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2002 Enzyme activities in soils as affected by management practices Mine Ekenler Iowa State University
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ProQuest Information and Learning 300 North Zeeb Road, Ann Arbor, Ml 48106-1346 USA 800-521-0600
Enzyme activities in soils as affected by management practices
by
Mine Ekenler
A dissertation submitted to the graduate faculty in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY
Major: Soil Science (Soil Microbiology and Biochemistry)
Program of Study Committee: M.A. Tabatabai, Major Professor Richard M. Cruse Thomas E. Fenton Anthony L. Pometto III Louisa B.Tabatabai
Iowa State University Ames, Iowa 2002 UMI Number: 3073445
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ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, Ml 48106-1346 ii
Graduate College Iowa State University
This is to certify that the doctoral dissertation of Mine Ekenler has met the dissertation requirements of Iowa State University
Signature was redacted for privacy.
Major Professor
Signature was redacted for privacy.
For the Major Program iii
DEDICATION To my parents: Muzaffer and Giillu Ekenler and My son: Deniz Jason Ekenler-Borges IV
TABLE OF CONTENTS
ABSTRACT viii
CHAPTER 1. GENERAL INTRODUCTION 1 Organization of Dissertation 5
CHAPTER 2. LITERATURE REVIEW 7 Soil Enzymes 7 Sources and States of Enzymes in Soils 8 Types of Enzymes in Soils 12
Glycosidases 14
Arylamidase 17
Amidohydrolases 19
Phosphatases 22
Arylsulfatase 24
Soil Health and Quality 25 Effects of Management Practices on Microbial Biomass and Enzyme Activities 29 Effects of liming on enzyme activities 29 Effects of cropping systems and N fertilization on enzyme activities 32 Effects tillage and residue management on enzyme activities 33 Nitrogen Mineralization 35 Indexes of available nitrogen 35 References 41 V
CHAPTER 3. EFFECTS OF LIMING AND TILLAGE SYSTEMS ON MICROBIAL BIOMASS AND GLYCOSIDASES IN SOILS 65 Abstract 65 Introduction 67 Materials and Methods 71 Results 76 Discussion 79 Conclusions 85 References 86
CHAPTER 4. ARYLAMIDASE AND AMIDOHYDROLASES IN SOILS AS AFFECTED BY LIMING AND TILLAGE SYSTEMS 107 Abstract 107 Introduction 108 Materials and Methods 113 Results 116 Discussion 119 Conclusion 123 References 125
CHAPTER 5. RESPONSES OF PHOSPHATASES AND ARYLSULFATASE IN SOILS TO LIMING AND TILLAGE SYSTEMS 139 Abstract 139 Introduction 141 Materials and Methods 145 Results 147 Discussion 151 vi
Conclusions 155 References 156
CHAPTER 6. (3-GLUCOSAMINIDASE ACTIVITY OF SOILS: EFFECT OF CROPPING SYSTEMS AND ITS RELATIONSHIP TO NITROGEN MINERALIZATION 172 Abstract 172 Introduction 173 Materials and Methods 176 Results 180 Discussion 182 References 186
CHAPTER 7. EFFECTS OF TRACE ELEMENTS ON p-GLUCOSAMINIDASE ACTIVITY IN SOILS 204 Abstract 204 Introduction, Materials and Methods, Results and Discussion 205 References 212
CHAPTER 8. TILLAGE AND RESIDUE MANAGEMENT EFFECTS ON p-GLUCOSAMINIDASE ACTIVITY IN SOILS 220 Abstract 220 Introduction, Materials and Methods, Results and Discussion 221 References 231
CHAPTER 9. p-GLUCOSAMINIDASE ACTIVITY AS AN INDEX OF NITROGEN MINERALIZATION IN SOILS 239 Abstract 239 Introduction 240 vii
Materials and Methods 243 Results and Discussion 245 References 248
CHAPTER 10. GENERAL CONCLUSIONS 260 APPENDIX. ADDITIONAL TABLES 267 ACKNOWLEDGEMENTS 279 viii
ABSTRACT
Studies were undertaken to investigate the long-term effects of lime application and tillage systems (no-till, ridge-till, and chisel plow) on soil microbial biomass C (Cmic) and N (Nmic) and the activities of glycosidases (a- and
P-glucosidases, a- and p-galactosidases and p -glucosaminidase); phosphatases
(acid and alkaline phosphatases and phosphodiesterase); amidohydrolases (L- asparaginase, L-glutaminase, amidase, urease, and L-aspartase); and arylamidase at their optimal pH values. With the exception of acid phosphatase, which was significantly but negatively correlated, all other enzyme activities were significantly and positively correlated with soil pH values at four sites in
Iowa. A activity/A pH values showed that among the enzymes studied p- glucosidase, L-glutaminase, and acid phosphatase are the most sensitive to pH changes and could be used as tools for monitoring ecosystems health and function.
The effect of crop rotations and N fertilization on p -glucosaminidase activity and its relationship to N mineralization were studied in soils of two long-term field experiments in Iowa. The activity of p-glucosaminidase was significantly affected by crop rotations and N fertilization, and was significantly
correlated with Corg and Norg, Cmic, and Nmic in soils, and with cumulative N mineralized during 24 weeks of incubation at 30°C. ix
Studies to evaluate the effects of 23 trace elements on the activity of p- glucosaminidase in three Iowa surface soils showed that at 5 mmol kg-1 soil, the activity of this enzyme was inhibited by 18, and activated by 5, of the trace elements tested, with Ag(I) and Hg(II) being the most effective inhibitors. Also, the activity of this enzyme was significantly affected by tillage systems (no-till, chisel plow, and moldboard plow) and four residue placements (bare, normal, mulch, and double mulch).
Other studies showed that the amounts of N mineralized in 56 surface soils, obtained from six states in the North Central region of the United States, by two biological and three chemical methods were significantly correlated with
P-glucosaminidase activity, and with organic C and total N. The amounts of N minerahzed during 14 days of incubation of field-moist soils under waterlogged conditions at 30°C were the most significantly correlated with p-glucosaminidase activity (r = 0.86***). 1
CHAPTER 1
GENERAL INTRODUCTION
Recent agricultural research priority has focused on sustaining soil health and quality, i.e., identifying key parameters or processes for monitoring changes in soils properties (physical, chemical, or biological) induced by management.
The quality of soil can impact sustainability, productivity, and environmental quality and, thus, human and animal health. Historically, much attention focused on impacts of agriculture on soils erosion and depletion of organic matter. Biological processes have received little attention because of the perceived difficulties associated with such work. More recently, attention has focused on the long-term impacts of agriculture on soil biology and biochemical
parameters (Dick, 1992; Turco et al., 1994; Doran and Parkin, 1994; Deng and
Tabatabai, 1996a; Moore et al., 2000). The term soil health or quality has been
defined as the capacity of a soil to function within ecosystem boundaries to sustain biological productivity, maintain environmental quality, and promote plant and animal health (Doran and Parkin, 1994).
Soil microorganisms and their processes are the major contributors to the
maintenance of soil quality because they control the decomposition of organic
matter, biogeochemical cycling including No fixation, the formation of soil
structure, and the fate of organics applied to soils (Dick, 1992; Turco et al.,
1994). One of the most commonly used parameter to study biological processes 2 in soils is measurement of total microbial bio mass. The soil microbial biomass is a small but the most labile C and N pools in soils. It is the impetus behind nutrient transformation and cycling in soils. Consequently, the fertility status of soils depends upon both the size and activity of microorganisms. Whereas the soil organic matter is relatively stable, the microbial biomass is very dynamic and responds quickly to changes in management practices. The total microbial biomass C has been used as early indicator of long-term trends in the total soil organic C quality (Powlson and Jenkinson, 1981). However, measurement of total soil microbial biomass does not give the indication of the total biological activity (Dick, 1992).
Measurement of enzyme activities has been used as specific indexes of microbial activity. Soil enzymes are the biological catalyst of innumerable chemical reactions necessary for life processes of microorganisms in soils, decomposition of organic residues, cycling of nutrients, and formation of organic matter and soil structure (Dick, 1994). Thus, information on enzyme activity provides insight into biochemical processes in soils (Frankenberger and Dick,
1983), and enzymes are the potential indicators of soil quality because of their relationship to soil biology, ease of measurement, and their rapid response to changes in soils management (Dick, 1994; Dick et al., 1996); thus, they provide a
useful tool to long-term monitor the changes in soil health and quality.
Soil management practices influence soil microorganisms and soil
microbial processes through changes in the quantity and quality of plant 3 residues entering the soil, their seasonal distribution, changes in nutrient inputs, and the ratio between above and below ground (Christensen, 1996).
Application of lime to soils most often leads to a significant increase in pH and, thus, a change in microbial biomass (Edmeades et al., 1981), microbial dynamic and diversity, and therefore, enzyme activities (Zelles et al., 1990). Maintenance of soil health and quality and improvement in the production of crop yields requires liming as well as the adoption of conservation tillage systems (Simard et al., 1994). Different soil disturbances and residue placements lead to significant changes in soil physical and chemical properties, especially pH and organic matter content. These treatments may also lead to significant changes in the composition, distribution, and activities of soil microbial communities and enzymes; consequently, affect nutrient cycling, fertility, other chemical, microbiological, and biochemical processes in soils (Deng and Tabatabai, 1996b).
Soil organic matter is more thoroughly distributed in soils under conventional tillage system compared to reduce or no-till systems where crop residues are concentrated on the soil surface.
Because crop residues are primary sources of organic matter, cropping systems and fertilizer regime may exert a significant influence on soil quality
(Campbell et al., 1991). Soils under monocultural systems, in general, contain significantly lower concentration and qualities of soil organic matter, less soil structural stability, and reduced amounts of microbial biomass and activities compared with systems involving crop rotations. Crop rotations lead to 4 significant effects on soil physical, chemical, and biological properties by providing higher C inputs and diversity of plant residues returned to soils
(Havlin et al., 1990; Varvel, 1994; Robinson et al., 1996). Systems with high organic matter inputs and easily available soil organic matter compounds tend to have higher microbial biomass (Vaughan and Malcoln, 1985) and enzyme activities (Deng and Tabatabai, 1996; Klose et al., 1999; Klose and Tabatabai,
2000) because they are preferred energy source for microorganisms. Studies on the effects of inorganic N fertilizer applications on soil biological and biochemical properties have shown contradictory results. While some researchers reported increased microbial biomass (Fraser et al., 1994) and enzyme activities (Khan,
1970), others reported the opposite (Ladd et al., 1994; Dick et al., 1988) or no effect on the enzyme activities (Klose et al., 1999).
Any changes in management practices is reflected in the microbial biomass and soil enzymes in a short-period of time; long before measurable changes in soil chemical properties can occur (Powlson et al., 1987; Dick, 1994).
Therefore, enzyme activities have been suggested as early indicators of changes
in soil properties induced by agriculture practices (Carter, 1986; Powlson et al.,
1987). Thus, it is crucial to obtain information about the effects of those soil
management practices on microbial biomass and their biochemical processes for
better management of our soils. Therefore, the objectives of this study were: (1)
to assess the effects of liming and tillage systems on selected soil chemical
properties, microbial biomass, and the activities of 15 enzymes involved in C, N, 5
P, and S cycling in soils, (2) to determine whether these response are consistent across four long-term management sites in Iowa, (3) to assess the sensitivity of the 15 enzymes to liming (changes in soil pH) and different tillage systems, (4) to study the relationship between microbial biomass C (Cmic) and N (Nmic) and the activities of the enzymes, (5) to study the impacts of crop rotations and N fertilization on the activity of P-glucosaminidase, an enzyme involved in C and N cycling in soils, (6) to assess the relationships between the activity of P- glucosaminidase and N mineralization in soils, (7) to asses the relationship between the activity of P-glucosaminidase and Cmic and Nmic, (8) to study tillage and residue management practices effects on p -glucosaminidase activity, (9) to determined the effects of trace elements on the activity of this enzyme, and (10) to study the relationships between p-glucosaminidase activity and selected biological and chemical indexes of available N in soils.
Organization of Dissertation
This dissertation is arranged in 10 chapters. Chapter one is general introduction and chapter two is the literature review. Chapters 3-9 are the papers prepared as independent articles submitted for publication in refereed journals. Chapters 3-5 contain the results obtained on the effects of liming and tillage systems on Cmic and Nmk and activities of glycosidases (a -and P- glucosidase, a- and p-galactosidase), arylamidase and amidohydrolases (L- asparaginase, L-glutaminase, amidase, urease, and L-aspartase), and 6 phosphatases (acid and alkaline phosphatases, phosphodiesterase) and arylsulfatase, respectively. Chapters 6-8 report, respectively, the effects of cropping systems and N fertilizer, trace elements, and tillage and residue management on the P-glucosaminidase activity. The article in chapter 6 also assessed the relationship between p-glucosaminidase activity and Cmic, Nmic, N mineralization, and selected soil properties. The article in chapter 9 investigated the relationship between N availability indexes and p- glucosaminidase activity in 56 surface soils from the North Central region of the
U.S. Summaries of the results are shown in Chapter 10, and additional tables and figures are presented in the Appendix. 7
CHAPTER 2
LITERATURE REVIEW
Soil Enzymes
Enzymes are proteins that catalyze chemical reactions to proceed at faster rates without undergoing permanent alterations. They lower the activation energy of the reaction thus, cause the reaction to proceed at faster rates. Soil enzymes are similar to those in other biological systems. However, the difference between enzyme activities in soils and those of other systems is that the source may be associated with living and nonliving components. They are specific for the types of chemical reactions in which they participate. For example, even tough the only differences between the substrates maltose and cellubiose is that maltose is an a-glucoside and cellubiose is P-glucoside, maltase
hydrolyzes maltose to glucose, whereas cellubiose hydrolyzes cellubiose to
glucose but nor vice versa. Enzyme specificity is mostly dictated by the nature of
the groups attached to the susceptible bonds (Tabatabai, 1994). Enzymes are
the specific activators since they combine with the right substrate in a
stereoscopic way that they cause changes in the electronic configuration around
the susceptible bond.
The physicochemical state of the enzymes and their effects on the
reactions are dependent on the pH, ionic strength, temperature, and the 8 presence or absence of inhibitors or activators (Tabatabai, 1994). Extremes of the pH and higher temperature denature the enzymes.
Sources and States of Enzymes in Soils
The total activity of enzymes in soils is composed of the activities of accumulated enzymes and enzymes of proliferating microorganisms.
Accumulated enzymes in soils are regarded as enzyme present and active in a soil in which no microbial proliferation occurs. These enzyme present in nonproliferating cells (spores, cysts, seeds, endospores), released from lysed cells, attached to dead cells and cell debris, and absorbed to clay and humic colloids
(Burns, 1982). Both microorganisms and plants release enzymes into the soil environment. Major sources of soil enzymes are microbial biomass, but they can also be derived from plant and animal residues. Alkaline phosphatase, for example, is released by Bacillus subtilis under some conditions (Cashel and
Freese, 1964), and is therefore mainly extracellular in nature with periplasmatic counterparts in Gram-negative bacteria (Burns, 1982). Acid phosphatase may exist extracellularly on the surface of cell walls of Saccharomyces mellis
(Weimberg and Orton, 1964). Dick et al (1983) noted that alkaline phosphatase in soils is not produced by plants, its activity is completely derived from microorganisms. Plants are also considered as extracellular enzyme sources in soils. A study of Juma and Tabatabai (1988) showed that sterile corn (Zea mays) and soybean (Glycine max) roots contain acid phosphatase but not alkaline 9 phosphatase. Speir and Ross (1978), however, reported that even thought both microorganisms and plant release enzyme into the soil environment, microorganisms are the major source of the soil enzyme activity, because of their large biomass, high metabolic activity and short lifetime, which allow them to produce and release relatively large amounts of extracellular enzymes than can plants or animals. The effect of microorganisms in supplying phosphatase activity to soils, however, temporary and short-lived (Dick and Tabatabai, 1992).
The increased activity of phosphatase produced during incubation of soil with glucose and nitrate was mostly lost during the cycle of prolifération-dying-lysing and many cycles of microbial activity are required to obtain a permanent increase in the exctracellular phosphatase activity (Speir and Ross, 1978). An important amount of enzymes released into the soils, both from microorganisms or plant roots, are inhibited by soil constituents, degraded by soil protease, or both (Dick and Tabatabai, 1992). A small percentage of active enzymes introduced into soil may become stabilized in the soil. The study of Dick and
Tabatabai (1987) on enzyme kinetic showed that clay-enzyme complexes are formed in soil and the activity of the enzyme is greatly reduced, but not totally eliminated. Plants have the capability to produce many enzymes. These enzymes are added to soil in plant residues and may remain active.
To differentiate between the source of intracellular and extracellular enzyme activity is important. A large amount of information has accumulated since 1950 about enzymatic reactions in soil. Many theoretical approaches and 10 methods have been developed to determine the state of enzyme in soil, such as exposing the soil samples to elevated temperatures, gamma-irradiation, treatment of the soil with plasmolytic agents, and electron beams and fumigants
(chloropicrin, methyl bromide, and Chloromycetin) (McLaren, 1969; Cawse, 1975;
Frankenberger and Johanson, 1986). The use of bacteriostatic agents to inhibit intracellular enzymatic activity is not applicable because of some factors such as cell wall permeability, the possible induction of ploasmolysis with release of intracellular enzymes, the promotion of microbial growth, and their variable effects depending on soil and enzyme assay condition (Ladd, 1985;
Frankenberger and Johanson, 1986; Tabatabai, 1994). The techniques using high-energy irradiation and antibiotics have not been satisfactory to clearly differentiate between enzyme activities of viable cells and abiotic enzymes.
Recently Klose and Tabatabai (1999a, b) were successful in estimating the urease and arylsulfatase activities associated with the microbial biomass. Those studies reported significant increases in the activities of urease and arylsulfatase upon chloroform fumigation of soil samples, thus differentiating the total activity of each enzyme into intracellular and extracellular origin. The
method proposed by Klose and Tabatabai (2002) also used to differentiate
between various pools of phosphomonoesterase activities in soils by the same
method that they used for urease and arylsulfatase. They concluded that this
method is not applicable in separating between different pools of acid and
alkaline phosphatase in soils. 11
The term 'state of enzyme in soils' was first time used by Skujins (1967,
1976) to describe the phenomena whereby enzymes exist in soils. Burns (1986) indicated that describing the state of an enzyme in soil is to attempt to describe the location and microenvironment in which it functions and how the enzyme is bound or stabilized within that microhabitat. Enzyme activities in soils are derived from free enzymes, such as exoenzymes released from living cells, endoenzymes released from disintegration cells, and enzymes bound to cell constituents (enzyme present in disintegrating cells, in cell fragments, and in viable but non proliferating cell). Free enzymes are very small amount in the soil solution. A major part of free enzymes in soil are adsorbed on organic and mineral constituents or they may have a complex with humic substances.
It is well established that free enzymes are readily decomposed or inactivated very rapidly when they are added to soils. Conrad (1940) suggested
that organic soil constituent protected native urease against microbial degradation or other process that inactivate the enzymes in soils. His result has
been supported by many other studies showing thai enzyme activities of soils are significantly correlated with the organic matter content of the soils.
Several mechanisms have been suggested to explain the protective affect
of soil constituents on enzyme activity. Weetall (1975) proposed the most
dominant mechanisms of immobilization and stabilization on solid supports.
Those mechanisms most likely apply to extracellular enzymes stabilization by
soils organic and inorganic constituents. These mechanisms are adsorption, 12 microencapsulation, cross-linking, copolymerization, entrapment, ion exchange, adsorption and cross-linking, and covalent attachment.
Types of Enzymes in Soils
Many enzymes have been detected in soils, but only some of those have been studied in details. Gianfreda and Bollag (1996) classified soil enzymes based on their location and function. To classify the activities based on function is more preferred because of the understanding of enzyme activity based on location in the soils is still unclear.
Many of the enzymes detected and quantified in soils are hydrolases, but the activities of some oxidoreductases, transferases, and lyases also have been studied. Gianfreda and Bollag (1996) and Dick and Tabatabai (1992) have provided a listing of approximately total of 35 enzymes that has been partially characterized in soils, however, more than 100 enzymes has been identified in soils.
Hydrolases catalyze the reaction by cleavage of bonds and the splitting of water molecules. These enzymes are responsible for many bioremediation transformations in soils including the functional groups amides, carbamates, esters, nitriles, thiocarbamates, alkyl, aryl, imines, epoxides, halides, silanols, and organometalls (Gianfreda and Bollag, 1996). This group of enzymes is mostly smaller in size, they do not require cofactor for the activity, and they are more resistant to inactivation compared to other group of enzymes. In addition 13 to the activity of individual hydrolases, the succession of enzymes involved in the transformation of a substrate in soils also has been studied. Cellulose hydrolysis in soils requires the activity of endo-l,4-(3-glucanase (EC 3.2.1.4), which attacks cellulose chains at random, exo-l,4-p-glucanase (EC 3.21.91) removes glucose or cellobiose from the non-reducing end of the cellulose chains, and (3-D-glucosidase
(EC 3.21.21) hydrolyses cellobiose and other water soluble cellodextrines to glucose. The other example is the release of amino acids from soil organic matter by arylamidase (EC 3.4.11.2) and the release of NHa from amino acids by amidohydrolases [i.e., L-asparaginase (EC 3.5.1.1); L-glutaminase (EC 3.5.1.2)] which act on C-N bonds other than peptide bonds in linear amides.
Oxidoreductases catalyze the transfer of electrons from one molecule to another. Examples of oxidoreductases are glucose oxidase, catechol oxidase and catalase with the substrates glucose, catechol, and hydrogen peroxide, respectively. Enzymes in this group are responsible for the polymerization of phenolic compounds (Nannipieri and Bollag, 1991). The most commonly studied oxidoreductase in soils is dehydrogenases. This enzyme transfers hydrogen from substrates to acceptors. Dehydrogenase is frequently assayed to estimate
microbial activity because it is an intracellular enzyme and is present in every living organism. Dehydrogenases function within the cytoplasm of proliferating cells and cannot function outside of living cells because they depend on cofactors;
some physiological specific properties of the cells or on being located close to the other. Transferases are involved in the group transfer from one molecule to another. The activities of transaminase as affected by aspartic acid, leucine, valine, glutamic acid or pyruvate in toluene-treated soils were reported by
Hoffman (1959, 1963). An important enzyme in this group is thiosulphate S- transferase (rhodanase) that performs the intermediate step in oxidation of elemental S, which is found in small amounts in soils or is added as a fertilizer.
Lyases are involved in the cleavage of bonds other than by hydrolysis or oxidation. Examples are the deamination or decarboxylation of amine groups or carboxyl groups in amino acids, respectively.
Glycosidases
The term glycosidases has been used for the group of enzymes that act on the glycosyl compounds including the glycoside hydrolases (EC 3.2.1): a- glucosidase (EC 3.2.1.20), P-glucosidase (EC 3.2.1.21), a-galactosidase (EC
3.2.1.22), P-galactosidase (EC 3.2.1.23), and ZV-Acetyl-P-D-glucosaminidase
(NAGase, EC 3.2.1.30). These enzymes are widely distributed in nature (Bahl and Agrawal, 1972; Dey and Pridham, 1972) including soils (Skujins, 1967,
1976). They play an important role in the breakdown of low molecular-weight carbohydrates, and their hydrolysis products are the main energy source to soil microorganisms.
This group of enzymes has been named according to the type of bond that they hydrolyze. For example, a-glucosidase, also named maltase, catalyzes the 15 hydrolysis of a-D-glucopyranosides (maltose). P-Glucosidase, also named gentibiase or cellobiase, catalyzes the hydrolysis of (3-D-glucopyranosides
(cellobiose). The enzyme a-galactosidase, also named melibiase, catalyzes the hydrolysis of a-D-galactopyranosides, and P-galactosidase, also named lactase, catalyzes the hydrolysis of P-D-galactopyranosides. Among all the glycosidases,
P-glucosidase was reported as the most predominant enzyme in soil (Eivazi and
Tabatabai, 1988, 1990).
P-Glucosaminidase (NAGase) is the enzyme that catalyzes the hydrolysis of 2V-acetyl-P-D-glucosamine residue from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also classified as p-hexosaminidase (EC
3.21.52) by the International Union of Biochemistry because it cleaves the amino sugar iV-acetyl-p-D-galactosamine (Webb 1984). Some of the amino sugars in soils may exist in the form of alkali-insoluble polysaccharide referred to as chitin. Chitin, which consists of iV-acetyl-P-D-glucosamine residues in P-1,4 linkages, is a major structural component in insects and comprises the cell wall, structural membranes and skeletal component of fungal mycelia, where it plays a structural role analogous to the cellulose of higher plants (Stevenson 1994).
This substance is considered one of the most abundant biopolymers in nature and an important pool of organic C and N in soils (Stryer 1988; Wood et al.
1994). It contains recalcitrant C and N; thus, it is most likely that the NAGase catalyzes the hydrolysis chitin producing amino sugars in soils, which constitute
5-10% of the organic N in the surface of most soils (Stevenson 1994). 16
The activity of NAGase has been detected in microorganisms, human tissues, insects, plant, and soils (Neufeld 1989; Trudel and Asselin 1989;
Martens et al. 1992; Sinsabaugh et al. 1993). Recently Parham and Deng (2000) reported on its quantification and characterization in soils, and developed a method for its assay. The procedure involves extraction and colorimetric determination of p-nitrophenol released when soil is incubated with buffered p- nitrophenyl-AT-acetyl-P-D-glucosaminide. The reaction involved is as follows:
HO -R + HiO HO P-Glucosaminidase NH I c=o I CH3 p-Nitrophenyl-N-acetyl-P-D-glucosaminide
c=o
CH3 AT-acetyl-p-D-glucosaminide p-Nitrophenol 17
Work by Smucker and Kim (1987) showed that chitin is an inducer of chitinase activity and it is a major source of organic N in soils. Not much work is available about the role of this enzyme in decomposition of crop residues in soils, but work by Sinsabaugh et al. (1993) showed that p -glucosaminidase is one of the enzymes required for chitin degradation; its activity was negatively correlated with N immobilization and decomposition rate for birch sticks decomposing at eight contrasting sites. Most likely soil management practices, such as liming, crop rotations, tillage and residue management systems, N fertilization, affect the activity of this enzyme in soils. But, little information is available of the effects of those practices on this important enzyme in soils.
Arylamidase
Arylamidase (a-aminoacyl-peptide hydrolase, EC 3.4.11.2) is the enzyme that catalyzes the hydrolysis of an N-terminal amino acid from peptides, amides or arylamides. This enzyme is widely distributed in nature and it has been detected in plants, animals and microorganisms (Hiwada et al., 1980; Appel,
1974). Arylamidase was recently detected in soils, and a method was developed for its assay (Acosta-Martinez and Tabatabai, 2000). It is believed that this enzyme catalyzes one of the initial reactions in N mineralization because it is involved in the release of amino acids from soil organic matter. Those amino acids released by the activity of arylamidase are the substrates for amidohydrolases involved in N mineralization: 18
Amino acid Ammonification by Nitrification amidohydrolases: e.g.. Arylamidase Amino acid NH/ X- NO, — NO, + energy activity (RNHJ L-Glutaminase activity L-Asparaginase activity L-Aspartase activity
Thus, understanding the environmental controls on the activity of this enzyme in soils is important for better understanding the N cycling process.
The studies of Acosta-Martinez and Tabatabai (2001) demonstrated that the activity of this enzyme in soils was significantly correlated with the activities of
L-asparaginase (r = 0.91***), L-aspartase (r = 0.90***), urease (r = 0.87***), and
L-glutaminase (r = 0.84***). Other recent studies by Dodor and Tabatabai
(2002) on the effects of crop rotation and N fertilization on arylamidase activity showed the activity of this enzyme was significantly correlated with the cumulative N mineralized in soils of the plots receiving 0 or 180 kg N ha1, with r values 0.88*** P<0.001 and 0.55 P<0.01, respectively.
Arylamidase is capable of hydrolyzing the neutral amino acids (3- naphthylamides and p-nitroanilides according to the following reaction (using the amino acid L-leucine as an example): 19
o IH!(JCHCH,CH(CHJ), /NH; hooc. I X NHjCl .CHCH,CH(CH ), + HO Arylamidase Z 3 OTO nh;
L-Leucine p-Naphthylamide P-Naphthylamine Leucine
Amidohydrolases
The amidohydrolases, L-asparaginase, L-aspartase, L-glutaminase, amidase and urease, are group of enzyme involved in the hydrolysis of organic N in soils. The chemical nature of N in soils is such that a large proportion (15-
25%) of organic N is released as NH4+ by 6 M HC1 hydrolysis. It was reported that a portion of release NH4"1" comes from the hydrolysis of amide (asparagines, glutamine) residues in soil organic matter (Sowden, 1958). With the exception of urease, the rest of the amidohydrolases play a major role in N mineralization in soils. The amidohydrolases are specific to their substrates as is for all other enzymes.
L-asparaginase (L-asparaginase amidohydrolase, EC 3.5.1.1) catalyzes the hydrolysis of L-asparagine producing aspartic acid and NH3. This enzyme has a major role in N mineralization in soils (Tabatabai, 1994). Bremner (1955) showed that after hydrolysis of humic preparations, 7.3-12.6% of the total N was in the form of amide-N. It was also reported that a percentage of the NH41" released during acid hydrolysis was equal to the sum of aspartic acid-N and 20 glutamic acid-N derived from asparagine and glutamine. The reaction is as follows:
COOH COOH
HC-NH, +H20 L-Asparaginase ^ HC-NH, + NH3.
CH, CH,
CO COOH
NH2
The reaction catalyzed by L-glutaminase (EC 3.5.1.2) involves the hydrolysis of L-glutamine to L-glutamic acid and NHA as follows:
COOH COOH
H C-NH H C-NH, + NH3 2 H,0 L-Glutaminase CH ÇH, 2
CH2 CH2
CO COOH
NH, 21
Amidase (acylamide amidohyrolase, EC 3.5.1.4) catalyzes the hydrolysis of amides and produces NHa and the corresponding carboxylic acid according to the following reaction:
AmidaSe RCONH, + H20 • NH3 + RCOOH
Urease (urea amidohydrolase, EC 3.5.1.5) is the enzyme that catalyzes the hydrolysis of urea to CO2 and NHa. It is not involved in N mineralization in soils. This enzyme catalyzes the hydrolysis of urea, added to soils as a fertilizer.
It breaks the C-N bonds other than peptide bonds in linear amides and releases
NHa (Ladd and Jackson, 1982; Tabatabai, 1994); thus, belongs to a group of enzymes that include glutaminase and amidase. The reaction of this enzyme is as follow:
O 11 Urease NHJCNH, • C02 + 2NH3 H2O
L-Aspartase (L-aspartate ammonia-lyase, EC 4.3.1.1) catalyzes the hydrolysis of L-aspartate to fumarate and NHa. This enzyme is widely distributed in nature. It has been detected in bacteria, plants, and certain 22 animal tissues (Tabatabai, 1994). The study of Frankenberger and Tabatabai
(1991) suggested that L asparagine in soils is hydrolyzed to aspartic acid and
NHa and that the product aspartic acid is not further hydrolyzed to NHa during two hours of incubation used for assay of L-asparaginase. The study of Senwo and Tabatabai (1996) showed that soils contain aspartase activity but its reaction is very slow to be assayed. However, incubation of soil with L-aspartate for 6 to 72 h could be detected quantitatively. The reaction is as follows by using its K form:
COOK COOK
HC-NH, + H-,0 L-Aspartase—^ c H + NH3. I " II
ÇH2 ÇH COOH COOH
Phosphatases
Phosphatases have been used to describe a large group of enzymes that catalyzes the hydrolysis of esters and anhydrides of H3PO4 (Tabatabai, 1994).
This enzymes play a major role in the mineralization of soil organic P. There are five groups of phosphatases according to the commission on enzymes of the
International Union of Biochemistry; phosphoric monoester hydrolases (EC
3.1.3), phosphoric diester hydrolases (EC 3.1.4), triphosphoric monoester hydrolases (EC 3.1.5), enzymes acting on phosphoryl-containing anhydrides (EC 23
3.6.1). and enzymes acting on P-N bonds (EC 3.9), such as the phosphoamidase
(EC 3.9.1.1). The most extensively studied group among the phosphatases in soils is the phosphomonesterases, acid phosphatase (EC 3.1.3.2), alkaline phosphatase (EC 3.1.3.1), and phosphodiesterase (EC 3.1.4) (Tabatabai and
Bremner, 1969; Eivazi and Tabatabai, 1977; Browman and Tabatabai, 1978).
The enzymes are classified as acid and alkaline phosphatases, because they show optimum activities in acid and alkaline ranges, respectively. Several studies showed that alkaline phosphatase activity in soils is totally derived from microorganisms (Dick et al., 1983; Juma and Tabatabai, 1988a,b,c). The work of
Eivazi and Tabatabai (1977) on the activities of phosphatases at different buffer pH suggested that acid phosphatase was predominant in acid soils while alkaline phosphatase was predominant in calcareous soils. They concluded that the production, stability, and distribution of acid and alkaline phosphatase activities are correlated to soil pH. These findings might be useful in predicting the type of soil enzymes that are induced under given management practices and soil fertility management decisions, because soil fertility and crop production are affected by biochemical process and these processes, including enzyme activities, are significantly affected by pH. Dick et al. (2000) studied the potential of using acid and alkaline phosphatase activities to determine the optimum pH for crop production and the amount of lime required to achieve this optimum. They concluded that for the cropping systems that rely heavily on natural biological processes to maintain productivity, measuring the alkaline phosphatase/acid 24 phosphatase ratio might be preferable to chemical approaches to evaluate effective soil pH and liming needs. The reaction involved in hydrolysis of phosphomonoesters by acid and alkaline phosphatases at their optimal buffer pH of 6.5 or 11 in soils, respectively, are as follows:
fj Acid or alkaline ^ RO P O + H20 phosphatase HO PO ~ + ROH O ~ " j)-
The reaction catalyzed by phosphodiesterase (orthophosphoricdiester phosphohydrolase, EC 3.1.4.1) is as follows; where the Ri and Ra represent either alcohol or phenol groups or nucleosides (Privât de Garilhe, 1967):
/0H yOH
O = P— OR, + H,0 Phosphodiesterase Q = p 0H + R OH V ' V
Arylsulfatase
There are several types of sulfatases; they are classified according to the type of bond they hydrolyze. Enzymes under this group include arylsulfatase, alkylsulfatase, steroid sulfatase, glucosulfatase, chondrosulfatase, and myrosulfatase (Fromageot, 1950; Roy, 1960). Tabatabai and Bremner (1970) detected the enzyme arylsulfatase in soils and developed a method for measurement of its activity. Arylsulfatase is important in S cycling because it 25 releases plant available SO4- It was also suggested that the activity of this enzyme could be used as an indirect indicator of fungi as only fungi (not bacteria) contain ester sulfate, the substrate of arylsulfatase (Bandick and Dick,
1999). Many studies have been done on the role of arylsulfatase in S mineralization showing that total S in surface soils of temperate regions is reduced to H2S by HI and is converted to inorganic SO4 with hot alkali
(Freney, 1961; Tabatabai and Bremner, 1972; Neptune et al., 1975).
