for determining the
potential of a soil to be a buffer of pollutants in the environment.
Studies from long- or short-term experiments indicate that management
practices such as crop rotation or minimum tillage can improve soil physical, chemical
and biological properties (Doran et al., 1994). These studies suggest there is potential
5
to develop soil quality indicators that would be useful to guide landmanagers in
adopting more sustainable management systems.
Crop rotation
Long-term continuous monocropping often results in an increase of soilborne
diseases, a decrease in fertility, yields and a general depletion of soil quality (lower
organic matter and increased compaction) (Paul et al., 1997). Short- or long-term
rotations of diverse crops have improved soil properties (Campbell, 1978; Haynes et al.,
1991). Crop rotation and associated residues returned to soil influence soil organic
matter (SOM). Various studies have shown the beneficial effect of crop rotation,
especially with legumes, on SOM and crop productivity (Odell et al., 1984; Johnston,
1986; Omay et al., 1997). These studies generally demonstrated that rotation of nonlegume grain with forage legumes increased organic C and nitrogen (N) over
continuous grain crop produced without manure or fertilizer-N inputs after several
decades of cropping. Crop rotations including high-residue-producing crops, such as
corn, and addition of fertilizer increased soil organic C and N (Omay et al., 1997).
Other studies reported that crop rotation has significantly greater levels of
microbial biomass (McGill et al., 1986) and soil enzyme activities (Khan, 1970; Dick,
1984) than do continuous cropping systems.
Tillage
The industrial revolution had a big impact on agricultural systems. The
mechanization of agriculture and use of external inputs, especially fertilizers, led to
considerable increases in yields, but this also has had negative impacts on the soil
6
resource base. The conventional tillage (CT) system uses the moldboard plow to bury
plant residues, and additional tillage operations are performed for seedbed preparation
and weed control. Because of the loss of SOM and increased soil erosion due to
conventional tillage systems, a range of conservation tillage practices has been
developed for specific cropping systems and regions. In the no-tillage (NT) systems,
tillage is limited to the opening of a small slot for seed placement and fertilizer
application. Minimum tillage or NT practices are used more by commercial growers
because it reduces soil erosion while maintaining profitable yields. Soil cultivation,
besides affecting its structure and chemistry, also affects its biology.
Conventional cultivation destroys macroaggregates' structure and produces loss
of structural stability due to the greater exposure of the formerly protected soil organic
matter fraction (Tisdall and Oades, 1982). Biological activity is reduced because
macroaggregates provide important microhabitats to soil microbes. Retention of crop
residues, however, increases the population of heterotrophic bacteria, fungi and
actinomycetes (Doran, 1980; Gupta and Roper, 1992).
In a short-term tillage experiment, Carter (1986) showed that microbial biomass
carbon (MBA) quickly changes in response to tillage before any change in total C was
measured. Higher microbial populations often are associated with higher nutrient
availability for plant growth. Greater potentially mineralizable N and microbial
biomass were reported in the 0-7.5 cm depth under NT system compared to CT (Doran,
1980).
7
COVER CROPS
Cover crop and soil quality
Sustainable agricultural systems favor the integration of winter cover crops into
vegetable production systems, especially in the Pacific Northwest. Cover crops were
used years ago by farmers as green manure and a nutrient source for a subsequent crop.
The increasing use of inorganic fertilizer since the 1950's led to a decrease in use of
cover crops. The positive effect of planting a cover crop during the winter fallow
season has been recognized by many scientists. When well established, winter cover
crops reduce soil erosion (Smith et al., 1987; Reeves, 1994) and nitrate leaching (Lamb
et al., 1985; Hargrove, 1991; Brandi-Dohrn et al., 1997), and enhance SOM and N
storage (Stivers and Sherman, 1991; Harper et al., 1995). Winter cover crops can
reduce soil erosion by returning residues to soil which improves its physical properties
such as water stable aggregation and water infiltration (Tisdall and Oades, 1982;
Roberson et al., 1991). Cover crops increase cropping intensity and add organic
residues to soil, which enhances microbial activity (Powlson et al., 1987).
Cover crops can be legume, non-legume or a combination of both. Legume
cover crops are preferred because of their ability to fix N and supply substantial
amounts of N for the subsequent crop (Heichel and Barnes, 1984; Smith et al., 1987;
Burket et al., 1997).
Non-legume cover crops such as rye have been found effective in capturing
residual N after harvest of the summer crops (Martinez and Giraud, 1990; Ditsch et al.,
1993). A more controlled study recently showed that a cereal rye cover crop recovered
an average of 40 kg N ha -1 but there was no evidence that the N in the cereal cover
8
crop tissue would reduce inorganic N requirements for the subsequent summer crops
(Burket et al., 1997).
Disadvantages
Winter cover crops may provide a significant benefit to soil and water quality
but also require good management practices to avoid possible disadvantages. Cover
crops can have various effects on soilborne diseases by hosting pathogens (Dillard and
Grogan, 1985). Plowing cover crop residues may provide organic C encouraging
pathogen growth (Phillips et al., 1971). High C:N ratio cover crop residues enhance N
immobilization, which may delay crop growth (Wyland et al., 1995). Cover crops have
monetary and labor costs that must be compared to traditional soil management
systems. Additional management factors must be considered such as fall planting and
spring biomass management of cover crops.
SOIL BIOLOGICAL PROPERTIES
Microbial biomass
Soil microbial biomass is an important component of SOM because it regulates
the transformation and storage of nutrients (Horwath and Paul, 1994). Although
accounting for only 1-3% of soil organic C, it is the "eye of the needle" through which
nearly all soil organic inputs must pass (Jenkinson, 1977). During this process, organic
materials are transformed by microorganisms to generate energy and produce new
cellular metabolites to support their growth (Martens, 1995), and simultaneously release
plant-available forms of nutrients. The soil microbial population is actively involved in
9
the mineralization of organic substances or immobilization of inorganic forms of
nutrients and therefore plays a key role in nutrient cycling.
Soil microbial biomass can be estimated by plate counts, direct observation,
chloroform fumigation-incubation, chloroform fumigation-extraction or substrate-
induced respiration (SIR) methods. Because of its simple procedure and lower
equipment costs, the fumigation-incubation technique (Jenkinson and Powison, 1976)
often has been used. The activity of the microbial biomass, however, must be measured
by other methods, such as respiration or soil enzyme assays.
Management practices, such as fertilization, cover crops, crop rotation and
conventional intensive tillage, may have a direct impact on soil MB. Martyniuk and
Wagner (1978) reported higher microbial populations on fertilized soil than on
unfertilized soil. Bolton et al. (1985) showed soil MB and enzyme activity increased in
soil amended with legume green manure over unfertilized soil. Microbial biomass also
is affected by soil pH, soil water content and texture. Soil texture may have a greater
influence on MB because of its importance in aggregate formation. Aggregates protect
organic C and indirectly enhance buildup of organic matter in soil. Several authors
demonstrated that aggregate stability is strongly related to MBc (Haynes and Swift,
1990; Roberson et al., 1991). Comparing a native grassland soil with an adjacent site
that has been cultivated for 69 years, Gupta and Germida (1988) showed that the
amount of MBc and MBN was greater in the macroaggregates than microaggregates.
Decreased tillage intensity increases the proportion of MBc to organic C (Lynch
and Panting, 1980; Carter, 1991). Because soil MB has a rapid turnover, integrates both
physical and chemical properties, and responds quickly to changes in management
10
practices, it may be considered a suitable biological indicator of soil quality (Rice et al.,
1996).
Soil enzymes
Enzymes are biological catalysts and specific to the reactions they catalyze. Soil
enzyme activities result from the activity of accumulated enzymes and from the
enzymatic activity of proliferating microorganisms (Kiss et al., 1975). Soil enzymes
originate from microorganisms and also from plants and animals. Enzymes strongly
influence nutrient cycling because of their ability to catalyze biochemical reactions
mediated by microorganisms, plant roots and animals. They are located intracellularly
within viable cells, extracellularly at the surface of cells or excreted into soil solution or
are associated with humic colloids in soil. Free enzymes result from exo- and endo­
enzymes originating from living cells and disintegrated cells, respectively. Soil
enzymes are found in solution, in suspension, adsorbed or complexed to humic
substances. Enzymes are believed to be relatively stable (Burns, 1982) and have been
suggested as a potential indicator of soil quality because of their rapid response to soil
management (Dick, 1994).
Soil enzymes have been related to soil biology for a long time, but more recent
studies have shown that soil enzymes are strongly correlated to soil microbial activity
(Frankenberger and Dick, 1983). A comparison of on-farm cropping systems in an
alternative system that included a legume green manure showed improvement of soil
structure (Reganold, 1988) and an increase of soil enzyme activities (Bolton et al.,
1985). Other studies have shown soil enzyme activities to be responsive to tillage
11
management. Gupta and Germida (1988) compared cultivated soils (69 yrs) with
adjacent grassland and reported that cultivation depressed arylsulfatase activity by 65%.
Frankenberger and Dick (1983) analyzed the relationship of 11 enzymes in 10
unamended, diverse soils with microbial respiration, biomass and viable plate counts.
They found that alkaline phosphatase, amidase and catalase were highly correlated
with both microbial respiration and total biomass in soil but not with microbial plate
counts. These results indicate that enzyme activities were associated with active, soil
microorganisms whose number were not reflected accurately by the viable plates
counts. In addition, enzyme activities are strongly correlated with soil organic C and
total N content (Dick et al., 1988).
Cotton strip decomposition
Litter bags have been one of the practical methods used to investigate organic
residue decomposition. Due to the important effects of substrate quality on litter
decomposition (Coulson and Butterfield, 1978; Swift et al., 1979), use of a standard
substrate was more appropriate for comparing decomposition among different sites
(Heal et al., 1981).
The history of the cotton strip assay started in the textile industry to test the
efficiency of fungicides. The test consisted of burying a strip of textile in a tank of soil
over a period of time (Wade, 1947; Barr, 1988). After the burial period, the tensile
strength loss of the strip was used to assess decomposition of the fabric and, thus, the
efficiency of the fungicide. As the test was used more widely, it became necessary to
develop a cotton standard that would minimize variations among textile fabrics.
Several standard substrates were previously used (Rovira, 1953; Heal et al., 1974)
12
before the Shirley Institute (Manchester, England) conceived the "Shirley Soil Burial
Test Fabric" in
1976.
It is a 100% unbleached combed cotton with colored threads.
Decomposition of plant residues is a biological process of great interest to
scientists studying soil processes. Cellulose provides an important source of energy and
C to a wide variety of organisms. Thus, the cotton strip assay was very attractive in
comparing the rate of cellulose breakdown in different soil types because of its
uniformity. Walton and Allsopp
(1977)
reported this technique to be a powerful tool
for field research if used for comparative assessments of biological activity in soils.
The International Biological Programme (Latter and Walton,
1988)
later used the cotton
strip assay to assess decomposition rates in a variety of soil systems. To justify the use
of cotton strip assay in ecological studies, French (1988) related two concepts:
1.
Cotton strip tensile strength loss (CTSL) is a good index of cellulose decomposition
rate in an ecosystem and,
2. Cellulose decomposition rate can be used as a general index of the processes of
organic matter decomposition and nutrient release.
The cotton strip assay has been used in several ecological studies, but little
information is available on its use as an index of microbial activity and management
effects in agricultural soils. In their study assessing cellulose decomposition in heavy
metal-contaminated sewage sludge-amended soils, Obbard and Jones (1993) found
lower decomposition rates in plots with elevated metal concentration than with
uncontaminated sludge-amended and unsludged controls. Williamson (1994) showed a
positive relationship between cotton strip decomposition and soil pH, cation exchange
capacity, water-stable aggregates and MBc. Further, Trettin et al. (1996) reported that
13
cellulose decomposition increased at each soil depth with increasing soil disturbance
with bedding, trenching and whole-tree harvest compared to the control. Temperature,
moisture and available N have been recognized to affect decomposition of cellulose
(Lande, 1969; Donnelly et al., 1990; Trettin et al., 1996).
LITERATURE CITED
Barr, A.M.R. 1988. The colonization and decay of cotton by fungi in soil burial tests
used in the textile industry. p 50-54. In A.F. Harrison, P.M. Latter and D.W.H.
Walton et al. (eds.) Cotton strip assay: an index of decomposition in soils.
Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Bolton, J., L.F. Elliott, P.R. Papendick, and D.F. Bezdicek. 1985. Soil microbial
biomass and selected soil enzyme activities: Effect of fertilization and
cropping practices. Soil Biol. Biochem. 17: 297-302.
Brady, N.C., and R.R. Weil. 1996. The nature and properties of soils. 11 ed. Prentice
Hall Upper Saddle River, New Jersey.
Brandi-Dohrn, F.M., R.P. Dick, M. Hess, S.M. Kauffman, D.D. Hemphill, Jr., and J.S.
Selker. 1997. Nitrate leaching under cereal rye cover crop. J. Environ. Qual.
26:181-188.
Burket, J.Z., D.D. Hemphill, Jr., and R.P. Dick. 1997. Winter cover crops and nitrogen
management in sweet corn and broccoli rotations. HortSci. 32: 664-668.
Burns, R.G. 1982. Enzyme activity in soil: location and a possible role in microbial
ecology. Soil Biol. Biochem. 14: 423-27.
Campbell, C.A. 1978. Soil organic carbon, nitrogen and fertility. Dev. Soil Sci. 8:173­
271.
Carter, M. R. 1986. Microbial biomass as an index of tillage induced changes in soil
biological properties. Soil Till. Res. 7: 29-40.
Carter, M.R. 1991. The influence of tillage on the proportion of organic carbon and
nitrogen in the microbial biomass of medium textured soils in a humid climate.
Biol. Fertil. Soils. 11:135-139.
Coulson, J.C. and J. Butterfield. 1978. An investigation of the biotic factors determining
the rates of plant decomposition on blanket bogs. J. Ecol. 66:631-650.
14
Dick, R.P., P.E. Rasmusssen, and E.A. Kerle. 1988. Influence of long-term residue
management on soil enzyme activities in relation to soil chemical properties of
a wheat-fallow system. Biol. Fertil. Soil. 6:159-164.
Dick, R.P. 1994. Soil enzyme activities as indicators of soil quality. p. 107-124. In J.W.
Doran et al. (eds.) Defining soil quality for a sustainable environment. SSSA
Spec. Publ. 35. Madison, WI.
Dick, W.A. 1984. Influence of long-term tillage and crop rotation combinations on
soil enzyme activities. Soil Sci. Soc. Am. J. 48:569-574.
Dillard, H.R. and R.G. Grogan. 1985. Influence of green manure crop and lettuce on
sclerotial populations of Sclerotinia minor. Plant Dis.. 69:579-582.
Ditsch, D.C., M.M. Alley, K.R. Kelley, and Y.Z. Lei. 1993. Effectiveness of winter
rye for accumulation residual fertilizer N following corn. J. Soil and Water
Cons. 48:125-132.
Donnelly, P.K., J.A. Entry, D.L. Crawford, and K. Cromack Jr. 1990. Cellulose and
lignin degradation in forest soils: response to moisture, temperature and acidity.
Microb. Ecol. 20: 289-295.
Doran, J.W. 1980. Soil microbial and biochemical changes associated with reduced
tillage. Soil Sci. Soc. Am. J. 44:765-771.
Doran, J.W., D.C. Coleman, D.F. Bezdicek, and B.A. Stewart. 1994. Defining soil
quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
Doran, J.W. and T.B Parkin. 1994. Defining and assessing soil quality. p 3-21.
In J.W. Doran, et al. (eds.) Defining soil quality for a sustainable environment.
SSSA Spec. Publ. 35. Madison, WI.
Frankenberger Jr., W.T. and W.A. Dick. 1983. Relationships between enzyme
activities and microbial growth and activity indices in soil. Soil Sci. Soc. Am.
J. 47:945-951.
French, D.D. 1988. Patterns of decomposition assessed by the use of litter bags and
cotton strip assay on fertilized and unfertilized heather moor in Scotland.
p. 100-108. In A.F. Harrison, P.M. Latter and D.W.H. Walton et al. (eds.)
Cotton strip assay: An index of decomposition in soils. Institute of Terrestrial
Ecology, Grange-Over-Sands, Cumbria, England.
Gupta, V.V.S.R. and J.J. Germida. 1988. Distribution of microbial biomass and its
activity in different soil aggregate size classes as affected by cultivation. Soil
Biol. Biochem. 20:777-786.
15
Gupta, V.V.S.R. and M.M. Roper. 1992. Influence of crop residues management
practices on microbial biomass and nutrient availability. Australian
Microbiology. 1:3-21.
Hargrove, W.L. 1991. Cover crops for clean water. Soil Water Conserv. Soc.
Ankeny, IA.
Harper, L.A., P.F. Hendrix, G.W. Langdale, and D.D. Coleman. 1995. Clover
management to provide optimum nitrogen and soil water conservation. Crop
Sci. 35:176-182.
Haynes, R.J and R.S. Swift. 1990. Stability of soil aggregates in relation to organic
constituent of soil water content. J. Soil Sci. 41:73-83.
Haynes, R.J., R.S. Swift, and R.C. Stephen. 1991. Influence of mixed cropping
rotations (pasture-arable) on organic matter content, water stable aggregation
and clod porosity in a group of soils. Soil Tillage Res. 19:77-87.
Heal, O.W., P.W. Flanagan., D.D. French and S.F. Maclean, Jr. 1981. Decomposition
and accumulation of organic matter. p. 587-634. In L.C. Bliss, O.W. Heal, and
J.J. Moore (eds.) Tundra ecosystems: a comparative analysis. Cambridge
Univer. Press, Cambridge, England.
Heal, O.W., G. Howson, D.D. French and J.N.R. Jeffers. 1974. Decomposition of
cotton strips in tundra. p. 341-362. In A.J. Holding, O.W. Heal, S.F. MacLean
and P.W. Flanagan (eds.) Soil organisms and decomposition in tundra.
Stockholm: Tundra Biome Steering Committee.
Heichel, G.H., and D.K. Barnes. 1984. Opportunities for meeting crop nitrogen needs
from symbiotic nitrogen fixation, In D. Bezdicek and J. Power (eds.) Organic
farming: current technology and its role in a Sustainable Agriculture. Spec.
Publ. 46. Madison. WI.
Horwath, W.R. and E.A. Paul. 1994. Microbial Biomass. p. 753-773. In R.W. Weaver
et al. (eds.) Methods of Soil Analysis, Part 2. Microbiological and Biochemical
Properties. SSSA Book Ser. 5. SSSA, Madison, WI.
Jenkinson D.S. 1977. The soil biomass. NZ Soil News 25.
Jenkinson, D.S. and D.S. Powlson. 1976. The effect of biocidal treatments on metabol­
ism in soil-V. A method for measuring soil biomass. Soil Biol. Biochem.
8:209-213.
Johnston, A.E. 1986. Soil organic matter, effects on soils and crops. Soil Use Manage.
2:97-105.
16
Khan, S.U. 1970. Enzymatic activity in gray wooded soil as influenced by cropping
systems and fertilizers. Soil Biol. Biochem. 2:137-139.
Kiss, I., M. Dragan-Burlarda and D. Radulescu. 1975. Biological significance of
enzymes in soil. Adv. Agron. 27:25-87.
Lande, E. 1969. Biological activity in some natural and drained peat soils with special
reference to oxidation-reduction conditions. Acta For. Fenn. 94:1-69.
Lamb, J.A., G.A. Peterson, and C.R. Fenster. 1985. Fallow nitrate accumulation in a
wheat-fallow rotation as affected by tillage system. Soil Sci. Soc. Am. J.,
49:1441-1446.
Larson, W.E., and F.J. Pierce. 1991. Conservation and enhancement of soil quality. In
Evaluation of sustainable land management in developing world. Vol. 2.
IBSRAM Proc. 12(2). Bangkok, Thailand. Int. Board for Soil Res. Management.
Latter, P.M. and D.W.H. Walton. 1988. The cotton strip assay for cellulose decomposi­
tion studies in soil: history of the assay and development. p. 7-10. In A.F.
Harrison, P.M. Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of
decomposition in soils. Institute of Terrestrial Ecology, Grange-Over-Sands,
Cumbria, England.
Lynch, J.M., and L.M. Panting. 1980. Cultivation and the soil biomass. Soil Biol.
Biochem. 12:29-33.
Martens, R. 1995. Current methods for measuring microbial biomass C in soil:
Potentials and limitations. Biol. Fertil. Soils. 19:87-89.
Martinez, J. and G. Giraud. 1990. A lysimeter study of the effect of a ryegrass catch
crop during a winter wheat/maize rotation, on nitrate leaching and on the
following crop. J. Soil Sci. 41:5-16.
Martyniuk, S., and G.H. Wagner. 1978. Quantitative and qualitative examination of
soil microflora associated with different management systems. Soil Sci.
125:343-350.
McGill, W.B., K.R. Cannon, J.A. Robertson and F.A. Cook. 1986. Dynamics of soil
microbial biomass and water soluble organic C in Breton L after 50 years of
cropping to two rotations. Can. J. Soil Sci. 66:1-19.
National Research Council. 1993. Soil and water quality: An agenda for agriculture.
Natl. Acad. Press, Washington, D.C.
17
Obbard, J.P., and K.C. Jones. 1993. The use of the cotton-strip assay to assess
cellulose decomposition in heavy metal-contaminated sewage sludge-amended
soils. Environmental Pollution. 81:173-178.
Odell, R.T., S.W. Melsted, and W.M. Walker. 1984. Changes in organic carbon and
nitrogen of Morrow Plots soils under different treatments, 1904-1973. Soil Sci.
137:160-171.
Omay, A.B., C.W. Rice, L.D. Maddux and W.B. Gordon. 1997. Changes in soil
microbial and chemical properties under long-term crop rotation and
fertilization. Soil Sci. Soc. Am. J. 61:1672-1678.
Paul, E.A., K. Paustian, E.T. Elliot, C.V. Cole (eds.). 1997. Soil organic matter
intemperate agroecosystems: long-term experiments in North America. CRC
Press, Inc.
Phillips, D.J., A.G. Watson, A.R. Weinhold, and W.C. Snyder. 1971. Damage of lettuce
seedlings related to crop residue decomposition. Plant Dis. Rep. 55:837-841.
Powlson, D.S., P.C. Brookes, B.T. Christensen. 1987. Measurements of soil microbial
biomass provides an early indication of changes in total soil organic matter due
to straw incorporation. Soil Biol. Biochem. 19:159-164.
Reeves, D.W. 1994. Cover crops and rotations. p. 125-172. In J.C. Hatfield and B.A.
Steward (eds) Advances in Soil Science: Crop residues management. Lewis
Publishers, Boca Raton, Florida.
Reganold, J.P. 1988. Comparison of soil properties as influenced by organic and
conventional farming systems. Am. J. Ahern. Agric. 3:144-155.
Rice, C.W., T.B. Moorman and M. Beare. 1996. Role of microbial biomass carbon
and nitrogen in soil quality p. 203-215. In J.W. Doran and A.J. Jones (eds.)
Methods of assessing soil quality. SSSA, Spec. Publ. 45. Madison, WI.
Roberson, E.B., S. Sang and M.K. Firestone. 1991. Cover crop management of
polysaccharide-mediated aggregation in an Orchard soil. Soil Sci. Soc. Am. J.
55:734-739.
Rovira, A.D. 1953. A study of the decomposition of soil organic matter in red soils of
the Lismore district. Aust. Conf. Soil Sci., Adelaide, 1, 3.17, 1-4.
Smith, M.S., W.W. Frye and J.J. Varco. 1987. Legume winter cover crops. Adv. Soil
Sci. 7:95-139.
Stivers, L.J., and C. Shennan. 1991. Meeting the nitrogen needs of processing
tomatoes through winter cover cropping. J. Prod. Agric. 4:330-335.
18
Swift, M.J., O.W. Heal and J.M. Andersen. 1979. Decomposition in terrestrial eco­
ystems. University of California Press, Berkley.
Tisdall, J.M., and J.M. Oades. 1982. Organic matter-stable aggregates in soils. J. Soil
Sci. 33:141-163.
Trettin, C.C. M. Davidian, M.F. Jurgensen, and R. Lea. 1996. Organic matter decompo­
sition following harvesting and site preparation of forest wetland. Soil Sci. Soc.
Am. J. 60:1994-2003.
Wade, G.C. 1947. Effect of some microorganisms on the physical properties of cotton
duck. J. Coun. Scient. Ind. Res. Aust. 20:459-467.
Walton, D.W.H. and D. Allsopp. 1977. A new test cloth for burial trials and other
studies on cellulose decomposition. Int. Biodeterior. Bull. 13:112-115.
Williamson, J.R. 1994. Calico cloth decomposition as an indicator of topsoil health
p.163-165. In Pankhurst et al. (eds.) Soil Biota: Management in Sustainable
Farming Systems, Poster Workshop Proc., March 15-18, 1994 Adelaide, S.
Australia, CSIRO Inform. Services, E. Melbourne, Victoria, Australia.
Wyland, L.J., L.E. Jackson, and K.F. Schulbach. 1995. Soil-plant dynamics following
incorporation of a mature cereal rye cover crop in a lettuce production system.
J. Agr. Sci. 124:17-25.
19
CHAPTER 2
WINTER COVER CROP EFFECTS ON MICROBIAL BIOMASS,
SOIL ENZYME ACTIVITIES, AND COTTON STRIP DECOMPOSITION
20
ABSTRACT
To promote agriculture sustainability, there is a growing interest in developing a
soil quality index that could be used by growers, extension agents and researchers as an
early indicator of changes in management. Soil samples were taken from six
commercial growers' fields and two research stations in western Oregon. The primary
comparison at each site was a winter cover crop and winter fallow in rotation with
summer vegetable crops. Microbial biomass carbon (MBc), mineralizable N, soil
enzyme activity (arylsulfatase and B-glucosidase), and cotton strip decomposition were
analyzed to monitor changes in soil quality after treatments were done. Results showed
that cover cropping significantly affected MBc and soil enzyme activity. Mineralizable
N and cotton strip decomposition did not respond to winter cover crop treatment.
Because MBc and B-glucosidase activity responded faster to treatment and were less
variable than the other measurements, they showed the most potential as soil quality
indicators.
INTRODUCTION
Soil, water and air are three natural resources essential for life on earth.
However, unlike air and water for which quality standards have been established, soil
quality has been difficult to define and quantify (Doran and Parkin, 1994). Soil quality
is none-the-less a very important concept; because world soil resources are not
renewable it is crucial to minimize the effect of all factors that contribute to its
degradation. In order to reach this goal, soil quality indicators are needed that are
sensitive to recent changes in soil management. To be practical, indicators should
21
integrate soil physical, chemical and biological properties and processes (Doran and
Parkin, 1994).
Winter cover crops often are planted to protect soil against erosion and to reduce
nutrient leaching, especially in zones receiving high precipitation during the winter.
Important benefits when cover crops are incorporated in early spring include increased
input of organic residues in the surface soil horizon, production of energy sources for
microorganisms, and enhancement of microbial activity (Powlson et al., 1987).
Management effects on soil biological properties and microbial community structure are
estimated through different methods such as direct counts with microscopy, microbial
plate counts and biomass, fatty acid extraction techniques, microbial respiration and
enzyme activity. To be a practical indicator of the soil biological component, indicators
must be simple and have low labor costs to be adopted by commercial labs, and not
have high seasonal variability. Microbial biomass carbon (MBA) and enzyme assays
would appear to meet these criteria for being practical soil biological indicators.
Microbial biomass (MB) represents only a small portion of soil organic matter
but it is a dynamic, living component that is involved in various biogeochemical
processes (Jenkinson and Ladd 1981; Voroney and Paul 1984). It plays an important
role in the cycle of nutrients and organic matter transformations. Microbial biomass is
sensitive to management effects such as tillage (Lynch and Panting, 1980), fertilization
and crop rotation. No-till systems have been shown to stimulate microbial biomass
nitrogen (MBN) and MBA (Doran, 1980; Carter and Rennie 1982) and mineralizable
carbon (C) and nitrogen (N) (Doran, 1980; Carter and Rennie 1982). Microbial
biomass C and MBN were found to be greater under a non-fertilized wheat system than
22
under wheat that received N fertilizer (Biederbeck et al., 1984). Greater MB and
organic matter also were found on a pasture site treated with low fertilization than on a
more highly fertilized site (Tate et al., 1991). Brookes (1995) found that MB, soil
enzyme activity and soil respiration respond more quickly to crop management
practices or environmental conditions than does soil organic matter. Microbial biomass
C has been reported better than MBN as an indicator of soil quality (Jordan and Kremer,
1994).
Enzymes catalyze most biological and biochemical reactions mediated by
microorganisms, plant roots and animals. Soil enzymes are involved in nutrient
cycling, organic matter decomposition, and conversion of inorganic to organic
substances, as well as in energy transformation. Frankenberger and Dick (1983)
analyzed the relationship of 11 enzymes in 10 unamended soils of diverse origin with
microbial respiration, biomass and viable plate counts. Their results indicated that
enzyme activities were associated with active microorganisms in soil but these findings
were not reflected accurately by the viable plate counts.
Soil enzymes originally were studied to measure soil fertility (Kuprevich and
Shcherbakova, 1971), but recently have been suggested as potential indicators of soil
quality (Dick, 1994) because of their rapid response to soil management. A number of
studies on long-term research sites have shown that enzyme assays can be sensitive to
soil management (Bolton et al., 1985; Perruci, 1992; Goyal et al., 1993; Miller and
Dick, 1995). Gupta and Germida (1988) reported a 65% decrease in arylsulfatase
activity in cultivated soils (69 yrs) in comparison with adjacent grassland.
23
Beta-glucosidase (EC 3.2.1.21) hydrolyzes B-D-glucopyranosides and
arylsufatase (EC 3.1.6.1) catalyzes the hydrolysis of the O-S bond (Spencer, 1958).
These two enzymes play an important role in nutrient cycling: the end product of
B-glucosidase activity is a simple sugar which can be utilized by microorganisms and
therefore influence the C cycle, and arylsufatase releases sulfate, often limited in soil
environment, to make it available to plants and microorganisms. These two enzyme
assays were chosen for this study because of the critical role they play as hydrolytic
enzymes in decomposition and mineralization, and because of the previous work which
has shown these assays to be sensitive to soil management (Bandick and Dick, 1998).
Residue decomposition provides an important source of C to organisms and
releases nutrients for plant uptake. Decomposition of organic residues in soil is
controlled by biological processes, which are modified by environmental conditions.
Several approaches have been investigated to assess soil decomposition potential, one
of them called "the cotton strip technique" (Harrison et al., 1988), is based on the
decrease in tensile strength of cotton strips buried in the soil for a variable period of
time. The standard cotton strip technique was developed originally for use in the
International Biological Programme (Latter and Walton, 1988) to assess decomposition
rates in a variety of soils.
Several studies have reported strong correlations between cotton strip
decomposition and nutrient release and turnover (Brown, 1988; French, 1988a, b). The
cotton strip assay was found to be a useful indicator of decomposition potential across
several wetlands, both natural and altered (Harrison et al., 1988). It has an advantage
over the litter bag technique because of its uniformity, direct contact with soil and easy
24
use. The cotton strip technique has been widely tested on various forest sites (Heal et
al., 1981; Hill et al., 1985; Maltby, 1988, Trettin et al., 1996) but little is known on how
effective this could be for agricultural systems. A few studies of cotton strip
decomposition have shown it to be sensitive to agricultural soil management.
Williamson (1994) showed a positive relationship between cotton strip decomposition
and a soil health score represented by five soil properties: soil pH, cation exchange
capacity, water stable aggregates and MBc. Lower cotton strip decomposition was
found in soils treated with high concentration of heavy metal relative to the
uncontaminated sludge-amended and unsludged control plots (Obbard and Jones, 1993).
There is relatively little information on the sensitivity of MBc, enzyme assays, and
cotton strip decomposition as early indicators after a change in soil management.
We hypothesize that addition of organic C through winter cover crop residues
would enhance soil biological properties such as MB and soil enzyme activities. The
objectives of this study were: (1) to examine the response of some biological indexes to
winter cover cropping, and (2) to explore and identify early indicators of changes in soil
quality that are useful to agroecosystem analysis and farm management.
MATERIALS AND METHODS
Research sites
Two research sites were at Oregon State University (OSU) experiment research
stations: the North Willamette Research and Extension Center (NWREC) and the
Vegetable Farm (V.F.). Both sites are located in western Oregon.
The OSU V.F., initiated in 1993, is a long-term experiment on a silt loam soil
(Cumulic Ultic Haploxerolls) in the southern Willamette Valley. The design is a
25
randomized block design with 4 replications of winter cover crop and winter fallow
treatments. Dimensions of each plot were 30 x 60 m. The cover crop treatment was a
80:20 weight mixture of annual ryegrass (Lolium multiflorum) and red clover (Trifolium
pratense) with small amounts of oats ( Avena sativa L.) and buckwheat (Fagopyrum
sagittatum) to attract benefical insects.
The NWREC, initiated in 1989, is on a Willamette (Pachic Ultic Argixeroll)
soil at Aurora, OR. The experiment is a sweet corn (Zea mays L.cv. Jubilee) and
broccoli (Brassica oleracea L. var Italica cv. Gem) rotation. The design is a
randomized split-plot complete block with four replications of six cover crop treatments
and three N rates subplots within each main plots. Main plots were 18 x 9 m and were
divided into three equal subplots areas of 54 m2 each. Nitrogen fertilizer rates of zero,
56 (medium) or 224 (recommended) kg N ha-I were applied for sweet corn and rates for
broccoli were zero, 140 (medium) or 280 (recommended) kg N ha-1. One replication of
winter fallow subplot was removed from our experiment because of frequent flooding
during the winter.