Arylsulfatase (arylsulfate sulfohydrolase, EC 3.1.6.1) catalyzes the hydrolysis of an arylsulfate anion by fission of the O S bond (Spencer, 1958).
The reaction is as follows:
2 RO SO 3- + H20 A ry Isu [fatase „ ROH+ H + + SO 4 ".
Soil Health and Quality
There is a growing interest in developing a universal soil quality index that can be used to assess the health of the soils. Soil quality has been defined as the capacity of a soil to function within its ecosystem boundaries to sustain biological activity and productivity, maintain environmental quality, promote plant and animal health, and habitation (Doran and Parkin, 1994; Gregorich et al., 1994). Even though water and air quality standards are well established, soil as a vital natural source is still without comprehensive quality standards. 26
The quality of soils impacts farm productivity, land use capability, agronomic sustainability and the environmental buffering capacity. An assessment of soil quality that includes measurement of some biological, chemical, and physical properties can provide valuable information to determine the status of such a soil parameters, their influence on the ecosystem boundaries, and the sustainability of soil management (Doran et al., 1994). Physical, chemical, and
biological parameters of the soil are significantly correlated with each other, and
thus any change of one of these soil parameters will affect another. For example, changes in physical and chemical properties of soil affect the organic
matter and moisture contents, which, in turn, affect microbial dynamics (Turco et al., 1994). The increase in soil organic matter is connected to the desirable
soil physical, chemical, and biological properties that are associated with soil
productivity (Stevenson, 1982). Organic matter of soils provides the capacity to
supply nutrients to plants, and modulate chemical and water availability
(Nelson and Sommers, 1982). Organic matter improves soil structure and
aggregation and is related to microbiological parameters such as the size of the
microbial biomass (Goyal et al., 1992) and soil enzyme activities (Frankenberger
and Dick, 1983).
Agricultural practices including liming, tillage, crop rotation, fertilization,
and application of organic amendments to soils affect the interaction of key soil
physical, chemical, and biological parameters. Even though each soil parameter
is a valuable indicator of soil quality, their response to management practices is 27 different. In agricultural ecosystems that have maintained cropping and management practices for a long period of time (ca.100 years), valuable information related to the quality of soils can be obtained from their physical and chemical properties. However, in those systems where changes are recent, the soil biological and biochemical parameters can be more sensitive indicators and provide earlier indications of change in soil quality, long before changes in total soil organic C and N can be detected by chemical analysis.
Studies have shown the soil biological and biochemical processes are an integral part of soil quality (Dick, 1992; Turco et al., 1994). Nannipieri (1983) found a faster response to changes in soil organic matter, indicated by increases in microbial biomass, than with the actual changes in the soil organic C content.
Powlson et al. (1987) demonstrated that small changes in total soil C and N are difficult to detect against the large background levels of C and N in soils.
Soil microorganisms and their associated processes are potentially one of the most sensitive and earliest biological indicators of soil quality. The microbial biomass C only constitutes 1-4% of total soil organic C (Anderson and Domsch,
1989) but it plays a key role in the decomposition of organic materials in soils and it is also the major source of soil enzymes. Soil enzymes activities are considered indicative of specific biochemical reactions of the entire microbial community in soils, because they are involved in the transformations of the main soil nutrients C, N, S, and P (Frankenberger and Dick, 1983; Nannipieri et al.,
1990). To establish a soil quality standard is difficult because soils are inherently variable. The maintenance of soil organic matter is considered beneficial for agricultural purposes and would be a likely measure of soil quality; however, optimal concentrations of organic matter vary among soils. Because many physical and chemical properties will hardly change over the next century, the focus to search for the soil quality indictors should be on those components expected to change more rapidly (Tscherko and Kandeler, 1997). For example, the effects of tillage system on total organic matter is experimentally detectible in soils of temperate areas only over extended time periods, whereas changes in the microbial biomass and microbial processes may become manifest over a shorter time scale (Christensen, 1996).
Recently, soil quality investigations, including the measurement of the soil enzyme activities, has shown to provide information needed for management and regulatory decision making in soil systems, including agricultural, forestry or turfgrass, and any of those systems in disturbed or contaminated conditions.
The activities of the soil enzymes have shown to be sensitive to soil management practices, including cropping systems (Khan, 1970; Dick, 1984; Bolton et al.,
1985; Klose et al., 1999), and tillage treatments (Dick, 1984; Gupta and
Germida, 1988; Deng and Tabatabai, 1996a,b; 1997) and to the applications of organic amendments (Yaroshevich, 1966; Khan, 1970; Verstraete and Voets,
1977; Dick et al., 1988; Martens et. al., 1992). 29
Effects of Management Practices on Microbial Biomass and Enzyme
Activities
Effects of liming on enzyme activities
Soil acidification occurs due to several factors, including: the accumulation of H2CO3 formed from high concentrations of CO2 in the soil atmosphere produced by plant roots and microbial respiration, the mineralization of organic
N and S, fixation of N by leguminous plants, NH4+- forming fertilizers, nitrification of NHa and NH4+ -producing fertilizers, organic acids produced from crop residues and litter decomposition, and the addition of wet and dry atmospheric deposition, especially near the point sources (Tabatabai, 1985).
Acid precipitation resulting from the combustion of fossil fuels is a widespread phenomenon that acidifies soils in central Europe (Zelles et al., 1987b).
Increased acidification is not desired because it lead to the release and leaching of nutrients and mobilization of toxic Al3+ in soils. Bacterial populations are decreased and fungi populations are increased by soil acidification, and the whole soil microbial community structure is also altered (Abrahamsen et al.,
1980; Alexander, 1980; Baath et al., 1980). Soil quality is affected by acidification, because microorganisms play a key role in the turnover of organic matter and minerals in soils, which, in turn, is very important to soil development and plant growth. Adverse effects to the soil microbial populations 30 by soil acidification are correlated to the changes that occur to soil process such as nitrification, denitrification, and glucose mineralization (Harmsen and van
Schreven, 1955; Tabatabai, 1985).
Lime, ash fertilization, and prescribed burning are practices that are used to increase the acid neutralizing capacity of soils. Application of lime to soils as limestone (CaCOs) and dolomite (Ca-Mg COa) are the most common practice in the United States and some areas rely on the addition of lime coupled with fertilizers for the production of adequate crop yields and maintenance of soil quality. Coal combustion residues have been also shown to be useful liming materials (McCarty et al. 1994). The addition of lime materials increases the soil pH, and causes significant changes in the chemical and biochemical reactions and in microbiological processes, with modifications in the solubility of many chemical compounds (Naftel, 1965).
Recent studies have focused on the problem of acidification of forest soils, because these systems have not been as well studied and managed as their agricultural counterparts (Zelles et al., 1987a, 1990; Wanner et al., 1994). Long term field experiments with repeated lime practices cause increases in the soil organic matter in the humus layer of Norway Spruce stands (Derome et al.,
1986; Derome and Patila , 1990; Derome, 1990, 1991 ). Badalucco et al., (1992) reported their findings on the effect of liming on some chemical, biochemical, and microbiological properties of acid soils under spruce. They showed that liming soils with pulverized CaCOs, added on the basis of the lime requirement 31 estimated by the Schoemaker—MacLean-Pratt buffer method (Schoemaker et al.,
1962) caused an increase in the cumulative CO2 evolution, the microbial biomass
C and N values, and the activity of dehydrogenase in the soil. Others have also reported increases in the microbial biomass and activity upon liming (Zelles et al., 1987, 1990; Badalucco et al., 1992). Zelles et al. (1990) demonstrated that there are changes in the microbial communities structure by soil liming, and showed that fungi populations are decreased upon liming, because they are unable to function optimally at the higher pH developed and because their ecological role is taken over by the prokaryotes. These shifts in the type of soil microorganisms, that are predominant at different soil pH, significantly affect the level and type of enzyme activities in soils.
Liming effects on enzyme activities have focused mostly on the changes of the activity of acid phosphatases in forest soils, because of the positive correlation between phosphate availability and soil pH (Haynes and Swift, 1988;
Cepeda et al., 1991; Illmer and Scchinner, 1991). Phosphatase synthesis is repressed when available P concentrations are increased in soil (Nannipieri et al., 1978). The others have reported the decreases of phosphatase activity are not related to an increase in the available P (Haynes and Swift, 1988; Badalucco et al., 1992). Studies on the effect of liming on the activities of different enzymes in agricultural soils are needed to obtain a better assessment of the effects of such practices on the status of C, N, P, and S cycling in soils. 32
Effects of cropping systems and N fertilization on enzyme activities
Crop rotations can provide higher input and diversity of organic materials to the soil and generally contain higher enzyme activities than under monoculture. Studies have shown that enzyme activities are sensitive to the positive effects of crop rotations compared to monoculture (Khan, 1970; Dick,
1984; Bolton et al., 1985; Klose et al., 2000). Cropping systems that return elevated levels of green manure/crop residues significantly increase the activities of several enzymes (Verstraete and Voets, 1977; Bolton et al., 1985). The increases in soil enzyme activities are not due to the addition of more enzymes with the plant residues, because that free enzymes are readily decomposed or are inactivated when added to the soil environment. It is more accepted that the observed increases in enzyme activity upon crop residues incorporation are due to the stimulation of the soil microbial biomass (Martens et al., 1992).
One study comparing 4-year rotations, including oats and meadow, with monoculture reported positive effects of the rotation on the microbial biomass.
They concluded that higher microbial biomass C and N in the 4-year rotation compared with continuous crops and soybean systems and the 2-year rotation may be due to several reasons: (1) enhanced soil structure, (2) greater amounts and diversity of residues produced, (3) the nearly year-around rhizosphere and plant cover, (4) stabilized microclimate, (5) and a higher root density under diverse crop rotations (Moore et al., 2000). Other studies found all the pools of arylsulfatase and urease activities (intracellular, extracellular and total) were 33 greater in cereal-meadow rotations, taken under oats or meadow, than under continuous corn; and reported that both enzymes were significantly correlated to microbial biomass C (Klose et al., 1999; Klose and Tabatabai, 2000). Angers et al., (1997) reported greater levels of alkaline phosphatase activity in rotations including barley-red clover compared to monoculture systems, and that soils under no-till with a corn-oats-alfalfa rotation contained the largest alkaline phosphatase activity. These results demonstrate that the enzyme activities are modulated by the C sources available in the crop residues, therefore, can be an indication of the decomposition rates of the C sources in the crop residues and, thus, of its structural complexity.
Effects of tillage and residue management on enzyme activities
Tillage and crop-residue management practices may lead to significant changes in biological, chemical, and microbiological properties, including the biochemical reactions in soils. Previous research has shown that no-till reduces soil erosion and water evaporation, and increases water infiltration and soil organic-matter levels (Doran 1980a,b; Tracy et al., 1990). Studies have also shown there is organic C and N accumulated at the soil surface (0—2.5 cm)
under reduced tillage involving crop residue placement (Havlin et al., 1990).
Accumulation of soil nutrients at the soil surface such as S and P, due to no-
tillage practices with residue placement, has been also reported (Tracy et al.,
1990). The use of such management practices will also alter the composition, 34 distribution, and activities of soil microbial communities and soil enzymes
(Doran, 1980a,b; Dick, 1984; Magnan and Lynch, 1986). Dick (1984) and
Kandeler et al. (1999) reported about the potential of measuring soil enzyme activities as early indicators of changes in soil properties induced by tillage practices. Studies have shown the microbial biomass and soil enzyme activities are greater in surface no-till soils than in plowed soils, while the reverse trend is observed in deeper layers (Angers et al., 1993; Kandeler and Bohm, 1996;
Kandeler et al., 1999). Dick (1984) reported the activities of acid phosphatase, alkaline phosphatase, arylsulfatase, invertase, amidase, and urease in 0-7.5 cm surface soils were significantly greater in soils from no-till plots as compared with those from conventional tillage plots. Jordan et al. (1995) reported higher activities of alkaline and acid phosphatases in continuous corn under no-till than in continuous corn under conventional tillage. Many researchers have also reported the activities of phosphatases are generally greater in soils under no-till than in soils under conventional tillage (Doran, 1980b; Angers et al., 1993).
Studies by Kandeler et al. (1999) with different tillage treatments applied to the top 10 cm of soil showed that the activities of alkaline phosphatase, xylanase, and protease responded faster than other tests (e.g., nitrification, N mineralization). The activities of the enzymes showed the following pattern of response to the different tillage practices used: minimum > reduced > conventional tillage. Studies with Iowa soils showed the activities of the glycosidases (a- and |3- glucosidase and a- and (3- galactosidase), 35 amidohydrolases (urease, amidase, L-asparaginase, and L-glutaminase), phosphatases (alkaline and acid phosphatases, phosphodiesterase and inorganic pyrophosphatase), and arylsulfatase were significantly affected by tillage and residue placement (Deng and Tabatabai, 1996a,b; 1997). Results from such work showed the activities of all the enzymes studied were greater in no- till/double mulch treatments than in the other treatments, including no-till/bare, no-till/normal, chisel/normal, chisel/mulch, moldboard/normal, and moldboard/ mulch.
Nitrogen Mineralization
Indexes of available nitrogen
Nitrogen (N) is the most frequently deficient nutrient in crop production and the fourth most common element in plant composition, being outranged only by carbon (C), hydrogen (H), and oxygen (O). The N atom presents in different oxidation and physical states. More than 95% of the total in most soils is organic in nature. The shifts between those states are mediated by soil organisms. The ease with which shifts occur in the oxidation states cause to the formation of different inorganic forms that are readily lost from the ecosystem. Thus, most
nonlegume crops require N inputs. To understand the behavior of N in the soils is crucial for maximizing agricultural productivity and to reduce the impacts of
N fertilization on the environment. Organic N in soils is not available to plants; 36 it has to be mineralized to inorganic forms for plant uptake. There are three major biological forms of N in soils that are proteins, nucleic acids, and microbial cell wall constituents such as chitin and peptidoglycans. Studies on the distribution of the major organic N compounds in soils showed that despite the variations in total N content among the soils, the proportions of total N that could be hydrolyzed by hot HCl (6 M) were quite similar (84.2 to 88.9 %) (Sowden et al., 1977). In general, it has been shown that 20 to 50 % of the total N in soils is in the form of bound amino acids, 5-10 % is in the form of combined hexosamines (combined aminosugars), < 1% purines and pyrimidine bases, and the rest of organic N compounds in soils remain unidentified.
Mineralization of organic N refers to the degradation of proteins, nucleic acids, and amino sugars to NH^. The inorganic N intermediates produced by this process are: NO2, NOa, dinitrogen gas (N2), and nitrous oxide (N2O). Soil heterotrophic microorganisms play the major role in such biological N transformations. An example of the succession of reactions involved in N mineralization under aerobic conditions is as follows:
Aminization Ammonification Nitrification / X Proteins » RNH2 » NH/ •NO2- •NO3+ energy 37
Heterotrophic bacteria and fungi are responsible for one or more steps in the reactions in organic matter decomposition. One of the final stages is the decomposition of proteins and release of amines, amino acids, and urea, which is called aminization. The amines and amino acids produced during aminization are decomposed by other microorganisms and NH4+ is released. This step of the
N mineralization is called ammonification. Nitrification of NH^ to NOs and
N03 is carried out by the obligate autotrophic aerobic soil bacteria including the species Nitrosomas and Nitrobacter, respectively.
Several factors may influence N mineralization in soils as well as N immobilization. Because the activity of the microorganisms responsible for N mineralization (as with other microorganisms), the rates and products of N mineralization are affected by environmental conditions such as moisture, soil pH, and temperature. Moisture is the most important factor that affects microbiological and biochemical processes in soils. Under aerobic conditions, the
N03 form is the most predominant end-product of N mineralization, whereas under anaerobic conditions induced in waterlogged soils, NH^ is the predominant end-product. Moisture also affects the movement and, thus, the availability of NO3 in soils. Mineralization increases with a rise in temperature.
In frozen soils, the N mineralization becomes slower because of the restricted microbial activity. In addition, experimental evidence suggests that N mineralization proceeds differently under fluctuating temperature conditions than it does at constant temperature (Das et al., 1995). Soil pH is one of the 38 most important factor affecting nitrification, as well as other biochemical processes (e.g., denitrification, glucose degradation) in soils (Tabatabai, 1985).
Fu et al. (1987) reported that a lag period is developed before crop residues added to soil are mineralized. This lag period was decreased as the soil pH was increased from 4 to 8. Studies by Karmarkar and Tabatabai (1991) showed that certain organic acids either inhibit or activate nitrification in soils. They also reported that aliphatic acids and aromatic acids decreased the accumulation of
NO2* -N. Thus, liming soils and increasing soil pH may induce significant favorable changes to the soil environment of the microorganisms and the associated enzyme systems, and, thus, it is reflected in the N transformations. A study showed that liming increased the in situ mineralization of soil N over a 3- year period (Nyborg and Hoyt, 1978). Simard (1994) reported that lime had a temporary effect on N mineralization, which disappeared after the second year in a Luvisolic soil under tillage treatments
The mineralization of N in soils is also affected by management practices.
A study by Deng and Tabatabai (2000) showed that soybean growth depleted soil organic matter and reduced net N mineralization, while the inclusion of meadow in the cropping systems enriched organic N, which resulted in increased N
mineralization. Between the rotations used in the study the meadow-based
multiple crop systems were always advantageous over monocropping systems
with respect to increase N mineralization rates, and thus supplying N for plant
nutrition. The greater microbial biomass and particulate organic matter 39 accumulated in reduced tillage systems result in larger pools of easily mineralizable N (Jenkinson and Ladd 1981). Conservation tillage practices result in higher microbial biomass (Saffîgna et al., 1989; Carter, 1992; Angers et al., 1993a), and thus, in mineralization rates (El-Haris et al., 1983; Campbell et al., 1991) due to the accumulation of plant residues at or near the soil surface, which leads to greater organic C (Logan et al., 1991) and organic components, such as soluble and acid-hydrolysable carbohydrates (Angers et al., 1993).
Mineralization (i.e., rate of N release) of the crop residues left in soils not only depends on the environmental factors but also on the type of crop, the part and amounts of plant residues that are returned to the soil, and its contents of N, S, soluble C, lignin, and carbohydrates (Janzen and Kucey, 1988). The relative amounts of organic C to organic N (C:N ratio) of the crop residue returned to soil is a key factor influencing not only the mineralization process but also the immobilization process. Most reviews on C:N ratios indicate that plant residues, with wider C:N ratios (C:N > 30), incorporated into soils favor N immobilization and those with narrower ratios (C:N < 20) favor N mineralization (N release) early in the decomposition process.
Several biological and chemical methods have been proposed as indexes of plant available N in soils. In general, biological methods can be divided into two groups. The first group involves short-term (1-2 weeks) incubation of soils under aerobic or waterlogged conditions. The second group involves long-term (2-30 weeks) incubation of soils under aerobic conditions. A method was proposed by 40
Stanford and Smith (1972), which involves leaching of soil columns after incubation under aerobic conditions and determination of the inorganic N produced with steam distillation. A short-term incubation method developed by
Waring and Bremner (1964) that involves determination of the NH^ produced under anaerobic conditions when soil is incubated under waterlogged conditions at 30 °C for 2 weeks. Several chemical indexes have been proposed, and the degree of utility has been evaluated through correlation with biological measurements of available N in soils such as crop yield, mineralizable N, or plant uptake (Stanford, 1982). The activities of the enzymes involved in the N cycling in soils, such as the amidohydrolases and p-glucosaminidase, could be important tools assessing the potential of soils to release inorganic N.
Measurement of the activities of those enzymes and studies of the factors that affect them promise to provide a better understanding of the biochemical reactions involved in N mineralization in soils.
The work reported in this dissertation should provide the information needed to answer many of the question raised, and provide the basic knowledge needed to make better strategies in managing the soil resources in Iowa, and, perhaps, in other areas as well. 41
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CHAPTER 3
EFFECTS OF LIMING AND TILLAGE SYSTEMS ON MICROBIAL BIOMASS AND GLYCOSIDASES IN SOILS
A paper submitted to Biology and Fertility of Soils
Mine Ekenler and M. A. Tabatabai
Abstract This study was undertaken to investigate the long-term influence of lime application and tillage systems (no-till, ridge-till and chisel plow) on soil microbial biomass C (Cmic) and N (None) and the activities of glycosidases (a- and p-glucosidases, a- and P-galactosidases and (3- glucosaminidase) at their optimal pH values in soils at four research sites
(Southeast Research Center (SERC), Southwest Research Center (SWRC),
Northwest Research Center (NWRC), and Northeast Research Center
(NERC) in Iowa, U.S.A. Results showed that the pH values of the soils were significantly varied due to lime application; however, organic C and N concentration were not affected at each site. In general, Cmic was significantly
(P < 0.001) and positively correlated with soil pH, with r values ranging from
0.68 (P < 0.01) at the NWRC site under no-till (0-5 cm) to 0.81 (P < 0.001) at the NWRC site under no-till (0-15 cm). The Nmic values were also significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.66 (P < 0.001) at the SERC site to 0.92 (P < 0.001) at the
SWRC site. Lime application increased the activities of glycosidases 66 significantly (P < 0.001). Simple correlation coefficients between the activities of enzyme and soil pH values ranged from 0.51 (P < 0.05) for the activity of a-glucosidase at the NWRC site, no-till (0-5 cm) to 0.98 (P < 0.001) at the SWRC site. A activity/ A pH values were calculated to determine the sensitivity of the enzymes to changes in soil pH. Lime application did not significantly affect the specific activities (g p-nitrophenol released kg1
Corg h"1) of the enzymes. In general, their order of sensitivity to changes in soil pH was consistent across the study sites and it was as follow: P- glucosidase > (3-glucosaminidase > P-galactosidase > a-galactosidase > a- glucosidase. The enzyme activities were greater in soils under chisel plow
management than under the other tillage systems studied, and they were
greater in 0-5 cm samples than those in the 0-15 cm samples under no-till
system. Most of the enzyme activities were significantly and positively
correlated with Cmic and Nmic. Among the glycosidases studied, P-glucosidase
and p-glucosaminidase were the most sensitive to soil management practices.
Therefore, the activities of those enzymes may provide a reliable-long term
monitoring tools as early indicators of changes in soil properties induced by
liming and tillage systems.
Keywords Glycosidases Cmic, Nmic Liming - Tillage systems 67
Introduction
A major agricultural research priority has been recently focused on sustaining soil productivity and identifying key parameters or processes that are affected by long-term agricultural management practices. Long-term monitoring of soil biochemical processes is essential because it allows for identification of current changes in soil and for prediction of future changes
(Kandeler et al. 1999). Because many soil physical and chemical properties will hardly change over the next century monitoring should focus on those components, such as microbial biomass and microbial processes, expected to change more rapidly (Christensen 1996; Tscherko and Kandeler 1997).
Therefore, the microbial biomass and enzyme activities are mostly used as early indicators of changes in soil properties induced by liming and different tillage managements (Carter 1986; Powlson et al. 1987; Acosta-Martinez and
Tabatabai 2000).
Soil management practices influence soil microbial community structure and its biochemical processes through changes in the quantity and quality of plant residues entering the soil, their seasonal and spatial distribution, the ratio between above and below ground inputs, and through changes in nutrient inputs (Christensen 1996). Lime application to soils most often causes a significant increase in pH and, thus, a change in microbial biomass (Edmeades et al. 1981), microbial dynamic and diversity, and therefore, enzyme activities (Zelles et al. 1990; Acosta-Martinez and
Tabatabai 2000). Production of adequate crop yields and maintenance of soil 68 health required the addition of liming as well as the adoption of conservation tillage systems (Simard et al. 1994). Tillage systems result in changes in soil physical and chemical properties and enzyme activities (Deng and Tabatabai
1996). Soil organic matter is more thoroughly distributed in soils under conventional cultivation compared with that under reduced tillage, where crop residues are concentrated on the soil surface (Arshad et al. 1990).
Consequently, microbial biomass and processes in surface soils under no-till system are greater than those under other tillage systems.
Soil microbial biomass is involved in the decomposition of organic matter and, thus, the nutrient cycling in soils. Although Cmic constitutes 1-
3% of the total soil organic C and Nmic up to 5% of the total soil organic N, they are the most labile C and N pools in soils (Jenkinson and Ladd 1981;
Dick 1992). The turnover time for C and N immobilized in the microbial biomass has been reported to be about 10-times faster than that derived from plant materials (Smith and Paul 1990). Thus, nutrient availability and productivity of soils mostly depends on both the size and the activity of microbial biomass (Friedel et al. 1996). Microbial biomass measurement has been mostly used as an early indicator of changes in soil chemical and physical properties resulting from soil management and environmental stress in agricultural systems (Brookes 1995; Jordan et al. 1995). However, microbial biomass measurements cannot indicate the activity. Therefore, enzyme activities are often used as indices of microbial activity, because enzymatic activities in soils are important for catalyzing a web of reactions 69 necessary for life processes of microorganisms, decomposition of organic residues, cycling of nutrients, and formation of organic matter and soil structure (Frankenberger and Dick 1983; Dick 1994). It is suggested that studying soil enzyme activities provides insight into biochemical processes in soils and is very sensitive as biological indexes (Frankenberger and Dick
1983), and thus, it provides a useful tool for long-term monitor the changes in soil health and quality. Changes in microbial biomass and certain enzymes often occur within months or a year from the time a soil treatment is applied.
Therefore, any change in management is reflected in the microbial biomass and soil enzymes in a short period of time and long before there are
measurable changes in soil organic matter (Powlson et al. 1987; Dick 1994).
Glycosidases or glycoside hydrolases is a group of enzymes that catalyze the hydrolysis of different glycosides. The reaction involved is as follow;
glycosides + H2O—• sugar + aglycons
The terms glycoside and glucoside have been used interchangeably.
Glycosidases have usually been named based on the type of bond that they
hydrolyze. Among the glycosidases, a-glucosidase (EC 3.2.1.20), which
catalyzes the hydrolysis of a-D-glucopyranosides, and (3-glucosidase (EC
3.2.1.21), which catalyzes the hydrolysis of (3-D-glucopyranosides, are
involved in hydrolysis of maltose and cellobiose, respectively. a-Galactosidase
(EC 3.2.1.22) and (3-galactosidase (EC 3.2.1.23) catalyze the hydrolysis of 70 melibiose and lactose, respectively. These four enzymes in the group of glycosidases play a major role in the degradation of organic C in soils.
Another important glycosidase, P-glucosaminidase (NAGase, EC
3.2.1.30), is the enzyme that catalyzes the hydrolysis 2V-acetyl-P-D- glucosamine residue from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also classified as p-hexosaminidase (EC
3.21.52) by the International Union of Biochemistry because it cleaves the amino sugar iV-acetyl-p-D-galactosamine (Webb 1984). Some of the amino sugars in soils may exist in the form of alkali-insoluble polysaccharide referred to as chitin, which is considered one of the most abundant biopolymers in nature and an important pool of organic C and N in soils
(Stryer 1988; Wood et al. 1994). Thus, this enzyme is involved in C and N cycling in soils. Recent work showed that the activity of this enzyme is significantly affected by long-term copping systems at two sites and is related to cumulative N mineralized at 30°C during 24 weeks (r = 0.84 and 0.79, P<
0.001) (Ekenler and Tabatabai 2002).
The response of the soil microbial biomass and activities of glycosidases to liming and different tillage practices has mainly been investigated on soil samples in studies that have taken on only from short-term management sampling sites (Bardgett and Leemans 1995) or taken only from one site
(Deng and Tabatabai, 1996; Curci et al. 1997; Acosta-Martinez and Tabatabai
2000) or that studied only one enzyme to represent glycosidases (Dick et al.
1988; Bergstrom et al. 1998), but little information is available on the effects 71 of those practices on the activities of the glycosidases involved in C and N cycling in soils and on Cmic and Nmic in many long-term management sites.
Before a soil property or process can be used as soil quality index, systematic studies across many long-term soil management sites are needed to identify whether the results are site specific. Thus, the first purpose of the present study was to determine whether the response of the microbial biomass and activities of five glycosidases are consistent across four long-term management sites.
It is well established that the microbial biomass and the activities of glycosidases can be used as early indicators of changes in soil properties induced by liming and different tillage systems (Carter 1986; Powlson et al.
1987; Acosta-Martinez and Tabatabai 2000). However, to be useful, the activity of an enzyme as a soil quality index needs to be responsive to soil management practices or restoration efforts in a relatively short time (Dick
1994). Thus, the second purpose of our study was to assess the sensitivity of the glycosidases to liming of agricultural soils under different tillage systems.
Materials and methods
Experimental design and sampling
The studies were conducted on soils from four different sites in Iowa,
USA: Southeast Research Center (SERC site), Southwest Research Center 72
(SWRC site), Northwest Research Center (NWRC site), and Northeast
Research Center (NERC site).
SERC site: The experimental site was established in 1989 at
Crawfordsville, Iowa. The soil is a Mahaska-Taintor-Kalona Association.
Agricultural limestone was applied at the following rates: 0, 1120, 2240, 4480,
6720 kg of effective calcium carbonate equivalent (ECCE) ha1 as dolomite, or
560, 1120, or 2240 kg ECCE ha-1 as calcite. The treatments were arranged in a randomized complete block design with three replications. The size of each plot was 6 x 14 m. Corn (Zea mays L.) and soybean (Glycine max L.) were grown in alternate years. Sufficient levels of N, P, and K were applied to maintain high nutrient levels.
Surface soil samples (0 to 15 cm) were taken in June 1999 under soybean from all the field replicates, 10 years after lime application by pooling 14 cores (1.8 cm dia.) from each plot
SWRC site: The experimental site was established in 1995 at Atlantic,
Iowa. The soil is a Marshall silty clay loam (fine-silty, mixed, mesic superactive Typic Hapludoll). Agricultural limestone was applied at the following rates: 0, 2167, 6373, 19118, or 57355 kg ECCE ha*1. The treatments were arranged in a randomized complete block design with four replicates.
The size of each plot was 6 x 11.5 m. Corn and soybean were grown in alternate years. Sufficient levels of N, P, and K were applied to maintain high nutrient levels. 73
Surface soil samples (0 to 15 cm) were taken in May, 2001, before planting from two field replicates, six years after lime application by pooling
20 cores of samples as described above.
NWRC site. Aglime and tillage treatments were established in 1992 at
Sutherland, Iowa. The predominant soils are fine-silty, mixed, mesic superactive Typic Hapludolls (Galva), fine-silty, mixed, mesic superactive
Typic Endoaquolls (Marcus), and fine-silty, mixed, mesic Aquic Hapludolls
(Primghar). Agricultural limestone was applied at the following rates: 0, 560,
1120, 2240, 4480 or 6720 kg ECCE ha1. The tillage treatments were no-till, ridge till and chisel plow. The entire site was treated with a moldboard plow in fall 1991. Chisel plow was applied to approximately 15 cm depth per year with disk and field cultivation for seedbed preparation. Ridge-till cultivated twice per year to reform ridges. The experimental design was split plot with
treatments assigned in a randomized complete block with three field
replicates. The size of each plot was 6 x 14 m. Corn and soybean were grown
in alternate years, and 134 to 156 kg ha1 N fertilizer was applied as surface
broadcast ammonium nitrate to corn. Sufficient levels of P and K were
applied to maintain high nutrient levels.
Soil surface samples (0 to 15 cm) were taken in May 2000 under
soybean from three field replicates, eight years after lime application by
pooling 14 cores as described above. In addition, soil samples were taken
from 0-5 cm under the no-till system. 74
NERC site: This experimental site was established in 1984 at Nashua,
Iowa. The soil of the site is a Kenyon loam (fine-loamy, mixed, mesic Typic
Hapludoll). Agricultural limestone was applied at the following rates: 0,
1120, 2240, 4480, 6720, 8960, 13,440 or 17,920 kg (ECCE) ha1. The experimental design was randomized complete block design with four field replicates. The size of each plot was 6 x 15 m. Corn and soybean were grown in alternate years, with periodic applications of N, K, P fertilizers.
Soil surface samples (0 to 15 cm) were taken after corn harvest from this site, seven years after lime application by pooling 10 cores (7.6 cm dia.).
The properties and the activities of 15 enzymes of the samples obtained from this site were previously reported (Acosta-Martinez and Tabatabai, 2000).
Laboratory analyses
The field-moist soil samples from the SERC, SWRC, and NWRC sites were sieved through a 2-mm screen and a portion of this was air-dried. The air-dried portion of the samples was ground to pass an 80-mesh (180-p.m) sieve. Organic C and total N were determined on the <180-jim samples by dry combustion method using LECO CHN 600 determinator (St Joseph, MI,
USA). Organic N was determined from the difference between total and inorganic N (Keeney and Nelson 1982), pH was determined on the <2-mm mesh air-dried samples by using a combination glass electrode (soikO.Ol M
CaCl2 ratio = 1:2.5). When not in use, the field-moist samples were stored at
4°C. All results reported are averages of duplicate analyses and are 75 expressed on an oven-dry basis. Moisture content was determined from loss in weight after drying at 105°C for 48 h.
Microbial biomass C and N
The Cmic and Nmic in soils were determined on the <2-mm mesh field- moist soil samples within 24 h after sampling by the chloroform fumigation- extraction method described by Vance et al. (1987) and the chloroform fumigation-incubation method described by Horwath and Paul (1994), respectively.
Enzyme activities
The activities of the enzymes were assayed on the <2 mm field-moist samples at their optimal pH values in duplicates and one in control, and are expressed on a moisture-free basis.
The assay methods used are summarized in Table 1. For all data points reported in the figures, the differences between the laboratory duplicates were smaller than the point size. 76
Results
Effect of liming and tillage on soil chemical properties
The application of lime affected the pH values at each site, not the concentrations of organic C and N in soils. At the SERC site, the pH values ranged from 4.87 to 5.56; at the SWRC site, it ranged from 4.65 to 6.72; at the
NWRC site, it ranged from 4.38 to 5.77 in soils under no-till (0-5 cm) and 4.46 to 5.54 (0-15 cm), from 4.63 to 5.32 in soils under ridge till, and from 4.71 to
5.36 in soils under chisel plow; at NERC site, it ranged from 4.90 to 6.90
(Tables 2 and 3).