Farmer's field sites
At the beginning of the experiment in 1996, six commercial growers agreed to
split their field for treatment application. There were two treatments on each field; a
winter fallow (one part of the field left bare during the winter) and a winter cover crop
(winter cover crop planted on the other part) treatment during winter 1996-97 and
1997-98. However, due to farm management constraints, two of the growers (Hamlin
and Hendricks) did not plant their cover crop in winter 1996-97 and in 1998 the
Pearmine field was under a grass rotation. The design is a randomized complete block
26
(RCB) and each field was a block. Table 2.1 shows the exact cover crop treatments at
each experimental site.
Sampling procedure
Ten sampling points, five on each side of the split of growers' field, were
established with the Global Positioning System (GPS) to allow researchers to relocate
them throughout the experiment. Each field was mapped in detail for soil type and
matched pairs according to soil type and texture from each treatment were identified.
The soil type on each site is shown in Table 2.1. Soil samples were taken at canopy
closure in 1996 (around 6 weeks after planting summer vegetable crop), before
treatment application in the farmers' fields for chemical and biological analysis. After
treatments were applied, soil samples were taken in spring (before cover crops were
plowed down), at crop canopy closure and at harvest (about one week before crop
harvest) in 1997 and 1998 for chemical and biological analyses. A random composite
sample was removed from the area of each sampling point by homogenizing 36 soil
cores taken to a 7.5-cm depth. At the farmers' field sites, soil samples were taken
within a 2 m radius of the designated sampling points.
At the experiment research sites soil samples were taken at the same times as
described above but the 1996 sampling was after treatments had been in place
(7 years at NWREC and 3 years at V.F.). Soil cores were taken from the inner
two-thirds of the winter fallow and cover crop at these research sites. At the NWREC,
soil samples were taken from winter fallow plots receiving the recommended N rate and
from the winter cover crop plots at the medium N and recommended N rate. Cover
Table 2.1. Winter cover crop and crop rotation on-farm research sites and experiment research stations.
Site
Location
Soil
1997
1998
summer crop cover crop
summer crop
cover crop
Corn
Oats/barley
Dickman
Silverton
Argiaquic Xeric Arfialboll
Cauliflower
Annual rye
Grover
Salem
Fluventic Haploxeroll
Corn
Barley+ vetch Corn
Barley+vetch
Hamlin
Corvallis
Cumilic Ultic Haploxeroll
Beans
None
Grass
Rye+vetch
Hendricks
Scio
Fluventic Haplumbrept
Corn
None
Beans
Oats+vetch
Lucht
Molalla
Aquic Xerochrept
Corn
Oats
Cauliflower
Barley+vetch
Pearmine
Gervais
Aquultic Argixeroll
Corn
Vetch
Grass
Grass rotation
NWREC
Aurora
Aquultic Argixeroll
Broccoli
Triticale/AWP Corn
Triticale/AWP
Veg.Farm
Corvallis
Cumulic Ultic Haploxeroll
Corn
Mixed §
Mixed §
§ Annual rye grass + red clover + oats + buckwheat
AWP = Austrian winter pea
Beans
28
crops were triticale (Triticosecale X) and a mix of triticale and Austrian winter pea
(Pisum sativum L.).
When crops were present, soil samples were removed to equally represent
rhizosphere and nonrizhozphere soil. Soil samples were passed through a 2-mm sieve
and divided in two parts. One part was kept field moist at 4°C and the other half was
air-dried at room temperature for chemical and enzyme analyses.
Chemical analyses
Chemical characteristics of soils from on-farm and experiment research sites at
canopy closure in 1996 were done according to standardized methods of the Soil Survey
Laboratory (USDA/NRCS, 1996). Air dried samples were characterized for total N
(Leco FP-438 analyzer, method No. 6B4a) (Leco Corp., St. Joseph, MI); total C (Leco
SC-444 Carbon analyzer, method No. 6A2e) (Leco Corp., St. Joseph, MI); cation
exchange capacity (CEC) (sum of cations, method No. 5A3a); extractable calcium (Ca)
and potassium (K) (extractable bases, method No. 5B5a/6N2e) and pH (1:2 CaC12,
method No. 8C1f) (Table 2.2).
Enzyme assays
J3- glucosidase activity. Enzyme activity was measured following the method of
Tabatabai (1994). One gram of air-dried soil was placed in a 50-mL Erlenmeyer flask
along with 0.25 mL of toluene (Fisher certified reagent), 4 mL of modified universal
buffer (MUB) at pH 6.0, and a 1 mL of p-nitrophenyl-B-D-glucoside (PNG) solution.
Flasks were stoppered, swirled for few seconds and placed in an incubator for 1 hr at
37°C. After incubation, stoppers were removed; 1 mL of CaC12 and 4 mL of 0.1 M of
29
Table 2.2. Chemical characteristics of soils from on-farm and experiment research sites
at canopy closure in 1996.
On-Farm sites
NWREC
Soil Characteristics
Fallow Cover Crop
Fallow Cover Crop
Total N (g kg -1)
0.16 at
Total C (g kg -1)
1.95 a
CEC (Cmolc kg -1)
0.17 a
V.F.
Fallow Cover Crop
0.13 a
0.12 a
0.15 a
0.14 a
1.93 a
1.82 a
1.60 a
1.53 a
1.74 a
26.32 a
27.04 a
23.40 a 22.96 a
33.97 a
31.45 a
Ca (Cmolc kg -1)
11.82 a
11.63 a
6.30 a
7.55 a
16.08 a
16.55 a
K (Cmolc kg -1)
0.69 a
0.65 a
0.45 a
0.49 a
0.60 a
0.73 a
pH (CaC12)
5.74 a
5.74 a
4.90 a
5.30 b
5.50 a
5.53 a
t Means within a row and research site followed by the same letter are not significantly
different at P < 0.10 LSD level.
tris (hydroxymethyl) aminomethane (TRAM) buffer at pH 12 were added. The gentlymixed soil suspension was filtered through a Whatman no. 2 v-folded filter paper. The
intensity of the yellow filtrate was measured on a spectrophotometer at 420 nm. For
each sample three analytical replicates and a control were used.
Arylsulfatase activity. One gram of air-dried soil was placed into a 50 mL
Erlenmeyer flask, 0.25 mL of toluene was added, 4 mL of acetate buffer pH 5.8 and 1
mL of p-nitrophenyl sulfate solution. Flasks were stoppered, swirled for few seconds,
and placed in an incubator for 1 hr at 37°C. After incubation, stoppers were removed
and 1 mL of CaC12 and 4 mL of 0.5 M NaOH were added. The gently-mixed soil
suspension was filtered through a Whatman no. 2 v-folded filter paper. The intensity of
the yellow color of the filtrate was measured at 410 nm with a spectrophotometer.
Three analytical replicates and one control were used per sample. The results for both
assays were expressed as the activity of pig p-nitrophenol (PNP) g-1 soil hr.'.
30
Microbial biomass
Microbial biomass carbon was measured on field-moist soil by chloroformfumigation incubation method (Jenkinson and Powlson, 1976) modified as follows.
Ten grams of soil were weighed into scintillation vials. The vials were placed in a
dessicator containing wet paper towels at the bottom and a 50-mL beaker containing 30
mL of ethanol-free chloroform with a few boiling chips. The dessicator was evacuated
and the soil exposed to chloroform vapor for 24 hours. Soil was then transferred into
72-mL acrylic tubes (20 cm long, 2 cm diameter) stoppered with rubber septa at each
end to allow sampling for gas chromatography. Samples were incubated in the dark at
25°C for ten days. After incubation, CO2 produced was measured by gas chroma­
tography. A lcc of 0.41 (Voroney and Paul, 1984) was used to calculate MBA without
subtraction of the control.
Mineralizable N (N,) was measured as ammonium nitrogen (NH4+-N) under
waterlogged conditions as described by Keeney (1982). Five grams of field-moist soil
were placed in a 16 x150 mm test tube and 12 mL of distilled water were added.
Samples were incubated for 7 days at 40°C and the amount of NH4+-N formed was
determined by direct distillation of the soil-mixture in 2 M KC1 and MgO and
subsequent titration (Keeney, 1982). The amount of NH4+-N present in soil before
incubation was not deducted from after incubation because it was always very low.
Three analytical replicates and a control were used for each sample.
Cotton strip decomposition
The experiment was conducted, once a year, at summer crop canopy closure of
each sampling year on three of the farmers' fields and the two experiment research
31
sites. On-farm sites were Dickman, Lucht and Pearmine (Table 2.1). The Shirley Soil
Burial Test Fabric (Shirley Institute, Manchester, England) cut to14 x 7.5 cm strips was
used in this field study. At canopy closure in 1996, before treatment application, cotton
strips were inserted horizontally in the soil with minimal disturbance at 4 to 6 cm depth
for 28 days. At canopy closure of the first cover crop year (1997), cotton strips were
buried the same way for 14 and 21 days, and for 7 and 14 days at canopy closure of the
second cover crop year (1998). Each piece of cotton strip was weighed before and after
field burial period. The number of strips at each sampling point was one in 1996, two
in 1997 and three in 1998.
Cotton strips were retrieved after each incubation period, gently washed with
deionized water, laid on paper towels, oven-dried at 85°C for 45 min and weighed.
Tensile strength of each cotton strip was measured using the Instron tensiometer
(Instron Corporation, Canton, Massachusetts). Decomposition was measured as Cotton
Breaking Ratio (CBR) based on the loss in tensile strength (Williamson, 1994)
according to the following equation:
CBR = (TSUC-TSDC) / TSUC
where TSUC = Tensile Strength Undecomposed Cloth (kN m kg-1); and
TSDC = Tensile Strength Decomposed Cloth (kN m kg-1).
Statistical analysis
The design was a randomized complete block. Data from the on-farm sites were
analyzed by analysis of variance over the first (Yr 1) or second cover crop year (Yr 2) at
crop canopy closure (Can), in spring (Spg) and at harvest (Hav). Data from the
experimental research sites were analyzed the same way as the on-farm sites with the
32
exception that in 1996 the NWREC plots received their seventh winter cover crop
treatment and the V.F. plots their third one. Main effects of means were separated with
LSD at the 95% level. Data were analyzed using SAS Statistical software package
(SAS, Institute, Cary, NC).
RESULTS
Microbial biomass
Microbial biomass and soil enzyme results on each on-farm site are reported in
appendices A, B and C. When the Grover data were included in analysis of variance for
on-farm results, it significantly affected the outcome of all measurements. This seemed
to be due to the unique history of being under forest vegetation for 80 years prior to
1995. In 1996, the forest was clear-cut and the field was prepared for planting of the
first summer vegetable crop. A significant difference in 13-glucosidase activity was
found at time zero (canopy closure in 1996) between the winter fallow and winter cover
crop plots of the field although no treatment had been applied (Appendix B). For this
reason the following results of this chapter did not include data from Grover's field in
the statistical analysis.
Microbial biomass C at on-farm research sites was significantly (P < 0.05)
greater in the winter cover crop plots than the winter fallow ranging from 190 to
215 mg C kg soil -I the first cover crop year and from 175 to 300 mg C kg soir1 the
second year (Fig. 2.1). Microbial biomass C was significantly affected by winter cover
crop the first year showing it is a sensitive early indicator of change in soil
management.
33
525
450
375
IDA
0
300
225
G4
150
75
0
Ci3
\V"
Cie"
gse''
SAMPLING PERIOD
Fig 2.1. Winter cover crop effects on MB at on-farm research sites
(* indicates significant treatment effect at P < 0.05).
34
At the NWREC, the mixed cereal/legume cover crop (Celia triticale and
Austrian winter pea) had a greater effect than the cereal on MBA (Fig. 2.2). Microbial
biomass C measured on the V.F. plots significantly increased between spring 1997 and
canopy closure in closure 1998 (Fig. 2.2). A seasonal trend was detected for both
treatments, with maximum Ml3c occurring in the spring and a minimum at canopy
closure with one exception.
Mineralizable N was variable across sites (Table 2.3). Greater Nmin was
measured at crop canopy closure at on-farm sites the first cover crop year, however,
significant treatment effects were at harvest of the same year and in spring of the second
year cover crop year. At the NWREC, there was a significant winter fallow effect at
three sampling times with a maximum of 88 ps Nmin soil 7 days -1measured at canopy
closure in 1997. Results were reversed on the V.F. plots with greater Nmit, recorded on
the cover crop soils (Table 2.3).
Soil respiration
Winter cover crop had no significant effect on soil respiration at on-farm sites.
During the first cover crop year soil respiration was relatively stable, but the second
year showed more variations with neither year showing significant treatment effect
(Fig. 2.3).
Soil respiration rates at the NWREC generally were higher on the winter fallow
plots and on plots that received the mixed cereal/legume cover crop treatment (Fig. 2.4).
A significant effect of these treatments was measured at canopy closure in 1998.
Respiration rates measured at the V.F. were relatively more stable over time. There
was, however, a significant cover crop effect on soil in spring 1998 (Fig. 2.4).
35
525
NWREC
450
I-- Winter fallow
375
--.6-- Triticale
---- Triticale & AWP
300
225
.--.
v... 4
0ct
150
'gyp
75
v
I
C...)
tO
E
c.)
0
VEGETABLE FARM
Er Winter fallow
450
A.-- Winter cover crop
*
375
300
225
150
75
0
cro,'CN
0
46,
S )\
$,1 V
.0 ,S )
Crtk-
Cc\
\50,44...
0
%4;? T.,
co,
SAMPLING PERIOD
Fig 2.2. Winter cover crop effects on MB on experiment research sites
(* indicates significant treatment effect at P < 0.05).
Table 2.3. Treatment effects on mineralizable N at on-farm and experiment research sites.
Mineralizable N
tYear 2
11Year 1
Sites
Treatment
Canopy 1996
Spring
Canopy
Harvest
Spring
Canopy
Harvest
1.tg NT-14+-N g-1 soil 7 days -1
Winter fallow
20.02 al
26.46 a
69.53 a
23.35 a
19.25 a
38.35 a
19.24 a
Winter cover crop
19.66 a
27.62 a
98.25 a
25.88 b
25.88 b
26.82 b
20.42 a
Winter fallow
21.51 a
26.20 a
88.12 a
62.22 a
16.49 a
27.95 a
37.23 a
Cereal
31.30 a
23.81 a
35.65 b
32.69 b
13.82 a
21.05 a
16.21 b
Cereal+legume
36.62 a
23.75 a
28.89 b
27.50 b
18.29 a
19.20 a
19.76 b
Vegetable
Winter fallow
33.04 a
21.75 a
14.00 a
44.41 a
25.80 a
25.15 a
18.92 a
Farm
Winter cover crop
37.06 a
35.55 a
20.78 a
51.58 a
28.05 a
36.69 b
36.65 b
On-farm
NWREC
¶ and t correspond to year 1997 and 1998 for both NWREC and Vegetable farm, respectively.
t Means followed by the same letter within a column at each site are not significantly different at P < 0.05.
37
1.0
0.8
ct
rzs
O
e.4
0.6
ti)
0.4
0.2
0.0
CP
0
ca
gs`z$
S'4%
SAMPLING PERIOD
Fig 2.3. Winter cover crop effects on soil respiration at on-farm research sites
(* indicates significant treatment effect at P < 0.05).
38
1.0
0.8
NWREC
--I-- Winter fallow
Triticale
Triticale & AWP
0.6
O
bq
0.4
ON 0.2
zO
0.0
VEGETABLE FARM
0.8
--a-- Winter fallow
Winter cover crop
E-21
cip
O
c1)
0.6
0.4
0.2
0.0
SAMPLING PERIOD
Fig. 2.4. Winter cover crop effects on soil respiration on experiment research sites
(* indicates significant treatment effect at P < 0.05).
39
Soil enzymes
Soil enzyme activities responded similarly to treatments as MBA. Arylsulfatase
and13-glucosidase had higher activity in cover crop than winter fallow plots at the
on-farm sites. A greater treatment effect, however, was measured during the second
cover crop year. Ranges of13-glucosidase activity were about two-fold higher than of
those of arylsulfatase activity (Fig. 2.5). Similarly, arylsulfatase and 13-glucosidase
activities were overall significantly higher on the winter cover crop plots at the
NWREC and the V.F. (Fig 2.6 and Fig 2.7). Activity of both assays, however, was
consistently greater on the V.F. plots than those at the NWREC.
Cotton strip decomposition
The cotton strip decomposition data is shown in Figures 2.8 to 2.10. The data
are presented as the cotton breaking ratio (CBR) which by definition means that the
higher the ratio the greater the decomposition.