Effect of liming and tillage on soil microbial biomass C and N
Results showed that the Cmic values were significantly increased, except at the SWRC site, with increasing soil pH due to lime application, with r values 0.68 (P < 0.01) at NWRC site in soils under no-till (0-5 cm), 0.81(P <
0.001) no-till (0-15 cm), -0.84 (P < 0.01) at SWRC site, and 0.78 (P < 0.001) at
SERC site (Fig. 1). Similarly, the Nmic values were significantly (P < 0.001) and positively correlated with the pH values of the soils, with r values 0.89 (P
< 0.001) at NWRC site in soils under no-till (0-5 cm), 0.81 (P < 0.01) no-till
(0.15 cm), 0.92 (P < 0.001) at SWRC site, and 0.66 (P < 0.01) at SERC site
(Fig. 2). 77
Expressed in mg kg-1 soil, the average Cmic value was 218 at 0-5 cm depth, while it was 183 at 0-15 cm depth in soils under no-till systems. The average Nmic value was 12.6 at 0-5 cm depth and 8.0 at 0-15 cm depth under that tillage system. Therefore, this study indicated that Cmic and Nmic values in surface soils at 0-5 cm depth are greater than at 0-15 cm depth under no- till systems (Figs.l and 2).
Effects of liming and tillage on the glycosidases
The activities of the glycosidases were significantly and positively correlated with soil pH (Table 4). The correlation coefficient between soil pH and a-glucosidase ranged from 0.51 (P< 0.05) to 0.76 (P< 0.001), |3- glucosidase from 0.57 (P< 0.05) to 0.90 (P < 0.001), a-galactosidase 0.55 (P<
0.05) to 0.90 (P< 0.001), P-galactosidase from 0.59 (P< 0.01) to 0.83 (P< 0.001), and P glucosaminidase from 0.52 (P< 0.05) to 81 (P< 0.001).
Enzyme activities were mostly greater in soils at 0-5 cm depth compared with those in the 0-15 cm samples of the no-till system. For example, expressed in mg p-nitrophenol kg-1 soil h*1, p-glucosidase activity was 251 at highest lime application rate in the 0-5 cm depth, while it was 177 in the 0-15 cm sample.
The activity of P-glucosidase was also greater in the 0-15 cm samples under chisel plow compared with other systems at the same depth. For example, expressed in mg p-nitrophenol kg-1 soil h*1, the activity value was 78
214 in the 0-15 cm samples at the highest lime application rate in soil under chisel plow, whereas it was 177 and 160 in soils under no-till and ridge-till systems, respectively; at the same depth and rate of lime application.
To determine the sensitivity of the glycosidases to changes in soil pH, A activity/ A pH values for the individual glycosidase activities were calculated
(Table 5). The values for a-glucosidase ranged from —1.42 to 10.4, for p- glucosidase ranged from 14.0 to 75.7, for a-galactosidase from 4.50 to 24.9, for
P-galactosidase from 3.60 to 48.5, and for P-glucosaminidase from 2.99 to 27.9.
Thus, in general, the order of the sensitivity was P-glucosidase > P- glucosaminidase > P-galactosidase > a-galactosidase > a-glucosidase. This study also showed that the activity of p-glucosidase was the greatest among the glycosidase studied, followed by p-glucosaminidase, P-galactosidase, a- galactosidase, and a-glucosidase (Fig. 3).
Activities of glycosidases in relation to Cmic and Nmic
The Cmic values were significantly and positively correlated with glycosidase activities, with r values ranging from 0.49 (P <0.05) for P- galactosidase to 0.60 (P <0.01) for a-galactosidase and P-glucosaminidase at
SERC site, except for those obtained for soils from the SWRC site, which were negatively correlated, with r values ranging from -0.40 (n.s.) for a- galactosidase to -0.91 (P <0.001) for a-glucosidase (Fig. 4). Other correlation coefficients for those relationships were from -0.20 (n.s.) for a-galactosidase to 79
0.83 (P <0.001) for P-galactosidase in samples from the NWRC site no-till (0-5 cm), and from 0.31(n.s.) for a-glucosidase to 0.58 (P <0.05) for P-galactosidase for samples from the NWRC site no-till (0.15 cm) (Figs. 4 and 5).
The Nmic values were also significantly and positively correlated with the glycosidase activities, with r values ranging from 0.19 for a-glucosidase to
0.64 (P< 0.001) for p-galactosidase in samples from the SERC site, from 0.64
(P <0.05) for P-glucosaminidase to 0.91 (P <0.001) for P-galactosidase in samples from the SWRC site (Fig. 6). Those relationships in soils from the other sites ranged from -0.35 (n.s.) for a-galactosidase to 0.72 (P <0.001) for P- galactosidase in samples from the NWRC site no-till (0-5 cm), and from 0.33
(n.s.) for a-glucosidase to 0.62 (P<0.01) for P-glucosidase in samples from the
NWRC site no-till (0.15 cm) (Fig. 7).
The specific activity values, expressed in g p-nitrophenol released kg-1
Corgh1, were not significantly affected by lime application (soil pH), but were markedly different among the enzymes within each site and for the same enzyme among the sites (Tables 6 and 7).
Discussion
To show the effect of soil pH on enzyme activities, it is important that the concentration of organic matter and N remain constant. Results of the present study showed that soil pH was significantly increased after lime 80 application at all sites; however, the concentrations of organic C and N remained constant. Thus, this condition was satisfied.
Tillage systems affected the concentrations of organic C and N slightly, whereas lime application did not change those chemical components. Our result was in agreement with the results of other authors showing that chemical properties hardly change over a long period of time by soil management practices (Carter and Rennie 1982; Tscherko and Kandeler
1997). Soils under chisel plow had slightly greater organic C and N contents compared with those under no-till and ridge till systems; also their concentrations were greater in 0-5 cm samples than those in the 0-15 samples under no-till. The result of this study supported the results of Doran et al
(1998) indicating that organic C and N reserves were concentrated near the soil surface (0-7.6 cm) with no-till.
Results showed that Cmic (except for at SWRC site) and Nmic were significantly (P< 0.001) increased with increasing soil pH due to lime application (Figs. 1 and 2). Soil management practices, including liming and tillage, can influence soil biological activities through their effects on the quantity, structure, and distribution of soil organic matter (Doran and Power
1983; Doran et al. 1998). Liming may have increased the quality of C sources available to microbes, so that a greater proportion of the microbes may be active, while the total biomass is not affected. Systems with high organic matter inputs and easily available soil organic matter compounds have higher microbial biomass contents and activities because they are preferred energy 81 sources for microorganisms. The enhanced CO2 production, content of ATP, and heat output in lime applied soils showed that liming resulted a global increase in microbial activity. (Zelles et al. 1987; 1990). Contrary to the results obtained for the other research sites, the Cmic values decreased with increasing soil pH in soils taken from the SWRC site. This site received considerably higher amount of lime application, with the highest rate being
57355 kg ECCE ha1 , than the other sites. Thus, pH values were also highly increased in those plots. It is commonly accepted that acid or slightly acid soil pH favors to fungi, while bacterial biomass is increased under alkaline soil pH conditions (Baath et al. 1992). Zelles et al. (1990) reported that both ergosterol and glucosamine are good indicators of fungal growth, and that liming decreased the glucosamine and ergostereol. These results suggest that fungi are unable to function optimally at higher pH. The fungal biomass has higher C content compared with that of bacterial biomass and, thus, increasing soil pH decreased the amount of fungi in the environment; consequently, decreased Cmic Work by Campbell et al. (1991) showed that
Cmic is more sensitive to changes in soils management practices than is Nmic.
They concluded that Cmic may be more labile in soil than is Nmic. In pot experiments, liming usually increases the microbial biomass C (Carter 1986;
Illmer and Schinner 1991). However, results from field experiments have been variable (Priha and Smolander 1994). Simard et al. (1994) reported that liming did not affect Cmic- Baath et al. (1980) found that microbial biomass 82 was not affected 5-6 years after liming. However, Kratz et al. (1991) showed an increment in microbial biomass three years after liming.
Liming may affect fungal and bacterial biomass differently. Shah et al.
(1990) determined fungal and bacterial biomass separately and showed that liming increased the bacterial biomass for only a short term, while fungal biomass was not affected. Zelles et al. (1990) suggested that liming slightly increases the bacterial biomass, but significantly decreases the fungal biomass.
The biological environment of soils is greatly modified by the type and degree of tillage. Our work showed that Cmic and Nmic values in surface soils at 0-5 cm depth were greater than those in the 0-15 cm depth samples under no-till systems (Figs. 1 and 2). It is often reported that biological environment near the soil surface of no-till system is cooler and wetter than that of tilled soils, resulting in concentration of microbial activity and organic
C and N near the soil surface (0-7.6 cm) of the no-till system (Carter and Rennie 1982; Doran et al. 1998).
Glycosidases activities were significantly and positively correlated with soil pH (Table 4). This finding confirms the observations of Acosta-Martinez and Tabatabai (2000) reporting that enzyme activities of soils are increased by liming. The significant increase in soil pH by lime applications may stimulate the microbial population and diversity resulting in an increase in the soil enzyme activities and, thus, affecting nutrient cycling. It has been suggested that either the rates of synthesis and release of enzymes by soil 83 microorganisms are related to soil pH or that the stability of these enzymes in soils is related to soil pH (Deng and Tabatabai 1996). Although Cmic values decreased in soils from the SWRC site by liming, glycosidases enzyme activities were increased. This may be because soil pH improved the stability of the previously released enzyme in soils or liming may have increased the quality of C sources available to microbes, so that a great proportion of the microbes were active, while total Cmic was decreased.
Even though the concentrations of soil organic C and N remained constant, lime application significantly (P < 0.001) increased the activities of glycosidases (Table 4). This observation suggested that not only the quantity of soil organic matter, the chemical nature of organic matter has an impact on soil microbial activities and processes. Doran et al. (1998) reported that in addition to long-term changes in quantity of soil organic matter and related soil physical characteristics, quality of soil organic matter varies with
management practices as well
To determine the sensitivity of the glycosidase to changes in soil pH, A activity/ A pH values for the individual glycosidase activities were calculated
(Table 5). In general, the order of the sensitivity was: P-glucosidase > p- glucosaminidase > P-galactosidase > P-galactosidase > a-glucosidase across the research sites studied. Bergstrom et al. (1998) studied the activities of six enzymes, dehydrogenase, urease, glutaminase, phosphatase, arylsulfatase,
and P-glucosidase under different management practices and concluded that
soil enzyme activities are sensitive to changes in soil quality resulting from 84 soil management practices. However, enzymes differ in their sensitivity to those management practices. Two years after imposing various cropping systems, a significant effect on p-glucosidase activity was observed even thought there were no measurable difference in total C content (Dick 1994).
-1 Expressed in g p-nitrophenol released kg Corg h \ the specific activities varied among the five enzymes, but lime application having no significant effect (Tables 6 and 7), suggesting that the pH values neither induced the production of the enzymes nor affected the population of the microorganisms responsible for their production in soils.
Glycosidase activities were greater in the 0-5 cm depth samples than those in the 0-15 cm depth samples under no-till systems, which were similar to values found in the 0-15 cm depth samples under chisel plow (Fig. 3). This is probably the result of induction of microbial activities in surface soils (0-5 cm) under no-till system because a greater microbial biomass was measured at those soils. The greater glycosidase activities in soils treated with chisel plow compared to those values of the same depth of other tillage systems probably resulted from increased organic C and microbial biomass. Higher level of organic C stimulates microbial activity and, thus, enzyme synthesis.
Result showed that p-glucosidase has the highest activity values in soils at all sites (Fig. 3). The predominance of p-glucosidase in soils has been well documented (Eivazi and Tabatabai 1988; Tabatabai 1994). Recently, however, Klose and Tabatabai (2002) studied the concentrations of glycosidase proteins in 10 Iowa surface soils and showed that the average 85 enzyme protein concentrations in soils were 3.72, 0.014, 0.027 and 1.56 mg protein kg-1 soil for a- and p-glucosidases and a and P-galactosidases, respectively. Although P-glucosidase protein concentration was the lowest among the four glycosidases, it yielded the greatest activity values. The result suggested that this enzyme is associated with location different from those of other three glycosidases, or that this enzyme has greater catalytic efficiency than others. They finally concluded that enzyme activity rates do not necessarily correspond to the amounts of enzyme proteins present in a soil.
Most of the enzyme activities were significantly and positively correlated with Cmic and Nmic (Figs. 4-7). This result was in agreement with the study of Curci et al. (1997), reporting significant correlation between Cmic and a glucosidase or p-galactosidase activities. The strong positive relationship between enzyme activities and soil microbial biomass has been well established for other enzymes involve in N and S cycling in soils (Klose and Tabatabai 1999; Klose et al. 2000). This conclusion is not surprising because microbial biomass is the most important source of enzymes.
Conclusions
The pH values of soil were significantly varied by lime application,
while the concentration of organic C and N were not affected. Microbial
biomass of soils, in general, was significantly increased with increasing soil 86 pH. The increase in soil pH by lime application may stimulate the microbial population and diversity, resulting in an increase in glycosidase activities and, thus, affecting the C and N cycling in soils. The sensitivity of the glycosidases to soil pH varied considerably, but the order of the sensitivity has, in general, remained constant across the four research sites. The order of the sensitivity was as follows: P-glucosidase > P-glucosaminidase > P- galactosidase > a-galactosidase > a-glucosidase. Soil microbial biomass and the glycosidase activities were greater in the 0-5 cm depth sample than those in the 0-15 cm samples under no-till systems. Those values also were greater in the 0-15 cm depth samples under chisel plow than those treated with other tillage systems.
This study demonstrated that Cmic, Nmic, and the P-glucosidase activity could be used for monitoring changes in agricultural ecosystems resulted from human-imposed management and tillage systems.
Acknowledgements This work was supported in part by the
Biotechnology By-products Consortium of Iowa.
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Class/EC Recommended Reaction Assay conditions Assay number name * Substrateb conditions Optimum pH Glvcosidases
3.2.1.20 a-Glucosidase Glucoside-R + HsO -> Glucose + R-OH />-Nitrophenyl-a-D glucopyranoside (10 mAf ) 6,0
3.2.1.21 |)-Glucosidase Glucoside-R + H20 -> Glucose + R-OH p -Nitrophenyl-[)-D glucopyranoside (10 mAf) 6,0
3.2.1.22 a-Galactosidase Galactoside-R + H ,0 -> Galactose + R-OH p -Nitrophenyl-a-D galactopyranoside (10 mM) 6,0
3.2.1.23 (l-Galactosidase Galactoside-R + H;>0 -> Galactose + R-OH p Nitrophenyi p D galactopyranoside (10 mM) 6.0
3.2.1.30 P-Glucosaminidasec TV acetyl P-D glucosamine R + H%0 P -Nitrophenyl-N acetyl [)-D glucosaminide (2 mM) 5.5 N acetyl p D glucosamine + R-OH
* For the methods used, see Tabatabai (1994). b Figures in parentheses are the substrate concentrations under assay conditions. c Purham and Deng (2000), 94
Table 2 Effect of lime application rates on selected soil properties of soils at the SERC and SWRC sites
Lime treatment a XT . Type Rate pH Core No,P Cmic 1>4mic Cmjj/Corp Nmic/No, kg ECCE ha'1 ...gkg"1 soil... ..mg kg'1 soil. .%
SERC site
0 4.87b 26.4 2.26 153 7.4 5.8 3.3 Fine calcite 560 5.08 25.7 2.51 181 7.9 7.0 3.1 Dolomite 1120 4.95 26.1 2.17 165 7.5 6.3 3.5 Calcite 1120 4.88 26.5 2.27 171 5.3 6.5 2.3 Dolomite 2240 5.13 25.2 2.37 171 9.3 6.8 3.9 Calcite 2240 5.06 25.9 2.27 181 8.1 7.0 3.6 Dolomite 4480 5.27 25.9 2.27 181 8.7 7.0 3.8 Dolomite 6720 5.56 26.1 2.17 216 11.0 8.3 5.1
LSD (P < 0.05) 0.19 1.08 0.46 27.6 1.63 0.12 0.12
SWRC site
0 4.65b 18.6 3.33 221 8.1 11.9 2.4 Dolomite 2167 5.34 19.8 3.04 192 13.0 9.7 4.3 Dolomite 6373 5.61 19.2 3.09 189 14.4 9.8 4.7 Dolomite 19118 6.39 20.0 3.09 169 15.7 8.5 5.1 Dolomite 57355 6.72 22.3 3.04 144 14.8 6.5 4.9
LSD (P < 0.05) 0.56 1.96 0.61 84.7 2.85 10.3 1.7
°Soil:0.01 M CaCl2 solution ratio, 1:2.5. 'Values are averages of soil samples from three and two replicated field plots at the SERC and SWRC sites, respectively. 95
Table 3 Effect of lime application rates and tillage systems on selected soil properties at the NWRC site
Lime treatment pH* Core Nore Cmic Nmic C|ni(/Corg Nmic/N0,e kg ECCE ha1 g kg'1 soil.... mg kg'1 soil... % No-till (0-5 cm) 0 4.38b 25.1 3.67 195 11.0 7.8 3.0 560 4.87 27.2 3.87 203 10.7 7.5 2.8 1120 4.68 25.4 3.75 191 8.6 7.5 2.3 2240 4.86 27.1 3.78 220 12.2 8.1 3.2 4480 5.36 26.1 3.49 205 15.4 7.9 4.4 6720 5.77 25.7 3.49 295 17.6 11.5 5.0 LSD (P < 0.05) 0.8 4.5 0.6 93 14 3.4 3.8 No-till (0-15 cm) 0 4.46b 25.0 3.42 116 5.8 4.6 1.7 560 4.83 24.7 3.42 107 7.6 4.3 2.2 1120 4.69 25.8 3.55 101 6.8 3.9 1.9 2240 4.87 26.1 3.36 164 5.5 6.3 1.6 4480 5.25 26.2 3.56 230 8.3 8.8 2.3 6720 5.54 24.6 3.39 382 14.0 15.5 4.1 LSD (P < 0.05) 0.44 1.77 0.3 80.1 5.0 3.2 1.5 Ridge-till
0 4.63b 26.6 3.48 364 9.6 13.7 2.8 560 4.63 26.1 3.41 172 9.6 6.6 2.8 1120 4.85 26.7 3.54 181 4.1 6.8 1.2 2240 4.88 25.7 3.18 284 8.5 11.1 2.7 4480 5.05 25.4 3.28 165 6.1 6.5 1.9 6720 5.32 24.6 3.28 167 9.2 6.8 2.8 LSD (P < 0.05) 0.2 1.3 0.3 116 1.1 4.3 0.4 Chisel plow
0 4.71b 26.7 3.47 163 5.0 6.1 1.4 560 4.75 27.4 3.44 172 10.3 6.3 3.0 1120 4.85 27.8 3.55 169 10.8 6.1 3.0 2240 4.89 27.5 3.64 186 12.4 6.8 3.4 4480 5.12 27.9 3.68 178 5.8 6.4 1.6 6720 5.36 28.0 3.54 198 7.8 7.1 2.2 LSD (P < 0.05) 0.6 1.9 0.4 82 3.7 3.2 1.1 *Soil:0.01 M CaCl2 solution ratio, 1:2.5. ""Values are averages of soil samples from three replicated field plots. Table 4 Simple correlation coefficients for the relationships between enzyme activities" and soil pHh at the sites specified
NWRC
No-till Glycosidase SERC SWRC 0-5 cm 0-15 cm Ridge-till Chisel NERCc a-Glucosidase n.s. 0.98*** 0.51* n.s. n.s. 0.76*** 0.56*** P-Glucosidase n.s. 0.90*** 0.69** 0.57* 0.57* 0.81*** 0.87*** a-Galactosidase 0.82*** 0.90*** n.s. 0.64** 0.55* 0.72*** 0.53** P-Galactosidase 0 72*** 0.83*** 0.72*** 0.59** 0.63** 0.82*** 0.81*** P-Glucosaminidase 0.67*** 0.59* 0.52* 0.81*** 0.55* 0.81*** 0.55*
" All activities are expressed in mg p -nitroplienol released kg'1 soil h \ b Soil:0,01 M CaCl2 (ratio= 1:2,5). cAcosta-Martinez and Tabatabai (2000), Vs., Not significant, Table 5 A Enzyme activity"/ A soil pHh for the sites and tillage systems studied
NWRC
No-till Glycosidase SERC SWRC 0-5 cm 0-15 cin Ridge-till Chisel NERCc a-Glucosidase 4.4 -1.4 1.9 0.05 2.9 10 4.4 P-Glucosidase 39 14 46 35 75 76 39 a-Galactosidase 4.5 5.1 -9.3 12 13 25 4.5 p-Galactosidase 8.8 3.6 26 20 23 49 8.8 p-Glucosaminidase 25 28 3.0 7.7 5.9 15 25
" All activities are expressed in mg p -nitroplienol released kg1 soil h b Soil;0,01 M CaCl2 (ratio= 1:2.5), "Acosta Martinez and Tabatabai (2000), 98
Table 6 Specific activitya of glycosidases as affected by liming at the SERC and SWRC sites
Lime treatment Specific Activity Type Rate a-Glu P-Glu a-Gal P-Gal NAGase 1 1 1 kg ECCE ha" ..g of p -Nitrophenol kg" Corg h" ...
SERC site
0 0.24" 4.55 0.52 0.57 1.72 Fine calcite 560 0.22 4.40 0.57 0.56 2.06 Dolomite 1120 0.23 4.54 0.54 0.57 1.61 Calcite 1120 0.23 4.53 0.53 0.53 1.80 Dolomite 2240 0.24 4.83 0.60 0.65 1.84 Calcite 2240 0.21 4.53 0.58 0.59 1.97 Dolomite 4480 0.26 4.72 0.62 0.63 2.27 Dolomite 6720 0.24 5.22 0.70 0.68 2.28
LSD (P < 0.05) 0.07 0.76 0.08 0.07 0.63
SWRC site
0 0.40b 6.33 1.81 2.53 1.89 Dolomite 2167 0.48 7.11 1.97 2.96 2.00 Dolomite 6373 0.57 7.97 2.19 3.41 2.11 Dolomite 19118 0.63 7.78 2.09 3.24 1.91 Dolomite 57355 0.59 7.14 1.96 2.91 1.92
LSD (P < 0.05) 0.15 0.96 0.33 0.49 0.41 "Values are averages of samples from three replicated field plots at the SERC site. 'Values are averages of samples from two replicated field plots at the SWRC site. 99
Table 7 Specific activitya of glycosidases as affected by liming and tillage systems at the NWRC site Specific activity Lime treatment a-Glu (3 Glu a-Gal |0-Gal NAGase 1 1 1 kg ECCE ha g of p-Nitrophenol kg' Core h" ... No-till (0-5 cm)
0 0.48" 7.38 2.02 3.06 2.47 560 0.54 7.31 1.99 3.09 2.61 1120 0.53 7.40 1.49 3.11 2.60 2240 0.58 7.99 1.44 3.46 2.86 4480 0.59 8.46 1.48 3.73 2.69 6720 0.59 9.74 1.70 4.37 2.65 LSD (P < 0.05) 0.11 1.91 0.89 1.10 0.46 No-till (0-15 cm) 0 0.39 5.46 1.90 2.78 2.28 560 0.35 4.91 2.04 2.47 2.18 1120 0.34 5.16 2.17 2.60 2.29 2240 0.32 5.92 2.18 3.06 2.31 4480 0.36 6.00 2.28 3.13 2.56 6720 0.42 7.20 2.49 3.71 2.61 LSD (P < 0.05) 0.09 1.25 0.54 0.67 0.34 Ridge-till 0 0.18 4.80 1.15 2.14 1.54 560 0.22 4.51 1.23 1.99 1.52 1120 0.30 5.69 1.56 2.63 1.71 2240 0.31 5.91 1.60 2.71 1.72 4480 0.29 5.67 1.50 2.59 1.75 6720 0.31 6.48 1.70 2.92 1.83 LSD (P < 0.05) 0.16 0.89 0.37 0.42 0.15 Chisel plow 0 0.22 5.83 1.70 2.66 1.78 560 0.19 5.79 1.60 2.65 1.74 1120 0.29 6.22 1.76 2.92 1.85 2240 0.39 6.98 2.01 3.37 1.98 4480 0.37 6.95 1.92 3.46 1.98 6720 0.49 7.66 2.31 3.83 2.11 LSD (P <0.05) 0.15 1.04 0.39 0.62 0.23 "Values are averages of soil samples from three replicated field plots. Y = -135 + 70,8 X -890 + 217 X NWRC site 500 NWRC site r = 0,68** No-till (0-5 cm) r = 0,81*** No-till (0-15 cm)
400
300
200
100
-L X ri U X Y = 392 - 36,4 X SWRC site . Y = 193 +72.6 X SERC site m 500 r = -0.84*** (015 cm) T - 0,78*** (0-15 cm) S f 400 O 300
200 O 100
0 0.0 4.0 4.5 5,0 5,5 6,0 6.5 7.0 0,0 4.0 4,5 5,0 5.5 6,0 6.5 7,0
Soil pH
Fig, 1 Relationships between soil pH and Cmic at indicated research sites, tillage and depths. Open symbols are results obtained for individual soil samples and solid symbols are averages obtained for the field replicates. In calculating the regression equations, the results of the individual soil samples were used (** and *** indicate significant at P < 0.01 and 0,001, respectively) -7^ -7^ 30 Y = -34.9 + 9.36 X NWRC site Y = -27.4 +7,17 X NWRC site No-till (0-5 cm) No-till (0 15 cm) r- 0.89*** r = 0,81*** 25
20 8 15
10
5
0 -7/ -I I I 1 1_ L_ » ' ' • 30 Y = -99.9 + 37.2 -2.99 X SWRC site Y =18.6 +5,24 X SERC site <0-15 cm) r = 0,92*** (0 15 cm) r = 0.66*** 25
20
15
10
5 0 -7^ _1 I I -7^" i i i i i i— 0. 3 4.0 4.5 5.0 5.5 6.0 6.5 7,0 0.0 4.0 4.5 5.0 5,5 6.0 6.5 7,0
Soil pH
is b< itween soil pH and Nmic at indicated research sites, tillage and depths. Open symbols are results lual soil samples and solid symbols are averages obtained for the field replicates. In calculating the is, t ie results of the individual soil samples were used (*** indicates significant at P < 0.001) 102
-/A 350 P-glucosidase SERC site P-glucosidase 300 r = 0.26 r = 0.90*** SWRC site 250 200 150 \ Soil pH between, soil pH and activities of glycosidases at i 275 O u-glucosidase A P-galactosidase SERC site SWRC site 250 # P-glucosidase 0 p-glucosaminidase A a-galactosidase 225 200 175 150 r = 0.50** r = -0.82** 125 100 75 r = 0,60** r = -0.80** 50 r = 0.49* r= -0.86** 25 A r = 0.60** r = n.s, r = ,-0.91*** 0 LVZ- ~Q r.= i 0 100 150 200 250 100 150 200 250 Cmic (mg kg"1 soil) Fig. 4 Relationships between Cmic and activities of glycosidases at the SERC and SWRC sites (*, **, and *** indicate significant at P< 0.05, 0,01, and 0.001, respectively) 350 FVZ o a-glucosidase p-galactosidase NWRC site a NWRC site • P-glucosidase o p-glucosaminidase No-till (0-5 cm) 300 - No-till (0-15 cm) A a-galactosidase e $ r = 0,82*** 1 250 3 I 200 r = 0,56** 0) es 150 r = 0.83*** 100 r = 0.58** Io, r= 0.49* r= n.s, !ho 50 r = n.s. E A A A A r = n.s. oocc^bQr&CP—ooo° r = 0.56* -oo r= n.s. 0 I I I 1 -poop-p—Qr 0 50 100 150 200 250 300 350 400 450 500 550 0 100 150 200 250 300 350 400 450 500 550 Cmic (mg kg'1 soil) Fig. 5 Relationships between Cmic and activities of glycosidases at the NWRC site for the 0-5 cm and 0-15 cm depth in soils under no-till system (*, **, and *** indicate significant at P< 0.05, 0,01, and 0,001, respectively) 275 A O a-glucosidase p-galactosidase SERC site SWRC site 250 • p-glucosidase D p-glucosaminidase A a-galactosidase 225 200 175 r = 0.88*** 150 r = 0.46* 125 100 • 75 r = n.s. D • r = 0.91*** 50 CÙ • O r = 0,64*** r = 0.64* 25 r = 0.51** r = 0.89*** r = n.s. tpo— Nmic (mg kg"1 soil) Fig. 6 Relationships between Nmic and activities of glycosidases at the SERC and SWRC sites (*, **, and *** indicate significant at P< 0.05, 0.01, and 0.001, respectively) 350 o a-glucosidase A P-galactosidase • p-glucosidase d p-glucosaminidase 300 A a-galactosidase NWRC site NWRC site 43 No-till (0-5 cm) No-till (0-15 cm) 1 250 r = 0.69** ho î 200 r = 0.62** Q) Cti TJ 150 'm r = 0.71*** • A r = 0,61** 100 r = 0.47* I1 O. r = 0.53* ho a 50 r = n.s. r = n.s. r = 0.64** r = 0.33 _l i i l_ 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 340 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 Nmic (mg kg'1 soil) Fig, 7 Relationships between Nmic and activities of glycosidases at the NWRC site for the 0-5 cm and 0-15 cm depth in soils under no-till system (*, **, and *** indicate significant at P< 0.05, 0.01, and 0.001, respectively) 107 CHAPTER 4 ARYLAMIDASE AND AMIDOHYDROLASES IN SOILS AS AFFECTED BY LIMING AND TILLAGE SYSTEMS A paper submitted to Soil & Tillage Research Mine Ekenler and M. A. Tabatabai Abstract This study investigated the long-term effect of lime application and tillage systems (no-till, ridge-till and chisel plow) on the activities of arylamidase and amidohydrolases involved in N cycling in soils at four long-term research sites in Iowa, U S A. The activities of the following enzymes were assayed: arylamidase, L-asparaginase, L-glutaminase, amidase, urease, and L- aspartase at their optimal pH values. The activities of the enzymes were significantly (P< 0.001) and positively correlated with soil pH, with r values ranging from 0.42* to 0.99*** for arylamidase, 0.81*** to 0.97*** for L- asparaginase, 0.62*** to 0.97*** for L-glutaminase, 0.61*** to 0.98*** for amidase, 0.66** to 0.96*** for urease, and 0.80*** to 0.99*** for L-aspartase. The A activity/ A pH values were calculated to assess the sensitivity of the enzymes to changes in soil pH. The order of the sensitivity of enzymes was as follows: L-glutaminase > L-asparaginase > amidase > arylamidase > urease > L-aspartase. The enzyme activities were greater in the samples of the 0-5 cm 108 depth than those of the 0-15 cm samples under no-till treatment. Most of the enzyme activities were significantly (P< 0.001) and positively correlated with microbial biomass C (Cmic) and N (Nmic). Lime application significantly affected the specific activities of the six enzymes studied. Results showed that soil management practices, including liming and type of tillage significantly affect soil biological and biochemical properties, which may lead to changes in nitrogen cycling, including N mineralization and nitrification, in soils. Keywords: Arylamidase; Amidohydrolases; Microbial biomass; Liming; Tillage systems 1. Introduction Recent interest in defining soil quality focused on identifying soil properties that reflect the effects of agricultural management practices on soil health and quality and long-term sustainability of terrestrial ecosystems. Because many physical and chemical properties of soils will hardly change over a century (Tscherko and Kandeler, 1997), studies are needed to find standard measurements that can be used as soil quality indicators. Such measurements should focus on soil components that expected to change more rapidly, such as microbial biomass and biochemical processes (Christensen, 109 1996). It has been reported that any change in soil management is reflected in the microbial biomass and soil enzymes in a short-period of time and long before there are measurable changes in soil organic matter (Powlson et al., 1987; Dick, 1994). Enzyme activities have been suggested as early indicators of changes in soil properties induced by liming and tillage (Carter, 1986; Powlson et al., 1987; Acosta-Martinez and Tabatabai, 2000a), because of their rapid response to change in soil management practices, their relationship to soil biology, and ease of assay. A significant increase in soil pH by liming affects microbial biomass (Edmeades et al., 1981), microbial dynamic and diversity, and therefore, enzyme activities (Zelles et al., 1990; Acosta-Martinez and Tabatabai, 2000a). Recent work showed that lime treatment and cropping systems significantly affect the rates of nitrification and N mineralization in soils (Tabatabai et al., 1992; Deng and Tabatabai, 2000; Senwo and Tabatabai, 2002). In addition, the biological environment of soil is strongly modified by type and degree of tillage systems. The different soil disturbances lead to significant changes in the composition, distribution, and the activities of soil microbial communities and enzymes through changes in the quantity and quality of plant residues entering the soil, their seasonal and spatial distribution, and the ratio between above and belowground inputs (Christensen, 1996; Deng and Tabatabai, 1996). Soil organic matter is more thoroughly distributed in soils under conventional tillage compared with that under reduced tillage, where crop residues are concentrated on the soil surface (Arshad et al., 1990). 110 Consequently, microbial biomass and microbial processes in surface soils under no-till system are greater than those under other tillage systems. Enzymatic activities are important for catalyzing a complex web of chemical reactions necessary for life processes in soils, decomposition of organic residues, cycling of nutrients, formation of organic matter, and soil structure (Dick, 1994). It is suggested that enzyme activities provide insight into the biochemical processes in soils and are useful sensitive biological indexes (Frankenberger and Dick, 1983), and thus, they provide a useful tool for long-term monitoring changes in soil health and quality. Arylamidase [EC 3.4.11.2] catalyzes the hydrolysis of an N-terminal amino acid from peptides, amides, or arylamidase according to the following reaction using L-leucine as amino acid moiety: o .ch(chj), hooc, Arylamidase •chch:ch(ch,): L-Leucine (3-naphthylamide P-Naphthylamine Leucine The activity of arylamidase was recently detected in soils and a method was developed for its assay (Acosta-Martinez and Tabatabai, 2000b). This enzyme catalyze one of the initial reactions in N mineralization, because it is Ill involved in the release of amino acids from soil organic matter and these released amino acids are the substrate for amidohydrolases involved in N mineralization: ino acid Ammoaification by Nitrification amidohydrolases: e.g., Arylamidas^ Amino acid NH/ X NO, X. NO, + energy activity (RNHJ L-Glutaminase activity L-Asparaginase activity L-Aspartase activity Amidohydrolases are enzymes involved in the hydrolyses of organic N compounds in soils. These enzymes are widely distributed in nature and have been detected in plants, animals, and microorganisms (Tabatabai, 1994); however, the most important source of these enzymes is soil microorganisms. The enzyme L-asparaginase (EC 3.5.1.1) catalyzes the hydrolysis of L- asparagine to L-aspartic acid and NH3. The enzyme L-glutaminase (EC 3.5.1.2) catalyzes the hydrolysis of L-glutamine to L-glutamic acid and NH3. These enzymes have an important role in N mineralization in soils and thus, N supply for plants, because the chemical nature of N in soils is such that a large proportion (15-26%) of the organic N is often released as NHa"1" by acid hydrolysis (6 M HC1), a portion of which is derived from the hydrolysis of amides such as asparagines and glutamine residues in soil organic matter (Sowden, 1958). Amidase (EC 3.5.1.4) hydrolyzes aliphatic amides to NH3 and the corresponding carboxylic acid, and it has microbial origin in soils. Amidase 112 activity in soils is particularly important for crop production because its substrates, aliphatic amides, are potential N fertilizer. N mineralization in soils is predominantly controlled by biochemical processes and active N pools in soils are strongly correlated with amidase activity (Deng et al., 2000). The studies on acid hydrolysis of humic preparation showed that 7.3 to 12.6% of the total N was present in the form of amide-N, and that the percentage of + NH4 released during acid hydrolysis was equal to the sum of aspartic acid-N plus glutamic acid-N derived from asparagines and glutamine (Bremner, 1955; Sowden, 1958). Urease (EC 3.5.1.5) catalyzes the hydrolysis of urea to CO-2 and NH3. It acts on C-N bonds other than peptide bonds in linear amides. Therefore, it belongs to a group of enzymes that include glutaminase and amidase. Urease is very widely distributed in nature. It has been detected in microorganisms, plants, and animals (Tabatabai, 1994). L-Aspartase (EC 4.3.1.1) is the enzyme that catalyzes the hydrolysis of L-aspartate to fumarate and NH3. The product of this enzyme, L-aspartic acid, is a major source of N mineralization in soils (Senwo and Tabatabai, 1996). Although liming and tillage systems are very common soil management practices effecting soil health and quality, their role in modifying the enzymes involved in N cycling in soils has not been demonstrated clearly. Therefore, the first objective of this study was to asses the effect of liming and various tillage systems on microbial biomass and the activities of six enzymes 113 involved in N cycling in soils, and to assess whether the response of those activities are similar in four long-term management sites in Iowa. To be useful, the activity of the enzyme as a soil quality index needs to be more responsive to soil management practices in a relatively short time (Dick, 1994). Thus, the second objective of this study was to determine the order of the sensitivity of the enzymes to lime treatment of agricultural soils and under different tillage systems. Even though enzyme activities in soils are associated with the microbial biomass (Klose and Tabatabai, 2000), and response of the microbial activity to soil disturbance is measured by enzyme activity (Dick, 1992), little is known about relationships between soil microbial biomass and enzyme activities under those treatments. Therefore, the third objective of this study was to assess the relationship between microbial biomass and the activities of arylamidase and amidohydrolases. 2. Materials and methods 2.1. Experimental sites and soil sampling The study was conducted using surface soils from four long-term management sites in Iowa, USA; Southeast Research Center (SERC site) at Crawfordsville, Southwest Research Center (SWRC site) at Atlantic, Northwest Research Center (NWRC site) at Sutherland, and Northeast 114 Research Center (NERC site) at Nashua. The experimental sites and designs, soil sampling and preparation, and analysis of the soil samples are described with details by Ekenler and Tabatabai (2002). 2.2. Soil analyses The field-moist soil samples from the SERC, SWRC, and NWRC sites were sieved through a 2-mm screen and a portion of this was air-dried. The air-dried portion of the samples was ground to pass an 80-mesh (180-p.m) sieve. Organic C and total N were determined on the <180-^im samples by dry combustion method using LECO CHN 600 determinater (St Joseph, MI). Organic N was determined from the difference between total and inorganic N (Keeney and Nelson, 1982), pH was determined on the <2-mm mesh air-dried samples by using a combination glass electrode (soil: 0.01 M CaCl2 ratio = 1:2.5). When not in use, the field-moist samples were stored at 4°C. All results reported are averages of duplicate analyses and are expressed on an oven-dry basis. Moisture content was determined from loss in weight after drying at 105°C for 48 h. 2.3. Microbial biomass C (Cmic) and N (Nmic) The Cmic and Nmic in soils were determined on the <2-mm mesh field- moist soil samples within 24 h after sampling by the chloroform fumigation- 115 extraction method described by Vance et al. (1987) and the chloroform fumigation-incubation method described by Horwath and Paul (1994), respectively. 2.4. Enzyme activities The activities of the enzymes were assayed on < 2 mm field-moist samples at their optimal pH values in duplicates and one in control, and are expressed on a moisture-free basis, except for samples from the NERC site, which was done on air-dried soils. The effects of liming of agricultural soils on the activities of 14 enzymes at the NERC site are reported by Acosta-Martinez and Tabatabai (2000a), and are summarized here for comparison. Moisture content was determined from loss in weight after drying at 105 °C for 48 h. The assay methods used are summarized in Table 1. 2.5. Specific activities From the activity values and organic C, the specific activity values were 1 -1 calculated as g of NH4+-N produced kg" Corg h for the amidohydrolases and 1 l as g (3-naphthylamine produced kg- Corg h for arylamidase. 2.6. Statistical analyses 116 Statistical analyses, including regression analyses and means separation by the least significant difference were performed according to general linear model procedure of the SAS system (SAS Institute, 1996). For all data points reported in the figures, the differences between the laboratory duplicates were smaller than the point size. 3. Results 3.1. Effects of liming and tillage on arylamidase activity Arylamidase activity was significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.42* to 0.99*** (Table 2). To assess the sensitivity of these enzymes to changes in soil pH, A activity/ A pH values were calculated for each enzyme at each site and the values ranged from 8.48 to 37.7 (Table 3). Results showed that arylamidase activity was greater in samples of the 0- 5 cm depth than of the 0-15 cm samples under no-till (Fig. 1). For example, expressed in mg (3-naphthylamine kg-1 soil h1, the activity value of arylamidase was 30 in the 0-5 cm sample, whereas it was 25 in the 0-15 cm sample of the control plots of the no-till system. The values were also slightly higher in soils under chisel plow than under no-till and ridge-till in the 0-15 cm samples. 117 3.2. Effects of liming and tillage on the activities of amidohydrolases The activities of amidohydrolases were significantly (P < 0.001) and positively correlated with soil pH, with r values ranged from 0.81*** to 0.97*** for ^ asparaginase, 0.62*** to 0.97*** for L-glutaminase, 0.61*** to 0.98*** for amidase, 66*** to 0.96*** for urease, and 0.80*** to 0.99*** for L- aspartase (Table 2). The sensitivity of the enzymes to changes in soil pH varied significantly (P < 0.001) (Table 3). A enzyme activity / A pH values for the individual amidohydrolases ranged as follows: 11.8 - 47.6 for L- asparaginase, 43.1 - 164 for L-glutaminase, 14.4 - 45.8 for amidase, 2.64 - 7.3 for urease, and 1.0 - 3.76 for L-aspartase (Table 3). Results showed that the order of the sensitivity of enzymes to change in soil pH were consistent at SERC, SWRC, and NERC site and it was as follows: L-glutaminase > L- asparaginase > amidase > arylamidase > urease > L-aspartase. L- glutaminase was not only the most sensitive enzyme to changes in soil pH, but it was also the most sensitive to different tillage systems, whereas L- aspartase was the least sensitive enzyme among the enzyme studied. Results also showed that L-glutaminase was the most predominant of the N cycling enzymes, followed by amidase, L-asparaginase, arylamidase, urease, and L-aspartase in soils. For example, expressed in NH4+-N kg*1 soil h"1, the L-glutaminase activity was 171, whereas the L-aspartase activity was only 3.8 in the highest lime application rate (57355 kg ECCE ha-1) at the SWRC site (Fig. 2) 118 The activities of the amidohydrolases were greater in the 0-5 cm samples than in the 0-15 cm depth samples under no-till system. For example, expressed in mg NH^-N kg*1 soil h l, the amidase activity was 53 in the 0-5 cm of the control plots under no-till, whereas it was 44 in the 0-15 cm samples. The activity values were similar in soils under no-till and chisel plow of the 0-15 cm depth and those values were higher than those for similar samples under ridge-till. 3.3 Arylamidase activity in relation to Cmic and Nmic The activity of arylamidase was significantly (P< 0.001) and positively correlated with Cmic at the NWRC site under no-till, with r values of 0.52* and 0.75*** for the 0-5 cm and the 0-15 cm depths, respectively. The correlation value was -0.77** at the SWRC site and there was no significant correlation among those variables at the SERC site (Table 4). Except for samples from the SERC site, the activity values were also significantly (P < 0.001) and positively correlated with Nmic, with r values 0.92***, 0.73***, and 0.64** for the samples from the SWRC site, NWRC site no-till (0-5 cm), and (0-15 cm), respectively (Table 5). 119 3.4 Activities of amidohydrolases in relation to Cmic and Nmic The enzyme activities were mostly significantly (P < 0.001) and positively correlated with Cmic, with r values ranging 0.48*- 0.84*** for L-asparaginase, 0.44* - 0.85*** for L-glutaminase, 0.59** - 0.72*** for amidase, 0.51* - 0.70*** for urease, and n.s.- 0.86*** for L-aspartase (Table 4). The correlation coefficient values were negative for the samples from the SWRC site. The activities were also significantly (P < 0.001) and positively correlated with Nmic, with r values ranging 0.54** - 0.83** for L-asparaginase, n.s. — 0.83** for L-glutaminase, 0.57** - 0.88*** for amidase, 0.45* - 0.85** for urease, and 0.57** - 0.80** for L-aspartase (Table 5). The specific activity values, expressed in g p-nitrophenol released kg*1 Corg h1, were significantly affected by lime application (soil pH) and markedly differed for the enzymes within each site, and for the same enzyme among the sites (Tables 6 and 7). 4. Discussion The significant, positive correlations between enzyme activities and soil pH (Table 2) support the previous findings showing that those enzymes are significantly increased with increasing soil pH, with correlation coefficient values 0.55** for arylamidase (Acosta-Martinez and Tabatabai, 2001a), 120 0.74***, 0.77***, and 0.72*** for L-asparaginase, L-glutaminase, and urease (Deng and Tabatabai, 1996). L-Glutaminase was the most sensitive of N cycling enzymes, followed by L-asparaginase, amidase, arylamidase, urease, and L-aspartase (Table 3). Bergstrom et al (1998) studied the activities of six enzymes, including urease and L-glutaminase, under different management practices and showed that soil enzyme activities are sensitive to changes in soil quality resulting from soil management practices; however, they differ in their sensitivity to those management practices. Our results showed that L-glutaminase was the most predominant and sensitive to liming, followed by amidase, L-asparaginase, arylamidase, urease, and L-aspartase (Figs, land 2). These support the conclusion of Sowden (1958) showing that most of the released in soils is derived from hydrolysis of amide (asparagine, aspartate and glutamine). Estimation of the enzyme protein concentrations in 10 Iowa surface soils by Klose and Tabatabai (1999, 2002), however, showed that the averages were 0.50, 0.73, 3.38, 5.3, 2.61 mg protein kg1 soil for L-glutaminase, L-asparaginase, amidase, urease, and L-aspartase. Even thought L-glutaminase and L- asparaginase protein concentrations were the lowest among 5 amidohydrolases studied, they yielded the greatest activity values. This suggests that these enzymes have greater catalytic efficiency than others. Even though the concentrations of soil organic C and N remained constant within each research site (Ekenler and Tabatabai, 2002), lime application 121 significantly (f<0.001) improved the activities of arylamidase and amidohydrolases (Figs. 1 and 2). This suggests that not only the quantity of soil organic matter, but the chemical nature of organic matter has an impact on soil microbial activities and processes. Doran et al (1998) reported that in addition to long-term changes in quantity of soil organic matter and related soil physical characteristics, quality of soil organic matter varies with management practices as well. Indeed, lime application significantly affected the specific activity values of the six enzymes studied (Tables 6 and 7). Enzyme activities were greater in the 0-5 cm samples than those in the 0- 15 cm depth under no-till systems. The variation in enzyme activities of soils between different tillage management and the depth may be due to differences in quality and distribution of crop residues that generally lead to a greater concentration of organic material in surface soils (0-5 cm) under not- till or reduced till compared with those under other tillage systems and, thus, to higher microbial biomass and enzyme activities. In fact, those soils showing greater enzyme activities had higher organic C and total N (Ekenler and Tabatabai, 2002), which are substrates for soil organisms near the soil surface. Consequently, soil microbial biomass and various soil microbial processes tend to increase in surface soils. This finding confirmed the observations of others reporting that the types of reduced tillage enrich the soil surface with organic C and N reserves, microbial biomass, microbial activity, and soil enzymes (Doran et al., 1998; Kandeler et al., 1999). The studies by Deng and Tabatabai (1996) showed that the activities of 122 amidohydrolases were strongly correlated with organic C in the 40 soil samples studied and the activities decreased markedly with increasing soil depth and this decreased was associated with a decrease in organic C content. Arylamidase was also significantly correlated with organic C (r = 0.80***) and total N (r = 0.71***) (Acosta-Martinez and Tabatabai, 2001b). Other studies have shown that the activities of amidase and urease in a 0-7.5 cm sample were significantly greater in soils from no-till plot than those under conventional tillage plots (Dick, 1984). The enzyme activities were significantly and positively correlated with the Cmic and Nmic values (Tables 4 andô). Similar findings were reported by Klose and Tabatabai (1999) for the relationships between total urease activity and Cmic (r = 0.84**) and Nmic (r = 0.76**) for 10 Iowa surface soils. Strong positive correlations were also reported between L-glutaminase, L- asparaginase, L-aspartase and Cmic and Nmic (Klose and Tabatabai, 2002). The strong correlation between the enzyme activities and Cmic or Nmic support the well-recognized hypothesis that urease in soils is mainly of microbial origin. Soil management practices, including liming and tillage systems, influence soil microbial biomass and their activities through their effects on the quantity, structure and distribution of soil organic matter (Doran and Power, 1983; Doran et al., 1998). In soils taken from the SWRC site, liming decreased Cmic, while increased the activities of the enzymes. This site received considerably higher rates of lime (up to 57355 kg ECCE ha*1) than the other sites. It is commonly 123 accepted that acid or slightly acid soil pH favors growth of fungi, whereas neutral or alkaline pH values favor growth of bacterial biomass (Baath et al., 1992). Fungal biomass contains higher C concentrations than bacterial biomass (Campbell et al., 1991) and thus, increasing soil pH decreased the amount of fungi in the soil environment; consequently, decreased Cmic. It has been reported that liming significantly decrease the fungal biomass (i.e., Cmic); but because of the increase of the bacterial biomass, the total enzyme activity increase (Zelles et al., 1987; 1990). The increase in enzyme activities at the SWRC site may also be the result of improvement in the stability of the previously released enzymes in soils or liming may have increased the quality of C sources available to microbes so that a great proportion of microbes remained active, while fungal population, consequently, total Cmic, was decreased. 5. Conclusions The enzymes arylamidase and amidohydrolases are sensitive to changes in soil quality resulting from liming and tillage practices. The activities of these enzymes significantly increased with increasing lime treatments. The significant increase in soil pH by liming seems to have stimulated the microbial biomass and diversity resulting in an increase in the activities of the enzymes and, thus, perhaps affecting N cycling in soils. Enzyme activities differed in their sensitivity to changes in soil pH; however, the order 124 of the sensitivity remained consistent across three research sites. The order of the sensitivity was as follows: L-glutaminase > L-asparaginase > amidase > arylamidase > urease > L-aspartase. Enzyme activities were greater in the 0-5 cm samples than those in the 0-15 cm depth samples under no-till systems. Most of the enzyme activities were strongly correlated with Cmic and Nmic. Results suggested that soil microbial processes, such as activity measurement of arylamidase and amidohydrolases, hold potential as a soil quality indicator because the enzyme assay procedures are very accurate, relatively simple, rapid, and can be done on a routine basis. 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Table 1 The methods used for assay of enzyme activities in soils Class/EC Recommended Reaction Assay conditions Assay conditions number name* Substrate1* Optimum pH Arylamidase and Amidohydrolases 3.4,11,2 Arylamidase' L leucine |)-naphthylamine -* L leucine + (i-Naphthylam L leucine ^-naphthylamine (2,0 niAf ) 8.0 3.5,1,1 L Asparaginase L-Asparagine + H20 —> L aspartate + NHs L -Asparagine (50 inM ) 10.0 3,5,1,2 L-Glutaminase L-Glutamine + H%0 —> L glutamate + NH3 L-Glutamine (50 mM) 10.0 3,5.1,4 Amidase R-CONHz + H20 -> NH3 + R-COOH Formamide (50 mM ) 8.5 3,5,1,5 Urease Urea + H%0 —» COa + 2NH3 Urea (20 mM) 9.0 d 4.3.1.1 L-Aspartase L Aspartate + H O —» L-aspartic acid + NHa L-Aspartate (200 mAf ) 8,5 * For the methods used, see Tabatabai (1994), b Figures in parentheses are the substrate concentrations under assay conditions. c Acosta-Martinez and Tabatabai (2000). d Senwo and Tabatabai (1996). Table 2 Simple correlation coefficients for the relationships between enzyme activities" and soil pH1' at the research sites indicated NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm Ridge-till Chisel NERCc Arylamidase 0.42* 0.99*** 0.84*** 0.86*** 0.89*** 0.90*** 0.74*** L-Asparaginase 0.86*** 0.97*** 0.81*** 0.91*** 0.96*** 0.95*** 0.84*** L-Glutaminase 0.62*** 0.97*** 0.85*** 0.90*** 0.89*** 0.93*** 0.73*** Amidase 0.87*** 0,98*** 0.93*** 0.76*** 0.81*** 0.93*** 0.61*** Urease 0.81*** 0.96"" 0.66** 0.92*** 0.71*** 0.68** 0.71*** L-Aspartase 0.87*** 0.99*** 0.80*** 0.92*** 0.91*** 0.89*** 0.80*** * Except for arylamidase activity, which is expressed in mg p-naplithylamine produced kg 1 soil h all other enzyme activities are expressed in mg NH/-N produced kg'1 soil li'\ *, **, and *** indicate significance at P< 0,05, 0,01, and 0,001, respectively. b Soil:0,01 M CaCl2 (ratio= 1:2,5), "Acosta-Martinez and Tabatabai (2000), Table 3 A Enzyme activity"/ A soil pH1' for the sites and tillage systems studied NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm Ridge-till Chisel NERCc Arylamidase 8.48 9.44 16.1 19.8 25.1 37.7 9.0 L-Asparaginase 47.6 13.5 11.8 21.5 30.7 36.6 15.8 L-Glutaminase 86.4 43.1 54.6 93.3 124 164 107 Amidase 45.8 17.0 17.5 31.1 23.7 45.1 14.4 Urease 5.55 2.64 4.16 6.40 5.75 7.3 6.2 L-Aspartase 3.76 1.28 1.04 1.97 2.11 2.8 1.0 " Except for arylamidase activity, which is expressed in mg p-naplithylamine produced kg 1 soil h\ all other enzyme activities are expressed in mg NH/-N produced kg1 soil h \ b SoiliO.Ol M CaCl2 (ratio= 1:2.5). "Acosta-Martinez and Tabatabai (2000). Table 4 Simple correlation coefficients for the relationships between enzyme b activities" and Cmi(, at the research sites indicated NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm r Arylamidase n.s. -0.77** 0.52* 0.75*** L-Asparaginase 0.71*** 0.75* 0.48* 0.84*** L-Glutaminase 0.44* n.s. 0.55* 0.85*** Amidase 0.72*** -0.75* 0.59** 0.69** Urease 0.70*** -0.75* 0.51* 0.64** L-Aspartase 0.73*** -0.82** n.s. 0.86*** 8 Except for arylamidase activity, which is expressed in mg p naphthylamine produced kg'1 soil hall other enzyme activities are expressed in mg NH/-N produced kg 1 soil h '.For the asterisks, see footnote of Table 2. b Cmic = Microbial biomass C, " n.s. = Not significant. Table 5 Simple correlation coefficients for the relationships between enzyme activitiesa and N.,.;..1' at the resaerch sites indicated NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm r Arylamidase n.s. 0.92*** 0.73*** 0.64** L-Asparaginase 0.54** 0.83** 0.70** 0.65** L-Glutaminase n.s. 0.83** 0.72*** 0.66** Amidase 0.57** 0.85** 0.88*** 0.76*** Urease 0.45* 0.85** 0.61** 0.75*** L-Aspartase 0.57** 0.80** 0.70** 0.70** " Except for arylamidase activity, which is expressed in mg p naphthylamine produced kg1 soil h all other enzyme activities are expressed in mg NH/-N produced kg'1 soil h'1, b Nmio = Microbial biomass N, "n.s. = Not significant. 135 Table 6 Specific activity of arylamidase and amidohydrolases as affected by liming at the SERC and SWRC sites Lime treatment Specific activity Type Rate Arl" Asp g Glu .Amid Ure Aspt 1 + 1 kg ECCE ha ...g of NH4 -Nkg- Corgh1.. SERC site 0 0.78b 3.36 1.83 0.38 0.39 0.03 Fine calcite 560 0.96 4.22 1.95 0.46 0.43 0.04 Dolomite 1120 2.06 4.02 2.49 0.48 0.93 0.08 Calcite 1120 3.13 6.08 3.00 0.55 1.49 0.12 Dolomite 2240 3.95 5.80 3.37 0.59 1.86 0.15 Calcite 2240 1.88 4.11 2.27 0.47 0.80 0.07 Dolomite 4480 1.87 4.85 2.42 0.43 0.93 0.08 Dolomite 6720 2.10 4.74 2.42 0.51 0.91 0.08 LSD (P < 0.05) 0.42 1.59 0.13 0.04 0.07 0.01 SWRC site 0 1.43b 0.72 4.16 2.48 0.71 0.05 Dolomite 2167 1.70 1.24 5.61 2.89 0.82 0.09 Dolomite 6373 1.95 1.54 6.43 3.33 0.85 0.13 Dolomite 19118 2.18 1.93 7.91 3.84 0.92 0.16 Dolomite 57355 2.10 1.93 7.70 3.70 0.86 0.17 LSD (P < 0.05) 0.19 0.26 1.03 0.24 0.11 0.03 a 1 1 Specific activity for arylamidase is g of P-naphthylamine kg" Corg h" . b Values are averages of soil samples from three and two replicated field plots at the SERC and SWRC sites, respectively. 136 Table 7 Specific activity of arylamidase and amidohydrolases as affected by liming (dolomite) and tillage systems at the NWRC site. Specific activity Lime treatment Arl* Aspr Glu Amid Ure Asyt kg ECCE ha1 g of NH,*-N kg1 Coreh1 No-till (0-5 cm) 0 1.21b 0.62 3.01 2.12 0.63 0.00 560 1.71 1.24 4.74 2.53 0.64 0.01 1120 1.65 1.18 4.37 2.75 0.63 0.01 2240 1.73 1.17 4.77 2.62 0.65 0.01 4480 2.12 1.39 6.07 2.96 0.75 0.01 6720 2.39 1.65 7.44 2.29 0.74 0.01 LSD (P < 0.05) 0.52 0.28 1.17 1.59 0.31 0.00 No-till (0 •15 cm) 0 0.98 0.57 2.81 1.74 0.59 0.00 560 1.46 0.78 3.66 2.02 0.64 0.01 1120 1.22 0.73 3.41 1.84 0.63 0.01 2240 1.57 0.98 4.54 1.03 0.64 0.01 4480 1.55 1.39 6.42 2.61 0.76 0.01 6720 2.06 1.61 7.34 3.11 0.82 0.01 LSD (P < 0.05) 0.39 0.15 0.56 0.34 0.19 0.01 Ridge-till 0 0.85 0.50 2.25 1.51 0.46 0.04 560 0.89 0.51 2.42 1.77 0.48 0.04 1120 0.95 0.74 2.66 1.54 0.49 0.06 2240 1.01 0.81 2.82 1.74 0.53 0.07 4480 1.19 1.15 2.84 2.18 0.56 0.09 6720 1.70 1.45 2.87 2.45 0.62 0.11 LSD (P < 0.05) 0.22 0.19 0.74 0.24 0.14 0.02 Chisel plow 0 1.00 0.53 2.65 1.74 0.56 0.05 560 1.02 0.61 2.83 1.97 0.54 0.05 1120 1.14 0.75 4.05 2.17 0.60 0.07 2240 1.36 0.94 4.68 2.40 0.62 0.08 4480 1.51 1.22 5.65 2.62 0.64 0.10 6720 1.97 1.46 6.95 2.86 0.64 0.12 LSD (P < 0.05) 0.32 0.18 0.66 0.35 0.18 0.02 1 l * Specific activity for arylamidase is g of (3-naphthylamine kg' Corg h" . b Values are averages of soil samples from three replicated field plots. 137 80 - Arylamidase SERC site . Arylamidase SWRC site r = 0.42* r = 0.99*** 60 40 ; 20 0 80 . Arylamidase NWRC site • Arylamidase NWRC site r = 0.84*** No-till r = 0.86*** No-till 0-5 cm 0-15 cm 60 40 O 20 : 0 80 . Arylamidase NWRC site . Arylamidase NWRC site r = 0.89*** Ridge-till r = 0.90*** Chisel plow 60 O 40 : y : y 20 0 rj_i 1 1 t 1 1 :— 4.0 4.5 5.0 5.5 6.0 6.5 7.0 4.0 4.5 5.0 5.5 6.0 6.5 7.0 Soil pH Fig. 1. Effect of soil pH on arylamidase activity at the research sites indicated. 138 80 - ^-Asparaginase SERC site L-Asparaginase SWRC site r = 0.86*** r = 0.97*** 60 40 20 0 250 • L-Glutaminase L-Glutaminase 200 . r = 0.62** r = 0.97*** 150 100 50 0 125 Amidase • Amidase r = 0.98*** 100 . r = 0.87*** 75 CXzS0 qffSb 50 4 (DO O/O 2 qCdD fly 0 4.5 5.0 5.5 6.0 6.5 7.0 4.5 5.0 5.5 6.0 6.5 7.0 Soil pH Fig. 2. Effect of soil pH on activities of amidohydrolases at the research sites indicated. 139 CHAPTER 5 RESPONSES OF PHOSPHATASES AND ARYLSULFATASE IN SOILS TO LIMING AND TILLAGE SYSTEMS A paper submitted to Journal of Plant Nutrition and Soil Science Mine Ekenler and M. A. Tabatabai Key words: acid phosphatase / alkaline phosphatase / phosphodiesterase / Arylsulfatase / microbial bio mass Summary - Zusammenfassung This study was carried out to investigate the long-term influence of lime application and tillage systems (no-till, ridge-till and chisel plow) on the activities of phosphatases and arylsulfatase in soils at four research sites in Iowa, USA. The activities of the following enzymes were studied: acid and alkaline phosphatases, phosphodiesterase, and arylsulfatase at their optimal pH values. With the exception of acid phosphatase, which was significantly (P < 0.001) but negatively correlated with soil pH (r ranged from -0.65** to - 0.98***), the activities of other enzymes were significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.65** to 0.99*** for alkaline phosphatase, from 0.79*** to 0.97*** for phosphodiesterase, and from 0.66*** to 0.97*** for arylsulfatase. The A activity/ A pH values were calculated to determine the sensitivity of each 140 enzyme to changes in soil pH. Acid phosphatase was the most sensitive and arylsulfatase the least sensitive to changes in soil pH. Activities of the enzymes were greater in the 0-5 cm depth samples than those in 0-15 cm samples under no-till treatment. Enzyme activities were mostly significantly (P < 0.001) and positively correlated with microbial biomass C (Cmic), with r values ranging from 0.28 (not significant) to 0.83*** and with microbial biomass N (Nmic), with r values ranging from 0.31 (not significant) to 0.94***. Liming and tillage systems significantly affected the activities of some enzymes but not others, as was evident from the specific activity values (g of 1 l p-nitrophenol released kg* Corg h ). Reaktionen von Phosphatase!! und Arylsulfatasen in Boden auf Kalkung und differenzierte Bodenbearbeitung In vier langjàhrigen Feldversuchen in Iowa, USA, wurde der Einfluss von Kalkung und differenzierter Bodenbearbeitung (Direktsaatverfahren, reduzierte Bearbeitung und Grubberverfahren) auf die Aktivitâten von Phosphatasen und Arylsulfatase in Bôden untersucht. Die Aktivitâten von saurer und alkalischer Phosphatase, Phosphodiesterase und Arylsulfatase wurden unter dem optimalen pH-Wert fiir das jeweilige Enzym bestimmt. Mit Ausnahme der sauren Phosphataseaktivitàt, welche signifikant negativ (f<0.001) mit dem pH-Wert des Bodens korreliert war (r = -0.65** bis — 0.98***), waren die Aktivitâten der anderen Enzyme signifikant (P<0.001) 141 positiv mit dem Boden-pH korreliert. Dabei variierten die Korrelationskoeffîzienten zwischen r = 0.65** und 0.99*** fur die alkalische Phosphatase, zwischen r = 0.79*** und 0.97*** fur die Phosphodiesterase und zwischen r = 0.66*** und 0.97*** fur die Arylsulfatase. Die Verhâltnisse von A Aktivitât / A pH-Wert wurden berechnet, um die Empfindlichkeit der untersuchten Enzyme gegeniiber pH-Wertveranderungen im Boden festzustellen. Dabei erwies sich die saure Phosphatase als das emfindlichste und die Arylsulfatase als das am wenigsten emfindlichste Enzym. In der Direktsaatvariante waren die Enzymaktivitâten in 0-5 cm Bodentiefe hôher als in 0-15 cm Tiefe. Die Enzymaktivitâten waren signifikant positiv mit dem mikrobiell gebundenen C (Cmik) und N (Nmik) korreliert. Die Korrelationskoeffîzienten variierten dabei zwischen r = 0.28 (nicht signifikant) und 0.83*** fur Cmik und zwischen r = 0.31 (nicht signifikant) und 0.94*** fur Nmik. Die spezifischen Enzymaktivitâten (g p-Nitrophenol kg1 Corg h l) zeigten, dass die Aktivitâten von einigen Enzymen signifikant von Kalkung und Bodenbearbeitungssystem abhângig waren. 1. Introduction There has been a need to develop sensitive indicators of soil health and quality that reflect the effects of land management on soil and long-term sustainability of terrestrial ecosystems. It is difficult to establish soil quality indicators with a single biological or chemical measurement, because soils are inherently variable and they have multiple functions. Since many physical 142 and chemical properties of soils will hardly change over a century (Tscherko and Kandeler, 1997), studies to establish standard measurements as soil quality indicators should focus on those components, such as microbial biomass and microbial processes, expected to change more rapidly ( biomass and soil enzymes in a short -period of time and long before there are measurable changes in soil organic matter (Powlson et al., 1987; Dick, 1994). Enzyme activities have been suggested as early indicators of changes in soil properties induced by liming and tillage (Carter, 1986; Powlson et al., 1987; Acosta-Martinez and Tabatabai, 2000), because of their rapid response to change in soil management practices, their relationship to soil biology, and ease of assay. This index would integrate physical, chemical and biological characteristics and could be used for long-term monitoring for both the identification of current changes in the soil and the prediction of future changes (.Kandeler et al., 1999). Soil management practices influence soil microorganisms and soil microbial processes through changes in the quantity and quality of plant residues entering the soil, their seasonal and spatial distribution, the ratio between above and belowground inputs, and through changes in nutrient inputs (Christensen, 1996). Lime application to soils most often causes a significant increase in pH and thus, a change in microbial biomass (Edmeades et al., 1981), microbial dynamic and diversity, and therefore, enzyme activities {Zelles et al., 1990; Acosta-Martinez and Tabatabai, 2000). Tillage 143 practices result in changes in soil physical and chemical properties and the composition, distribution, and the activities of soil microbial communities and enzymes (Deng and Tabatabai, 1997). Organic matter is more thoroughly distributed in soils under conventional cultivation compared with that in soils under reduced tillage plots where crop residues are concentrated on the soil surface. Consequently, microbial biomass and microbial processes in surface soils under no-till system are mostly greater than in soil treated with tillage systems. Enzymatic activity in soil is important for catalyzing a complex web of chemical reactions necessary for life processes of microorganisms, decomposition of organic residues, cycling of nutrients, and formation of organic matter and soil structure {Dick, 1994). It is suggested that studying soil enzyme activities provides insight into biochemical processes in soils and is very sensitive as biological indexes (Frankenberger and Dick, 1983), and thus, it provides a useful tool to long-term monitor the changes in soil health and quality. Phosphatase is a general name used to describe a broad group of enzymes that catalyze the hydrolysis of both esters and anhydrides of phosphoric acid (Schmidt and Laskowski, 1961). Among the phosphatases, acid phosphatase (orthophosphoric monoester phosphohydrolase, EC 3.1.3.2) is the most studied in nature, whereas alkaline phosphatase (orthophosphoric monoester phosphohydrolase, EC 3.1.3.1) and phosphodiesterase (orthophosphodiester phosphohydrolase, EC 3.1.4.1) are the least studied. 