Winter cover crop effect on cotton strip decomposition at on-farm research sites was
inconclusive, although there was a trend towards a treatment effect after 7 days burial
period in the second cover crop year (Fig. 2.8).
At the NWREC, cover crop mix cereal/legume had a greater effect on cotton
strip decomposition at both 7 and 14 days burial period at canopy closure in 1998
(Fig. 2.9). Results of cotton strip decomposition at the V.F. were inconclusive from
year to year (Fig. 2.10).
40
240
ARYLSULFATASE
200
IN-- Winter fallow
Winter cover crop
160
120
ci)
80
40
1-4
0 bf)
Z a.,
13-GLUCOSIDASE
0
200
E-1
160
120
80
40
0
0
cizfr""­
%4';
\
\
\-8$1
c,,vc
SAMPLING PERIOD
Fig. 2.5. Winter cover crop effects on arylsulfatase and B-glucosidase activities at
on-farm research sites (* indicates significant treatment effect at P < 0.05).
41
200
NWREC
1- Winter fallow
160
Triticale
Triticale & AWP
120
VEGETABLE FARM
--s-- Winter fallow
&-- Winter cover crop
80
40
0
9
cAC
4\ ;vs9'\
C.)3'cs
99'
c64%
ei%
9%
\5"
SAMPLING PERIOD
Fig 2.6. Winter cover crop effects on arylsulfatase activity on experiment research
sites (* indicates significant treatment effect at P < 0.05).
42
200
NWREC
--IF Winter fallow
160
Triticale
Triticale & AWP
120
*
C-T4
CA
At
80
40
-7to
0
C.4
VEGETABLE FARM*
P-4
0 tl)
160
120
80
40
Winter fallow
161''''' Winter cover crops
0
C,
9b
I
I
0;\
0,;\
Crl)''
04. CO
01\
-tS.
I
095
ofb
I
opicb
s$N1
SAMPLING PERIOD
Fig 2.7. Winter cover crop effects on B-glucosidase activity on experiment research
sites (* indicates significant treatment effect at P < 0.05).
Winter fallow
VA Winter cover crop
0.6
PC1
a
0.4
Z
0
H
H 0.2
0
U
0.0
Day 21
Can-Yr 0
Day 14
Day 21
Can-Yr 1
Day 7
Day 14
Can-Yr 2
Fig. 2.8. Winter cover crop effects on cotton strip decomposition at on-farm research sites. Within each day,
treatment bars followed by the same letters are not significantly different (P < 0.05).
Winter fallow
Triticale
Triticale & AWP
b
C7
0.6
PC1
Z
0.4
0
H
H
0.2
0
U
0.0
Day 21
Can-Yr 96
Day 14
Day 21
Can-Yr 97
Day 7
Day 14
Can-Yr 98
Fig 2.9. Winter cover crop effect on cotton strip decompositon at NWREC. Within each day,
treatment bars with the same letter are not significantly different (P < 0.05).
t
1.0
2
E-4
0.8
0.6
0.0
Day 21
Can Yr-96
Day 14
Day 21
Can-Yr 97
Day 7
Day 14
Can-Yr 98
Fig 2.10. Winter cover crop effect on cotton strip decomposition at the Vegetable Farm. Within each day,
treatment bars with the same letter are not significantly different (P < 0.05).
46
DISCUSSION
Temporal effects
Responses of MI3c and soil enzyme activities were relatively stable from year
one to year two after treatment application at the on-farm sites, although a slight
decrease in activity was measured the second year. There were, however, significant
seasonal biological responses. Soil enzyme activities and MBA were generally highest
during the spring season for both on-farm and experiment research sites. These findings
could be attributed to a rhizosphere effect. Root growth and density of cover crops in
spring may have released exudates that are potential sources of C and nutrients for
microbes.
Similar seasonal patterns of MBA have been reported (Collins et al., 1992;
Buchanan and King, 1992) and are thought to result from increased availability of root
exudates as root growth and turnover increases in spring (Lynch and Panting, 1980).
Temperature tends to be a critical factor because there was a negative correlation
between MBA and temperature at crop canopy closure (Table 2.4). However, this runs
contrary to the widely observed phenomenon where increasing soil temperature
promotes biological growth and activity. It seems likely that other factors may be
controlling this seasonal biological response. Canopy closure in our experiment refers to
the early stage of vegetable growing season. During this period, nutrients are needed to
support plant growth. It is possible that as crops mature and utilize large quantities of
water and nutrients, microbial growth and activity is depressed because of substrate
limitation. Soil water content at sampling time also may have influenced the activity of
microorganisms because MBc and water content were significantly correlated (r = 0.34;
Table 2.4. Matrix of Pearson correlation coefficients for 11 of the measured soil biochemical parameters.
Variables
MBc
Mineralizable N (MN)
Soil respiration (SR)
MBc
MN
-0.03
SR
-0.06
-0.01
GI
0.75
***
0.68
0.04
-0.02
-0.14
**
13-Glucosidase (GI)
Ar
***
-0.16
**
0.79
***
Arylsulfatase (Ar)
CBR
WL
14d
14d
-0.24
***
-0.45
***
0.21
***
-0.03
0.098
-0.02
0.20
0.40
CC
***
0.12
0.24
*
**
-0.22
***
-0.21
***
-0.51
***
0.27
-0.50
0.22
***
0.03
Cotton strip Decomposition at 14 d
(CBR 14 d)
Weight loss at 14 days (WL 14 d)
***
0.57
***
-0.16
**
***
TC
***
-0.02
0.05
0.08
0.14
**
-0.10
0.06
Te
Wa
14d
14d
-0.43
***
0.34
0.13
-0.09
0.57
***
-0.46
-0.21
***
**
**
*
0.26
-0.36
0.39
***
***
0.14
-0.14
0.40
0.03
***
Clay content (CC)
Total C (TC)
Temperature at 14 days (Te 14 d)
Water content
14 d (Wa 14 d)
*, **, *** Significant at P 5 0.05, 0.01, and 0.001, respectively for number above.
0.51
***
-0.52
***
-0.17
0.08
-0.01
0.12
48
P = 0.01). Furthermore, sampling in the summer was often after extensive tillage,
which may have caused a reduction in biological activity from spring to summer due to
destruction of habitats in aggregates.
Cover crop effects
Several trends were observed in our experiment. Microbial biomass C, and
arylsufatase and 13-glucosidase activities generally were higher on the winter cover crop
plots. These measurements were uncorrelated with total C (Table 2.4). This supports a
previous study showing a significant increase in enzymatic activity 2 years after
initiation of cover crop treatment whereas total C remained unchanged (Dick, 1994).
This would indicate that these biological responses are occurring before there is any
significant accumulation of soil organic matter. These results suggest that the early
responses within one or two years at the on-farm plots predict soil quality trajectory
because the research sites, which had been in place longer (seven years at NWREC and
three years at the V.F. in 1996) still showed significant winter cover crop effects.
Beta-glucosidase and arylsulfatase activities were significantly correlated with
MBc (Table 2.4); which suggests that activities of these enzymes are related to the
viable microbial population. Beta-glucosidase and arylsulfatase activities were
significantly correlated which may confirm their similar origins.
Effects of treatment on Nn,in was highly variable between seasons. There was no
correlation between potentially Nnfir, and MBc, which may be due partially to different
seasonal patterns of these two biological indexes. Nitrogen mineralization potential
was low in spring and at a maximum at canopy closure whereas MBc was just the
opposite. This lack of correlation of N mineralization potential with biological
49
properties may be due to the likelihood that the organisms involved in N mineralization
are a subset of the microbial community represented by MBc and enzyme assays. Also,
N mineralization potential represents both the biological potential and labile N pool in
soil organic matter that is readily mineralized. For these reasons we might expect N
mineralization potential not to correlate well with integrative biological indexes like
MBc and enzyme assays. This indicates that N mineralization would not be a good
candidate for a soil quality indicator of soil biology.
Soil respiration results were variable across on-farm research sites. Greater soil
respiration generally was associated with smaller microbial biomass. This is consistent
with Campbell et al. (1991) who found the specific respiration (SR) during a growing
season to be higher in systems with a smaller MB.
The mix of a legume cover crop, AWP, with triticale at the NWREC plots had a
greater effect than the cereal or winter fallow on microbial and enzyme activity., Our
results confirm the importance of leguminous cover crops in agricultural systems and
agree with those of Bolton et al. (1985) who reported an increase in soil MBA and
enzyme activity on soil that received legume green manure compare to unamended
soils. Legumes are known to fix N and may provide substantial amounts of N to the
subsequent crop (Heichel and Barnes, 1984; Burket et al., 1997). Their narrow C:N
ratio (< 20:1) accelerates residue decomposition and cycling of nutrients which in turn
could stimulate microbial activity. The lower response of the cereal cover crop at the
NWREC may be due to the poor cover crop stands caused by excessive rains and late
planting for the years 1994 to 1996 (Minshew, 1998). Conversely, the cereal/legume
50
mix tended to recover in the spring with some legume growth that may have maintained
the higher biological response of this treatment.
Cotton strip decomposition did not respond to winter cover crop treatment
possibly due to variations of abiotic factors such as moisture and temperature within
sites and treatments. Results of our laboratory incubation experiment presented in
Chapter 4 confirm the influence of N, moisture and possibly temperature on cotton strip
decomposition. Significant correlation (P < 0.001) was obtained between CBR and
cotton strip weight loss after a 14 day burial period (r = 0.57; Table 2.4). This indicates
that the simpler weighing method could be used to measure the effect of microbial
activity on cotton strip decomposition.
Sensitive indicators of soil quality
Sensitive soil quality indicators should respond quickly to changes in
management, be easy to use, have high sensitivity and low seasonal variability.
Significant year-to-year or seasonal variations in activity would make it difficult to
assess changes due to management. Among the indicators tested in our study,
B-glucosidase activity was found to be very responsive to winter cover crop, to be less
variable from year to year and within each treatment across experimental sites.
Coefficients of variation across all treatments were approximately 32% at the on-farm
research sites, ranged from 35 to 50% at the V.F.; and from 20 to 32% at the NWREC.
Microbial biomass C also responded quickly to management but had greater
variability than B-glucosidase activity. Coefficient of variation at on-farm sites was
about 42% but was lower at the NWREC (30%).
51
Arylsulfatase activity and particularly Nmin had the greatest variability within
year and treatment. Coefficients of variation were about 50% for arylsufatase activity
and as high as 183% for mineralizable N.
Based on variability and treatment responses, it appears that 13- glucosidase
activity and MBc are good choices as soil quality indicators of the biology. Overall, we
did see a positive effect of winter cover cropping on MBc and enzyme activity accross
sites. Soil respiration and cotton strip decomposition were found not to be as sensitive
soil enzyme activity and MBc. The cotton strip method, however, still could be used as
a semi-quantitative indicator of microbial activity by just weighing cotton strips after 14
days burial period under field conditions.
LITERATURE CITED
Bandick, A.K. and R.P. Dick. 1998. Field management effects on soil enzyme
activities. Soil Biol. Biochem. In Press.
Biederbeck, V. 0., C.A. Cambpell, and R.P. Zenter. 1984. Effect of crop rotation and
fertilization on some biological properties of a loam in southwestern
Saskatchewan. Can. J. Soil. Sci. 64:355-367.
Bolton, J., L.F. Elliott, P.R. Papendick, and D.F. Bezdicek. 1985. Soil microbial
biomass and selected soil enzyme activities: Effect of fertilization and
cropping practices. Soil Biol. Biochem. 17: 297-302.
Brown, A.H.F. 1988. Discrimination between the effects of soils of 4 tree species in
pure and mixed stands using cotton strip assay. p. 80-85. In A.F. Harrison, P.M.
Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition
in soils, ITE Symposium 24, Institute of Terrestrial Ecology, Grange-OverSands, Cumbria, England.
Brookes, P.C. 1995. The use of microbial parameters in monitoring soil pollution
by heavy metals. Biol. Fert. of Soils. 19:269-279.
Buchanan, M., and L.D. King. 1992. Seasonal fluctuations in soil microbial biomass
carbon,phosphorus, and activity in no-till and reduced-chemical-input maize
agroecosystems. Biol. Fertil. Soils. 13:211-217.
52
Burket, J.Z., D.D. Hemphill, Jr., and R.P. Dick. 1997. Winter cover crops and nitrogen
management in sweet corn and broccoli rotations. HortSci. 32:664-668.
Campbell, C.A., M. Schnitzer, G.P. Lafond, R.P. Zentner, and J.E. Knipfel. 1991.
Thirty year crop rotations and management practices effects on soil and amino
nitrogen. Soil Sci. Soc. Am. J. 55:739-745.
Carter, M.R. and D.A. Rennie, 1982. Changes in soil quality under zero tillage
farming systems: Distribution of microbial biomass and mineralizable C and N
potentials. Can. J. Soil Sci. 62:587-597.
Collins, H.P., P.E. Rasmussen, and C.L. Douglas, Jr. 1992. Crop rotation and residue
management effects on soil carbon and microbial biomass. Soil Sci. Soc. Am. J.
56:783-788.
Dick, R.P. 1994. Soil enzyme activities as indicators of soil quality. p. 107-124. In J.W.
Doran, D.C. Coleman, D.F. Bezdicek and B.A. Stewart (eds.) Defining soil
quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
Doran, J.W. 1980. Soil microbial and biochemical changes associated with reduced
tillage. Soil Sci. Soc. Am. J. 44:765-771.
Doran J.W. and T.B. Parkin 1994. Defining and assessing soil quality. p 3-21.
In J.W. Doran, D.C. Coleman, D.F. Bezdicek and B.A. Stewart (eds.)
Defining soil quality for a sustainable environment. SSSA Spec. Publ. 35.
Madison, WI.
Frankenberger Jr., W.T. and W.A. Dick. 1983. Relationships between enzyme
activities and microbial growth and activity indices in soil. Soil Sci. Soc. Am.
J. 47:945-951.
French, D.D. 1988a. Some effects of changing soil chemistry on decomposition ofplant
litters and cellulose on a Scottish moor. Oecologia 75:608-618.
French, D.D. 1988b. Patterns of decomposition assessed by the use of litter bags and
cotton strip assay on fertilized and unfertilized heather moor in Scotland.
p. 100-108. In A.F. Harrison, P.M. Latter and D.W.H. Walton (eds.) Cotton
strip assay: An index of decomposition in soils. Institute of Terrestrial
Ecology, Grange-Over-Sands, Cumbria, England.
Goyal, S., M.M. Mishra, S.S. Dhankar, K.K. Kapoor, and R. Batra. 1993. Microbial
biomass turnover and enzyme activities following the application of farmyard
manure soil field oils with and without previous long-term applications. Biol.
Fert. Soils. 15:777-786.
53
Gupta, V.V.S.R. and J.J. Germida. 1988. Distribution of microbial biomass and its
activity in different soil aggregate size classes as affected by cultivation. Soil
Biol. Biochem. 20:777-786.
Harrison A.F., P.M. Latter and D.W.H. Walton 1988. Cotton strip assay: an index of
decomposition in soils. ITE Symposium 24, Institute of Terrestrial Ecology,
Grange-Over-Sands, Cumbria, England.
Heal, 0.W., P.W. Flanagan, D.D. French and S.F. Maclean, Jr. 1981. Decomposition
and accumulation of organic matter. p. 587-634. In L.C. Bliss et al. (eds.)
Tundra ecosystem: A comparative analysis. Cambridge Univer. Press,
Cambridge, England.
Heichel, G.H., and D.K. Barnes. 1984. Opportunities for meeting crop nitrogen needs
from symbiotic nitrogen fixation. In D. Bezdicek and J. Power (eds.) Organic
farming: current technology and its role in a Sustainable Agriculture.
ASA Spec. Publ. 46. Madison, WI.
Hill, M.O., P.M. Latter and G. Bancroft. 1985. A standard curve for inter-site
comparison of cellulose degradation using the cotton strip method. Can. J. Soil
Sci. 65:609-619.
Jenkinson, D.S. and J.N. Ladd. 1981. Microbial biomass in soil: Measurement and
turnover. p.415-471. In E.A. Paul and J.N. Ladd (eds.) Soil Biochem.Vol. 5.
Marcel Dekker, New York.
Jenkinson, D.S. and D.S. Powlson. 1976. The effect of biocidal treatments on
metabolism in soil-V. A method for measuring soil biomass. Soil Biol.
Biochem. 8:209-213.
Jordan, D. and R.J. Kremer. 1994. Potential use of soil microbial activity as an
indicator of soil quality. p. 243-249. In C.E. Pankhurst et al. (eds.) Soil Biota:
Management in sustainable farming systems. CSIRO, Australia.
Keeney, D.R. 1982. Nitrogen availability indices. p. 711-733. In A.L. Page, R.H.
Miller, and D.R. Keeny (eds.) Method of soil analysis Part 2. 2nd ed. Agron.
Monogr. 9. ASA and SSA, Madison, WI.