144 Arylsulfatase (EC 3.1.6.1) is the enzyme that catalyzes the hydrolysis of organic-sulfate esters releasing inorganic sulfate. The reaction is as follow: + 2 R.O.SOa +H2O —• R.OH + H + S04 Thus, those enzymes play an important role in the process whereby organic phosphorus and sulfur are mineralized in soils and made available to plants. The response of the activities of some of the enzymes to liming and different tillage practices has mainly been investigated on a limited number of sites or enzymes (Dick et al., 1988; Curci et al., 1997; Bergstrom et al., 1998; Bandick and Dick, 1999; Acosta-Martinez and Tabatabai, 2000), or on soil samples of short-term management sites (Bardgett and Leemans, 1995). Less information is available, however, on comparison of the effects of liming and tillage systems on phosphatases and arylsulfatase, and on CnUc and Nmic in soils of long-term management sites. Before a soil property or process can be used as soil quality index, systematic studies across many long-term soil management sites are needed. Thus, the first objective of this study was to assess the responses of phosphatases and arylsulfatase to liming and tillage systems at four long-term management sites in Iowa. To be useful as a soil quality index, the activity of the enzyme should be responsive to soil management practices or restoration efforts in a relatively short time (Dick, 1994). Thus, the second objective was to determine the sensitivity of those enzymes to limed agricultural soils under different tillage systems. 145 Although the effects of liming and tillage practices on enzyme activities in soils were studied (Deng and Tabatabai, 1997; Acosta-Martinez and Tabatabai, 2000), little is known about relationship between soil microbial biomass and enzyme activities under those management systems. Therefore, the third objective was to determine the relationship between Cmic and Nmic and the activities of the phosphatases and arylsulfatase in soils. 2. Materials and methods 2.1. Experimental design and soil sampling The study was carried out on soils from four sites in Iowa: Southeast Research Center (SERC site) at Crawfordsville, Southwest Research Center (SWRC site) at Atlantic, Northwest Research Center (NWRC site) at Sutherland, and Northeast Research Center (NERC site) at Nashua. Description of the sites, experimental designs and soil sampling were reported by Ekenler and Tabatabai (2002). 2.2. Soil characterization The pH values were determined on < 2-mm mesh air-dried samples by using a combination glass electrode (soihO.Ol M CaClz ratio = 1:2.5), organic C and total N by a LECO CHN 600 determinator (St. Joseph, MI), inorganic N by steam distillation (Keeney and Nelson, 1982), and organic N by the difference between total N and inorganic N. Microbial biomass C (Cmic) and N (Nmic) were determined by the methods described by Vance et al. (1987) and 146 Horwath and Paul (1994). All results are expressed on a moisture-free basis. Moisture content was determined from loss in weight after drying at 105 °C for 48 h. 2.3. Enzyme assays The activities of the enzymes in soils of the first three sites were assayed on < 2 mm field moist samples. The activities of the enzymes in samples of the fourth sites were performed on air-dried samples and reported by Acosta- Martinez and Tabatabai (2000). They are summarized here for comparison. The enzyme assays (Tab. 1) were performed at their optimal pH values as described by Tabatabai (1994). 2.4. Specific activities From the enzyme activity values and organic C, the specific activity values 1 1 were calculated as g of p-nitrophenol released kg* Corg h* . 2.5. Statistical analyses Statistical analyses, including regression analyses and means separation by the least significant difference were performed according to general linear model procedure of the SAS system (SAS Institute, 1996). For all data points 147 reported in the figures, the differences between the laboratory duplicates were smaller than the point size. 3. Results 3.1 Effects of liming and tillage systems on phosphatases With the exception of acid phosphatase, which was significantly (P < 0.001) but negatively correlated with soil pH (r ranged from -0.65** to - 0.98***), the activities of the other phosphatases were significantly (P < 0.001) and positively correlated with soil pH, with r values ranged from 0.65** to 0.99*** for alkaline phosphatase and 0.79*** to 0.97*** for phosphodiesterase (Tab. 2). To determine the sensitivity of the enzymes to change in soil pH, A activity/ A pH values for the individual enzymes were calculated (Tab. 3). The values ranged from -35 to -139 for acid phosphatase, from 25.6 to 143 for alkaline phosphatase, and from 12.1 to 76.6 for phosphodiesterase. Results showed that acid phosphatase was the most sensitive enzyme and phosphodiesterase was the least sensitive to changes in soil pH, and in soils under ridge-till and chisel plow. In general, acid phosphatase activity was greatest among the soil phosphatases, followed by alkaline phosphatase and phosphodiesterase. For example, expressed in mg p-nitrophenol kg-1 soil h1, the acid phosphatase activity values ranged from 386 in the control plots to 323 in the plots with 148 the highest lime application rate. The corresponding values for phosphodiesterase activity ranged from 25 in the control plots to 47 in the plots with the highest lime application rate at the SERC site (Fig 1). Similarly, phosphatase activities were greater in soils at 0-5 cm depth samples than in the 0-15 cm samples of soil under the no-till system. For example, alkaline phosphatase activity was 229 mg p-nitrophenol kg1 soil h1 at highest lime application rate in the 0-5 cm depth samples, whereas it was only 170 in the 0-15 cm sample at the NWRC site. There was no consistent trend for the effects of studied tillage systems on the activity values of the phosphatases. 3.2 Effects of liming and tillage systems on arylsulfatase Arylsulfatase activity was significantly (P < 0.001) and positively correlated with soil pH, with r values ranged from 0.66*** to 0.97*** (Tab. 2). As for the phosphatases, the A activity/ A pH values were calculated, and those values ranged from 11.2 to 70.5 (Tab. 3). Results indicated that arylsulfatase was less sensitive than the phosphatases to changes in soil pH and to tillage systems. The activity of this enzyme was greater in the 0-5 cm depth samples than those of the 0-15 cm samples under no-till system. Expressed in mg p- nitrophenol kg*1 soil lrL, the values were 62 and 49 in the control plots and 123 and 93 in samples of highest lime application rate of the 0-5 cm and 0-15 149 cm depths, respectively. The activity values were also greater in soil under chisel plow than the other tillage system studied. 3.3 Activities of phosphatases in relation to Cmic and Nmic Phosphatase activities were significantly (P < 0.001) and positively, except for the SWRC site, correlated with Cmic, with r values ranging from 0.67*** for phosphodiesterase to 0.73*** for acid and alkaline phosphatases at the SERC site, from 0.77** for acid phosphatase to 0.83** for alkaline phosphatase at the SWRC site, from 0.33 (not significant) for alkaline phosphatase to 0.79*** for phosphodiesterase at the NWRC site in soils of the 0-5 cm depth under no-till, and from 0.53* for alkaline phosphatase to — 0.75*** for acid phosphatase at the NWRC site in soils of the 0-15 cm depth under no-till (Figs 2 and 3). The activity values were also significantly (P < 0.001) and positively correlated with Nmic, with r values ranging from 0.54** for alkaline phosphatase to -0.70*** for acid phosphatase at the SERC site, from 0.85** for acid and alkaline phosphatases to 0.94*** for phosphodiesterase at the SWRC site, from 0.33* for alkaline phosphatase to 0.90*** for phosphodiesterase in 0-5 cm depth samples of the no-till plots at the NWRC site, and from 0.64** for alkaline phosphatase to —0.84*** for acid phosphatase in the 0-15 cm samples under no-till at the NWRC site (Figs 4 and 5). 150 3.4 Arylsulfatase activity in relation to Cmic and Nmic Except for the samples from the SWRC site, arylsulfatase activity of soils was significantly (P < 0.001) and positively correlated with Cmic. with r values of 0.66***, 0.28, 0.77***, and 0.75*** for the samples from the SERC, SWRC, NWRC no-till (0-5 cm), and NWRC no-till (0-15 cm), respectively (Figs 2 and 3). The activity values were also mostly significantly and positively correlated with Nmic, with r values of 0.31, 0.75*, 0.86***, and 0.76*** at SERC, SWRC, NWRC no-till (0-5 cm), and NWRC no-till (0-15 cm), respectively (Figs 4 and 5). The effects of the tillage systems studied on Cmic and Nmic values varied; the effect of no-till on those variable was consistent (i.e., Cmic and Nmic increased with increasing lime application), but the effects of ridge till and chisel plow on the those properties were not consistent, due to disturbance of the surface soil by tillage (Ekenler and Tabatabai, 2002). Therefore, the lack of relationships between Cmic or Nmic values and the activities of the enzymes presented here in soils under ridge till and chisel plow are not presented. 3.5 Specific activities Expressed in g p-nitrophenol released kg -1 Core h'L> the specific activity values were affected by soil pH, depending on the enzyme and the site (Tabs. 4 and 5). 151 4. Discussion The finding of significant and positive correlation, with the exception of acid phosphatase, between enzyme activities and soil pH confirmed the observations of other studies reporting that enzyme activities of soils are often increased with increasing soil pH (Bardgett and Leemans, 1995; Deng and Tabatabai, 1997; Acosta Martinez and Tabatabai, 2000; Klose and Tabatabai, 2002). It has been reported that the rate of synthesis, release, and stability of acid and alkaline phosphatases by soil microorganisms are dependent on soil pH (Eivazi and Tabatabai, 1977; Juma and Tabatabai, 1977; Tabatabai, 1994). The present study also demonstrated that acid phosphatase was predominant in the control plots and decreased with increasing pH by liming, and that liming increased the activities of the alkaline phosphatase and phosphodiesterase (Fig 1). This result was in agreement with the finding of Juma and Tabatabai (1978) showing that acid phosphatase is predominant in acid soils and that alkaline phosphatase is predominant in alkaline soils. Increase in the activity of alkaline phosphatase by liming may indicate the effect of lime treatment on the size of the soil microbial population, as this enzyme is not produced by plants; its activity is completely derived from microorganisms (Dick et al., 1983; Juma and Tabatabai, 1988 a, b, c). Recent work by Dick et al. (2000) showed that the ratio of acid phosphatase/ alkaline phosphatase in soils could be used as pH adjustment indicator. This ratio responded immediately to the change in 152 soil pH caused by calcium carbonate addition, suggesting possible enzyme induction. The results showed that acid phosphatase was the most sensitive enzyme and arylsulfatase was least sensitive to changes in soil pH (Tab. 3). Bergstrom et al. (1998) studied the activities of six enzymes, including phosphatase and arylsulfatase, under different management practices and showed that soil enzyme activities are sensitive to changes in soil quality resulting from soil management practices; however, the enzymes differ in their sensitivity to those management practices. Different response of acid and alkaline phosphatases to liming supported the previous finding that phosphatases are inducible enzymes and the intensity of their release by microorganisms and plants is determined by their requirement for orthophosphate, which is strongly affected by soil pH (Skujins, 1976). Acid phosphatase activity, in general, was greatest followed by alkaline phosphatase, phosphodiesterase, and arylsulfatase. The relatively greater acid phosphatase activity values may be the result of greater acid phosphatase enzyme protein concentration in soils. Recent studies by Klose and Tabatabai (1999, 2002) on phosphomonoesterases and arylsulfatase enzyme protein concentrations in ten Iowa surface soils showed that the average acid phosphatase protein concentration was much greater (22.5 mg kg"1 soil) than that of alkaline phosphatase (2.1 mg kg1 soil) and arylsulfatase (7.0 mg kg"1 soil). Their result may also suggest that alkaline phosphatase is much more efficient in the hydrolysis of its substrate than is arylsulfatase, 153 because the activity of alkaline phosphatase is greater than that of arylsulfatase. Even though the concentrations of soil organic C and N were constant at each site (Ekenler and Tabatabai, 2002), with the exception of acid phosphatase, lime application significantly (P<0.001) improved the activities of the enzymes (Fig. 1). This result suggested that not only the quantity of soil organic matter, but also the chemical nature of organic matter has an impact on soil microbial activities and processes. This conclusion is supported by the effect of liming and tillage systems on the specific activity values obtained for the enzymes studied (Tabs 4 and 5). Doran et al (1998) reported that in addition to long-term changes in quantity of soil organic matter and related soil physical characteristics, the quality of soil organic matter varies with management practices as well. The greater enzyme activities in the 0-5 cm depth samples than those in the 0-15 cm samples under no-till systems is probably resulted from the increased organic C and total N; thus, induced microbial activities in surface soils (0-5 cm) under no-till system. Soils subjected to reduced and minimum tillage accumulate crop residues and organic C, which are substrates for soil organisms near the soil surface. Consequently, soil microbial biomass and various soil microbial processes tend to increase in surface soils. This finding confirmed the observations of other authors reporting that the types of reduced tillage enrich the soil surface with organic C and N reserves, microbial biomass, microbial activity, and soil enzymes (Deng and Tabatabai, 154 1997; Doran et al., 1998; Kandeler et al., 1999). The study by Deng and Tabatabai (1997) showed that the activities of phosphatases and arylsulfatase were strongly correlated with organic C in the 40 soil samples studied and the activities of these enzymes decreased markedly with increasing soil depth and this decreased was associated with a decrease in organic C content. The finding of significant and positive correlation between enzyme activities and Cmic and Nmic (Figs. 2-5) is in agreement with the study of Klose and Tabatabai (2002) showing that alkaline phosphatase is strongly correlated with Cmic of the soil. The strong positive relationship between alkaline phosphatase or phosphodiesterase and Cmic and Nmic is indicative of the importance of microorganisms as sources of soil enzymes. Plants have also been considered a source of extracellular enzymes in soils. Juma and Tabatabai (1988b) showed that sterile corn and soybean roots contain acid phosphatase, but not alkaline phosphatase activity. Even thought both microorganisms and plants release enzymes into the soil environment, microorganisms are the logical choice for supplying most of the soil enzyme activity, because of their large biomass, high metabolic activity and short lifetimes, which allow them to produce and release relatively larger amounts of extracellular enzymes than can plants or animals Speir and Ross (1978). In soils taken from the SWRC site, Cmic decreased by liming, while alkaline phosphatase, phosphodiesterase, and arylsulfatase activities increased. This site received considerably greater rates of lime application 155 compared to other sites, with the highest rate being 57355 kg ECCE ha*1. Thus, pH values were also greatly increased in those plots. It is commonly accepted that acid or slightly acid soil pH favors the growth of fungi, while neutral or alkaline pH favor the growth of bacterial biomass (Baath et al., 1992). Fungal biomass contains higher C concentration than bacterial biomass (Campbell et al., 1991) and thus, increasing soil pH decreased the amount of fungi in the environment; consequently, decreased Cmic It has been suggested that liming significantly decrease the fungal biomass, but the total microbial activity increases (Zelles et al., 1987, 1990). The increase in enzyme activities with decreasing Cmic at the SWRC site may also be because soil pH improved the stability of the previously released enzyme in soils or liming may have increased the quality of C sources available to microbes, so that a great proportion of microbes remained active, while fungal population decreased, consequently, leading to decrease in the total Cmic 5. Conclusion Liming significantly decreased the activity of acid phosphatase, while significantly increased the activities of alkaline phosphatase, phosphodiesterase, and arylsulfatase. Increase in soil pH by lime application seems to have stimulated the microbial population and activity, resulting in an increased alkaline phosphatase, phosphodiesterase, and arylsulfatase activities; thus, affecting the P and S cycling in soils. The sensitivity of enzyme activities to change in soil pH varied considerably; however, the order 156 of the sensitivity remained similar among the sites. Among the enzyme studied, arylsulfatase was the least sensitive enzyme, while acid phosphatase was the most sensitive to change in soil pH. Therefore, this enzyme may provide a reliable long-term monitoring tool as early indicators of changes in soil properties induced by liming. Enzyme activities were greater in the 0-5 cm depth samples than in the 0-15 cm samples under the no-till system. However, no regular trend was found in sensitivity of enzyme activities to studied tillage systems. A highly significant relationship was found between soil enzyme activities and microbial biomass. This study demonstrated that soil microbial processes, such as phosphatases and arylsulfatase activities involved in P and S cycling, may provide reliable tools with which to estimate early changes in the dynamics and distribution of microbial processes in soils. Acid phosphatase may be the best enzyme to be used as an indicator of the changes in soil health and quality resulted from agricultural practices because it is the most sensitive to soil disturbance, while arylsulfatase was the least sensitive enzyme. Acknowledgements This work was supported in part by the Biotechnology By-products Consortium of Iowa. References Acosta-Martinez, V., and M. A. Tabatabai (2000): Enzyme activities in a limed agricultural soil. Biol. Fertil. Soils. 31, 85 -91. 157 Baath, E., A. Frostegard, and H. Fritze (1992): Soil bacterial biomass, activity, phsopholipid fatty acid pattern, and pH tolerance in an area polluted with alkaline dust deposition. Appl. Environ. Microbiol. 58, 4026 -4031. Bandick, A.K., and R. P. Dick (1999): Field management effects on soil enzyme activities. Soil Biol. Biochem. 31, 1471 - 1479. Bardgett, R.D., and D. K. Leemans (1995): The short-term effects of cessation of fertilizer applications, liming, and grazing on microbial biomass and activity in a reseeded upland grassland soil. Biol. Fertil. Soils. 19, 148 - 154. Bergstrom, D.W., C. M. Monreal, and D. J. King (1998): Sensitivity of soil enzyme activities to conservation practices. Soil Sci. Soc. Am. J. 62, 1286 -1295. Campbell, C.A., G. P. LaFond, A. J. Leysohon, R. P. Zentner, and H. H. Janzen (1991): Effect of cropping practices on the initial rate of N mineralization in a thin Black Chernozem. Can. J. Soil Sci. 71, 43 - 45. Carter, M.R. (1986): Microbial biomass as an index for tillage-induced changes in soil biological properties. Soil Till. Res. 7, 29 - 40. Christensen, B.T. (1996): Matching measurable soil organic matter fractions with conceptual pools in simulation models of carbon turnover: revision of model structure. In D. S. Powlson, P. Smith, P., and J. U. 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Tabatabai (1988c): Comparison of kinetic and thermodynamic parameters of phosphomonoesterases of soils and of corn and soybean roots. Soil Biol. Biochem. 20, 533 - 539. Keeney, D. R., and D. W. Nelson (1982): Nitrogen-Inorganic forms. In A. L. Page, R. H. Miller, and D. R. Keeney (eds): Methods of Soil Analysis, Part 2, 2nd edn, Agronomy Monograph. No. 9, American Society of Agronomy, Madison, WI, pp. 642 -698. Kandeler, E., D. Tscherko, and H. Spiegel (1999): Long-term monitoring of microbial biomass, N mineralization and enzyme activities of Chernozem under different tillage management. Biol. Fertil. Soils. 28, 343 - 351. Klose, S., and M. A. Tabatabai (1999): Arylsulfatase activity of microbial biomass in soils. Soil Sci. Soc. Am. J. 63, 569 - 574. Klose, S., and M. A. Tabatabai (2002): Response of phosphomonoesterases in soils to chloroform fumigation. J. Plant Nutr. Soil Sci. 165, 429 - 434. Powlson, D.S., P Brookes, and B. T. Christensen (1987): Measurement of soil microbial biomass provides an early indication of changes in total soil organic matter due to straw incorporation. Soil Biol. Biochem. 19, 59 - 164. SAS Institute (1996): SAS Procedure Guide. Version 6.12 ed. SAS Institute Inc., Cary, NC. Schmidt, G.,and M. Laskowski, Sr. (1961): Phosphate ester cleavage (survey). In: Boyer, P.D. et al. (Eds.), The Enzymes, 2nd edn. Academic Press, New York, pp. 3 - 35. 161 Skujins, J. (1976): Enzymes in soil. In A. D. McLaren, and G. H. Peterson (eds.): Soil Biochemistry, vol 1. Dekker, New York, pp. 371 - 414. Speir, T.W., and D. J. Ross (1978): Soil phosphatase and sulphatase. In R. G. Burns (ed.): Soil Enzymes. Academic Press, London, pp. 197 - 250. Tabatabai, M.A. (1994): Soil enzymes. In R. W. Weaver, J. S. Angel, and P. S. Bottomley (eds.): Methods of Soil Analysis, Part,2, Book Series: 5, Soil Science Society of America, Madison, WI, pp. 775 - 833. Tscherko, ZZ, and E. Kandeler (1997): Biomonitoring of soils-denitrification enzyme activity and soil microbial processes as indicators for environmental stress. In K. H. Becker, and P. Wieser (eds): Proceedings of the 7th International Workshop on Nitrous Oxide Emissions, Cologne, April 21-23, 1997. Physikalische Chemie, Bericht 41, Wuppertal, pp. 373 -- 381. Vance, E. D., P. C. Brookes, and D. S. Jenkinson (1987): An extraction method for measuring soil microbial biomass C. Soil Biol. Biochem. 19, 703 - 707. Zelles, L., I. Scheunert, and K. Kreutzer (1987): Bioactivity in limed soil of spruce forest. Biol. Fertil. Soils. 3, 211 - 216. Zelles, L., I. Stepper, and A. Zsolnay (1990): The effect of liming on microbial activity in spruce (Picea abies L.) forests. Biol. Fertil. Soils. 9, 78 - 82. Table 1; The methods used for assay of enzyme activities in soils. Class/EC Recommended Reaction Assay conditions Assay conditions number name8 Substrate1" Optimum pH Phosnhatases 3,1.3,1 Alkaline Phosphatase RNa,PO< + H20 -> R OM + Na2HP04 p-Nitrophenyl phosphate (10 mM) 11,0 3,1,3,2 Acid Phosphatase RNa2P04 + H20 -» R OH + Na2HP04 p-Nitrophenyl phosphate (10 mM) 6.5 3,1,4,1 Phosphodiesterase RgNaPOj + HoO -> R-OH + RNaHPO., Bis -p -Nitrophenyl phosphate (10 mM) 8.0 Sulfatase + 2 3,1,6,1 Arylsulfatase ROSOa' + H20 -» R-OH + H + S04 p -Nitrophenyl sulfate (10 mM ) 5.8 * For the methods used, see Tabatabai (1994). b Figures in parentheses are the substrate concentrations under assay conditions. Table 2: Simple correlation coefficients for the relationships between enzyme activities3 and soil pHb at the sites indicated. NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm Ridge-till Chisel NERCc Phosphatases Alkaline Phosphatase 0.88*** 0.99*** 0.65** 0.78*** 0.82*** 0.93*** 0.89*** 0.95***" Acid Phosphatase -0.91*** -0.98*** -0.81*** -0.86*** -0.65** 0.92*** -0.69*** Phosphodiesterase 0.88*** 0.97*** 0.92*** 0.79*** 0.85*** 0.94*** 0.89***d 0.91***" Sulfatase Arylsulfatase 0.72*** 0.97*** 0.89*** 0.89*** 0.88*** 0.93*** 0.66*** 8 All activities are expressed in mgp -nitrophenol released kg'1 soil h'\ b Soil:0,01 M CaCl2 (ratio= 1:2.5), cAcosta-Martinez and Tabatabai (2000), d Linear regression for soils with pH>6.0, ** and *** indicate significance at P< 0.0land 0.001, respectively. 0 Second linear regression values. Table 3: A Enzyme activity8/ A soil pHb for the sites and tillage systems studied. NWRC No-till Enzymes SERC SWRC 0-5 cm 0-15 cm Ridge-till Chisel NERCc Phosphatases Alkaline Phosphatase 93 32 33 37 78 143 97d 26e Acid Phosphatase 137 -59 80 -77 101 139 -35 Phosphodiesterase 77 -28e 64 41 64 74 39d 12e Sulfatase Arylsulfatase 33 "28e 56 40 43 71 11 a All activities are expressed in mg p -nitrophenol released kg'1 soil h \ b Soil:0.01 M CaClg (ratio= 1:2.5). °Acosta-Martinez and Tabatabai (2000). d Linear regression for soils with pH>6,0, 0 Second linear regression values. 165 Table 4: Specific activities of phosphatases and arylsulfatase as affected by liming at the SERC and SWRC sites. Lime treatment Specific activity Type Rate Acid P Alk P Phosphodi Arylsul 1 1 1 kg ECCE ha' g of p -Nitrophenol kg' Corg h* ".... SERC site 0 12.3" 2.10 0.62 1.94 Fine calcite 560 10.7 3.06 1.52 1.83 Dolomite 1120 11.9 2.26 0.74 2.05 Calcite 1120 12.5 2.89 0.72 2.51 Dolomite 2240 13.2 3.31 1.68 2.64 Calcite 2240 12.7 3.31 1.42 2.05 Dolomite 4480 12.3 4.36 2.54 2.21 Dolomite 6720 11.8 5.11 2.83 2.28 LSD (P < 0.05) 1.35 0.29 0.25 0.70 SWRC site 0 11.8b 6.52 5.14 2.78 Dolomite 2167 9.11 7.47 7.29 3.75 Dolomite 6373 8.13 8.39 8.80 3.77 Dolomite 19118 5.37 9.14 8.57 3.44 Dolomite 57355 4.47 8.54 7.90 3.14 1.07 LSD (P < 0.05) 1.07 0.97 1.21 "Values are averages of soil samples from, three replicated field plots at SERC site. bValues are averages of soil samples from two replicated field plots at SWRC site. 166 Table 5: Specific activities of phosphatases and. arylsulfatase as affected by liming and tillage systems at the NWRC site. Specific activity Lime treatment AcidP Alk P Phosphodi Arylsul 1 1 1 kg ECCE ha' g® of ~p -Nitrophenolf to kg' Core h' — No-till (0-5 cm) 0 20.3" 6.99 2.47 1.27 560 16.5 7.89 3.86 2.80 1120 16.2 7.64 3.44 2.47 2240 14.8 7.98 4.31 2.80 4480 14.8 8.63 5.01 3.85 6720 14.2 8.95 6.31 4.76 LSD (P < 0.05) 3.27 1.79 1.80 1.35 No-till (0-15 cm) 0 18.05 5.41 2.47 1.95 560 17.05 5.87 2.39 2.32 1120 16.72 5.54 2.58 2.18 2240 16.59 5.85 3.10 2.60 4480 15.57 6.01 3.67 3.35 6720 14.52 6.92 4.27 3.76 LSD (P < 0.05) 2.16 1.25 0.91 0.64 Ridge-till 0 15.3 5.10 3.75 2.01 560 15.8 5.66 4.33 2.38 1120 13.8 6.22 4.83 2.68 2240 14.0 6.54 4.92 2.79 4480 12.6 6.43 4.92 2.80 6720 12.2 7.97 6.11 3.58 LSD (P <0.05) 2.22 1.32 1.01 0.61 Chisel plow 0 15.03 5.20 3.95 2.15 560 13.83 5.45 4.06 2.23 1120 13.11 5.84 4.35 2.46 2240 12.82 5.56 4.21 2.25 4480 11.69 7.08 5.50 3.09 6720 10.27 8.43 6.52 3.72 LSD (P < 0.05) 1.28 1.34 1.02 0.61 "Values are averages of soil samples from three replicated field plots. 167 —rA—— 350 Alkaline Phosphatase SERC site Alkaline Phosphatase SWRC site 300 Y = -390 + 93.3X ' Y = -25.5 + 32.3X 250 r = 0.88*** . r = 0.99*** 200 150 100 50 0 600 Acid Phosphatase Acid Phosphatase 500 Y = 1035 - 137X r= -0.91*** " Y = 490 - 58.9X r = -0.98*** 400 300 200 100 0 200 Phosphodiesterase Phosphodiesterase Y = -344 + 76.6X Y = -944 + 355X2 -28. IX r = 0.88*** ' r = 0.97*** 150 100 50 2S8 Arylsulfatase Arylsulfatase 200 Y = -113 + 33.3X - Y = -944 + 355X" -28.1X r = 0.72*** r = 0.97*** o 0 150 100 50 0 —fit 1 1 1 1 1— 4.5 5.0 5.5 6.0 6.5 7.0 4.5 5.0 5.5 6.0 6.5 7.0 Soil pH g. 1 of soil pH on activities of phosphatases or arylsulfatase at research y// 500 • Acid Phosphatase # Phosphodiesterase O Alkaline Phosphatase • Arylsulfatase 400 SERC site SWRC site % CO M 300 ii03 ti r = -0.73** r = 0.78** I 200 il r = 0.73*** Ô, r = 0.83** 100 .8 r = 0,67*** r = 0.77** ° D-DQO-O—9-^! r = 0.66*** r = 0.28'" 0 —y/~ vz- 0 125 150 175 200 225 250 275 300 0 100 150 200 250 300 Cmic (mg kg*1 soil) Fig. 2. Relationships between the activities of phosphatases or arylsulfatase and Cmic in soils from the SERC and SWRC sites. S Acid Phosphatase # Phosphodiesterase 600 O Alkaline Phosphatase • Arylsulfatase NWRC site NWRC site 500 - s • • (No till 0-5 cm) (No-till 0-5 cm) (0 %CO 400 i> r = -0,75*** <8 C r = -0,54* 300 1 r = 0.33 IIM % 200 r = 0.53* o, bo r = 0.79*** a r = 0.70*** 100 - r = 0.75*** r = 0,77*** 400 500 100 200 300 400 500 Cmic (mgkg^ soil) Fig. 3. Relationships between the activities of phosphatases or arylsulfatase and Cmic in soils of the 0-5 cm and 0-15 cm depths under no-till system at the NWRC site 500 • Acid Phosphatase • Phosphodiesterase O Alkaline Phosphatase Q Arylsulfatase SERC site SWRC site 400 300 r = -0.70*** 200 r = 0.85** O Oo r = 0.94*** O CD = 0.54** 100 r = -0,86** r = 0.64*** r = 0.75* r = 0.31"s 10 15 20 10 15 20 Nmic (mg kg'1 soil) Fig, 4. Relationships between the activities of phosphatases or arylsulfatase and Nmic for soil samples from the SERC and SWRC sites. • Acid Phosphatase • Phosphodiesterase 600 O Alkaline Phosphatase • Arylsulfatase NWRC site NWRC site 500 • » • (No-till 0-5 cm) (No-till 0-15 cm) 400 r = -0,75*** r = -0.84*** 300 r = 0.58** 200 r = 0,90*** r = 0.64** r = 0.86*** r = 0.69** 100 r = 0,76*** 10 15 20 25 30 0 10 15 20 25 30 Nmic (mg kg'1 soil) Fig. 5. Relationships between the activities of phosphatases and arylsulfatase and Nmic in soils of the 0-5 cm and 0-15 cm depths under no-till system at the NWRC site. 172 CHAPTER 6 P-GLUCOSAMINIDASE ACTIVITY OF SOILS: EFFECT OF CROPPING SYSTEMS AND ITS RELATIONSHIP TO NITROGEN MINERALIZATION A paper in press in Biology and Fertility of Soils Mine Ekenler and M. A. Tabatabai Abstract The enzyme P-glucosaminidase (EC 3.2.1.30) is involved in C and N cycling in soils. The effects of crop rotations and N fertilization on p- glucosaminidase activity and its relationship to N mineralization were studied in soils of two long-term field experiments involving different cropping systems at the Northeast Research Center (NERC) and Clarion- Webster Research Center (CWRC) in Iowa that were initiated in 1978 and 1954, respectively. Surface soil samples (0 -15 cm) were taken in 1996 and 1997 in corn (Zea mays L.), soybean (Glycine max (L) Merr.), oats (Avena sativa LJ, or meadow (alfalfa) (Medicago sativa L.) plots that received 0 or 180 kg N ha i before com and an annual application of 20 kg P and 56 kg K ha % The P-glucosaminidase activity in the soils was assayed at optimal pH (acetate buffer, pH 5.5); microbial biomass C (Cmic) and N (Nmic) were determined by chloroform-fumigation methods; N mineralization was studied 173 in leaching columns under aerobic conditions at 30°C for 24 weeks. The activities of P-glucosaminidase were significantly affected by crop rotation (p < 0.001) and N fertilization (p ranging from 0.05 to 0.001). Generally, the highest enzyme activity was obtained in soils under 4-year corn-oats-meadow rotations taken under meadow, and the lowest under continuous mono-cropping systems. The activity of this enzyme was significantly correlated with Corg (r ranging from 0.42** to 0.76***), Norg (ranged from not significant at one site one year to r = 0.76***), Cmic (r ranging from 0.44** to 0.71***), and Nmic (r ranging from 0.33* to 0.76***) in soils, and with cumulative N mineralized (r > 0.84*** and r > 0.79*** at the NERC and CWRC sites, respectively). The results suggest that p- glucosaminidase plays a major role in N mineralization in soils and is affected by cropping systems; i.e., ecosystem function and health. Keywords: Soil enzymes, N mineralization, Cropping systems, Cmic, Nmic Introduction iV-Acetyl-p-D-glucosaminidase (NAGase, EC 3.2.1.30) is the enzyme that catalyzes the hydrolysis AT-acetyl-P-D-glucosamine residue from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also classified as P-hexosaminidase (EC 3.21.52) by the International Union of Biochemistry because it cleaves the amino sugar AT-acetyl-P-D-galactosamine (Webb 1984). 174 Some of the amino sugars in soils may exist in the form of alkali-insoluble polysaccharide referred to as chitin. This substance, which consists of NAG residues in (3-1,4 linkages, is a major structural component in insects and comprises the cell wall, structural membranes and skeletal component of fungal mycelia, where it plays a structural role analogous to the cellulose of higher plants (Stevenson 1994). It is considered one of the most abundant biopolymers in nature and an important pool of organic C and N in soils (Stryer 1988; Wood et al. 1994). Because this fraction (chitin) of soil organic matter contains recalcitrant C and N, it is most likely that the enzyme NAGase catalyzes the hydrolysis of the amino sugars in soils, which constitute 5-10% of the organic N in the surface of most soils (Stevenson 1994). This hydrolysis is considered to be important in C and N cycling in soils because it participates in the processes whereby chitin is converted to amino sugars, which is one of the major sources of mineralizable N in soils. NAGase is one of the three chitinases (N-Acetyl-P-D-glucosaminidase, chitobiosidase and endochitinase) that degrade chitin (Tronsmo and Harman 1993). The activity of this enzyme has been detected in microorganisms, human tissues, insects, plant, and soils (Neufeld 1989; Trudel and Asselin 1989; Martens et al. 1992; Sinsabaugh et al. 1993). Recently Parham and Deng (2000) reported on its quantification and characterization in soils, and developed a method for its assay. Work by Smucker and Kim (1987) showed that chitin is an inducer of chitinase activity and it is a major source of organic N in soils. Not much 175 work is available about the role of this enzyme in decomposition of crop residues in soils, but work by Sinsabaugh et al. (1993) showed that (3- glucosaminidase is one of the enzymes required for chitin degradation; its activity was negatively correlated with N immobilization and decomposition rate for birch sticks decomposing at eight contrasting sites. Although the impact of long-term crop rotation and N fertilization practices on soil physical and chemical properties is well documented in terms of soil density, water infiltration, pH values, organic C, N and S transformations, and enzyme activities (Rasmussen et al. 1980; Odell et al. 1984; Havlin et al. 1990; Varvel 1994; Klose et al. 1999; Klose and Tabatabai 2000; Deng et al. 2000; Deng and Tabatabai 2000; Moore et al. 2000), the effects of these factors on the activity of NAGase and its relationship to N mineralization in soils are still unknown. Thus, information is needed to evaluate the contribution of the P-glucosaminidase activity to N cycling and ecological changes in soils. The objectives of this work were to: (i) study the impact of long-term cropping systems on P-glucosaminidase activity, (ii) assess the relationship between the activity of this enzyme and organic C, organic N, Cmic, and Nmic, and (iii) assess the relationship between the activity of this enzyme and N mineralization in soils. 176 Materials and methods Soils and their properties Soil samples were collected from two long-term cropping systems at the Northeast Research Center (NERC) in Nashua and the Clarion-Webster Research Center (CWRC) in Kanawha, Iowa. The NERC study was initiated in 1979 on Kenyon (fine-loamy, mixed, mesic superactive Typic Hapludoll) and Readlyn (fine-loamy, mixed, mesic superactive Aquic Hapludoll) loams, with a mean particle-size distribution of 32.0% sand, 45.6% silt and 22.4% clay; pH values of 6.9 (in water) and 6.4 (in 0.01 M CaClz); 21.2 g kg*1 organic C; and 1.4 g kg1 organic N. The study at the CWRC site was established in 1954 on a Webster clay loam (fine-loamy, mixed, mesic superactive Typic Endoaquolls), with a mean particle-size distribution of 21.9% sand, 44.9% silt and 33.2% clay; pH values of 6.2 (in water) and 5.6 (in 0.01 M CaCla); 33.7 g kg-1 organic C; and 2.2 g kg1 organic N. Surface soil samples (0 -15 cm) were taken in corn (Zea mays L.), soybean (Glycine max (L) Merr.), oats (Avena sativa L.J, or meadow [alfalfa (Medicago sativa L.) at the CWRC site, alfalfa alone or mixed with red clover (Trifolium pratense L.) at the NERC site] plots that received an annual application of 0 or 180 kg N ha-1 as an ammoniacal fertilizer before corn and an annual application of 20 kg P and 56 kg K ha*1. The experimental design at the NERC site was a split-plot design, replicated three times, with rotation-crop year as the main plot and N rate as the subplot (4.6 m by 15.2 m, 69.9 m2). The crop rotations at this site were 177 continuous corn (CCCC), corn-soybean (CSbCSb), corn-corn-oats- meadow (CCOM) and continuous soybean (SbSbSbSb). The experimental design at the CWRC site was similar to that at the NERC site, but was replicated two times, with rotation-crop year as the main plot and N rate as the subplot (6.1 m by 12.2 m, 74.4 m2). The crop rotations were CCCC, CSbCSb, CCOM, and COMM (Robinson et al. 1996). The soil samples from the NERC site were collected in May 1996 and June 1997, and the soil samples from the CWRC site were collected in July 1996 and 1997. The soil samples were collected by pooling at least 6-8 soil cores from each subplot. The field-moist soil samples were sieved through a 2-mm screen, a portion of which was air-dried, and a portion of this was ground to pass an 80- mesh (180-^im) sieve. In the soil properties reported (Table 1 and 2), pH was determined by using a combination glass electrode (soil: water or 0.01 M CaCl2 ratio = 1:2.5), organic C by the Mebius method (1960), total N by a semimicro-Kjeldahl procedure (Bremner and Mulvaney 1982), and particle- size distribution by a pipette analysis (Kilmer and Alexander 1949). Organic N was determined from the difference between total and inorganic N (Keeney and Nelson 1982). Organic C and N were determined on the <180-|im samples, and pH and particle-size distribution were determined on the <2- mm mesh air-dried samples. When not in use, the field-moist samples were stored at 4°C. All results reported are averages of duplicate analyses and are expressed on an oven-dry basis. Moisture content was determined from loss in weight after drying at 105°C for 48 h. 178 Microbial bio mass C and N Microbial biomass C (Cmic) and N (Nmic) in soils were determined on the <2- mm mesh field-moist soil samples within 24 h after sampling by the chloroform fumigation-extraction method described by Vance et al. (1987) and the chloroform fumigation-incubation method, described by Horwath and Paul (1994), respectively. Details of the procedures used were reported previously (Moore et al. 2000). P-Glucosaminidase activity (3-Glucosaminidase activity was assayed by the method described by Parham and Deng (2000). Soil (1.0 g <2-mm mesh field-moist) was placed into a 50-ml Erlenmeyer flask, treated with 4 ml of 0.1 M acetate buffer (pH 5.5) and 1 ml of 10 mM p-nitrophenyl-ZV-acetyl-(3-D-glucosaminide as a substrate. The flask was stoppered, swirled to mix the contents and incubated at 37°C. After 1 h of incubation, the mixture was treated with 1 ml of 0.5 M CaCla and 4 ml NaOH to stop the reaction and develop the yellow color of p-nitrophenol released. The soil-solution mixture was swirled for a few seconds and filtered through a Whatman No. 2v folded filter paper (Fisher Scientific Co., Itasca, IL). For the controls the substrate was added after the reaction was stopped. The enzymatic reaction involved is as follows: 179 H0 mh Hz° (3-Glucosaminidase^ ho^Z--\--\^oh • R-oh I ^ NH Ic=o IC=0 CH, | CHj p-Nitrophenyl-Af-acetyl-p -D- glucosaminide yv-acetyl-P-D-glucosaminide p-nitrophenol "• -v/n°° The intensity of the yellow filtrates was measured with a Klett-Summerson photoelectric colorimeter fitted with a blue No. 42 filter, and the p- nitrophenol content of the filtrate was calculated from a calibration graph prepared as described by Tabatabai (1994) for assay of acid phosphatase or arylsulfatase. From the values and organic C contents, the specific activity, expressed in g p-nitrophenol released per kg C0rg, was calculated. For all data points shown in figures 3-7, the differences between duplicate values obtained in the analysis or assays were smaller than the symbol size. N mineralization Nitrogen mineralization was studied in leaching columns under aerobic conditions at 30°C for 24 weeks (Stanford and Smith 1972). Details of the procedure used and the individual results were previously reported by Deng et al. (2000). 180 Statistical analysis Statistical analyses, including analysis of variance, contrast comparisons and separation of means by least significance differences were performed by using the general linear model procedure of the SAS system (Barr et al. 1976). Results Effect of cropping systems on P-glucosaminidase activity P-Glucosaminidase activity in soils was significantly affected by crop rotations and N fertilization in both sites and years (Figs. 1 and 2). In general, the highest p-glucosaminidase activities were obtained in soils under the 4-year corn-oats-meadow rotations (CCOM) taken under the 180 kg N ha-1 treatment. The lowest activities were found in soils under continuous soybean, followed by the 2-year corn-soybean (CSbCSb) rotation at the NERC site. Soils under com monoculture showed the lowest enzyme activity, followed by the 2-year corn-soybean (CSbCSb) rotation at the CWRC site. In general, the activity of P-glucosaminidase was significantly affected by N fertilization at the NERC site (p < 0.01 and p < 0.001) and at the CWRC site (p < 0.01 and p < 0.05)(Table 3). 181 P-Glucosaminidase activity in relation to organic C and N Linear regression analyses showed that, with the exception of the results for the soils sampled at the NERC site in 1997, P-glucosaminidase activity values were significantly correlated with organic C and N, with r values > 0.37* (Figs. 3 and 4). P-Glucosaminidase activity in relation to Cmic and Nmic » p-Glucosaminidase activity values were significantly correlated with Cmic for the soils from the NERC site, with r> 0.71*** and r> 0.49*** in 1996 and 1997, respectively (Fig 5). The enzyme activity values at this site were also significantly correlated with Nmic in 1996 (r = 0.76***) (Fig. 6). This relationship, however, was not significant for the soils sampled at the NERC site in 1997 (not shown). The results from the CWRC site showed a significant correlation between P-glucosaminidase and Cmic with r = 0.64*** and r = 0.44** in 1996 and 1997, respectively (Fig. 5). The enzyme activity values were significantly correlated (r = 0.33*) with Nmic at this side only in the samples obtained in 1996 (Fig. 6). 182 P-Glucosaminidase activity in relation to N mineralization Linear regression analyses showed that the P-glucosaminidase activity values were significantly correlated with the cumulative of N mineralized during 24 weeks at 30°C for the soils sampled at the NERC and CWRC sites in 1996, with r values of 0.84*** and 0.79***, respectively (Fig. 7). Discussion The pH values of soils from the two sites were not significantly affected by N fertilization. Organic C and N contents of soils from the NERC site were lower than those at the CWRC site, and crop rotations significantly affected those values at both sites (Tables 1 and 2). Analyses of variance showed that P-glucosaminidase activity was significantly affected by crop rotation and N fertilization (Table 3). Cropping systems significantly affected the specific activity values at both sites, and values were greater in the soils sampled in 1997 than in those sampled in 1996 at the NERC site (Table 4). In general, no such differences were observed for the soils sampled in both years at the CWRC site (Table 5). The specific activity values could be used as indexes of organic C quality. Differences in the activity of p-glucosaminidase under the various crop rotations may be caused by differences in quantity, quality and distribution of crop residues, and therefore by differences in the microbial community. This is because crop rotation generally provides a greater 183 concentrations and diversity of organic materials compared with the monocultural systems and, thus, stimulates microbial activity, which generates greater microbial biomass and enzyme activities (Miller and Dick 1995; Friedel et al. 1996; Klose et al. 1999; Klose and Tabatabai, 2000). The results showed greater enzyme activity in soils under CCOM rotation taken under meadow than those under the other rotations (Figs. 1 and 2). This supports the results of Bolton et al. (1985), who showed that long-term field experiments that include legume plants in crop rotation contribute greater amounts of organic C and N and thereby greater microbial activities to soils than other rotations. The greater P-glucosaminidase activity in the soils under COMM rotation sampled under meadow as compared with those in soils under continuous corn or soybean systems, may also be the result of improved structure of the soil, plant cover, stabilized microclimate and higher root density found in very diversified crop rotations. Soil microbial populations and activities are mostly limited by C source in soils; thus the amounts and the rates of easily decomposable organic material are important in supporting microbial growth (Friedel et al. 1996). The higher amount of decomposable organic C fraction in alfalfa residue compared to corn and soybean residue (Ajwa and Tabatabai 1994) may explain the finding showing greater p-glucosaminidase activity in cropping systems containing meadow (alfalfa) than in CCCC or SbSbSbSb, or 2-year CSbCSb rotations. The results obtained for p-glucosaminidase activity are in agreement with those reported, by Dick (1984) showing much greater urease, alkaline phosphatase, 184 arylsulfatase, invertase and amidase activities in soils under a 3-year crop rotation including alfalfa than in those under corn monoculture and CSbCSb rotation. Other studies by Klose et al. (1999) and Klose and Tabatabai (2000) showed greater total urease and arylsulfatase activities in the same soils under CCOM rotation than in soils under monoculture and CSbCSb rotation. They found that total, intracellular, extracellular, and specific activities of urease and arylsulfatase in soils of the NERC site were significantly affected by crop rotation, but not by N fertilization. Also, they reported that the greatest total urease and arylsulfatase activities were obtained under 4-year rotations including oats and meadow and reported lowest activities under continuous corn. Studies by Deng and Tabatabai (1996a,b) on the effects of tillage and residue management on the activities of amidohydrolases and glycosidases showed increased soil enzyme activities under various management practices with increasing the amounts of crop residue. In addition to differences in the amount of residue left on soils by different rotations, the decomposition rates of the residue material, and their contribution to an easily degradable soil organic matter pool, vary among the residue materials, depending on the contents of N, S, soluble C lignin, and carbohydrates (Janzen and Lucey 1988; Klose and Tabatabai 2000). In general, the soils from the CWRC site showed 6 - 33% greater (3- glucosaminidase activity than those from the NERC site. This could be the result of variation in properties of the soils from the two sites such as pH, 185 Corg, Norg and clay contents; factors that are known to affect microbial growth and enzyme production and stability. p-Glucosaminidase activity was significantly correlated with Corg' Norg, Cmic and Nmic, with the exception Norg at the NERC site in 1997 (Figs. 3 and 6). The results, in general, support the findings of Klose et al. (1999) and Klose and Tabatabai (2000) that showed significant correlation between Cmic or Nmic and urease or arylsulfatase activity. Also, a significant correlation has been reported between urease activity and Corg in soils (Deng and Tabatabai 1996a; Klose et al. 1999). The greater P-glucosaminidase activity in soils at the CWRC site than at the NERC site could be attributed to the greater Corg content of the soils at the CWRC site than at the NERC site. These results are expected because of increased plant biomass resulting from fertilizer additions to soils, which stimulated microbial growth (containing chitin in the cell wall) and microbial activities. This conclusion is supported by the finding of S mucker and Kim (1987) showing that chitin enhances the activity of this enzyme in an estuarine system. Other studies showed significant correlation between amidase, asparaginase, urease, dipeptidase, and protease activity values and N mineralization (Burton and McGill 1992; Burket and Dick 1998; Zaman et al. 1999; Deng et al. 2000). The significant, positive correlation between P- glucosaminidase activity values and Cmic and Nmic values (Figs. 5 and 6), together with the significant correlation with N mineralization (Fig. 7), suggest that this enzyme plays a major role in N mineralization. This finding 186 is important because the activity of this enzyme could be a rate-limiting step in organic N mineralization in soils; i.e., producing amino sugars, which, in turn, mineralize. Other recent work showed that the annual means of arginine ammonification assays were correlated with gross N mineralization values in four different agricultural fields in Denmark (Bonde et al. 2001). 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Biol Fertil Soils 29:178-186 192 Table 1 Properties of the soils at the Northeast Research Center (NERC) at Nashua, Iowa B Crop N PH rotation" treatment H2O CaClo Org. Cb Total Nb kg ha'1 — g kg I C-c-c-c 0 6.7 (6.4) 6.2 (5.9) 25.3 (21.2) 1.64 (1.67) 180 6.4 (6.5) 6.1 (6.0) 22.6 (22.6) 1.49 (2.07) C-sb-c-sb 0 7.1 (7.1) 6.6 (6.5) 17.3 (18.7) 1.21 (1.73) 180 6.6 (7.1) 6.2 (6.3) 17.7 (19.5) 1.32 (1.87) c-Sb-c-sb 0 7.1 (7.1) 6.6 (6.5) 19.8 (21.4) 1.37 (1.63) 180 7.1 (6.6) 6.5 (6.3) 19.2 (21.9) 1.40 (1.67) C-c-o-m 0 -C (6.9) " (6.9) - (22.7) - (1.93) 180 - (6.5) " (6.5) - (20.3) - (1.90) c-C-o-m 0 7.0 (7.1) 6.5 (7.1) 21.4 (21.6) 1.57 (1.70) 180 6.2 (6.8) 6.0 (6.8) 23.3 (22.4) 1.68 (1.67) c-c-O-m 0 7.2 (6.8) 6.6 (6.8) 22.0 (22.7) 1.40 (1.67) 180 7.0 (6.7) 6.3 (6.7) 21.4 (23.5) 1.46 (1.90) c-c-o-M 0 7.0 - 6.5 25.5 - 1.79 - 180 6.7 - 6.1 23.2 - 1.63 - Sb-sb-sb-sb 0 7.1 (7.0) 6.5 (7.0) 17.4 (18.6) 1.25 (1.77) LSD P < 0.05D 0.5 (0.6) 0.3 (0.5) 1.9 (2.2) 0.11 (0.17) LSD P < 0.05E 0.6 (0.4) 0.2 (0.5) 1.2 (2.5) 0.10 (0.20) a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold letters indicate crop in which the sample was taken in 1996. In 1997, the samples were taken in the crop following the capital bold letter. b Means of three field replicates for 1996. Figures in parentheses are data for 1997. c Indicates that sample was not taken. d Least significant difference due to crop rotation at 0 N. e Least significant difference due to crop rotation at 180 N. 193 Table 2 Properties of the soils at the Clarion-Webster Research Center (CWRC) at Kanawha, Iowa Crop N pHb b b rotation" treatment H2O CaCl2 Org. C Total N kg ha'1 g kg C-c-c-c 0 6.9 (7.0) 6.2 (6.2) 28.2 (29.8) 1.79 (2.00) 180 6.0 (6.0) 5.3 (5.4) 35.2 (35.6) 2.32 (2.20) C-sb-c-sb 0 7.1 (7.3) 6.5 (6.7) 31.2 (32.9) 2.08 (2.20) 180 6.7 (6.8) 6.2 (6.2) 29.6 (32.1) 1.96 (2.20) c-Sb-c-sb 0 6.7 (6.7) 6.2 (6.1) 30.6 (34.5) 2.03 (2.25) 180 6.5 (6.3) 5.9 (5.8) 31.8 (33.6) 2.15 (2.15) C-c-o-m 0 6.2 (6.5) 5.6 (5.7) 29.5 (31.2) 2.05 (1.60) 180 5.8 (6.0) 5.3 (5.3) 34.9 (36.4) 2.42 (1.80) c-C-o-m 0 6.4 (6.3) 5.8 (5.4) 32.1 (33.9) 2.15 (1.95) 180 5.9 (6.2) 5.4 (5.4) 35.1 (37.7) 2.40 (1.95) c-c-O-m 0 6.0 (6.0) 5.3 (5.2) 33.1 (35.7) 2.18 (2.35) 180 5.9 (5.8) 5.2 (5.1) 34.2 (36.4) 2.36 (2.30) c-c-o-M 0 5.9 (5.9) 5.4 (5.2) 38.4 (40.9) 2.50 (2.40) 180 5.7 (5.6) 5.2 (4.9) 37.1 (39.5) 2.45 (2.00) C-o-m-m 0 5.8 (6.2) 5.2 (5.6) 35.4 (36.1) 2.43 (2.25) 180 5.5 (5.7) 4.9 (4.9) 34.8 (34.5) 2.37 (2.30) c-O-m-m 0 6.0 (5.9) 5.3 (5.3) 34.5 (35.3) 2.34 (2.50) 180 6.4 (6.4) 5.8 (5.6) 37.0 (44.3) 2.52 (2.85) c-o-M-m 0 6.2 (6.2) 5.6 (5.5) 30.2 (30.5) 2.12 (2.20) 180 6.1 (5.8) 5.5 (5.1) 32.6 (34.9) 2.24 (2.35) c-o-m-M 0 5.9 (5.6) 5.3 (5.1) 37.7 (39.2) 2.54 (2.50) 180 5.8 (5.4) 5.2 (4.9) 37.8 (39.1) 2.67 (2.55) LSD P < 0.05° 1.1 (1.1) 1.1 (1.2) 4.9 (6.0) 0.33 (0.47) LSD P < 0.05D 1.3 (1.5) 1.4 (1.5) 5.5 (4.9) 0.28 (0.28) a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold letters indicate crop in which the sample was taken in 1996. In 1997, the samples were taken in the crop following the capital bold letter. b Means of two field replicates for 1996. Figures in parentheses are data for 1997. c Least significant difference due to crop rotation at 0 N. d Least significant difference due to crop rotation at 180 N. 194 Table 3 Analysis of variance of effects of cropping systems on P-Glucosaminidase in soils from the Northeast Research Center (NERC) and Clarion Webster Research Center (CWRC) Source ft-Glucosaminidase activity of variation NERC CWRC 1996 Crop rotation *** *** N fertilizer ** ** Contrasts'* CI *** C2 ** n.s. C3 n.s. n.s. C4 *** *** C5 *** *** 1997 Crop rotation *** *** N fertilizer *** * Contrasts1 CI *** _b C2 n.s. *** C3 *** n.s. C4 *** *** C5 * *** ***, ** and * are significant at P < 0.05,0.01 and 0.001 level, respectively; n.s. not significant. a Contrasts: CI: CCCC vs. SbSbSbSb, C2: CCCC vs. multicropping systems, C3: CCCC vs. 2-year rotation, C4: 2-year rotation vs. 4-year rotation, C5: CCCC vs. 4-year rotation. b No SbSbSbSb treatment at the CWRC site. 195 Table 4 Specific activity of (3-glucosaminidase in soils under different croping systems at the NERC site in!996 andl997 Specific activity Crop N 1996b 1997b- 11 rotation treatment Rt R2 R3 Mean Rt R2 R3 Mean kg ha1 C-c-c-c 0 1.47 1.22 1.73 1.47 2.03 2.94 2.07 2.34 180 1.41 1.97 1.32 1.57 2.37 1.85 2.73 2.32 C-sb-c-sb 0 1.29 1.51 1.55 1.45 1.91 2.10 2.07 2.03 180 1.85 1.85 1.48 1.73 2.34 1.81 1.77 1.97 c-Sb-c-sb 0 1.55 2.01 1.72 1.76 1.65 2.37 1.70 1.91 180 1.56 1.80 2.56 1.97 2.04 2.30 1.93 2.09 C-c-o-m 0 - - - - 2.04 2.28 2.08 2.13 180 - - - - 3.01 3.10 2.52 2.88 c-C-o-m 0 2.19 2.12 2.19 2.17 2.75 2.93 2.53 2.74 180 1.93 2.19 2.48 2.20 2.80 2.71 2.48 2.66 c-c-O-m 0 1.99 2.39 2.03 2.14 2.30 2.56 2.53 2.46 180 2.16 2.39 2.48 2.34 2.33 3.07 2.45 2.62 c-c-o-M 0 2.39 2.04 2.24 2.22 - - - - 180 2.52 2.49 2.70 2.57 - - - - Sb-sb-sb-sb 0 1.14 1.17 1.27 1.19 1.44 1.49 1.38 1.43 LSD P < 0.05' 0.28 0.33 0.40 0.28 0.18 0.74 0.43 0.37 LSD P < 0.05d 0.16 0.25 0.35 0.33 0.13 0.15 0.07 0.35 LSD P < 0.05e 0.30 0.30 0.11 0.54 0.11 0.27 0.13 0.61 a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold letters indicate crop in which the sample was taken in 1996. In 1997, the samples were taken in the crop following the capital bold letter. b R = field replicate; subscript = replication number. c Least significant difference due to crop rotation at both 0 and 180 N. d Least significant difference due to crop rotation at 0 N. e Least significant difference due to crop rotation atlSO N. 196 Table 5 Specific activity of p-glucosaminidase in soils under different cropping systems at the CWRC site in 1996 and 1997 Specific activity Crop N —1996b- .... i997b. rotation™ treatment Ri R2 Mean Ri Ro Mean kg ha ' -g of p -Nitrophenol / kg'1 of Organic c C-c-c-c 0 1.13 1.21 1.17 0.80 1.18 0.99 180 1.22 1.82 1.52 0.93 1.66 1.29 C-sb-c-sb 0 1.13 1.05 1.09 1.14 0.98 1.06 180 1.43 0.99 1.21 1.30 1.19 1.25 c-Sb-c-sb 0 1.09 1.13 1.11 0.79 1.31 1.05 180 0.98 1.33 1.16 1.22 1.11 1.16 C-c-o-m 0 1.55 1.20 1.38 1.39 1.31 1.35 180 1.49 1.32 1.41 1.36 1.42 1.39 c-C-o-m 0 1.39 1.35 1.37 1.42 1.72 1.57 180 1.58 1.42 1.50 1.64 1.40 1.52 c-c-O-m 0 1.39 1.57 1.48 2.03 1.47 1.75 180 2.06 1.63 1.85 1.69 1.97 1.83 c-c-o-M 0 2.14 2.06 2.10 1.36 1.48 1.42 180 1.88 2.04 1.96 1.56 1.25 1.40 C-o-m-m 0 1.67 1.12 1.40 1.04 1.52 1.28 180 1.73 1.31 1.52 1.51 1.48 1.49 c-O-m-m 0 1.41 1.31 1.36 1.63 1.63 1.63 180 1.53 1.60 1.57 1.81 1.80 1.81 c-o-M-m 0 1.72 1.72 1.72 2.26 2.43 2.34 180 2.42 1.46 1.94 1.80 2.10 1.95 c-o-m-M 0 1.95 1.39 1.67 1.32 1.18 1.25 180 1.98 2.10 2.04 1.26 1.44 1.35 LSD P < 0.05= 0.25 0.24 0.54 0.24 0.29 0.61 LSD P < 0.05d 0.04 0.10 0.39 0.12 0.15 0.51 LSD P < 0.05e 0.10 0.10 0.69 0.05 0.09 0.46 a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold letters indicate crop in which the sample was taken in 1996. In 1997, the samples were taken in the crop following the capital bold letter. b R = field replicate; subscript = replication number. c Least significant difference due to crop rotation at both 0 and 180 N. d Least significant difference due to crop rotation at 0 N. e Least significant difference due to crop rotation at 180 N. 0 kg N ha NERC site 180 kg N ha'1 100 V ^ § 4 40 Crop Rotation Crop Rotation Fig. 1 Effect of cropping systems on p-glucosaminidase activity at the NERC site. Capital letters indicate the crop in which samples were taken in 1996. In 1997, samples were taken in the crop following the capital letter. Results are averages of three field replicates. Within each N treatment, bars with the same letter are not significantly different at P<0.05 level using LSD (0 N treatment, lower case letters; 180 N treatment, upper-case letters). 198 1996 - CWRC site 0 kg N ha 180 N ha"1 1997 Crop rotation Fig. 2 Effect of cropping systems on P-glucosaminidase activity at the CWRC site. Capital letters indicate the crop in which samples were taken in 1996. In 1997, samples were taken in the crop following the capital letter. Results are averages of two field replicates. Within each N treatment, bars with the same letter are not significantly different at F<0.05 level using LSD (0 N treatment, lower case letters; 180 N treatment, upper-case letters). vz- 100 Y = -20.46 + 2.89 X NERC Y = -24.9 +3.45 X NERC r = 0.70 *** 1996 r = 0.58 *** 1997 80 60 40 20 0 //- 100 Y = -60.2 + 3.33 X CWRC Y =1.88+1.41 X CWRC r = 0.42 ** r = 0,76 *** 1996 1997 co 80 CD 60 40 20 • OkgNha'1 0 180 kg N ha'1 0 50 0 Organic C (g kg soil) hip between p-glucosaminidase activity and organic C of soils at the NERC and CWRC sites sampled 7. 100 Y = -29.7 + 47.8 X NERC Y = 44.8 +2.18 X NERC 90 r = 0.75 *** 1996 r = 0.03 1997 80 70 60 50 40 30 20 10 0 100 Y = -57.5 + 48.1 X CWRC Y = 17,4 + 15.7 X CWRC 90 r = 0.76 *** 1996 r = 0,37 * 1997 80 o o 6o 70 •o 8 • / 60 50 40 30 20 • 0 kg N ha-1 10 ° 180 kg N ha1 0 0, 0 0.5 1.0 1.5 2.0 2.5 3.0 3,5 4.0 4,5 0.0 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 Organic N (g kg'1 soil) between p-glucosaminidase activity and organic N of soils at the NERC and CWRC sites 100 Y =-2,53 +0.20 X NERC Y = 23.2 +0.12 X NERC r = 0,71 *** 1996 r = 0.49 *** 1997 80 60 40 20 0 100 Y = 1,86+ 0,13 X CWRC Y = 30.4 + 0.07 X CWRC r = 0.44 ** 1997 r = 0,64 *** 1996 Kl 80 o O 60 40 20 • 0 kg N ha'1 ° 180 kg N ha'1 0 100 200 300 400 500 600 700 0 100 200 300 400 500 600 700 cmic (mg C kg"1 soil) ship between p-glucosaminidase activity and microbial biomass C of soils at the NERC and CWRC sites 6 and 1997. 202 Y = 2.58 + 0.75 X NERC 90 r = 0.76 *** 1996 80 70 60 o Sb 50 40 30 20 10 0 .00 Y =32.9 +0.30 X CWRC 90 r = 0.33 * 1996 80 70 60 50 oe o * 40 30 20 0 kg N ha*1 10 180 kg N ha1 0 20 40 60 80 100 120 140 160 1 Nmic (mg N kg* soil) nship between P-glucosaminidase activity and microbial biomass N of SRC and CWRC sites sampled in 1996. 203 100 Y = -13.9 + 0.33 X NERC 90 r = 0.84 *** 1996 80 70 60 50 40 30 20 10 0 100 Y =3.55 +0.27 X CWRC r = 0.79 *** 90 1996 80 70 60 50 40 30 20 10 • 0 kg N ha"1 ° 180 kg N ha*1 0 ) 50 100 150 200 250 300 350 Cumulative N mineralized (mg N kg*1 soil) inship between p-glucosaminidase activity and cumulative N NERC and CWRC sites sampled in 1996 and 1997. 204 CHAPTER 7 EFFECTS OF TRACE ELEMENTS ON P-GLUCOSAMINIDASE ACTIVITY IN SOILS A paper in press in Soil Biology & Biochemistry M. Ekenler and M. A. Tabatabai Abstract The enzyme P-glucosaminidase (2V-acetyl-P-D-glucosaminidase, EC 3.2.1.30) hydrolyzes iV-acetyl-P-D-glucosamine residues from the terminal non-reducing ends of chitooligosaccharides in soils. This is one of the enzymes that play a major role in N mineralization in soils. Studies to evaluate the effects of salts of 23 trace elements on the activity of P-glucosaminidase in three Iowa surface (0-15 cm) soils showed that at 5 mmol kg*1 soil, the activity of P-glucosaminidase was inhibited by 18 and activated by 5 of the trace elements tested. The inhibition values ranged from 0 to 73%, depending on the trace element and the soil used. In general, Ag(I) and Hg(II) were the most effective inhibitors. Other metals that inhibited the p-glucosaminidase activity in soils were Cu(I), Ba(H), Cu(H), Niai), Pb(II), Sn(ID, Zn(II), As(IH), Cr(m), Se(IV), Ti(IV), V(IV), W(VI) and Mo(VI). At the concentration tested, A1(IH), Fe(II), and Fe(ill) also inhibited the activity of this enzyme. The elements Co (H), Mg (H), Mn (H), B 205 (III) and As (V) activated this enzyme by values ranging from 4 to 68% in three soils. Keywords: Soil enzymes; Heavy metals; Trace elements, N cycle; N mineralization P-Glucosaminidase (.ZV-acetyl-P-D-glucosaminidase, EC 3.2.1.30) is the enzyme that catalyzes the hydrolysis of 2V-acetyl-p -D-glucosamine residues from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also classified as P-iV-acetyl-D-hexosaminide iV-acetylhexosaminohydrolase (EC 3.2.1.52) because it acts on glucosides, galactosides and several oligosaccharides (Webb, 1984). This enzyme is one of the three chitinases (iV-acetyl-P-D- glucosaminidase, chitobiosidase and endochitinase) that degrade chitin (Tronsmo and Harman, 1993). Information on the effect of trace elements on p- glucosaminidase in soils is needed because chitin is one of the most abundant biopolymers on earth serving as an important transient pool of organic C and N in soils (Wood et al., 1994), and because amino sugars comprise about 5 to 10% of organic N in soils (Stevenson, 1994). Studies by Liang and Tabatabai (1977, 1978) using three Iowa surface soils showed that, at 5 mmol kg-1 soil, 19 metals inhibited N mineralization and nitrification in soils, and that the degree of inhibition varied with the trace elements and soils used. Other studies on the toxicity of heavy metals to microorganisms and their processes in soils employed ecological dose value, EDso, showing the critical concentrations or loading, 206 especially of trace elements, including heavy metals, in sewage sludges (for review, see Giller et al., 1998). By using this concept, Speir et al (1995) showed that EDso values of Cr(VI) for three soils varied from 63 to 730 mmol kg-1 soil. Another ecological concept that has been used in describing the effects of heavy metals on soil microbial diversity and activity is the sensitive-resistance index (Doelman et al., 1994). These concepts, however, are not used in enzyme chemistry. Recent studies in our laboratories showed that cropping systems affect the activity of P-glucosaminidase in soils, and that the activity of this enzyme is significantly correlated with N mineralization in soils of two long-term cropping systems in Iowa, with r values of 0.79*** and 0.84***. This suggested that the activity of P-glucosaminidase could be used as an index of N mineralization. The term trace element is used here to refer to elements that are, when present in sufficient concentration, toxic to living systems (Page, 1974). The relative effect of those elements and of A1 and Fe on the activity of p- glucosaminidase deserves investigation because these elements are naturally added to soils as fertilizers, as impurities in fertilizers or as components of municipal and industrial wastes (Berrow and Webber, 1972; Page, 1974; Tabatabai and Frankenberger, 1979, Charter et al., 1993, 1995). In addition, some of these elements, such as Pb, Hg, Cd and V, are present in industrial wastes added to soils or present in fuel oils and gasoline, and are emitted into 207 the atmosphere upon burning and deposited on soils, especially near urban areas (Lagerwerff, 1972). To our knowledge, no information is available on the effects of trace elements and heavy metals on the activity of (3-glucosaminidase in soils. Therefore, this study was conducted to evaluate the effect of 23 trace elements and A1 and Fe, some of which are at different oxidation states, on the activity of p-glucosaminidase in soils. The soils used were surface soils (0- 5 cm) selected to include a range of pH, organic C, total N, texture, Cmic, Nmic, and p-glucosaminidase activity (Tables 1 and 2). The soils were passed through a 2-mm mesh sieve, and a portion of this was air-dried. A portion of the air-dried soils was ground to pass an 80- mesh screen (180 |im). Soil pH was determined with a combination glass electrode and particle-size distribution by a pipette analysis (Kilmer and Alexander, 1949) on the 2-mm air-dried soils. Organic C and total N were determined on the <180-pm samples by dry combustion method using LECO CHN 600 determinator (LECO Corp., St Joseph, MI). The Cmic and Nmic values were determined in field-moist soils within 24 h after sampling as described by Moore et al (2000). The assay of P-glucosaminidase activity was performed on the field-moist (< 2-mm) samples as described by Parham and Deng (2000). The chemical elements used were Fisher certified reagent-grade chemicals (Fisher, Itasca, IL). Of these, Ag(I), Cd(II), Co(H), Cu(H), Fe(II), Zn(II) and V(IV) were added as the sulfate; Cu(I), Ba(II), Hg(II), Mg (II), Mn(II), Ni(H), Sn(H), A1(III), 208 Cr(m) and Fe(III) were added as the chloride; Pb(II) as the nitrate and as acetate; and As(III), B(III), Se(IV), As(V), Mo(VI), W(VI) and Ti(IV) were added as NaAsOs, NaoB^?, HgSeOa, NaaHAsC^, NaoMo04, NagW04 and TiOSÛ4, respectively. The effect of trace elements and of A1 and Fe on p-glucosaminidase activity was studied by treating 1 g of soil in a 50-ml Erlenmeyer flask with 1 ml of a solution containing 5 |imol of the trace element. The solution was added dropwise to moisten the whole soil sample. After 1 h of equilibration, the sample was treated with 3 ml of 0.1 M acetate buffer (pH = 5.5) and 1 ml of 10 mM p- nitrophenyl-N-acetyl-P-D-glucosaminide solution. Because the soil sample was treated with 1 ml of trace-element solution, 3 ml of the buffer was added instead of the 4 ml recommended. The p-glucosaminidase activity values obtained for the soil treated with trace elements were compared with those obtained with a 1- g soil sample treated with 1 ml of deionized water instead of the trace element solution. The percentage of inhibition or activation by each trace element was calculated from [(A-B)/A] x 100, where A is the p-glucosaminidase activity of non-treated soil and B is the P-glucosaminidase activity of the trace-element- treated soil. All results reported are averages of