Kuprevich, V.F., and T.A. Shcherbakova. 1971. Comparative enzymatic activity in
diverse types of soil. p. 167-201. In A.D. McLaren and J.J. Skujins (eds.) Soil
Biochem. Vol. 2. Marcel Dekker, New York.
54
Latter, P.M. and D.W.H. Walton. 1988. The cotton strip assay for cellulose decomposi­
tion studies in soil: history of the assay and development. p.7-10. In A.F.
Harrison, P.M. Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of
decomposition in soils. Institute of Terrestrial Ecology, Grange-Over-Sands,
Cumbria, England.
Lynch, J.M., and L.M. Panting. 1980. Cultivation and the soil biomass. Soil Biol.
Biochem. 12:29-33.
Maltby, E. 1988. Use of cotton strip assay in wetland and upland environments an
international perspective. p. 140-154. In A.F. Harrison, P.M. Latter and
D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition in soils.
Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Miller, M., and R.P. Dick. 1995. Dynamics of soil C and microbial biomass on whole
soil aggregates in two cropping systems differing in C-input. Appl. Soil Ecol.
2:253- 261.
Minshew, H.F. Nitrate leaching and model evaluation under winter cover crops. Oregon
State Univ. Personal communication. November 11, 1998.
Obbard, J.P., and K.C. Jones. 1993. The use of the cotton-strip assay to assess
cellulose decomposition in heavy metal-contaminated sewage sludge-amended
soils. Environmental Pollution. 81:173-178.
Perrucci, P. 1992. Enzyme activity and microbial biomass in field amended with
municipal refuse. Biol. Fert. Soils. 14:54-60.
Powlson, D.S., P.C. Brookes, B.T. Christensen. 1987. Measurements of soil microbial
biomass provides an early indication of changes in total soil organic matter due
to straw incorporation. Soil Biol. Biochem. 19:159-164.
Spencer, B. 1958. Studies on sulphatases: 20. Enzymatic cleavage of arylhydrogen
sulphates in the presence of H2 180. Biochem. J. 69:155-159.
Tabatabai, M.A. 1994. Soil enzymes. p. 775-833. In R.W. Weaver et al. (eds.)
Methods of Soil Analysis, Part 2. Microbiological and Biochemical Properties.
SSSA Book Ser. 5. SSSA, Madison,WT.
Tate, K.R., T.W. Speir, D.J. Ross, R.L. Parfitt, K.N. Whale and J.C. Cowling. 1991.
Temporal variations in some plant and soil P pools in two pasture soils of
widely different P fertility status. Plant and Soil 132: 219-232.
Trettin, C.C., M. Davidian, M.F. Jurgensen, and R. Lea. 1996. Organic matter
decomposition following harvesting and site preparation of a forested wetland.
Soil Sci. Soc. Am. J. 60:1994-2003.
55
USDA/NRCS. 1996. Soil survey laboratory methods manual. Soil Survey Investigation
Report No. 42, Ver. 3.0. USDA/NRCS/National Soil Survey Center,
Washington, D.C.
Voroney, R.P., and E.A. Paul. 1984. Determination of kc and kN in situ for calibration
of the chloroform fumigation-incubation method. Soil Biol. Biochem. 16:9-14.
Williamson, J.R. 1994. Calico cloth decomposition as an indicator of topsoil health
p.163-165. In Pankhurst et al. (eds.) Soil Biota: Management in Sustainable
Farming Systems, Poster Workshop Proc., March 15-18, 1994 Adelaide, S.
Australia, CSIRO Inform. Services, E. Melbourne, Victoria, Australia.
56
CHAPTER 3
TILLAGE EFFECT ON SOIL BIOLOGICAL PROPERTIES
57
ABSTRACT
Agricultural practices such as tillage, crop rotation or cover cropping
significantly affect soil physical, chemical and biological properties. Biological
indicators of soil quality have been proposed to monitor the sustainability of field
management. The effects of conventional and minimum tillage practices were
evaluated on a commercial grower's field. Winter cover crops were grown in rotation
with summer vegetable crops on the entire field for 2 years. Each tillage treatment was
applied on one-half of the field and soil samples were taken at a 7.5 cm depth from each
part of the field, microbial biomass carbon (MBA), mineralizable nitrogen, soil enzyme
activities (arylsulfatase and 13-glucosidase), and cotton strip decomposition were
analyzed to detect early changes due to management. Results showed overall a
significant effect of minimum tillage on soil biological properties. Biological
parameters measured were strongly correlated with total carbon.
INTRODUCTION
Tillage operations generally are used by farmers for seedbed preparation,
residue incorporation or weed suppression. The moldboard plow has been a primary
piece of equipment used for these cultural practices but it is being replaced by the chisel
plow or disk harrow because they minimize soil disturbance. Intensive tillage practices
have been shown to affect soil physical properties such as aggregate size distribution
and stability (Elliott, 1986; Gupta and Germida, 1988) and hence, modify soil chemical
and biological properties.
Reduced tillage increases aggregate stability and soil structure (Doran and
Smith, 1987; Campbell et al., 1989; Miller and Dick, 1995) as well as microbial
58
activities (Dick, 1984; Carter, 1986; Gupta and Roper, 1992; Angers et al., 1993).
Residues left on the soil surface protect soil from water and wind erosion and contribute
to maintenance of soil organic carbon. Conservation tillage practices in combination
with cover crops might increase carbon (C) sequestration to offset elevated carbon
dioxide (CO2) in the atmosphere and decrease nutrient leaching.
We hypothesized that minimum tillage systems favor biological activity of
microorganisms by preserving microbial habitat. The objective of this study was to
examine tillage effects of winter cover cropping on soil biological parameters.
MATERIALS AND METHODS
Experimental design
This experiment was conducted on one commercial grower's field located in
Albany, OR. This study was a paired design with two treatments: winter cover
crop/minimum-till and winter cover crop/conventional till. In summer 1996, prior to
treatment application, soil chemical and biological properties, with the exception of
microbial biomass carbon (MBc), were measured on each part of the field as a baseline.
There was no significant difference in soil physical and chemical properties of the two
portions of the fields in 1996 (Table 3.1). Soil samples were taken the same way and at
the same time as described in Chapter 2. Soil enzyme activity, MBA and cotton strip
decomposition were determined following the same methods as in Chapter 2.
Chemical analysis
Physical and chemical characteristics of soil in summer 1996 were done
according to the standardized methods of the Soil Survey Laboratory (USDA/NRCS,
59
1996). Air-dried samples were characterized for particle size analysis (air-dry samples
Method No. 3A1), for total N (Leco FP-438 analyzer, method No. 6B4a) (Leco Corp.,
St. Joseph, MI); total C (Leco SC 444 Carbon analyzer, method No. 6A2e) (Leco Corp.,
St. Joseph, MI); cation exchange capacity (CEC) (sum of cations, method No. 5A3a)
and pH (1:2 CaC12, method No. 8C1f) (Table 3.1).
Table 3.1. Physical chemical properties of soil in 1996.
Treatment
Clay
Silt
Minimum-till plot
24
53
22
g kg -1
1.79
0.19
No-till plot
25
58
16
2.06
Sand
%
Total C
Total N
0.21
CEC
pH
Cmo lc kg'
36.26
4.92
34.58
4.96
Tillage system
In March 1996, barley (Hordeum vulgare) was planted with a no-till planter on
the entire field. After crop harvest, a heavy cover crop disc was used to work the barley
stubble twice to a depth of 15 to 30 cm. Oats (Avena sativa) were planted as a cover
crop with a grain drill on the entire field but the cover crop did not survive the winter
flood in December 1996. Barley was planted as a second cover crop in April 1997
using a no-till planter and was chemically killed with glyphosate in May 1997. On the
conventionally-managed plot, the cover crop was mowed and disked twice at depth of
15-30 cm. A spring-tooth harrow was passed on the tilled plot once before the use of a
conventional planter to plant the summer crop snap bean (Phaseolus vulgaris L). On
the minimum-tilled plot, a no-till grain drill was used to plant the summer vegetable
crop directly into the barley stubble.
60
The second cover crop year the following changes in management were made:
a strip-till planter was used to plant both cover crop and vegetable crop on the entire
field. Surface band fertilizer application was applied on the minimum-tilled plot and
a broadcast application was done on the conventionally-managed plot.
Statistical analysis
Data were analyzed using a paired t-test and significant differences were
determined at P < 0.05 using the SAS statistical package (SAS Inst. 1996). Correlation
analysis used the Pearson correlation coefficient on this SAS Proc Corr subroutine.
RESULTS AND DISCUSSION
Microbial biomass, respiration and N mineralization
Microbial biomass carbon on the minimum-tilled plot was significantly greater
than on the conventionally managed treatment. Maxima of 400 and 430 mg C kg' soil
were measured in spring of the first and second cover crop years, respectively (Fig. 3.1).
The increase in MBc the first year after changing management is in agreement with
findings of Gupta et al. (1994) who found a response of MBc to modified tillage and
stubble management after one year. Similar results were found by Doran (1980) who
reported higher aerobic and facultative bacteria in two surface soils under zero tillage
compared to plowed soils. Microbial biomass was particularly affected by tillage at
harvest, on the tilled portion of the field, resulting in a consistent decrease in activity
after spring sampling. There was a general trend of less variation in MBc on the
minimum-tilled soil than the conventional-tilled soil (Fig. 3.1). Our results confirm the
61
525
MICROBIAL BIOMASS CARBON
450
7:7>
375
bA
300
L)
st).0
225
150
--o-- Cony. till
75
Min. till
0
SOIL RESPIRATION
0.2
0.0
cfr
\
a
1.
4A:s.
5c.
cA%
jr6,4
sea
SAMPLING PERIOD
Fig 3.1. Tillage effect on MBc and soil respiration at on-farm research site
(* indicates significant treatment effect at P < 0.05).
62
effect of reduced tillage practices on soil MBA as observed in other cropping systems
(Carter, 1986; Doran, 1987; Gupta and Germida 1988).
Soil respiration tended to increase with cultivation but there was no significant
treatment effect (Fig. 3.1). Greater respiration was measured on the conventional-tilled
plot. These results might be expected because cultivation breakes down
macroaggregates and releases protected organic matter that is available to microbial
degradation.
Mineralizable N generally was lower on the minimum-tilled portion of the field,
but a significant treatment effect was observed only at harvest of the second cover crop
year (Fig. 3.2). Under field conditions, zero tillage may possibly reduce net N
mineralization due to a decrease of physical disturbance of the soil (Van Veen and Paul,
1981). Furthermore, tillage differences may affect soil porosity and aeration possibly
resulting in N loss by denitrification.
Enzymes activities
Activities of both arylsulfatase and B-glucosidase were significantly greater in
the no-tilled site with a maximum of 165 µg PNP g-I soil h -1 and 214 lig PNP g-I
soil WI (Fig. 3.3), respectively. These results agree with those of Gupta and Germida
(1988) who found that enzyme activity and MBc were significantly lower after 69 years
of continuous cultivation. Dick (1984) measured significantly greater enzyme activity
in no-till than conventionally-tilled soils. In addition, other studies showed that residues
left on the surface increase the microbial biomass pool (Carter, 1986), C and N
mineralization (Campbell et al., 1989) and soil enzyme activities (Dick et al., 1988).
63
120
' cn
100
ct
80
O4
c.)
tO
60
z
40
z
ttO
20
0
coP.
,c\ck
SAMPLING PERIOD
Fig 3.2. Tillage effect on mineralizable N at on-farm research site
(* indicates significant treatment effect at P < 0.05).
64
240
ARYLSULFATASE
200
*
160
*
120
4t
80
4.4
40
014
074
II Cony. till
A.
bp
0
*
ao
0 bo
E--(
;14-4
Min. till
*
*
200
160
120
80
40
B-GLUCOSIDASE
0
rvc.
Cfr
g,.
,1,VC
^­
,csf
$4 $40 C b
SAMPLING PERIOD
Fig 3.3. Tillage effect on arylsulfatase and B-glucosidase activity at on-farm
research site (* indicates significant treatment effect at P < 0.05).
65
Conservation tillage practices reduce soil disturbance and have been shown to have
greater levels of enzyme activity (Klein and Koths, 1980; Angers et al., 1993).
The decrease in activity observed in the second cover crop year might be
attributed to the decrease in MB measured in the tilled plot probably due to a loss in soil
organic matter (Khan, 1970). Greater B-glucosidase activity was measured at harvest
1997 whereas greater arylsulfatase activity was measured in spring 1998 on the no-tilled
plot.
Cotton strip decomposition
Tillage effect on cotton strip decomposition was variable. Greater decomposi­
tion was found on the minimum tilled plot the first cover crop year, but during the
second year the trend was reversed and significant differences were found on the
conventionally tilled plot at the 14-day burial period (Fig. 3.4). Changes observed the
first cover crop year could be attributed to the greater microbial activity measured
on this plot following organic input. These conflicting results limit the potential to
recommend the cotton strip decomposition method as a soil quality indicator.
Correlations
The correlations among the various biological parameters were statistically
significant. Microbial biomass C was correlated with arylsulfatase activity (r = 0.72,
P < 0.0001) correlation also was found between these two soil enzymes (r = 0.71,
P < 0.0001). Biological parameters measured were significantly correlated with total C
(Table 3.2). A strong correlation was obtained between cotton breaking ratio (CBR)
.4
Cony. till
Min. till
a
= 0.4
z0
H
E-4
0
U
0.2
0.0
Day 14
Day 21
Can-Yr 1
Day 7
Day 14
Can-Yr 2
Fig. 3.4. Tillage effect on cotton strip decomposition at on-farm research site. Within each day,
treatment bars with the same letter are not significantly different (P < 0.05).
67
and cotton strip weight loss at seven days (r = 0.82; P = 0.0001), but not at the fourteen
days burial period (r = -0.083; P = 0.53).
Table 3.2. Pearson coefficients (r) for association between biological parameters and
Total C.
Arylsulfatase
0.72
r
p-value
0.0001
13-glucosidase
0.72
0.0001
CBR 7 days
0.37
0.0037
MBc
0.56
0.0001
Respiration
-0.44
0.001
Results from this study showed that the soil biological properties arylsulfatase,
13-glucosidase and MBc are sensitive to tillage within one year after tillage treatment
were imposed. This may confirm the potential use of these indexes as soil quality
indicators. Beta-glucosidase activity tended to be more stable than arylsulfatase and
MBc suggesting that it might be a better indicator because it seems to be less subject to
spatial or temporal variability which is an important factor in defining soil quality
indicators.
LITERATURE CITED
Angers, D.A., N. Bissonnette, A. Legere, and N. Samson. 1993. Microbial and
biochemical changes induced by rotation and tillage in soil under a barley
production. Can. J. Soil Sci. 73:39-50.
Campbell, C.A., V.O. Biederbeck, M. Schnitzer, F. Selles, and R.P. Zentner. 1989.
Effect of 6 years of zero tillage and N fertilizer management on changes in soil
quality of an orthic brown chernozem in southeastern Saskatchewan. Soil Till.
Res. 14:39-52.
Carter, M. R. 1986. Microbial biomass as an index of tillage induced changes in soil
biological properties. Soil Till. Res. 7: 29-40.
Dick, R.P., P.E. Rasmusssen, and E.A. Kerle. 1988. Influence of long-term residue
management on soil enzyme activities in relation to soil chemical properties of
a wheat-fallow system. Biol. Fertil. Soil. 6:159-164.
68
Dick, W.A. 1984. Influence of long term tillage and crop rotation combinations on soil
enzyme activities. Soil Sci. Soc. Am. J. 48:569-574.
Doran, J.W. 1980. Soil microbial and biochemical changes associated with reduced
tillage. Soil Sci. Soc. Am. J. 44:765-771.
Doran, J.W. 1987. Microbial biomass and mineralizable nitrogen distributions in
no-tillage and ploughed soil. Biol. Fert. Soils. 5:68-75.
Doran, J.W., and M.S. Smith. 1987. Organic matter management and utilization of soil
fertilizer nutrients. p. 53-72. In J.J. Mortvedt et al. (eds.) Soil fertility and
organic matter as critical component of production systems. SSSA Spec. Publ.
19, SSSA, Madison, WI.
Elliot, E.T. 1986. Aggregate structure and carbon, nitrogen and phosphorus in native
cultivated soils. Soil Sci. Soc. Am. J. 50:627-633.
Gupta, V.V.S.R. and J.J. Germida. 1988. Distribution of microbial biomass and its
activity in different soil aggregate size classes as affected by cultivation. Soil
Biol. Biochem. 20:777-786.
Gupta, V.V.S.R., and M.M. Roper. 1992. Influence of crop residues management
practices on microbial biomass and nutrient availability. Australian
Microbiology 1:3-21.
Gupta, V.V.S.R., M.M. Roper, J.A. Kirkegaard and J.F. Angus. 1994. Changes in
microbial biomass and organic matter levels during the first year of modified
tillage and stubble management practices on a red earth. Aust. J. Soil Res.