duplicate assays and one control, expressed on a moisture-free basis. Moisture was determined from weight loss after drying the soil at 105°C for 48 h. Difference due to trace element effect for each soil was evaluated by using the LSD (P< 0.05) values 209 calculated by the Statistical Analysis System (SAS) computer program (Barr et al., 1976). Results indicated that of the 23 elements, 18 inhibited and 5 activated (3- glucosaminidase activity in the three soils used (Tables 2 and 3). The inhibition values ranged from 0 to 73%, depending on the element and the soil used. In general, Ag(I) and Hg(II) were the most effective inhibitors. Other elements that inhibited the p-glucosaminidase activity in soils were Cu(I), Ba(II), Cu(II), Fe(II), Ni(II), Pb(ID, Sn(II), Zn(II), Al(III), As(III), Cr(III), Fe(III), Se(IV), Ti(IV), V(IV), W(VI) and Mo(VI). The trace elements Co(II), Mg(II), Mn(II), B(III) and As(V) activated this enzyme by values ranging from 4 to 68% in three soils. Information on the chemical nature of the active sites of soil P- glucosaminidase is not available, but studies on the catalytic active sites and amino acid sequencing of this enzyme purified from microorganisms indicate that the sites contain glutamate and tryptophan residues (Scigelova and Grout, 1999). This information is clearly supported by the much less than complete inhibition values obtained by Ag(II) and Hg(II), suggesting that the active sites of this enzyme are not totally controlled by amino acids containing sulfhydryl groups such as those in urease (Tabatabai, 1977). The form and concentration of the trace element, the soil investigated and the type of enzyme assayed affect the degree of enzyme inhibition (Ladd, 1985; Nannipieri, 1994). Effectiveness of the trace elements depends on the type and concentration added because a proportion of the —SH groups of enzyme-active sites reacts with the trace 210 element ions. The chemical and physical properties of the soil, such as organic matter content, kind and amount of clay and soil pH, influence the toxic effect of trace elements (Bunzl et al., 1976; Gadd and Griffiths, 1978; Sinha et al., 1978). Metal ions may inhibit enzyme reactions by complexing the substrate, by combining with the enzyme's active sites or by reacting with the enzyme- substrate complex. It is well known that different metal ions show different behavior in their ability to act as inhibitors of P-glucosaminidase (Ueda and Arai, 1992; Takahashi et al., 1993; Scigelova and Grout, 1999). In general, the inhibition by heavy metals follows the noncompetitive kinetics, i.e., there is no relationship between the amount of inhibitor and the concentration of the substrate. Inhibition depends on the concentration of the inhibitor; the formation of EI occurs at a locus which is not attacked by the substrate. E +1 = EI and ES+ 1 = ESI, where E is the enzyme, ES is the enzyme-substrate complex, and I is the inhibitor (White et al., 1959; p. 250). Metal ions may inactivate enzymes by reacting with sulfhydryl groups, a reaction analogous to the formation of a metal sulfide (Shaw and Raval, 1961). Sulfhydryl groups in enzymes may serve as integral parts of the catalytic active sites or as groups involved in maintaining the correct structural relationship of enzyme protein. The difference in the order of effectiveness of inhibition among soils may be related to the different soil-trace element interactions despite the fact that 211 the activity of the enzyme was assayed after 1 h of equilibration of the soil-trace element solution mixture. It is possible that because of the variation in the original soil pH values before addition of acetate buffer for assay of the enzyme activity, the ionic species of the trace elements varied among the soils. In addition, there are also other trace element reaction processes in soils, e.g., cation-exchange reaction with functional groups of organic matter, adsorption and precipitation reactions that could consume significant portions of the added trace elements. These processes could be responsible for the differences in the inhibition or activation observed among the soils used (Lindsay, 1979; Karmarkar and Tabatabai, 1993). Thus, the effectiveness of the trace elements in inhibition or activation of (3-glucosaminidase varied among soils. Control studies with 5 mmol of NaCl or K2SO4 kg-1 soil indicated that K+, Na+, CV and SO4- associated with the salts of the trace element studied did not affect p-glucosaminidase activity in soils. Also, NOa did not affect the activity of this enzyme in soils. The pH of the trace element solution varied considerably, ranging from 2.1 for FeCla.GHaO solution to 10.2 for the NaAsOg solution. However, the use of acetate buffer ensured a constant pH of 5.5 ± 0.2 in the reaction mixture. Therefore, the observed inhibition or activation of P-glucosaminidase in soils in the presence of the trace elements studied was not a result of changes in pH of the reaction mixture but of the reaction between the free trace element in solution and the functional groups of P-glucosaminidase. 212 Acknowledgements Journal Paper no. J-19667 of the Iowa Agric. and Home Econ. Exp. Stn., Ames. Projects no. 3338 and 4184. This work was supported in part by the Biotechnology By-products Consortium of Iowa. References A.F. Barr, J.H. Goodnight, J.P. Sail, J.I. Helwig, A User's Guide to SAS, SAS Institute, Raleigh, NC, 1976. M.C. Berrow, J. Webber, Trace elements in sewage sludges, Journal Science of Food and Agriculture 23 (1972) 93-100. K. Bunzl, W. Schmidt, B. Sansonik, Kinetics of ion exchange in soil organic matter: IV. Adsorption and desorption of Pb2+, Cu2+, Cd2+, Zn2+ and Ca2+ by peat, Journal of Soil Science 27 (1976) 32-41. R.A. Charter, M.A. Tabatabai, J.W. Schafer, Metal contents of fertilizers marketed in Iowa, Communications in Soil Science and Plant Analysis 24 (1993) 961-972. R.A. Charter, M.A. Tabatabai, J.W. Schafer, Arsenic, molybdenum, selenium, and tungsten contents of fertilizers and phosphate rocks, Communications in Soil Science and Plant Analysis 26 (1995) 3051-3062. 213 P. Doelman, E. Jansen, M. Michels, M. van Till, Effects of heavy metals in soils on microbial diversity and activity as shown by the sensitivity-resistance index, an ecological relevant parameter, Biology and Fertility of Soils 17(1994) 177-184. G.M. Gadd, A.J. Griffiths, Microorganisms and heavy metal toxicity. Microbial Ecology 4 (1978) 307-317. K.E. Giller, E. Witter, S.P. McGrath, Toxicity of heavy metals to microorganisms and microbial processes in agricultural soils: A review. Soil Biology & Biochemistry 30 (1998) 1389-1414. S.V. Karmarkar, M.A. Tabatabai, Adsorption of oxyanions by soils. Soil Science (Trends in Agricultural Sciences) 1 (1993) 63-101. V.J. Kilmer, J.T. Alexander, Methods of making mechanical analysis of soils. Soil Science 68 (1949) 15-24. J.N. Ladd, Soil enzymes, in: D. Vaughan, E. Malcom (Eds.), Soil organic matter and biological activity, Martinus Nijhoff, Dordrecht, 1985, pp 175- 221. J.V. Lagerwerff, Lead, mercury, and cadmium as environmental contaminants, in: J.J. Mortvedt, P.M. Giordano, W.L. Lindsay (Eds.), Micronutrients in Agriculture, Soil Science Society of America, Madison, WI, 1972, pp. 593- 636. C.N. Liang, M.A. Tabatabai, Effects of trace elements on nitrogen mineralization in soils, Environmental Pollution 12 (1977) 141-147. 214 C.N. Liang, M A. Tabatabai, Effects of trace elements on nitrification in soils, Journal of Environmental Quality 7 (1978) 291-293. W.L. Lindsay, Chemical equilibria in soils, Wiley, New York, NY, 1979. J.M. Moore, S. Klose, M.A. Tabatabai, Soil microbial biomass carbon and nitrogen as affected by cropping systems, Biology and Fertility of Soils 31 (2000) 200-210. P. Nannipieri, The potential use of soil enzymes as indicators of soil productivity, sustainability, and pollution, in: C.E. Pankhurst, B.M. Doube, V.V.S.R. Gupta, P R. Grace, (Eds.), Soil Biota: Management in Sustainable Farming Systems. CSIRO Australia, Victoria, 1994, pp 238- 244. A.L. Page, Fate and effect of trace elements in sewage when applied to agricultural land: A Literature Review, Environmental Protection Agency, Cincinnati, OH, 1974. J.A. Parham, S.P. Deng, Detection, quantification and characterization of P- glucosaminidase activity in soil, Soil Biology & Biochemistry 32 (2000) 1183-1190. M. Scigelova, D.H.G. Crout, Microbial P-iV-acetylhexosaminidases and their biotechnological applications, Enzyme and Microbial Technology 25 (1999) 3-14. W.H.R. Shaw, D.N. Raval, The inhibition of urease by metal ions at pH 8.9, Journal of the American Chemical Society 83 (1961) 3184-3187 215 M.K. Sinha, S.K. Dhillon, S. Dyamond, Solubility relationship of iron, manganese, copper, and zinc in alkaline and calcareous soils, Australian Journal of Soil Research 16 (1978) 19-26. T.W. Speir, H.A. Kettles, A. Parshotam, P L. Searle, L.N. Vlaar, A simple kinetic approach to derive the ecological dose value, ED50, for the assessment of Cr(VI) toxicity to soil biological properties, Soil Biology & Biochemistry 6 (1995) 801-810. F.J. Stevenson, Humus Chemistry: Genesis, Composition, Reactions, Wiley, New York, 1994. M.A. Tabatabai, Effects of trace elements on urease activity in soils. Soil Biology & Biochemistry 9 (1977) 9-13. M.A. Tabatabai, W.T. Frankenberger, Jr, Chemical composition of sewage sludges in Iowa, Iowa Agriculture and Home Economics Experiment Station Research Bulletin 586, 1979. M. Takahashi, T. Mashiyama, T. Suzuki, Purification and some characteristics of p-AT-acetylglucosaminidase produced by Vibrio sp, Journal of Fermentation and Bioengineering 76 (1993) 356-360. A. Tronsmo, G.E. Harman, Detection and quantification of iV-acetyl-P-D- Glucosaminidase, chitobiosidase and endochitinase in solutions and on gels, Analytical Biochemistry 208 (1993) 74-79. 216 M. Ueda, M. Arai, Purification and some properties of fi-N- acetylglucosaminidase from Aeromonas sp. 10S-24, Bioscience, Biotechnology, and Biochemistry 56 (1992) 1204-1207. E C. Webb, Enzyme Nomenclature, Academic Press, New York, 1984. A. White, P. Handler, E.L. Smith, D. Stetten, Principles of Biochemistry, Second éd., McGraw-Hill, New York, 1959. C.W. Wood, H A. Torbert, H.H. Rogers, G.B. Runion, S.A. Prior, Free-air CO2 enrichment effects on soil carbon and nitrogen, Agricultural and Forest Meteorology 70 (1994) 103-116. Table 1 Some selected properties of the soils used Soils Chemical Properties Biological Properties1' Particle-size Distribution Series Subgroup pH" Organic C Total N f^inic iN^mic Clay Sand g kg 1 mgkg1 soil g kg1 Clarion Typic Hapludoll 4.6 21.1 3.7 335 13.2 240 360 Nicollet Aquic Hapludoll 6.1 21.3 3.2 279 21.8 240 370 Canisteo Typic Endoaquoll 7.4 37.7 3.8 527 40.3 280 320 "Soil: water ratio (1:2.5). b Cinin = microbial biomass C, Nmi(. = microbial biomass N. 218 Table 2 Inhibition of p -glucosaminidase activity in soils by trace elements Trace element Inhibition of P-glucosaminidase activity (%) in soils specified" Element Oxidation Clarion Nicollet Canisteo state Ag I 20 73 50 Cu 5 33 32 Ba II 11 16 0 Cd 9 40 37 Cu 6 34 33 Fe 7 8 7 Hg 35 55 72 Ni 12 14 18 Pb 7 16 5 Pb 6 18 0 Sn 19 12 21 Zn 8 18 32 A1 III 33 17 13 As 0 21 16 Cr 16 12 9 Fe 13 30 18 Se IV 6 33 18 Ti 12 17 31 V 12 16 9 W VI 2 24 13 Mo 18 13 0 LSDb (P < 0.05) 1.1 0.6 2.1 a p-glucosaminidase activity of non-treated soils (mgp -nitrophenol kg"1 soil h"1); (Clarion = 90; Nicollet = 88; Canisteo = 36). b LSD = least significant difference. Table 3 Activation of p-glucosaminidase activity in soils by trace elements Trace element Activation of p-glucosaminidase activity (%) in soils specified" Element Oxidation state Clarion Nicollet Canisteo Co II 37 56 68 Mg 15 20 6 Mn 7 22 5 B III 31 46 As V 14 23 13 LSD (P < 0.05) 0 1.8 0 " P-glucosaminidase activity of non-treated soils (mg p -nitrophenol kg"1 soil h '); (Clarion = 90; Nicollet = 88; Canisteo = 36). LSD = least significant difference. 220 CHAPTER 8 TILLAGE AND RESIDUE MANAGEMENT EFFECTS ON (3- GLUCOSAMINIDASE ACTIVITY IN SOILS A paper in press in Soil Biology & Biochemistry M. Ekenler and M. A. Tabatabai Abstract There is increasing interest in assessing the effects of tillage systems and residue management on biochemical processes, especially enzyme activities, of soils. This study was carried out to investigate the effects of three tillage systems (no-till, chisel plow and moldboard plow) and four residue placements (bare, normal, mulch and double mulch) on the activity of AT-acetyl-p- glucosaminidase (NAGase, EC 3.2.1.30) involved in C and N cycling in soils, from three replicated field plots. Tillage and residue management practices significantly affected organic C contents of 21 surface (0-15 cm) soil samples. The highest organic C values were obtained for soils under no-till/double mulch plots, whereas the lowest values were found in soils under no-till/bare and moldboard/normal plots. Results indicated that the NAGase activity values were significantly affected by tillage and residue management practices, being greatest in soils with no-till/double mulch and least with no-till/bare and moldboard/normal. The activity of NAGase was greater in mulch-treated plots 221 under no-till and moldboard plow, whereas mulching did not show a significant effect on the enzyme activity in soils under chisel plow. Linear regression analyses showed that the activity of NAGase was significantly correlated with organic C in the surface soils, with correlation coefficient r = 0.89*** and with organic C content at different depths, with correlation coefficient r = 0.97***. The NAGase activity values were significantly (r = 0.63**) correlated with the arylamidase activity values of soils, suggesting that tillage and residue management practices have similar impacts on the activities of these enzymes and, thus, on N cycling in soils. As with organic C, the activity of this enzyme decreased markedly with increasing soil depth in no-till/ double mulch-treated soil. Keywords: Soil enzymes; Tillage systems; Residue management; N cycling; Soil N mineralization AT-Acetyl-P-D-glucosaminidase (NAGase, EC 3.2.1.30) catalyzes the hydrolysis of 2V-acetyl-|3-D-glucosamine residues from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also named P-N-acetyl-D- hexosaminide AT-acetylhexosaminohydrolase (EC 3.2.1.52) because it cleaves the amino sugar iV-acetyl-(3-D-galactosamine (Webb, 1984). NAGase is involved in chitin degradation in soils. Information on the effects of tillage and residue management on this enzyme is necessary because chitin is one of the most 222 abundant biopolymers on the earth serving as an important transient pool of organic C and N in soils (Wood et al., 1994), and amino sugars comprise about 5 to 10% of organic N in soils (Stevenson, 1994). Recent study in our laboratories showed that cropping systems have an important effect on NAGase activity, and that the activity of this enzyme is significantly correlated with N mineralization in soils of two long-term cropping systems in Iowa, with r values of 0.79*** and 0.84*** (Ekenler and Tabatabai, 2002). These results suggested that NAGase activity could be used as an index of N mineralization in soils. Soil management practices change the soil physical, chemical and biological environment, consequently affecting crop growth. It has been well documented that tillage causes soil disturbance by altering the distribution of organic matter and plant nutrients in soils (Dick, 1984; Dick, 1992; Christensen, 1996; Curci et al., 1997; Kandeler et al., 1999a). Because changes in soil physical and chemical properties are very slow processes, the modification in soil biological properties by soil disturbances should receive special attention because of more rapid response to ecological changes (Christensen, 1996). Tillage systems usually do not significantly alter total soil organic C and N (Carter and Rennie, 1982). Soil management practices, however, impact the microbial dynamics and their biochemically mediated processes by affecting the quality and quantity of plant residue in soils, their distribution through the soil profile and changing nutrient inputs (Christensen, 1996). Biological environment near the soil surface with no-till is cooler and wetter compared with 223 conventional tillage practices, and as a consequence, organic C and N reserves are concentrated close to the soil surface. Therefore, soil microbial population is stimulated by greater organic matter and water content (Dick, 1992; Doran et al., 1998). Because soil biological properties respond to soil disturbance more rapidly, enzyme activities can be used as early indicators of changes in soil properties affected by tillage (Frankenberger and Dick, 1983; Curci et al., 1997; Kandeler et al., 1999b). Kandeler et al. (1999a) investigated the influence of tillage on selected microbial properties in soils over a period of 8 years. They found that xylanase activity was significantly greater in the 0 to 10-cm soil surface of the reduced and minimum tillage systems within the first year of the experiment, whereas significant treatment effects on the microbial biomass, N mineralization and potential nitrification were observed after a 4-year period. The response of microbial biomass and enzyme activities to different tillage systems and residue management have been investigated by Deng and Tabatabai (1996a,b, 1997), Ahl et al (1998), and by Kandeler et al (1999a). Their results indicated that the activities of glycosidases (a-and (3-glycosidases and a- and (3-galactosidases), amidohydrolases (L-asparaginase, L-glutaminase, amidase, urease), phosphatases (acid and alkaline phosphatases and phosphodiesterase) and arylsulfatase were generally greater in soils under no-till/double mulch plots compared with those under other tillage and residue management systems investigated (Deng and Tabatabai, 1996a,b, 1997). Other results showed that 224 alkaline phosphates, xylanase, protease and invertase activities in particle-size fractions were mostly affected by the type of tillage and to a lesser extent by the date of sampling (Kandeler et al., 1999a). Because of the importance of the recently characterized P-glucosaminidase in soils (Parham and Deng, 2000), and because no information is available on the effects of soil management practices on the activity of this enzyme in soil, this work was carried out to investigate the impacts of three tillage systems (no-till, chisel plow and moldboard plow) and four residue placements (bare, normal, mulch and double mulch) on the P-glucosaminidase activity in soils. Unless otherwise specified, surface soil samples (0-15 cm) were obtained from the Lancaster Experiment Station in Wisconsin. The soil, Palsgrove silt loam (fine-silty, mixed, mesic superactive Typic Hapludalf), contained 17% clay and 6% sand. The experiment was initiated in 1981. The tillage systems studied were no-till, chisel plow and moldboard plow. The crop residue treatments were as follows: (1) Bare, with crop residue removed before planting (no prior tillage); (2) normal, with no removal or addition of crop residue; (3) mulch, with crop residue added after primary tillage; and (4) double mulch, with crop residue added to achieve double the normal level of crop residue. The crop residues in all cases were corn stalks. The area had been in continuous corn since 1971 (at least 10 year before the tillage experiments were initiated) and was maintained in continuous com during the study. The design and location of the experiments are described by Swan et al. (1987a, b). For this 225 study, 21 soil samples were taken from three replicated plots under different treatments in May 1991 (10 years after the tillage and crop residue treatments were initiated) by pooling at least eight cores (0-15 cm) of each plot. Samples of a fourth field replicate were highly acidic because of being the furthest from a gravel road, where nitrification of the ammoniacal N caused the acidity. The samples of this fourth replicate were not used in this study. In addition, nine soil samples were taken from 0-5, 5-10 and 10-15 cm of the no-till/ double mulch- treated plots. When not in use, the field-moist samples were stored at 4°C. Periodic assay of several hydrolases showed that enzyme activities were not changed after storing the soil samples in these conditions for several years. Organic C was determined by the Mebius (1960) method on < 80-mesh (180-^m) samples (Table 1). The results reported are averages of duplicate analyses and are expressed on a moisture-free basis. Moisture was determined by weight loss after drying at 105°C for 48 h. (3-Glucosaminidase activity was assayed by the method described by Parham and Deng (2000) on < 2-mm mesh field-moist surface samples and air- dried samples of the soil profiles. The method used for p-glucosaminidase activity involves colorimetric determination of the p-nitrophenol (CeHôNOs, FW = 139.1) produced when field-moist soil is incubated at 37°C for 1 h with acetate buffer (pH 5.5) andp-nitrophenyl iST-acetyl- P-D-glucosaminide solution. The enzymatic reaction involved is as follows: 226 HO -R + H20 HO p-Glucosaminidase NH *" I c=o I CH3 p-Nitrophenyl-AT-acetyl-P-D-glucosaminide c=o I CH3 N-acetyl-P-D-glucosaminide p-Nitrophenol For all data points shown in Fig. 2, the differences between duplicate values obtained in the analysis or assays were smaller than the symbol size. The procedure used for assay of arylamidase activity in soils was that described by Acosta-Martmez and Tabatabai (2000). The method involves quantitative extraction and colorimetric determination of the p -naphthylamine (C10H9N, FW = 143.2) produced when field-moist soil (on an oven-dried basis) is incubated at 37°C for 1 h with THAM buffer (pH 8.0) and L-Leucine P- 227 naphthylamide. Statistical analyses of the data were performed by using the Statistical Analysis System (SAS) (Barr et al., 1976). Results indicated that the activity of P-glucosaminidase in soils was significantly affected by the different tillage and residue management practices studied, with the highest activity values obtained under no-till/double mulch and least under no-till/bare and moldboard plow/normal (Fig. 1). Enzyme activity in soils with no-till/double mulch was significantly greater than those in other treatments, while the lowest activity was measured in soils under no-till/bare. The second highest activity was obtained in soils under moldboard plow/mulch treatment. Mulching had a significant impact on the NAGase activity in soils under the no-till and moldboard plow, whereas there was no significant difference between the effect of normal and mulch-treated plots with chisel plow. The enzyme activity in no-till/normal was comparable with those in chisel plow/normal. Change in soil organic C contents and distribution under different tillage and residue management practices influenced microbial populations and their activity through changes in substrate supply, water relations and associated soil physical changes (Doran et al., 1998). Mulching had a significant impact on NAGase activity in soils under no-till and moldboard plow. Work by Doran (1980) showed that the populations of bacteria, actinomycetes and fungi were 2 to 6 times greater as a result of mulch treatments. Reduced C is the main energy source for heterotrophic microorganisms; thus, it is not surprising to find higher microbial biomass and enzyme activities in soils with higher 228 organic C values. The number and activity of organisms in soils are not only governed by the availability of organic C and nutrients (McGill et al., 1986), but also by the physical and chemical environment (Elliot et al., 1980). Dick et al. (1988) reported a very significant effect of residue management on soil pH; thus, the catalytic efficiency and possibly the level of accumulation of a given enzyme could be changed by residue treatments. NAGase activity was highly significantly correlated with organic C contents of surface soils as follows: Y = - 51.1 + 78.3 X, with r value = 0.89***, where Y is NAGase activity in mg p-nitrophenol released kg*1 soil h l, and X is percentage of organic C, and samples from different depth of the no-till/double mulch plots as follows: Y = -11.0 + 27.2 X, with r value = 0.97***. Because the soil samples obtained from different depths of the no-till/double mulch plots were air dried before use, and because Parham and Deng (2000) showed that air drying reduces the NAGase activity, the results reported for the samples from different depths of the no-till/double mulch plots are perhaps under estimated. The results from the present study supported the finding of Deng and Tabatabai (1996 a, b; 1997), showing that glycosidases, phosphatases, amydohydrolases, and arylsulfatase enzyme activities were significantly correlated with soil organic C. Dick et al. (1988) suggested that higher levels of organic C stimulate microbial activity and, therefore, enzyme synthesis. In 229 addition, the higher organic matter levels in the residue treatment may provide a more favorable environment for accumulation of enzymes in the soil matrix, because organic constituents are thought to be important in the formation of stable complexes with free soil enzymes (Dick et al., 1988). Because it has been suggested that p-glucosaminidase and arylamidase are two of the major enzymes involved in N mineralization in soils (Acosta- Martinez and Tabatabai, 2000; Parham and Deng, 2000), we studied the relationship between the activities of those two enzymes. The activities of these two enzymes were highly correlated: Y = 18.4 + 0.82 X, with r value = 0.63**, where Y is NAGase activity in mg p-nitrophenol release kg-1 soil h*1, and X is arylamidase activity in mg P-naphthylamine produced kg*1 soil h l. Results of the no-till/double mulch treatment deviated from this relationship; proportionally greater P-glucosaminidase activity was found than arylamidase activity. These results suggest that tillage and residue management practices have similar effects on the activities of these enzymes. These findings support those reported by Acosta-Martinez and Tabatabai (2001a) showing that the activity of arylamidase was significantly correlated with the activity of amidohyrolases, other enzymes involved in N mineralization in soils. This is because arylamidase is involved in releasing amino acids that are hydrolyzed by amidohydrolases from soil organic matter (Acosta-Martinez and Tabatabai, 2001b). The significant correlations among the activities of 15 enzymes reported 230 by Deng and Tabatabai (1996a, b; 1997) and Acosta-Martinez and Tabatabai (2001a) in addition to the findings reported here suggest that tillage and residue management practices have similar trends on the activities of the enzymes involved in C, N, P and S cycling in soils. The activity of P-glucosaminidase decreased with increasing depth in the surface (0-15 cm) of the no-till/ double mulch treatment (Fig. 2) and reached the residual enzyme activity (least activity values) at 5-10 cm in the plow layer in soils of the three replicated plots. This decrease in NAGase actiivty was associated with a decrease in organic C content (Table 1). The differences in patterns of the NAGase activity among the soils of three replicated plots could be the result of differences in pH and organic C distribution in the plow layer. Organic C of soils usually decreases with soil depth, and the organic C content of the studied soils also followed the same pattern. Biological activity and organic C and N reserves were concentrated near the soil surface (0-7.6 cm) with the no-till systems, and accumulated crop residue and organic C are substrates for soil microorganisms (Doran et al., 1998; Kandeler et al., 1999). Our results support the previous information showing that the activities of arylamidase and amidohydrolases, glycosidases, phosphatase and arylsulfatase are significantly correlated with soil organic C content (Deng and Tabatabai, 1996a, b; 1997; Acosta-Martinez and Tabatabai, 2001b), suggesting that organic matter plays an important role in protecting soil enzymes, thus, further supporting the 231 hypothesis that enzymes are immobihzed in a three-dimensional network of clay and humus complexes (McLaren, 1975). In summary, this study showed that p -glucosaminidase activity is significantly affected by soil disturbance, tillage and residue management practices. Because this is one of the key enzymes that play a key role in N mineralization in soils, the activity of this enzyme can provide a potential tool as an N mineralization index of soils and for studies of ecosystem health, especially when N cycling is of interest. Acknowledgements Journal Paper no. J-19779 of the Iowa Agric. and Home Econ. Exp. Stn., Ames. Projects 3338, 4184 and 6518. This work was supported in part by the Biotechnology By-products Consortium of Iowa. References V. Acosta-Martinez, M.A. Tabatabai. Arylamidase activity of soils. Soil Science Society of America Journal 64 (2000) 215-221. V. Acosta-Martinez, M.A. Tabatabai. Arylamidase activity in soils: effect of trace elements and relationships to soil properties and activities of amidohydrolases. Soil Biology & Biochemistry 33 (2001a) 17-23. V. Acosta-Martinez, M.A. Tabatabai. Tillage and residue management effects on arylamidase activity in soils. Biology and Fertility of Soils 34 (2001b) 21-24. 232 A. Ahl, R.G. Joergensen, E. Kandeler, B. Meyer, V. Woehler. Microbial biomass and activity in silt and sand loams after long-term shallow tillage in central Germany. Soil & Tillage Research 49 (198) 93-104. A.J. Barr, J.H. Goodnight, J.P. Sail, J.I. Helwig. A user's guide to SAS. 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Simulating the effects of soil depth and climatic factors on corn yield. Soil Science Society of America Journal 51 (1987a) 1025-1032. J.B. Swan, E C. Schneider, J.F. Moncrief, W.H. Paulson, A.E. Peterson. Estimation of corn growth, yield, and grain moisture from air growing degree days and residue cover. Agronomy Journal 79 (1987b) 53-60. E.G. Webb. Enzyme Nomenclature. Academic Press, New York, 1984. C.W. Wood, H.A. Torbert, H.H. Rogers, G.B. Runion, S.A. Prior. Free-air CO2- enrichment effects on soil carbon and nitrogen. Agricultural and Forest Meteorology 70 (1994) 103-116. Table 1 Effect of tillage and residue management on soil organic C Treatments Rep I Rep II Rep III Mean LSD" P< 0.05 % Organic C No-till/Bare 1.29 1.03 1.12 1.15 0.06 No-till/Normal 1.57 1.42 1.36 1.45 0.05 No-till/2X Mulch 1.91 2.07 1.72 1.90 0.03 Chisel/Normal 1.60 1.37 1.42 1.46 0.03 Chisel/Mulch 1.45 1.41 1.48 1.45 0.07 Moldboard plow/Normal 1.21 1.09 1.07 1.12 0.04 Moldboard plow/Mulch 1.49 1.17 1.44 1.37 0.07 LSD P< 0.05 0.03 0.03 0.04 0.03 Depth of no-till/2X Mulch 0-5 cm 2.68 3.01 2.98 2.89 0.07 5-10 cm 1.91 1.29 1.03 1.41 0.02 10-15 cm 1.43 1.00 0.95 1.13 0.02 LSD P < 0.05 0.09 0.09 0.08 0.09 ° LSD, least significant difference. 237 120 S f îoo - NTB NTN NT2M CPN CPM MPN MPM Fig. 1. Effect of tillage and residue management on the P-glucosaminidase activity (averages of three replicated field-plots) in soils. Different letters indicate significantly different means at P < 0.05, least significant difference test. NTB, no-till/bare; NTN, no-till/normal; NT2M, no-till/double mulch; CPN, chisel plow/normal; CPM, chisel plow/mulch; MPN, moldboard plow/normal; MPM, moldboard plow/mulch. 238 P-Glucosaminidase activity (mg p-Nitrophenol kg"1 soil h*1) Fig. 2. Distribution of P-glucosaminidase activity with soil depth in the plow layer (0-15 cm), no-till/double mulch treatment. Rep, replicate. 