32:1339-1354.
Khan, S.U. 1970. Enzymatic activity in gray wooded soil as influenced by cropping
systems and fertilizers. Soil Biol. Biochem. 2:137-139.
Klein, T.M., and Koths, J.S. 1980. Urease, protease and acid phosphatase in soil
continuously cropped to corn by conventional or no-tillage methods. Soil Biol.
Biochem. 12:293-294.
Miller, M., and R.P. Dick. 1995. Dynamics of soil C and microbial biomass on whole
soil aggregates in two cropping systems differing in C-input. Appl. Soil Ecol.
2:253-261.
SAS. 1996. Statistical Analysis Software. Cary, NC, SAS Institute Inc.
Van Veen J.A. and E.A. Paul. 1981. Organic carbon dynamics in grass land soil. I.
Background information and computer simulation. Can. J. Soil Sci. 61:185-201.
69
CHAPTER 4
COTTON STRIP ASSAY AS A SOIL BIOLOGICAL INDICATOR:
A LABORATORY EXPERIMENT
70
ABSTRACT
A laboratory incubation experiment was conducted to determine the effect of
soil moisture and N content on cotton strip decomposition and to assess the degree of
correlation between cotton strip decomposition and cotton strip weight loss after
incubation. Two soil types, a Willamette silt loam and a Chehalis silt loam, were taken
at 0 to 20 cm depth in December 1997. After being passed through a 5-mm sieve, 2 kg
(dry weight basis) of soil were placed in plastic containers and adjusted with the proper
amount of deionized water. The treatments were N rate (0 or 100 lig N g-1 as urea) and
soil water content (0.16 or 0.25 m3 m-3). Containers were placed in an incubator at
18°C or 25°C, after insertion of cotton strips, for 7, 14 and 21 days. Decomposition
rates were measured as loss in tensile strength and weight loss of buried cotton strips.
Results indicated that high soil water and N content significantly accelerated cotton
strip decomposition. Cotton strip decomposition was positively correlated with cotton
strip weight loss and13-glucosidase activity. The results indicate that it is important to
use cotton strips under the same environmental conditions or season for it to provide
comparable results from year to year.
INTRODUCTION
Agricultural management systems are sustainable only when soil quality is
maintained or enhanced; it is therefore important to quantitatively assess changes in soil
quality and evaluate sustainable land and soil management systems (Doran and Parkin,
1994). Potential soil quality indexes integrating physical, chemical and biological soil
properties are being defined and tested by soil scientists. To be useful, soil quality
indicators should reflect recent changes in soil management. Among the potential
71
biological indicators of soil quality are respiration, microbial biomass, soil enzyme
activities and soil fauna. Most of these methods are laboratory techniques that may be
labor intensive or require expensive equipment.
Rates of organic matter decomposition are likely related to soil health, quality
and productivity, and are indicative of soil biological activity and biomass. Simple
techniques that may reflect organic matter decomposition are the buried litter bags or
standard cotton strips. The standard cotton strip technique originally was developed for
use in the International Biological Programme (Latter and Walton, 1988) to assess
decomposition rates in a variety of soils. The cotton strip assay, also known as calico
cloth, has been used effectively to characterize decomposition potential among different
forest soils (Heal et al., 1981; Hill et al., 1985; Maltby, 1988). It has potential
advantages over plant litter decomposition because cotton strips are uniform, easier to
manage, in direct contact with soil and decompose faster and therefore need a shorter
burial period (Rosek et al., 1996). This technique would be attractive for assessing soil
quality in agricultural soils, but relatively few studies have been done on cotton strips in
these systems.
Preliminary results have shown that cotton strip decomposition is sensitive to
changes in nitrogen (N) fertilizer source (chemical vs. manure), tillage or incorporation
of legume in the crop rotation (Rosek et al., 1996). Williamson (1994) showed that
cotton strip breakdown is able to reflect differences in the soil physical, chemical and
biological properties under minimum tillage in comparison to conventional tillage. Soil
moisture and aeration as well as temperature and available N (Lande, 1969) influence
cellulose decomposition (Williams and Crawford, 1983).
72
The hypothesis of our study was that increases in soil moisture and N increase
the rate of decomposition because of a more favorable environment for microbial
activity. The objectives of our study were to: (1) determine the effect of soil N and
water content on cotton strip decomposition under controlled laboratory conditions; (2)
assess the degree of correlation between tensile strength and cotton strip and (3) assess
relationships between cotton strips decomposition and microbial properties of soils.
MATERIALS AND METHODS
Soils
Soil samples were taken from the 0 to 20 cm depth of two locations in
December 1997: the Oregon State University (OSU) Vegetable Farm and the North
Willamette Research and Extension Center (NWREC) in Oregon. The study at the
OSU Vegetable Farm, initiated in 1993, is a long-term experiment on a Chehalis silt
loam soil (Cumulic Ultic Haploxeroll) in the southern Willamette Valley. Soil samples
were taken with a shovel from the winter fallow plots.
The NWREC, initiated in 1989, is on a Willamette silt loam (Pachic Ultic
Argixeroll) soil at Aurora, OR. Soil samples were taken with a shovel from the N zero
winter fallow plots.
Field-moist soil was homogenized and sieved moist through a 5-mm mesh sieve.
Two kg of soil (dry weight basis) were placed in plastic containers (34x20x11 cm) for
each treatment (3 replications). The treatments were water content (0.16 or
0.25 m3 m-3) and N rate (0 or 100 [tg NH2CONH2-N g -1). The containers were covered
with a lid and placed for one week in incubators maintained at 18 or 25°C before
73
insertion of cotton strips. Water content was adjusted with deionized water by
gravimetry every two days.
Cotton strip incubation study
For the burial test, a standardized cotton fabric (Sagar, 1988) was obtained from
the Shirley Institute in Manchester, England. Cotton strips were cut to 14x7.5 cm to
fit tensiometer jaws and weighed before and after incubation. A representative soil
subsample (15 g dry-weight) was obtained for each treatment for biological measure­
ments before insertion of cotton strip (day 0) and at 14 days after burial. Three cotton
strips were placed horizontally in each container so that about 1 cm of soil was beneath
the strips and 2 cm of soil covered them. At day 7, 14 and 21, one cotton strip was
removed randomly from each container and weighed. Soil water content was adjusted
gravimetrically every two days. Control strips and buried cotton strips were washed
gently, laid on paper towel and oven-dried at 85°C for 45 min. Tensile strength of each
cotton strip was measured using the Instron tensiometer (Instron Corporation, Canton,
Massachusetts). Cotton strip decomposition was measured as Cotton Breaking Ratio
(CBR) based on the loss in tensile strength of buried cotton strip (adapted from
Williamson, 1994) according to the following equation:
CBR = TSUC-TSDC) / TSUC
where TSUC = Tensile Strength Undecomposed Cotton strip (kN m kg-1); and
TSDC = Tensile Strength Decomposed Cotton strip (kN m kg-1).
74
Biological measurements
Microbial biomass C (MBc) and B-glucosidase activity were measured at day 0
(before insertion of cotton strips) and at 14 days after incubation. Microbial biomass C
was determined by chloroform-fumigation incubation (Jenkinson and Powlson, 1976)
method modified as follows. Ten grams of soil were weighed into scintillation vials.
The vials were placed in a dessicator containing wet paper towels at the bottom and a
50-mL beaker containing 30 mL of ethanol-free chloroform and few boiling chips. The
dessicator was evacuated and the soil exposed to chloroform vapor for 24 hours. Soil
was then transferred into 72-mL acrylic tubes fitted with rubber septa at each end to
allow sampling for gas chromatography. Samples were incubated at 25°C in the dark
for ten days. After incubation, CO2 produced was measured by gas chromatography. A
Kc of 0.41 (Voroney and Paul, 1984) was used to calculate MBc without subtracting the
control. Beta-glucosidase activity was measured on moist soil following the method
described by Tabatabai (1994) where the product measured was lig PNP g soil
Experimental design and statistical analysis
The design of the study was a split plot (three replications) with factorial
combinations (22x31) of two factors (water content, N rate) as main plot and one factor
(time) as a subplot (three sampling periods).
Statistical analyses were performed by analysis of variance using the Mixed
procedure in SAS (SAS Inst. 1996) with water and N treatments as a between-groups
factors and day as a within groups. Treatment means were separated by least significant
difference. Correlation analysis was used to evaluate the association between CBR and
75
cotton strip weight loss. A logarithmic transformation of the data response variable
(cotton strip weight loss) was applied before the fitting the regression model.
RESULTS
A strong influence of temperature and soil type on cotton strip decomposition
was observed during the experiment; however, no inference can be made because these
two factors were not replicated. The main effects of
N, soil
moisture and time on the
Chehalis soil and possible interactions are reported in Table 4.1.
Table 4.1. Analysis of variance of soil environment effects on cotton strip
decomposition on a Chehalis silt loam soil.
Source of variation
Nitrogen (N)
Moisture (M)
Df
F value Pr > F
1
1.86
1
22.73
0.187
0.0001
1
4.73
0.041
Day (D)
1
154.39
0.0001
D x N
1
0.18
0.836
D x M
1
0.42
DxNxM
1
7.88
0.659
0.001
Nx
M
The cotton breaking ratio (CBR) on the Chehalis soil increased from day 7 to
21, but this increase was most evident between day 7 and 14 (Fig. 4.1). Nitrogen
addition significantly accelerated cotton strip decomposition when water was at a lower
water content. Temperature tended to increase cotton decomposition except when N
was added to soil at the higher water content. Soil water content had a significant effect
(P < 0.05) on MBC only in the presence of N whereas temperature seemed to have little
or no effect (Fig. 4.2).
76
1.0
aa
Day 21
0.8
0.6
0.4
0.2
r_-(1
0.0
0.8
Day 14
aa
0.6
0.4
GC1
z 0.2
O
0.0
u
0.8
iii
N =0µgNg 1
_1
N=100 ggNg
Day 7
a
0.6
0.4
a
0.2
0.0
18°C
0.16
25°C
18°C
25°C
0.25
WATER CONTENT (m3 m-3)
Fig. 4.1. Effect of temperature, soil moisture and N rate on cotton strip decomposition
in a Chehalis silt loam soil. Bars followed by the same letter within a soil
moisture, temperature and sampling time were not significantly different
at P < 0.05.
77
300
250
200
150
.:5)"
100
50
U
aA
0
250
gt1
a
200
0
aDay
N 0 N g1
a ®N= l00µgNg
a
a
150
100
50
0
18°C
0.16
25°C
25°C
18°C
0.25
WATER CONTENT (m3 m-3)
Fig 4.2. Effect of temperature, soil moisture and N rate on MBC in a Chehalis silt loam
soil. Bars followed by the same letter within a soil moisture, temperature and
sampling time were not significantly different at P < 0.05.
78
Soil enzyme activity measured, as 13- glucosidase, was relatively stable over time
(Fig. 4.3). Maximum activity was measured on the Chehalis soil that had the lowest
water content (0.16 m3m-3) at both incubation times regardless of temperature. Overall,
there was a non-significant effect of soil water content on13-glucosidase activity.
Main effects of N and water on the Willamette soil are presented in Table 4.2.
Table 4.2. Analysis of variance showing source of effects on cotton strip
decomposition on a Willamette silt loam soil.
Source of variation
Nitrogen (N)
Moisture (M)
Df
F value
1
13.11
1
3.72
Pr > F
0.002
0.068
NxM
Day (D)
1
0.054
0.541
1
186.48
0.0001
DxN
DxM
1
3.93
0.027
1
0.27
0.765
DxNxM
1
0.09
0.91
Cotton strip decomposition on the Willamette soil was generally greater with N
addition. The temperature effect diminished from day 7 to 21, especially at high water
content (Fig. 4.4). Soil water content significantly (P < 0.05) increased CBR when no
N was added (Fig. 4.4). Microbial biomass carbon slightly increased with increasing
soil moisture, but overall, neither soil water content nor temperature (Fig. 4.5)
influenced MBc.
Although a slight increase in 13- glucosidase activity was noted from day 0 to day
14 at lower soil water content (Fig. 4.6), an increase in soil water content significantly
(P < 0.05) decreased 13- glucosidase activity at day 14 when no N was added.
79
"--, 200
:-.
5
160
120
P-4
Z
r=k
al)
=..
W
rA
80
40
0
aDay 0
.< 160
Q
'-ci-D'
a
N = 0 p.g N g1
f2Z2N = 100 pg 11 g-1
120
0
0 80
..-4
(....
40
0
18°C
0.16
25°C
18°C
25°C
0.25
WATER CONTENT (m3 m-3)
Fig 4.3. Effect of temperature, soil moisture and N rate on13-glucosidase activity
in a Chehalis silt loam soil. Bars followed by the same letter within a soil
moisture, temperature and sampling time were not significantly different
at P < 0.05.
80
1.0
aa
Day 21 a
0.8
0.6
0.4
E-2, 0.2
0.0
a
Day 14
0.8
0.6
At
0.4
z 0.2
0
E--4 0.0
H
C
L)
0.8
0.6
ggNg-1
iii N=0
N= 100 ggNg 1d
b
a
Day 7
a
a
0.4
0.2
0.0
18°C
0.16
25°C
18°C
25°C
0.25
WATER CONTENT (m3 m-3)
Fig. 4.4. Effect of temperature, soil moisture and N rate on cotton strip decomposition
in a Willamette silt loam soil. Bars followed by the same letter within a soil
moisture, temperature and sampling time were not significantly different
at P < 0.05.
81
300
250
200
150
.31
100
b.()
50
0
E 250
u 200
150
100
50
0
18°C
25°C
0.16
25°C
18°C
0.25
WATER CONTENT (m3 m-3)
Fig 4.5. Effect of temperature, soil moisture and N rate on Mc in a Willamette silt loam
soil. Bars followed by the same letter within a soil moisture, temperature and
sampling time were not significantly different at P < 0.05).
82
200
160
120
Ito
P-+
bA
80
40
0
160
7) 120
O
80
40
c
0
18 °C
25 °C
0.16
18 °C
25 °C
0.25
WATER CONTENT (m3 m-3)
Fig 4.6. Effect of temperature, soil moisture and N rate on 13-glucosidase activity
in a Willamette silt loam soil. Bars followed by the same letter within a soil
moisture, temperature and sampling time were not significantly different
at P < 0.05.
83
Correlation analysis at 14 days showed a significant association between CBR
and B-glucosidase activity (r = 0.53, P = 0.0001), and soil water content (r = 0.37;
P = 0.01). There was, however, a poor correlation between CBR and MBA or
respiration (r = -0.11, P = 0.47; r = 0.16, P = 0.3).
Cotton breaking ratio was significantly correlated with cotton strip weight loss
after adjusting for weight before incubation. There was a positive linear relationship
between CBR and cotton strip weight loss (r = 0.86; P = 0.0001). The estimated
relationship between weight loss and CBR is:
{log (weight loss)/CBR} = -3.4697 + 2.87 CBR
DISCUSSION
Nitrogen addition was found to increase cotton strip decomposition but was
dependent on soil water content. This could be expected because soils are often
N-limited for biological growth. A large number of studies have shown that
when crop residues with a wide C:N ratio are added to the soil, the rate and extent of
their decomposition can be increased by addition of inorganic N (Parr et al., 1970).
This is analogous to cotton strip that would have a wide C:N ratio. These findings
follow other studies on cellulose decomposition where it has been shown that micro­
organisms use the soil N, which drives decomposition (Hill, et al., 1985; French, 1988;
Ineson et al., 1988). Nutrient availability enhances microbial growth, therefore
increases decomposition of organic compounds in soil. Our results agree with those of
Widden (1986) in his study relating fungal community structure to cellulose decomposi­
84
tion rates in the field; he reported greater tensile strength losses of cotton strip in soil
amended with N and calcium (Ca) than with controls.
The Chehalis and Willamette soils, with or without N addition, seemed to
increase CBR when temperature increased. These observations would be similar to
those of Donnelly et al. (1990) who reported temperature as the dominant factor
affecting cellulose and soil organic matter decomposition (Dickinson, 1974; Oades,
1988). Because temperature has a big effect on the growth and activity of micro­
organisms, it would be expected to control the decomposition rate of residues and of
cellulose. Williamson (1994) reported that soil water content and temperature are the
most critical components governing cotton strip decomposition. Temperature is one of
the common site factors that increase cellulose decomposition in agricultural fields
(Trettin et al., 1996).
Smith and Maw (1988) found a relationship of tensile strength and an increase
in metabolic activity with an increase in FDA hydrolysis that occurred rapidly during
the first 7 days of incubation. Our results generally did not show a significant increase
in 13-glucosidase activity from day 0 to day 14. Generally biological activity in our
study was not affected by temperature. It may be that temperatures used in this study
were in the optimal range for biological activity. In contrast, soil moisture was more
important in affecting microbial biomass and activity; similar results were also found in
forest soils (Donnelly, 1990).
Our results showed that water content had an effect on N availability because
the N x M interaction was strongly significant. These observations would be expected
because microorganisms get their nutrients through water by mass flow or diffusion.
85
Therefore when water is limited, microorganisms need to spend more energy to
maintain water and carry out life processes. Soil water content generally had a
significant effect on MBA and soil enzyme activity, which would be expected because
water limits microbial growth. This provides evidence that soil moisture, by controlling
microbes, in turn affects cotton strip decomposition.
Our study showed that water and perhaps temperature are the main abiotic
factors influencing cotton strip decomposition; water availability, however, significantly
(P < 0.05) affected microbial activity. When water was sufficient, microorganisms
responded to an increase in temperature, but when water became limited, microbial
growth and activity was reduced.
In nature, lignin and other recalcitrant compounds often are associated with
cellulose, therefore decomposition rates will be slow especially at early stages of the
process. Water and temperature effects are more difficult to differentiate; few studies
have tried to determine the effect of each factor on cellulose decomposition but results
indicate that more research needs to be done to fully understand the major role of each
component.
Treatment differences generally were not significant at 21 days. This indicates
that given enough time, even under suboptimal condition moisture or temperature
regimes, decomposition will proceed. Decomposition rates were assessed in this study
with cotton strip made of 96% pure cellulose. This is an important finding for assessing
cotton strip as a soil biological indicator. Under optimal laboratory conditions, 7 days
are optimal for discriminating treatment effects. However, work from the field study
(Chapter 2) indicates 14 days may be best in conditions of western Oregon for showing
86
treatment effects. The longer time under field conditions maybe due to less ideal
conditions than in the laboratory. Field conditions may have dry periods or fluctuations
in temperature that may slow biological decomposition compared to the laboratory.
A strong association was found between CBR and cotton strip weight loss. This
is important because the weighing method could be readily adopted by farmers and
extension agents.
Besides temporal effects, this study showed the importance of moisture and N
availability. This has important ramifications for using cotton strips as a soil quality
indicator because there is seasonal variation in soil moisture and farmers apply varying
rates of N fertilizer according to crop recommendations. Therefore to have meaningful
and interpretable results from year to year, the cotton strip test should be run under
similar nutrient and environmental conditions.
LITERATURE CITED
Dickinson, C.H. 1974. Decomposition of litter in soil. p.633-658. In C.H. Dickinson
and G.J.F. Pugh (eds.) Biology of plant litter decomposition. Vol 2. Academic
Press, New York.
Donnelly, P.K., J.A.Entry, D.L. Crawford and K. Cromack Jr. 1990. Cellulose and
lignin degradation in forest soils: response to moisture, temperature and acidity.
Microbial Ecol. 20: 289-295.
Doran, J.W. and T.B. Parkin. 1994. Defining and assessing soil quality. p. 3-21. In
J.W. Doran, D.C. Coleman, D.F. Bezdicek and B.A. Stewart (eds.). Defining
soil quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
French, D.D. 1988b. Patterns of decomposition assessed by the use of litter bags and
cotton strip assay on fertilized and unfertilized heather moor in Scotland. p. 100­
108. In A.F. Harrison, P.M. Latter and D.W.H. Walton et al. (eds.) Cotton strip
assay: An index of decomposition in soils. Institute of Terrestrial Ecology,
Grange-Over-Sands, Cumbria, England.
87
Heal, 0.W., P.W. Flanagan., D.D. French and S.F. Maclean, Jr. 1981. Decomposition
and accumulation of organic matter. p. 587-634. In L.C. Bliss et al. (eds.)
Tundra ecosystem: A comparative analysis. Cambridge Univer. Press,
Cambridge, England.
Hill, M.O., P.M. Latter and G. Bancroft. 1985. A standard curve for inter-site
comparison of cellulose degradation using the cotton strip method. Can. J. Soil
Sci. 65:609-619.
Jenkinson, D.S. and D.S. Powlson. 1976. The effect of biocidal treatments on
metabolism in soil-V. A method for measuring soil biomass. Soil Biol.
Biochem. 8:209-213.
meson, P., P.J. Bacon, and D.K. Lindley. 1988. Decomposition of cotton strips in soil:
analysis of the world data set. p. 155-165. In A.F. Harrison, P.M. Latter and
D.W.H. Walton al. (eds.) Cotton strip assay: An index of decomposition insoils.
Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Lande, E. 1969. Biological activity in some natural and drained peat soils with special
reference to oxidation-reduction conditions. Acta For. Fenn. 94:1-69.
Latter, P.M. and D.W.H. Walton. 1988. The cotton strip assay for cellulose decomposi­
tion studies in soil: history of the assay and development. In A.F. Harrison, P.M.
Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition
in soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Maltby, E. 1988. Use of cotton strip assay in wetland and upland environments An
international perspective. p. 140-154. In A.F. Harrison and D.W.H. Walton
(eds.) Cotton strip assay: An index of decomposition in soils. Institute of
Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Oades, J.M. 1988. The retention of organic matter in soils. Biogeochem. 5:35-70.
Parr, J.F., S. Smith, and G.H. Willis. 1970. Soil Anaerobiosis: I. Effect of selected
environments and energy sources on respiratory activity of soil
microorganisms. Soil Sci. 110:37-43.
Rosek, M.J., J.C. Gardner and B.S. Miller. Influence of tillage and cropping system on
litter bag and cotton strip decomposition. Soil Quality Workshop, July 17-18,
1996 Ames, Iowa.
Sagar, B.F. 1988. The Shirley soil burial cloth test fabric and tensile testing as a
measure of biological breakdown of textiles. p.11-16. In A.F. Harrison, P.M.
Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition
in soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria,
England.
88
SAS. 1996. Statistical Analysis Software. Cary, NC, SAS Institute Inc.
Smith, R.N., and J.M. Maw.1988. Relationship between tensile strength and increase
in metabolic activity on cotton strips. p 55-59. In A.F. Harrison, P.M. Latter
and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition in
soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Tabatabai, M.A. 1994. Soil enzymes. p. 775-833. In R.W. Weaver (eds.) Methods of
Soil Analysis, Part 2. Microbiological and Biochemical Properties. SSSA Book
Ser. 5. SSSA, Madison,WI.
Trettin, C.C., M. Davidian, M.F. Jurgensen, and R. Lea. 1996. Organic matter
decomposition following harvesting and site preparation of a forested wetland.
Soil Sci. Soc. Am. J. 60:1994-2003.
Voroney, R.P., and E.A. Paul. 1984. Determination of IQ and KN in situ for calibration
of the chloroform fumigation-incubation method. Soil Biol. Biochem. 16:9-14.
Widden, P. 1986. Use of cotton strips to relate fungal community structure to cellulose
decomposition rates in the field. Short comm. Soil Biol. Biochem. 18:335-337.
Williams, E.S., and R.L. Crawford. 1983. Effects of various physiochemical factors on
microbial activity in peatlands: Aerobic biodegradation processes. Can. J.
Microbiol. 29:1430-1437.
Williamson, J.R. 1994. Calico cloth decomposition as an indicator of topsoil health
p.163-165. In Pankhurst et al. (eds.) Soil Biota: Management in Sustainable
Farming Systems, Poster Workshop Proc., March 15-18, 1994 Adelaide, S.
Australia, CSIRO Inform. Services, E. Melbourne, Victoria, Australia.
89
CHAPTER 5
SUMMARY
90
The first objective of this study was to identify sensitive biological indicators of
soil quality to changes in soil management by cover crops and to test potential of the
buried cotton strip as an indicator of soil biological activity and as a soil quality index.
We hypothesized that winter cover cropping would increase soil biological properties
such as microbial biomass, soil enzyme activities and decomposition of cotton strips
due to year-round vegetative cover and greater C input with cover cropping.
Our results showed overall that winter cover cropping significantly affected soil
enzyme activity and MBA. Surprisingly greater microbial activity was measured in
spring than in summer. Responses of mineralizable N and cotton strip decomposition to
winter cover crop were highly variable and often not significant. It was determined
under field conditions that the optimal time for cotton strip burial was 14 days.
The objective outlined in Chapter 3 was to examine the effect of conventional
and minimum tillage on soil biological properties under agricultural fields managed
with cover crops. The less-disturbed minimum-tillage system had a significant increase
in biological activity over conventional tillage. It is, however, necessary to repeat this
study on various soil types.
Our final objectives were to determine the effect of soil water and N content on
cotton strip decomposition buried in two different soils for 7, 14 and 21 days; and to
assess the degree of correlation between cotton strip decomposition and cotton strip
weight loss. This study showed moisture and N content accelerated decomposition of
cotton strip. There was a significant correlation between tensile strength loss and cotton
strip weigh loss. Although decomposition of cotton strip was found not to be a good
indicator of soil quality because the influence of abiotic factors, it still can be used by
91
growers as a quick and easy test of general biological activity by just weighing the
strips before and after removal from soil. Under optimal laboratory conditions, 7 days
were sufficient for discriminating treatment effects.
92
BIBLIOGRAPHY
Angers, D.A., N. Bissonnette, A. Legere, and N. Samson. 1993. Microbial and
biochemical changes induced by rotation and tillage in soil under a barley
production. Can. J. Soil Sci. 73:39-50.
Bandick, A.K. and R.P. Dick. 1998. Field management effects on soil enzyme activities.
Soil Biol. Biochem. In Press.
Barr, A.M.R. 1988. The colonization and decay of cotton by fungi in soil burial tests
used in the textile industry. p 50-54. In A.F. Harrison, P.M. Latter and D.W.H.
Walton et al. (eds.) Cotton strip assay: an index of decomposition in soils.
Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Biederbeck, V. 0., C.A. Cambpell, and R.P. Zenter. 1984. Effect of crop rotation and
fertilization on some biological properties of a loam in southwestern
Saskatchewan. Can. J. Soil. Sci. 64:355-367.
Bolton, J., L.F. Elliott, P.R. Papendick, and D.F. Bezdicek. 1985. Soil microbial
biomass and selected soil enzyme activities: Effect of fertilization and
cropping practices. Soil Biol. Biochem. 17: 297-302.
Brady, N.C., and R.R. Weil. 1996. The nature and properties of soils. 11 ed. Prentice
Hall Upper Saddle River, New Jersey.
Brandi-Dohrn, F.M., R.P. Dick, M. Hess, S.M. Kauffman, D.D. Hemphill, Jr., and J.S.
Selker. 1997. Nitrate leaching under cereal rye cover crop. J. Environ. Qual.
26:181-188.
Brookes, P.C. 1995. The use of microbial parameters in monitoring soil pollution by
heavy metals. Biol. Fert. of Soils. 19:269-279. Biol. Fert. of Soils. 19:269-279.
Brown, A.H.F. 1988. Discrimination between the effects of soils of 4 tree species in
pure and mixed stands using cotton strip assay. p. 80-85. In A.F. Harrison, P.M.
Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition
in soils, ITE Symposium 24, Institute of Terrestrial Ecology, Grange-Over
Sands, Cumbria. England.
Buchanan, M., and L.D. King. 1992. Seasonal fluctuations in soil microbial biomass
carbon,phosphorus, and activity in no-till and reduced-chemical-input maize
agroecosystems. Biol. Fertil. Soils. 13:211-217.
Burket, J.Z., D.D. Hemphill, Jr., and R.P. Dick. 1997. Winter cover crops and nitrogen
management in sweet corn and broccoli rotations. HortSci. 32: 664-668.
93
Burns, R.G. 1982. Enzyme activity in soil: location and a possible role in microbial
ecology. Soil Biol. Biochem. 14: 423-27.
Campbell, C.A. 1978. Soil organic carbon, nitrogen and fertility. Dev. Soil Sci.
8:173- 271.
Campbell, C.A., M. Schnitzer, G.P. Lafond, R.P. Zentner, and J.E. Knipfel. 1991.
Thirty year crop rotations and management practices effects on soil and amino
nitrogen. Soil Sci. Soc. Am. J. 55:739-745.
Campbell, C.A., V.O. Biederbeck, M. Schnitzer, F. Se lles, and R.P. Zentner. 1989.
Effect of 6 years of zero tillage and N fertilizer management on changes in soil
quality of an orthic brown chernozem in southeastern Saskatchewan. Soil Till.
Res. 14:39-52.
Carter, M. R. 1986. Microbial biomass as an index of tillage induced changes in soil
biological properties. Soil Till. Res. 7: 29-40.
Carter, M.R. 1991. The influence of tillage on the proportion of organic carbon and
nitrogen in the microbial biomass of medium textured soils in a humid climate.
Biol. Fertil. Soils. 11:135-139.
Carter, M.R. and D.A. Rennie, 1982. Changes in soil quality under zero tillage
farming systems: Distribution of microbial biomass and mineralizable C and N
potentials. Can. J. Soil Sci. 62:587-597.
Collins, H.P., P.E. Rasmussen, and C.L. Douglas, Jr. 1992. Crop rotation and residue
management effects on soil carbon and microbial biomass. Soil Sci. Soc. Am. J.
56:783-788.
Coulson, J.C. and J. Butterfield. 1978. An investigation of the biotic factors
determining the rates of plant decomposition on blanket bogs. J. Ecol.
66:631-650.
Dick, R.P., P.E. Rasmusssen, and E.A. Kerle. 1988. Influence of long-term residue
management on soil enzyme activities in relation to soil chemical properties of
a wheat-fallow system. Biol. Fertil. Soil. 6:159-164.
Dick, R.P. 1994. Soil enzyme activities as indicators of soil quality. p. 107-124. In J.W.
Doran, D.C. Coleman, D.F. Bezdicek and B.A. Stewart (eds.) Defining soil
quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
Dick, W.A. 1984. Influence of long term tillage and crop rotation combinations on soil
enzyme activities. Soil Sci. Soc. Am. J. 48:569-574.
94
Dickinson, C.H. 1974. Decomposition of litter in soil. p.633-658. In C.H. Dickinson
and G.J.F. Pugh (eds.) Biology of plant litter decomposition. Vol 2. Academic
Press, New York.
Dillard, H.R. and R.G. Grognan. 1985. Influence of green manure crop and lettuce on
sclerotial minor. Plant Dis. 69:579-582.
Ditsch, D.C., M.M. Alley, K.R. Kelly, and Y.Z. Lei. 1993. Effectiveness of winter rye
for accumulation residual fertilizer N following corn. J. Soil and Water Cons.
48:125-132.
Donnelly, P.K., J.A. Entry, D.L. Crawford and K. Cromack Jr. 1990. Cellulose and
lignin degradation in forest soils: response to moisture, temperature and acidity.
Microbial Ecol. 20: 289-295.
Doran, J.W. 1980. Soil microbial and biochemical changes associated with reduced
tillage. Soil Sci. Soc. Am. J. 44:765-771.
Doran, J.W. 1987. Microbial biomass and mineralizable nitrogen distributions in
no-tillage and ploughed soil. Biol. Fert. Soils. 5:68-75.
Doran, J.W., D.C. Coleman, D.F. Bezdicek, and B.A. Stewart. 1994. Defining soil
quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
Doran, J.W. and T.B Parkin. 1994. Defining and assessing soil quality. p. 3-21. In J.W.
Doran, D.C. Coleman, D.F. Bezdicek and B.A. Stewart (eds.) Defining soil
quality for a sustainable environment. SSSA Spec. Publ. 35. Madison, WI.
Doran, J.W., and M.S. Smith. 1987. Organic matter management and utilization of soil
fertilizer nutrients. In J.J. Mortvedt et al. (eds.) Soil fertility and organic matter
as critical component of production systems. p. 53-72. SSSA Spec. Publ. 19,
SSSA: Madison, WI.
Elliot, E.T. 1986. Aggregate structure and carbon, nitrogen and phosphorus in native
cultivated soils. Soil Sci. Soc. Am. J. 50:627-633.
Frankenberger Jr., W.T. and W.A. Dick. 1983. Relationships between enzyme
activities and microbial growth and activity indices in soil. Soil Sci. Soc. Am.
J. 47:945-951.
French, D.D. 1988a. Some effects of changing soil chemistry on decomposition ofplant
litters and cellulose on a Scottish moor. Oecologia 75:608-618.
95
French, D.D. 1988b. Patterns of decomposition assessed by the use of litter bags and
cotton strip assay on fertilized and unfertilized heather moor in Scotland. p. 100­
108. In A.F. Harrison, P.M. Latter and D.W.H. Walton (eds.) Cotton strip assay:
an index of decomposition in soils, ITE Symposium 24, Institute of Terrestrial
Ecology, Grange-Over-Sands, Cumbria. England.
Goyal, S., M.M. Mishra, S.S. Dhankar, K.K. Kapoor, and R. Batra. 1993. Microbial
biomass turnover and enzyme activities following the application of farmyard
manure soil field oils with and without previous long-term applications. Biol.
Fert. Soils. 15:777-786.
Gupta, V.V.S.R. and J.J. Germida. 1988. Distribution of microbial biomass and its
activity in different soil aggregate size classes as affected by cultivation. Soil
Biol. Biochem. 20:777-786.
Gupta, V.V.S.R., and M.M. Roper. 1992. Influence of crop residues management
practices on microbial biomass and nutrient availability. Australian
Microbiology 1:3-21.
Gupta, V.V.S.R., M.M. Roper, J.A. Kirkegaard and J.F. Angus. 1994. Changes in
microbial biomass and organic matter levels during the first year of modified
tillage and stubble management practices on a red earth. Aust. J. Soil Res.
32:1339-1354.
Hargrove, W.L. 1991. Cover crops for clean water. Soil Water Conserv. Soc. Ankeny,
IA.
Harper, L.A., P.F. Hendrix, G.W. Langdale, and D.D. Coleman. 1995. Clover
management to provide optimum nitrogen and soil water conservation. Crop
Sci. 35:176-182.
Harrison A.F., P.M. Latter and D.W.H. Walton 1988. Cotton strip assay: an index of
decomposition in soils. ITE Symposium 24, Institute of Terrestrial Ecology,
Grange-Over-Sands, Cumbria, England.
Haynes, R.J. and R.S. Swift. 1990. Stability of soil aggregates in relation to organic
constituent of soil water content. J. Soil Sci. 41:73-83.
Haynes, R.J., R.S. Swift, and R.C. Stephen. 1991. Influence of mixed cropping
rotations (pasture-arable) on organic matter content, water stable aggregation
and clod porosity in a group of soils. Soil Tillage Res. 19:77-87.
Heal, 0.W., P.W. Flanagan., D.D. French and S.F. Maclean, Jr. 1981. Decomposition
and accumulation of organic matter. p. 587-634. In L.C. Bliss et al. (eds.)
Tundra ecosystem: A comparative analysis. Cambridge Univer. Press,
Cambridge, England.
96
Heal, O.W., G. Howson, D.D. French and J.N.R. Jeffers. 1974. Decomposition of
cotton strips in tundra. p. 341-362. In A.J. Holding, O.W. Heal, S.F. MacLean
and P.W. Flanagan (eds.) Soil organisms and decomposition in tundra.
Stockholm: Tundra Biome Steering Committee.
Heichel, G.H., and D.K. Barnes. 1984. Opportunities for meeting crop nitrogen needs
from symbiotic nitrogen fixation. In D. Bezdicek and J. Power (eds.) Organic
farming: current technology and its role in a Sustainable Agriculture. ASA Spec.
Publ. 46. Madison, WI.
Hill, M.O., P.M. Latter and G. Bancroft. 1985. A standard curve for inter-site
comparison of cellulose degradation using the cotton strip method. Can. J. Soil
Sci. 65:609-619.
Horwath, W.R. and E.A. Paul. 1994. Microbial Biomass. p. 753-773. In R.W. Weaver
et al. (eds.) Methods of Soil Analysis, Part 2. Microbiological and Biochemical
Properties. SSSA Book Ser. 5. SSSA, Madison, WI.
Meson, P., P.J. Bacon, and D.K. Lindley. 1988. Decomposition of cotton strips in soil:
analysis of the world data set. p. 155-165. In A.F. Harrison, P.M. Latter and
D.W.H. Walton et al. (eds.) Cotton strip assay: An index of decomposition in
soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria,
England.
Jenkinson D.S. 1977. The soil biomass. NZ Soil News 25.
Jenkinson, D.S. and J.N. Ladd. 1981. Microbial biomass in soil: Measurement and
turnover. p.415-471. In E.A. Paul and J.N. Ladd (eds.) Soil Biochem.Vol. 5.
Marcel Dekker, New York.
Jenkinson, D.S. and D.S. Powlson. 1976. The effect of biocidal treatments on
metabolism in soil-V. A method for measuring soil biomass. Soil Biol.
Biochem. 8:209-213.
Johnston, A.E. 1986. Soil organic matter, effects on soils and crops. Soil Use Manage.
2:97-105.
Jordan, D. and R.J. Kremer. 1994. Potential use of soil microbial activity as an
indicator of soil quality. p. 243-249. In C.E. Pankhurst et al. (ed) Soil Biota:
Management in sustainable farming systems. CSIRO, Australia.
Keeney, D.R. 1982. Nitrogen availability indices. p. 711-733. In A.L. Page, R.H.
Miller, and D.R. Keeny (eds.) Method of soil analysis Part 2. 2nd ed. Agron.
Monogr. 9. ASA and SSA, Madison, WI.
97
Khan, S.U. 1970. Enzymatic activity in gray wooded soil as influenced by cropping
systems and fertilizers. Soil Biol. Biochem. 2:137-139.
Kiss, I., M. Dragan-Burlarda and D. Radulescu. 1975. Biological significance of
enzymes in soil. Adv. Agron. 27:25-87.
Klein, T.M., and Koths, J.S. 1980. Urease, protease and acid phosphatase in soil
continuously cropped to corn by conventional or no-tillage methods. Soil Biol.
Biochem. 12:293-294.
Kuprevich, V.F., and T.A. Shcherbakova. 1971. Comparative enzymatic activity in
diverse types of soil. p. 167-201. In A.D. McLaren and J.J. Skujins (eds.) Soil
Biochem. Vol. 2. Marcel Dekker, New York.
Lande, E. 1969. Biological activity in some natural and drained peat soils with special
reference to oxidation-reduction conditions. Acta For. Fenn. 94:1-69.
Lamb, J.A., G.A. Peterson, and C.R. Fenster. 1985. Fallow nitrate accumulation in a
wheat-fallow rotation as affected by tillage system. Soil Sci. Soc. Am. J.,
49:1441-1446.
Larson, W.E., and F.J. Pierce. 1991. Conservation and enhancement of soil quality. In
Evaluation of sustainable land management in developing world. Vol. 2.
IBSRAM Proc. 12 (2). Bangkok, Thailand. Int. Board for Soil Res.
Management.
Latter, P.M. and D.W.H. Walton. 1988. The cotton strip assay for cellulose
decomposition studies in soil: history of the assay and development. In A.F.
Harrison, P.M. Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of
decomposition in soils. Institute of Terrestrial Ecology, Grange-Over-Sands,
Cumbria, England.
Lynch, J.M., and L.M. Panting. 1980. Cultivation and the soil biomass. Soil Biol.
Biochem. 12:29-33.
Maltby, E. 1988. Use of cotton strip assay in wetland and upland environments- An
international perspectives. p. 140-154. In A.F. Harrison and D.W.H. Walton
(eds.) Cotton strip assay: An index of decomposition in soils. Institute of
Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Martens, R. 1995. Current methods for measuring microbial biomass C in soil:
Potentials and limitations. Biol. Fertil. Soils. 19:87-89.
Martinez, J. and G. Giraud. 1990. A lysimeter study of the effect of a ryegrass catch
crop during a winter wheat /maize rotation, on nitrate leaching and on the
following crop. J. Soil Sci. 41:5-16.
98
Martyniuk, S., and G.H. Wagner. 1978. Quantitative and qualitative examination of
soil microflora associated with different management systems. Soil Sci.
125:343-350.
McGill, W.B., K.R. Cannon, J.A. Robertson and F.A. Cook. 1986. Dynamics of soil
microbial biomass and water soluble organic C in Breton L after 50 years of
cropping to two rotations. Can. J. Soil Sci. 66:1-19.
Miller, M., and R.P. Dick. 1995. Dynamics of soil C and microbial biomass on whole
soil aggregates in two cropping systems differing in C-input. Appl. Soil Ecol.
2:253- 261.
Minshew, H.F. Nitrate leaching and model evaluation under winter cover crops. Oregon
State Univ. Personal communication. November 11, 1998.
National Research Council. 1993. Soil and water quality: An agenda for agriculture.
Natl. Acad. Press, Washington, DC.
Oades, J.M. 1988. The retention of organic matter in soils. Biogeochem. 5:35-70.
Obbard, J.P., and K.C. Jones. 1993. The use of the cotton-strip assay to assess
cellulose decomposition in heavy metal-contaminated sewage sludge-amended
soils. Environmental Pollution. 81:173-178.
Odell, R.T., S.W. Melsted, and W.M. Walker. 1984. Changes in organic carbon and
nitrogen of Morrow Plots soils under different treatments, 1904-1973. Soil Sci.
137:160-171.
Omay, A.B., C.W. Rice, L.D. Maddux and W.B. Gordon. 1997. Changes in soil
microbial and chemical properties under long-term crop rotation and
fertilization. Soil Sci. Soc. Am. J. 61:1672-1678.
Paul, E.A., K. Paustian, E.T. Elliot, C.V. Cole (eds.). 1997. Soil organic matter in
temperate agroecosystems: long-term experiments in North America. CRC
Press, Inc.
Parr, J.F., S. Smith, and G.H. Willis. 1970. Soil Anaerobiosis: I. Effect of selected
environments and energy sources on respiratory activity of soil micro­
organisms. Soil Sci. 110:37-43.
Perrucci, P. 1992. Enzyme activity and microbial biomass in field amended with
municipal refuse. Biol. Fert. Soils. 14:54-60.
Phillips, D.J., A.G. Watson, A.R. Weinhold, and W.C. Snyder. 1971. Damage of lettuce
seedlings related to crop residue decomposition. Plant Dis. Rep. 55:837-841.
99
Powlson, D.S., P.C. Brookes, B.T. Christensen. 1987. Measurements of soil microbial
biomass provides an early indication of changes in total soil organic matter due
to straw incorporation. Soil Biol. Biochem. 19:159-164.
Reeves, D.W. 1994. Cover crops and rotations. In J.C. Hatfield and B.A. Steward (eds).
Adv. Soil Sci.: Crop residues management. Lewis Publishers, Boca Raton,
Florida. p. 125-172.
Reganold, J.P. 1988. Comparison of soil properties as influenced by organic and
conventional farming systems. Am. J. Ahern. Agric. 3:144-155.
Rice, C.W., T.B. Moorman and M. Beare. 1996. Role of microbial biomass carbon
and nitrogen in soil quality p. 203-215. In J.W. Doran and A.J. Jones (eds.)
Methods of assessing soil quality. SSSA, Spec. Publ. 45. Madison, WI.
Roberson, E.B., S. Sarig and M.K. Firestone. 1991. Cover crop management of
polysaccharide-mediated aggregation in an Orchard soil. Soil Sci. Soc. Am. J.
55:734-739.
Rosek, M.J., J.C. Gardner and B.S. Miller. Influence of tillage and cropping system on
litter bag and cotton strip decomposition. Soil Quality Workshop, July 17-18,
1996 Ames, Iowa.
Rovira, A.D. 1953. A study of the decomposition of soil organic matter in red soils of
the Lismore district. Aust. Conf. Soil Sci., Adelaide, 1, 3.17, 1-4.
Sagar, B.F. 1988. The Shirley soil burial cloth test fabric and tensile testing as a
measure of biological breakdown of textiles. p.11-16. In A.F. Harrison, P.M.
Latter and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition
in soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria,
England.
SAS. 1996. Statistical Analysis Software. Cary, NC, SAS Institute Inc.
Smith, M.S., W.W. Frye and J.J. Varco. 1987. Legume winter cover crops. Adv. Soil
Sci. 7:95-139.
Smith, R.N., and J.M. Maw.1988. Relationship between tensile strength and increase
in metabolic activity on cotton strips. p. 55-59. In A.F. Harrison, P.M. Latter
and D.W.H. Walton (eds.) Cotton strip assay: an index of decomposition in
soils. Institute of Terrestrial Ecology, Grange-Over-Sands, Cumbria, England.
Spencer, B. 1958. Studies on sulphatases: 20. Enzymatic cleavage of arylhydrogen
sulphates in the presence of H2 180. Biochem. J. 69:155-159.
100
Stivers, L.J., and C. Sherman. 1991. Meeting the nitrogen needs of processing
tomatoes through winter cover cropping. J. Prod. Agric. 4:330-335.
Swift, M.J., O.W. Heal and J.M. Andersen. 1979. Decomposition in terrestrial
ecosystems. University of California Press, Berkley.
Tabatabai, M.A. 1994. Soil enzymes. p. 775-833. In R.W. Weaver (eds.) Methods of
Soil Analysis, Part 2. Microbiological and Biochemical Properties. SSSA Book
Ser. 5. SSSA, Madison,WI.
Tate, K.R., T.W. Speir, D.J. Ross, R.L. Parfitt, K.N. Whale and J.C. Cowling. 1991.
Temporal variations in some plant and soil P pools in two pasture soils of
widely different P fertility status. Plant and Soil 132: 219-232.
Tisdall, J.M., and J.M. Oades. 1982. Organic matter-stable aggregates in soils. J. Soil
Sci. 33:141-163.
Trettin, C.C., M. Davidian, M.F. Jurgensen, and R. Lea. 1996. Organic matter
decomposition following harvesting and site preparation of a forested wetland.
Soil Sci. Soc. Am. J. 60:1994-2003.
Van Veen J.A. and E.A. Paul. 1981. Organic carbon dynamics in grass land soil. I.
Background information and computer simulation. Can. J. Soil Sci. 61:185-201.
Voroney, R.P., and E.A. Paul. 1984. Determination of kc and kN in situ for calibration
of the chloroform fumigation-incubation method. Soil Biol. Biochem. 16:9-14.
Wade, G.C. 1947. Effect of some microorganisms on the physical properties of cotton
duck. J. Coun. Scient. Ind. Res. Aust. 20:459-467.
Walton, D.W.H. and D. Allsopp. 1977. A new test cloth for burial trials and other
studies on cellulose decomposition. Int. Biodeterior. Bull. 13:112-115.
Widden, P. 1986. Use of cotton strips to relate fungal community structure to cellulose
decomposition rates in the field. Short comm. Soil Biol. Biochem. 18:335-337.
Williams, E.S., and R.L. Crawford. 1983. Effects of various physiochemical factors on
microbial activity in peatlands: Aerobic biodegradation processes. Can. J.
Microbiol. 29:1430-1437.
Williamson, J.R. 1994. Calico cloth decomposition as an indicator of topsoil health
p. 163-165. In Pankhurst et al. (eds.) Soil Biota: Management in Sustainable
Farming Systems, Poster Workshop Proc., March 15-18, 1994 Adelaide, S.
Australia, CSIRO Inform. Services, E. Melbourne, Victoria, Australia.
101
Wyland, L.J., L.E. Jackson, and K.F. Schulbach. 1995. Soil-plant dynamics following
incorporation of a mature cereal rye cover crop in a lettuce production system.
J. Agr. Sci. 124:17-25.
102
APPENDICES
103
Appendix A. Winter cover crop effect on MiBc at each on-farm site
(* shows significance at P < 0.05).
525
Dickman
450
375
7.1
0
-7
Fallow
Cover crop
Lucht
300
t0
*
225
(..)
*
E 150
75
0
525
Ito fit\
"Vc\ "Vc-%
co ci4g
%
"k.sl°
cP -vs
Grover
co
"vc
cfr vs O'o cs"' vs
Hamlin
450
*
375
O
300
U 225
bA
5 150
75
0
";:c.
cz*:
c.,,P ,FS
\
rvc
$.0 cp.'0.
525
450
Hendricks
Pearmine
0 375
to 300
() 225
bA
5
150
75
0
%
GO
6-s
$9110
Cit,
.1;k:c
GO
sO
,tc.
104
Appendix B. Winter cover crop effects on B-glucosidase activity at each
on-farm site (* shows significance at P < 0.05).
300
Dickman
Lucht
- --- Winter fallow
240
Winter cover crop
cn 180
*
*
9,
`1.
120
aD
60
0
N.
cov coo
300
N.
6:4C'
_,N;VC
stro
co-­
Grover
4 ,t't
.45.
.45.
\
"VC
N.
;s
'N.
cfr .\*4
ce6V
Hamlin
240
0
taq
60
t0 4t\
0
c.
c,
cAql,
300
Hendricks
Pearmine
-7s_, 240
vii 180
t10
120
CL
60
0
-4{0
N
N.
-VC-
\
" 1.
CibY-
S'T°
Gam
-503"
105
Appendix C. Winter cover crop effects on arylsulfatase activity at each on-farm site
(* shows significance at P < 0.05).
300
Dickman
240
Lucht
-- Winter fallow
Winter cover crops
*
120
bn
*
=- 60
0
Cat'
300
-.Vc%
cif; c,0 AFP- co0;
.csAsfa.s(
Grover
4s
4sl)
css- cA5
.ccAs..b.'''
Hamlin
0
180
120
bn
60
tti
cV
54% c rt
1,
0
:N.
...VC
ct.v
\
-.VC
cP"
300
Hendricks
Pearmine
240
O
Cl)
180
120
on
60
0
0
co
SO
­
$0; CO .4csfa.4'
:4 240
0
n.
co
%NO
n.