239 CHAPTER 9 P-GLUCOSAMINIDASE ACTIVITY AS AN INDEX OF NITROGEN MINERALIZATION IN SOILS A paper submitted to Communications in Soil Science and Plant Analysis Mine Ekenler and M. A. Tabatabai ABSTRACT iV-Acetyl-p-D-glucosaminidase (NAGase, EC 3.2.1.30) is one of three enzymes that catalyze the hydrolysis of chitin. This hydrolysis is important in C and N cycling in soils because it participates in the processes whereby chitin is converted to amino sugars, which are major sources of mineralizable N in soils. This study investigated the relationship between N mineralization indexes and activity of P-glucosaminidase in soils from six agroecological zones of the North Central region of the United States. N mineralization was studied by using short-term laboratory incubations under aerobic and anaerobic conditions and by hydrolysis of soil organic N by steam distillation with chemical reagents. Enzyme activity was assayed at its optimal pH value. The amounts of N mineralized by all the biological and chemical methods studied were significantly correlated with P-glucosaminidase activity, with r values of 0.73*** and 0.76*** for the amounts of NH^-N released by steam distillation with PO4- B4O7 for 4 or 8 min, respectively; of 0.69*** and 0.74*** for the 240 amounts of NH^-N released with NazI^O? for 4 or 8 minutes, respectively; of 0.47*** for the amounts of inorganic N produced under aerobic incubation; of 0.80*** for amounts of inorganic N produced by incubation under anaerobic conditions of field-moist soils; and of 0.86*** for anaerobic incubation of air-dried soils. There was a significant correlation between P-glucosaminidase activity and organic C (r = 0.56***), total N (r = 0.60***), and fixed NH4"1" of the soils (r = 0.79***). The amounts of inorganic N released by the methods, with the exception of those under aerobic incubation, were significantly (P < 0.001) correlated with organic C and total N contents of the soils, with r values ranging from 0.65*** to 0.78*** and from 0.62*** to 0.80*** for organic C and total N, respectively. INTRODUCTION Nitrate contamination of surface and ground water is of environmental concern. One of the major sources is the high input of N fertilizers in intensive crop production (1). Therefore, there is increasing need to develop rapid method for estimating the N mineralization potential of soils. Many biological and chemical methods have been proposed as N mineralization indexes (2-4). Some of these methods were accepted and widely used as most reliable tools to estimate N mineralization in soils. However, most of those methods are not simple and rapid, and to our knowledge, little information is available on the relationships 241 between the N mineralization values obtained by those methods and assay of the enzymes that catalyze the reaction involved. Mineralization of organic N compounds in soils are mediated by enzymes, which are predominantly released by soil microorganisms (5). Studies of enzyme activities should provide insight into biochemical processes in soils by which the N is mineralized. Such assays, if successful, can be used as sensitive biochemical indexes (6). Enzyme activities have also been suggested as early indicators of changes in soil properties induced by soil management processes because of their rapid response to change in management practices, their relationship to soil biology and ease and accuracy of the assay (7, 5). A strong correlation between soil microbial biomass and gross N mineralization has been reported (8), and microbial biomass of soils was suggested to predict net N mineralization (9). Studies on the characterization of active N pools in soils under different cropping systems, however, suggested that N mineralization in soils is predominantly controlled by biochemical process such as enzyme activities other than total microbial activities (10). Those studies reported a significant correlation between active N pools and amidase activity, which has an important role in N mineralization in soils. iV-Acetyl-p-D-glucosaminidase (NAGase, EC 3.2.1.30) is the enzyme that catalyzes the hydrolysis of N-acetyi p-D glucosamine residue from the terminal non-reducing ends of chitooligosaccharides. This enzyme is also classified as P- hexosaminidase (EC 3.21.52) by the International Union of Biochemistry 242 because it cleaves the amino sugar iV-acetyl-P -D-galactosamine (11). P- Glucosaminidase involves in chitin degradation in soils. Evaluation of the relationship between the activity of this enzyme and indexes of N mineralization is necessary, because chitin is one of the most abundant biopolymer on the earth serving as an important transient pool of organic C and N in soils (12). Amino sugars, the hydrolysis product of this enzyme, are major sources of mineralizable N in soils (13-15). A recent study in our laboratories showed that cropping systems have important effect on P-glucosaminidase activity, and that the activity of this enzyme is significantly correlated with N mineralization in soils of two long-term cropping systems in Iowa, with r values of 0.79*** and 0.84*** (16). Tillage and residue management significantly affect the activity of this enzyme in soils (17). It has been reported that chitin induces chitinase activity (18). Not much work is available about the role of this enzyme in N mineralization, but studies indicated that p-glucosaminidase is one of the enzymes required for chitin degradation; its activity was negatively correlated with N immobilization and decomposition rate for birch sticks decomposing at eight contrasting sites (19). These results suggest that p-glucosaminidase activity could be used as an index of N mineralization in soils. Therefore, this study was carried out to investigate the relationships between p- glucosaminidase activity and N mineralization indexes in 56 surface soils from six agroecological zones of the North Central region of the United States. 243 MATERIALS AND METHODS Soils and Their Properties The soils used in this study were surface (0-15 cm) soils collected from cultivated fields in six states of the North Central region of the United States to include a wide range of chemical, physical and biological properties (Table 1). pH was determined on air-dried samples (<2mm) by using a combination glass electrode (soil:0.01 M CaCls ratio = 1:2.5). Organic C was determined on <180 |im samples by the Mebius method (20), total N by a Semimicro-Kjeldahl method (21), and inorganic N by a steam distillation method (22). Fixed NH4+ was determined by the method of Silva and Bremner (23). N Availability Indexes The biological and chemical methods used as indexes (4, 22, 24-26) of N mineralization are summarized in Table 2. P-Glucosaminidase Activity P-Glucosaminidase activity was assayed by the method described by Parham and Deng (27). It involves colorimetric determination of the p-nitrophenol released 244 when field-moist soil is incubated with p-nitrophenyl-N-acetyl-p-D- glucosaminide and acetate buffer pH 5.5 for 1 h. The enzymatic reaction is as follows: HO —"-°x rjA \ \ >0-R + HiO V " p-Glucosaminidase NH I C=0 I CH3 p-Nitrophenyl-iV-acetyl-P-D-glucosaminide HO O HO °H +R-OH R=—< '—N0 HO 2 NH I c=o I CH3 2V-acetyl-p-D-glucosaminide p-Nitrophenol Statistical Analyses Statistical analyses were performed according to the general linear model procedure of the SAS system (28). For all data points shown in Figs 1-4, the differences between duplicate values obtained in the analysis or assays were smaller than the point size. 245 RESULTS AND DISCUSSION P-Glucosaminidase Activity and N Mineralization Indexes Results showed that p-glucosaminidase activity was significantly correlated with the amounts of inorganic N produced upon incubation of soils at 30°C under aerobic conditions for 14 days (r = 0.47***) or anaerobic conditions (field-moist soils, r = 0.80***; and air-dried soils, r = 0.86***) (Fig. 1). The amounts of inorganic N produced on incubation of field-moist soils under anaerobic conditions were about 25% of those produced in air-dried soils incubated under the same conditions. It has been recommended that anaerobic incubation be used for estimation of plant available N in field-moist or air-dried soils (29, 30). The highly significant linear relationships between the amounts of inorganic N produced by those methods and the P-glucosaminidase activity suggest the significant role that this enzyme play in the reactions involved in N mineralization. The relationships between the activity of p-glucosaminidase and the amounts of NH4+-N produced on steam distillation of field-moist soils with the reagents described in Table 2 are shown in Figs. 2 and 3. The correlation coefficients were highly significant (P < 0.001), with the r values as follow: 0.73*** and 0.76*** for the amounts of NH^-N released with PO4-B4O7, 0.69*** and 0.74*** for amounts released with Na2B4Û7, at 4 and 8 min, respectively, and 0.37** for the amounts released with hot KC1 extraction. 246 The results reported here agree with those recent published showing that activity of this enzyme is significantly correlated with the amount of N mineralized in 24 weeks of incubation at 30°C in soils of two long-term cropping systems in Iowa, with r values of 0.79*** and 0.84*** (16). Other studies reported a highly significant relationships (r = 0.60*** and 0.52***) between amidase activity and the cumulative amounts of N mineralized in field-moist soils, obtained from two long-term cropping systems in Iowa, incubated at 30°C for 24 weeks (10). Our results also support the previous findings showing a significant, positive correlation between N mineralization and activity of arylamidase, which plays a major role as the initial reaction-limiting step in mineralization of organic N in soils by catalyzing the hydrolysis of N-terminal amino acid from arylamides (31). Protease was also recommended as potential enzyme to predict gross N mineralization because of the strong correlation with N mineralization rate in soils treated with dairy shed effluent (32). Other recent work has shown that the annual mean of arginine ammonification assays were correlated with gross N mineralization values in four different agricultural fields in Denmark (33). Results also showed that the correlation coefficients of the linear relationship between P-glucosaminidase activity and the amounts of NH^-N produced on steam distillation of soil samples with PO4-B4O7 was greater than those released with hot KC1 extraction or with NasB407, suggesting P-glucosaminidase and PO4-B4O7 hydrolyze the same bonds of organic N in soils. 247 Total organic C and N, the amounts of N mineralized by short-term incubation under aerobic and anaerobic conditions, and by chemical hydrolysis were significantly inter-correlated (Table 3). P-Glucosaminidase Activity in Relation to Soil Properties P-Glucosaminidase activity was significantly (P < 0.001) correlated with the organic C and N contents and the amount of fixed NH4+-N of soils, with r values 0.56*** and 0.60***, and 0.79***, respectively (Fig. 4). It is well established that enzyme activities are strongly correlated with organic C and N contents of soils (16, 17, 34). The significant positive correlation between p-glucosaminidase activity and + fixed NH4 was unexpected because it is generally accepted that fixed NH4+ in soils is in inorganic form. If true, then no such relationship should exist between p- + glucosaminidase activity and fixed NH4 . The fact that there is such a relationship (r = 0.79***) suggests that, if not all, at least part of the fixed NH4+ is organic in nature. This finding support the hypothesis that enzyme proteins in soils are stabilized by internal clay surfaces (35). ACKNOWLEGMENTS This work was supported, in part, by the Biotechnology By-products Consortium of Iowa. 248 REFERENCES 1. Johnson, S.L.; Adams, R M.; Perry, G.M. The On-Farm Costs of Reducing Groundwater Pollution. Am. J. Agric. Econ. 1991, 73, 1063-1073. 2. Keeney, D R. Nitrogen-Available Indices. In Methods of Soil Analysis, Part 2: Chemical and Microbiological Properties; Page, A.L., Miller, R.H., Keeney, D R., Eds.; ASA-SSSA: Madison, WI, 1982, 711-733. 3. 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Ammonium Production in Soils Under Waterlogged Conditions as an Index of Nitrogen Availability. Nature 1964, 201, 951-952. 26 Oien, A.; Selmer-Olsen, A.R. A Laboratory Method for Evaluation of Available Nitrogen in Soil. Acta Agric. Scand. 1980, 30, 149-156. 27. Parham, J. A.; Deng, S.P. Detection, Quantification and Characterization of P-Glucosaminidase Activity in Soil. Soil Biol. Biochem. 2000. 52:1183-1190. 28. SAS Institure. SAS Procedure Guide. Version 6.12 ed. SAS Institute Inc., Cary, NC. 29. Keeney, D R.; Bremner, J.M. Comparison and Evaluation of Laboratory Methods of Obtaining an Index of Soil Nitrogen Availability. Agronomy Journal. 1966, 58, 498-503. 30. Geist, J.M. Nitrogen Response Relationships of Some Volcanic Soils. Soil Sci. Soc. AM. J. 1977, 41, 996-1000. 252 31. Dodor, D E.; Tabatabai, M.A. Effects of Cropping Systems and Microbial Biomasson Arylamidase Activity in Soils. Biol. Fertil. Soils 2002; 35, 253- 261. 32. Zaman, M.; Di, H.J.; Cameron, K.C.; Frampton, CM. Gross Nitrogen Mineralization and Nitrification Rates and Their Relationships to Enzyme Activities and the Soil Microbial Biomass in Soils Treated with Dairy Shed Effluent and Ammonium Fertilizer at Different Water Potential. Biol. Fertil. Soils 1999, 29, 178-186. 33. Bonde, T. A., Nielsen T. H., Miler, M., Sorensen, J. Arginine Ammonification Assay as Rapid Index of Gross N Mineralization in Agricultural Soils. Biol. Fertil. Soils 2001, 34, 179-184. 34. Deng, S.P.; Tabatabai, MA. Effect of Tillage and Residue Management on Enzyme Activities in Soils: I. Amidohydrolases. Biol Fertil Soils 1996. 22:202-207. 35. Boyd, S. A., Mortland, M. M. Enzyme Interaction with Clays and Clay- Organic Matter Complexes. In Soil Biochemistry, Bollag, J-. M., Stotzky, G. Eds., Marcel Dekker, New York, 1990, Vol. 6, 1-28. Table L Selected chemical properties and p-glucosaminidase activity of the soils used P-Glucosaminidase No, of pH Total N Organic C Activity" State Soil series soils range mean range mean range mean range m g kg soil - IL Raub 4 5,7-6.4 6.0 1.4-1.5 1.48 11.9-24.0 18.7 37-44 41 KS Crete 7 5.3-6.3 5.9 1.0-1,2 1.08 11.7-15.6 13.3 18-57 44 KS Pratt1 6 4.6-4.9 4.7 0.6-1.0 0.75 3.7-12.5 8.8 14-34 23 KS Pratt2 5 5.1-5.2 5.2 1.1-1.6 1.38 12.8-18.0 15.5 34-47 42 MI Capacl 4 6.1-7,0 6.6 1.9-2.0 1.94 20.2-23.7 22.0 27-36 31 MI Capac2 4 6.1-6.9 6.7 1.6-2.1 1.86 20.2-23.7 21.8 31-36 34 MN Nicollet 1 4 5.2-5.4 5.3 1.5-2,6 1.99 26.5-29.5 28.3 62-78 70 MN Nicollet2 4 5.3-5.4 5.3 2.5-2.7 2.60 28,2-30.7 29.8 76-84 70 MO Mexico 4 6.2-6.8 6.5 1.4-1.6 1.50 14.9-16,6 15.6 34-39 36 NE Sharpsburg 4 5.3-5.5 5.4 1.5-1.6 1.59 16.8-17.6 17.2 69-95 84 NE Othello 4 6.3-7.2 6.7 1.1-1.7 1.36 8.9-15.8 11.9 32-86 52 NE Hasings 4 5.3-5,6 5.4 1.4-1.5 1.43 16.1-16.6 16.2 46-69 54 NE Othello 2 4.7-4.8 4.7 0.6-0,7 0.68 7.6-8.3 8.0 14-17 16 8 mgp -Nitrophenol kg'1 soil h'\ Table 2, Biological and chemical indexes used Method Measurement performed Reference Biological indexes 10 g soil + 30 g quartz sand + 6 ml distilled water NH/ and N03' in 20 ml of the extract (24) incubated at 30°C for 14 days and inorganic N were determined by steam distillation (22) produced is extracted with 100 ml of 2 M KC1 5 g soil (air dried or field moist) + 12,5 ml distilled NH/ and N03 in 20 ml of the extract (25) water incubated at 30°C for 14 days and were determined by steam distillation inorganic N produced is extracted with 4 M KC1 Chemical indexes 4 g of field-moist soil + 40 ml 0.26 M Na3P04xl2H20 NH/ and N03" released are steam (4) + 0,066 M Na2B4O7xl0H2O (pH = 11,2) distilled for 4 or 8 min 4 g of field moist soil + 40 ml 0.066 M NH/ and N03 released are steam Na2B4O7xl0H2O (pH = 11,5) distilled for 4 or 8 min 4 g of field-moist soil + 40 ml 2 M KC1 NH/ and N03 released are steam (26) heated for 20 h at 80°C distilled Table 3. Correlation between indexes of N mineralization and some soil properties N index8 PB AI ANIM ANID KC1 SB C N PB 1.00 AI 0,42* 1.00 ANIM 0.84*** 0.26"' 1.00 ANID 0,93*** 0.32* 0.91*** 1.00 KC1 0.56*** 0.25ns 0.31* 0,33* 1.00 SB 0.95*** 0,43* 0.83*** 0.88*** 0.62*** 1.00 ^org 0.78*** 0.05"" 0.71*** 0.75*** 0.65*** 0.76*** 1.00 Norg 0.80*** 0.25"' 0.68*** 0.75*** 0.62*** 0.76*** 0.88*** 1.00 0 PB, Phosphate borate N; AI, Aerobic incubation; ANIM, Anaerobic incubation (field moist soil); ANID, Anaerobic incubation (air-dried soil); KC1, Hot KC1 N; SB, Sodium-borate buffer N; Norg, organic nitrogen; * and *** indicate significant at P < 0,05 and 0.001, respectively; ns, indicates not significant. 256 120 110 . Y = 4.18 +1.49 X Aerobic Incubation 100 r = 0.47*** 90 QD O ° 80 - o 0 70 C O O 60 cP o 50 °o o • <29r 40 Aoa? o 30 ro $ 20 o to 10 .a: 0 i 3 10 20 30 40 50 60 Y = -6.14 +5.12 X Anaerobic Incubation I r = 0.80*** field moist 9. 60 E 5 > "5 «(A •ocs E IC8 o 120 . cs. Y = 4.26 +1.01 X 110 Anaerobic Incubation r = 0.86*** air dried 100 O 90 0 80 o 70 o o o 60 / 50 o 9bfS/ 40 On 30 20 \/é& 0 10 0 20 40 60 80 100 120 Amount of N mineralized (mg N kg.-i. soil) Fig. 1. Relationships between P-glucosaminidase activity and the amounts of N mineralized in soils by biological indexes. 120 Y =12.9+ 1.29 X P04-B407 . Y = 5,90 + 0.88 X P04-B407 110 = 0.73*** mins) r = 0.76*** (8 mins) 100 r (4 <5> 90 CP 80 O OPO 70 o° 60 50 40 30 20 °o^o 10 0 i 10 20 30 40 SO 60 70 0 10 20 30 40 SO 60 70 80 90 100 110 120 Na B 110 Y = 5.24 +4.19 X Na2B407 Y = -8.55 + 2.53 X 2 4°7 100 r = 0.69*** (4 mins) . r = 0.74*** (8 mins) o o °o 90 ° Q9° 80 ° O o ojy 0 70 0 0% O Ofx/ 60 oo SO O OrCfuQ ° oojppb _ 40 QO ® 30 o o ^ 20 10 0 I 5 10 15 20 0 5 10 15 20 25 30 35 40 45 Amount of N mineralized (mg N kg"1 soil) Re hips between P-glucosaminidase activity and the amounts of N mineralized in soils by the indicated. 258 120 110 - Y =26.6 +2.34 X Hot KCl extraction r = 0.37** 100 O 90 o ° o cP 80 0 o o°1 o 70 O o o 60 % o o 50 1 O n/ 40 ° ° Amount of N mineralized (mg N kg*1 soil) Fig. 3. Relationships between P-glucosaminidase activity and the amounts of N mineralized in soils by hot KCl extraction. Y = 15,7+ 17.8 X Y = 11.0 +237 X Y = 3.98 + 0.33 X r = 0.56*** r = 0.60*** r = 0.79*** & 40 50 100 150 200 250 + 1 Organic C (mg kg1 soil) Total N (mg kg1 soil) Fixed NH4 (mg kg' soil) Fig. 4. Relationships between p-glucosaminidase activity and selected soil properties. 260 CHAPTER 10 GENERAL CONCLUSIONS The objectives of this study were: (1) to assess the effects of liming and tillage systems on selected soil chemical properties, microbial biomass, and the activities of 15 enzymes involved in C, N, P, and S cycling in soils, (2) to determine whether these response are consistent across four long-term management sites in Iowa, (3) to assess the sensitivity of the 15 enzymes to liming (changes in soil pH) and different tillage systems, (4) to study the relationship between microbial biomass C (Cmic) and N (Nmic) and the activities of the enzymes, (5) to study the impacts of crop rotations and N fertilization on the activity of b-glucosaminidase, an enzyme involved in C and N cycling in soils, (6) to assess the relationships between the activity of b-glucosaminidase and N mineralization in soils, (7) to assess the relationship between the activity of b- glucosaminidase and Cmic and Nmic, (8) to study tillage and residue management practices effects on b-glucosaminidase activity, (9) to determined the effects of trace elements on the activity of this enzyme, and (10) to study the relationships between b-glucosaminidase activity and selected biological and chemical indexes of available N in soils. 261 The findings can be summarized as follows: 1. Studies revealed that the pH values of the soils were significantly varied due to lime application; however, organic C and N concentration were not affected at each site. In general, Cmic was significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.68 (P < 0.01) at the NWRC site under no-till (0-5 cm) to 0.81 (P < 0.001) at the NWRC site under no-till (0- 15 cm). The Nmic values were also significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.66 (P < 0.001) at the SERC site to 0.92 (P < 0.001) at the SWRC site. Lime application increased the activities of glycosidases significantly (P < 0.001). Simple correlation coefficients between the activities of enzyme and soil pH values ranged from 0.51 (P < 0.05) for the activity of a-glucosidase at the NWRC site, no-till (0-5 cm) to 0.98 (P < 0.001) at the SWRC site. Lime application did not significantly affect the 1 l specific activities (g p-nitrophenol released kg- Corg h ) of glycosidases. In general, their order of sensitivity (A activity/ A pH) to changes in soil pH was consistent across the study sites and it was as follow: p-glucosidase > (3- glucosaminidase > (3-galactosidase > a-galactosidase > a-glucosidase. The enzyme activities were greater in soils under chisel plow than under the other tillage systems studied, and they were greater in 0-5 cm samples than those in the 0-15 cm samples under no-till system. Most of the enzyme activities were significantly and positively correlated with Cmic and Nmic. Among the 262 glycosidases studied, P-glucosidase and p-glucosaminidase were the most sensitive to soil management practices. 2. With the exception of acid phosphatase, which was significantly (P < 0.001) but negatively correlated with soil pH (r ranged from -0.65** to - 0.98***), the activities of alkaline phosphatase, phosphodiesterase and arylsulfatase were significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.65** to 0.99*** for alkaline phosphatase, from 0.79*** to 0.97*** for phosphodiesterase, and from 0.66*** to 0.97*** for arylsulfatase. The A activity/ A pH values were calculated to determine the sensitivity of each enzyme to changes in soil pH. Acid phosphatase was the most sensitive and arylsulfatase the least sensitive to changes in soil pH. Activities of the enzymes were greater in the 0-5 cm depth samples than those in 0-15 cm samples under no-till treatment. Enzyme activities were mostly significantly (P < 0.001) and positively correlated with Cmic, with r values ranging from 0.28 (not significant) to 0.83*** and with Nmic, with r values ranging from 0.31 (not significant) to 0.94***. Liming and tillage systems significantly affected the activities of some enzymes but not others, as was evident from the specific 1 1 activity values (g of p-nitrophenol released kg- Corg h ). 3. The activities of arylamidase and amidohydrolases were significantly (P < 0.001) and positively correlated with soil pH, with r values ranging from 0.42* to 0.99*** for arylamidase, 0.81*** to 0.97*** for L- asparaginase, 0.62*** to 0.97*** for L-glutaminase, 0.61*** to 0.98*** for 263 amidase, 0.66** to 0.96*** for urease, and 0.80*** to 0.99*** for L-aspartase. The A activity/ A pH values were calculated to assess the sensitivity of the enzymes to changes in soil pH. The order of the sensitivity of enzymes was as follows: L-glutaminase > L-asparaginase > amidase > arylamidase > urease > L- aspartase. The enzyme activities were greater in the samples of the 0-5 cm depth than those of the 0-15 cm samples under no-till treatment. Most of the enzyme activities were significantly (P < 0.001) and positively correlated with Cmic and Nmic. Lime application significantly affected the specific activities of the six enzymes studied. 4. The activities of b-glucosaminidase were significantly affected by crop rotations (p < 0.001) and N fertilization (P ranging from 0.05 to 0.001). Generally, the highest enzyme activity was obtained in soils under 4-year corn- oats-meadow rotations taken under meadow, and the lowest continuous mono- cropping systems. The activity of this enzyme was significantly correlated with Corg (r ranging from 0.42** to 0.76***), Norg (ranged from not significant at one site one year to r = 0.76***), Cmic (r ranging from 0.44** to 0.71***), and Nmic (r ranging from 0.33* to 0.76***) in soils, and with cumulative N mineralized (r > 0.84*** and r > 0.79*** at the NERC and CWRC sites, respectively). 5. The activity of p-glucosaminidase was inhibited by 18 and activated by 5 of the trace elements tested. The inhibition values ranged from 0 to 73%, depending on the trace element and the soil used. In general, Ag(I) and Hg(H) were the most effective inhibitors. Other metals that inhibited the 264 (3-glucosaminidase activity in soils were Cu(I), Ba(H), Cu(II), Ni(II), Pb(II), SndD, Zn(ID, As(IID, Cr(m), Se(IV), Ti(IV), V(IV), W(VD and Mo(VI). At the concentration tested, A1(HI), Fe(II), and Fe(III) also inhibited the activity of this enzyme. The elements Co (II), Mg (II), Mn (II), B (III) and As (V) activated this enzyme by values ranging from 4 to 68% in three soils. 6. Tillage and residue management practices significantly affected organic C contents of the surface (0-15 cm) soils studied. The highest organic C values were obtained for soils under no-till/double mulch plots, whereas the lowest values were found in soils under no-till/bare and moldboard/normal plots. Results indicated that the b-glucosidase activity values were significantly affected by tillage and residue management practices, being greatest in soils with no-till/double mulch and least with no-till/bare and moldboard/normal. The activity of b-glucosidase was greater in mulch-treated plots under no-till and moldboard plow, whereas mulching did not show a significant effect on the enzyme activity in soils under chisel plow. The activity of this enzyme was significantly correlated with organic C in the surface soils, with correlation coefficient r = 0.89*** and with organic C content at different depths, with correlation coefficient r = 0.97***. The activity values were significantly (r = 0.63**) correlated with the arylamidase activity values of soils. As with organic C, the activity of this enzyme decreased markedly with increasing soil depth in no-till/ double mulch-treated soil. 265 7. The amounts of N mineralized by all the biological and chemical methods studied were significantly correlated with P-glucosaminidase activity, with r values of 0.73*** and 0.76*** for the amounts of NH4+-N released by steam distillation with PO4-B4O7 for 4 or 8 min, respectively; of 0.69*** and 0.74*** for the amounts of NH4+-N released with Na?B407 for 4 or 8 minutes, respectively; of 0.47*** for the amounts of inorganic N produced under aerobic incubation; of 0.80*** for amounts of inorganic N produced by incubation under anaerobic conditions of field-moist soils; and of 0.86*** for anaerobic incubation of air-dried soils. There was a significant correlation between b-glucosaminidase activity and organic C (r = 0.56***), total N (r = 0.60***), and fixed NH4+ of the soils (r = 0.79***). The amounts of inorganic N released by the methods, with the exception of those under aerobic incubation, were significantly (P < 0.001) correlated with organic C and total N contents of the soils, with r values ranging from 0.65*** to 0.78*** and from 0.62*** to 0.80*** for organic C and total N, respectively. 8. These studies showed that soil management practices including, liming, tillage and residue management, cropping systems and N fertilization significantly affect soil biological and biochemical properties, which may lead to changes in C, P, S and N cycling, including N mineralization and nitrification, in soils. 266 9. These studies also indicated that enzyme activities can provide a reliable-long term monitoring tool as early indicators of changes in soil properties induced by agricultural management practices. 10. The study suggested that p-glucosaminidase plays a major role in N mineralization in soils and is significantly affected by management practices; i.e., ecosystem function and health. 267 APPENDIX ADDITIONAL TABLES Table Al, Effect of liming on properties of soils (0-15 cm) at the Southeast Research Center, Crawfordsville, Iowa PH Inorganic N Microbial biomass Aglime Replication HzO CaClg Organic C Total N NH«-N 1 Z Organic N C„iic ^raic Cmic/org.C N,mic /org.N kg ECCE ha1 g kg'1 soil mg kg'1 soil- g kg ' —mg kg'1 soil— %— 0 1 5,56 4,84 26,1 2.4 5 35 2.36 137 7,8 0.52 0.33 0 2 5,68 4,91 26.8 2.4 4 33 2.36 184 8,9 0,69 0,37 0 3 5,64 4.87 26.2 2.1 4 31 2.07 139 5.5 0.53 0.26 5.63 4.87 26.4 2.3 4 33 2.26 153 7.4 0.58 0.32 1120 1 5,76 4,91 26,5 2.4 4 36 2.36 160 8.2 0.60 0.34 1120 2 5,78 4,97 26.4 2.3 4 30 2.27 170 8.4 0,64 0.37 1120 3 5.72 4,96 25.5 1.9 3 28 1.87 165 5.9 0,65 0.31 5.75 4.95 26.1 2.2 3 31 2.17 165 7.5 0.63 0.34 2240 1 5,96 5,05 24.7 2.7 4 29 2.67 165 9.9 0.67 0.37 2240 2 5.95 5,11 26.0 2.2 4 29 2.17 177 10.8 0.68 0.49 2240 3 6.07 5.24 24,9 2,3 2 23 2.28 170 7.2 0.68 0.31 5.99 5.13 25.2 2.4 3 27 2.37 171 9.3 0.68 0.39 4480 1 6,32 5,52 25.8 2.1 5 33 2.06 202 9.5 0.78 0.45 4480 2 5,98 5.16 26.4 2.5 5 27 2.47 172 9.1 0.65 0,36 4480 3 5,90 5.14 25,5 2.3 3 29 2.27 170 7,4 0.67 0.32 6.07 5.27 25.9 2.3 4 30 2.27 181 8.7 0.70 0.38 6720 1 6,50 5,68 24.8 2.2 4 28 2.17 239 11.2 0.96 0,51 6720 2 6,32 5.48 27.4 1.9 3 28 1.87 209 12.0 0.76 0.63 6720 3 6.39 5,53 26.2 2.5 3 28 2,47 201 9.7 0.77 0.39 6.40 5.56 26.1 2.2 3 28 2.17 216 11.0 0.83 0.51 1120 1 5.66 4,85 26.2 2,3 4 33 2.26 151 6.3 0,58 0.28 1120 2 5.50 4.80 26.6 2,5 5 33 2.46 194 5.1 0.73 0.20 1120 3 5.83 4.98 26.8 2.1 3 27 2.07 168 4.6 0.63 0.22 5.66 4.88 26.5 2.3 4 31 2.27 171 5.3 0.64 0.23 2240 1 5.94 5.10 25.2 1.9 5 28 1.87 178 8.9 0,71 0,47 2240 2 5.79 4.97 26.2 2.7 3 28 2.67 183 8,4 0.70 0,31 2240 3 5,86 5.10 26.2 2.3 2 28 2.27 183 7,0 0.70 0,30 5.86 5.06 25.9 2.3 3 28 2.27 181 8.1 0.70 0.36 560 1 5,78 5,16 26.4 2.4 3 31 2.37 189 8.0 0.72 0,33 560 2 5.90 5,10 25.2 2.7 2 23 2.68 178 7.0 0.71 0,26 560 3 5.81 4,99 25.6 2.5 3 25 2.47 175 8.9 0.68 0,35 5.83 5.08 25.7 2.5 3 26 2.51 181 7.9 0.70 0.32 * Mean values are in bold Table A2. Effect of liming on properties of soils (0-15 cm) at the Southwest Research Center, Atlantic, Iowa Microbial biomass Aglime Replication pH Organic C Organic N Croie N„,jc 0 1 4.55 18.1 3.18 217 6.5 1.20 0.20 0 2 4.75 19.0 3.47 224 9.6 1.18 0.28 4.65 18.6 3.3 221 8.1 1.19 0.24 2167 1 5.14 19.7 2.99 175 12.6 0.89 0.42 2167 2 5.53 19.8 3.09 209 13.4 1.06 0.43 5.34 19.8 3.0 192 13.0 0.97 0.43 6373 1 5.42 19.0 2.99 160 12.7 0.84 0.42 6373 2 5.80 19.4 3.18 218 16.1 1.12 0.51 5.61 19.2 3.1 189 14.4 0.98 0.47 19118 1 6.38 19.1 2.89 148 15.2 0.77 0.53 19118 2 6.41 20.8 3.28 189 16.2 0.91 0.49 6.40 20.0 3.1 169 15.7 0.84 0.51 57355 1 6.70 22.1 2.99 120 14.7 0.54 0.49 57355 2 6.75 22.4 3.08 167 14.9 0.75 0.48 6.73 22.3 3.0 144 14.8 0.64 0.49 * Mean values are in bold Table A3. Effect of liming on properties of soils under no-till system (0-5 cm) at the Northwest Research Center, Sutherland, Iowa PH Inorganic N Microbial biomass Aglime Replication H,0 CaCl2 Organic C Total N NH4-N NOa-N Organic N C^MIC ^INIC Cmic/org.C Nmic/org.N kg ECCE ha'1 g kg'1 soil -mg kg'1 soil- g kg1 —mg kg'1 soil— 0 1 5.03 4.36 25.4 3.8 15 82 3.70 242 3.3 0.95 0.09 0 2 4.91 4.40 24,0 3.6 32 119 3.45 132 26.3 0.55 0,73 0 3 4.93 4.37 25.8 4.0 23 121 3.86 210 3.5 0.81 0.09 4.96 4.38 25.1 3.8 23 107 3.67 195 11.0 0.78 0.30 560 1 4.95 4.38 25,6 4,1 21 113 3.97 131 3.1 0.51 0.08 560 2 5.10 4,57 25.2 3,3 15 103 3,18 146 5.3 0.58 0.16 560 3 6.21 5.65 30.9 4,5 7 45 4.45 331 23.6 1.07 0,52 5.42 4.87 27.2 4.0 15 87 3.87 203 10.7 0.72 0.25 1120 1 5.08 4,53 26,7 4,1 16 112 3,97 159 4,5 0.60 0.11 1120 2 5,24 4.66 25,1 3.7 8 96 3.60 194 8.1 0.77 0.22 1120 3 5.38 4.84 24.4 3,8 7 96 3,70 219 13.2 0,90 0.35 5.23 4.68 25.4 3.9 10 101 3.75 191 8.6 0.75 0.23 2240 1 5.40 4.94 23,1 3.4 7 100 3.29 169 12.9 0.73 0.38 2240 2 5,23 4.73 30.2 4,2 10 89 4.10 241 11.7 0.80 0.28 2240 3 5.70 4,90 28.0 4.0 8 57 3,94 251 11.9 0.90 0,30 5.44 4.86 27.1 3.9 8 82 3.78 220 12.2 0.81 0.32 4480 1 5.96 5.67 25.2 3.5 7 78 3.42 186 18,1 0,74 0,52 4480 2 6.16 5,78 23,5 3.3 6 78 3,22 225 16.1 0.96 0,49 4480 3 5.31 4,62 29.7 3,9 7 61 3.83 205 11.9 0.69 0,31 5.81 5.36 26.1 3.6 7 72 3.49 205 15.4 0.80 0.44 6720 1 6.04 5,75 24.3 3.6 6 78 3.52 293 18.0 1.21 0.50 6720 2 5.99 5.57 23.2 3.3 6 79 3.22 280 15,1 1,21 0,46 6720 3 6,30 5.99 29,7 3.8 7 59 3.73 311 19.8 1.05 0.52 6.11 5.77 25.7 3.6 6 72 3.49 295 17.6 1.15 0.49 * Mean values are in bold Table A4, Effect of liming on properties of soils under no-till system (0-15 cm) at the Northwest Research Center, Sutherland, Iowa PH Inorganic N Microbial biomass Aglime Replication H20 CaCI2 Organic C Total N NH4-N NO3-N Organic N ^inic ^mic CMIC/org.C Nmic/org.N kg ECCE ha 1 g kg'1 soil -mg kg'1 soil- g kg"' —mg kg 1 soil— 0 1 5.49 4,42 24.9 3.7 4 44 3.65 119 4.9 0.48 0.13 0 2 5.31 4.44 24.0 3.3 3 40 3.26 132 5.3 0.55 0.16 0 3 5.24 4.52 26.2 3.4 4 45 3.35 97 7.3 0.37 0.21 5.35 4.46 25.0 3.5 4 43 3.42 116 5.8 0.46 0.17 560 1 5,42 5.04 24.8 3.3 2 44 3.25 85 8.8 0.34 0.27 560 2 5.41 4.67 23.4 3.4 5 41 3.35 113 6.4 0.48 0.19 560 3 5.36 4.78 26.0 3.7 4 38 3.66 122 7.6 0.47 0.21 5.40 4.83 24.7 3.5 4 41 3.42 107 7.6 0.43 0.22 1120 1 5.27 4,56 26.5 3.5 4 47 3.45 106 6.8 0.40 0.19 1120 2 5,43 4,69 25.4 3.5 2 48 3.45 107 6.6 0.42 0.19 1120 3 5.45 4.81 25.4 3.8 4 45 3.75 90 7.0 0.35 0,18 5.38 4.69 25.8 3.6 3 47 3.55 101 6.8 0.39 0.19 2240 1 5.60 4.98 27.9 3.5 3 41 3.46 57 6.5 0.20 0.19 2240 2 5.44 4.76 24.8 3.3 4 45 3.25 202 4.2 0,81 0.13 2240 3 5.52 4.88 25.6 3.4 3 36 3.36 234 5.8 0.91 0.17 5.52 4.87 26.1 3.4 4 41 3.36 164 5.5 0.63 0.16 4480 1 5,76 5.10 28.2 3.6 5 34 3.56 213 9.5 0.76 0.26 4480 2 5.87 5.61 24.4 3.7 5 33 3.66 254 7.8 1.04 0,21 4480 3 5.71 5.05 26.1 3.5 4 33 3,46 223 7.5 0.85 0.21 5.78 5.25 26.2 3.6 5 33 3.56 230 8.3 0.88 0.23 6720 1 5.78 5,08 24,7 3.4 4 40 3.36 293 7.0 1.19 0,21 6720 2 6,17 5.75 23.2 3.5 5 39 3.46 419 17.1 1.81 0,49 6720 3 6,24 5.78 26.0 3.4 5 49 3.35 435 17.8 1.67 0,52 6.06 5.54 24.6 3.4 5 42 3.39 382 14.0 1.55 0.41 * Mean values are in bold Table A5, Effect of liming on properties of soil under ridge till system (0-15 cm) at the Northwest Research Center, Sutherland, Iowa gH Inorganic N Microbial biomass Aglime Replication H20 CaCI2 Organic C Total N NH4-N NQ3-N Organic N C„,ic N,nic Cmic/org.C N,nic/org.N 1 1 1 1 1 kg ECCE ha' g kg soil - -mg kg soil g kg' —mg kg soil— •••••••••••• »%•« 0 1 5.34 4.81 26.4 3.4 4 49 3.35 450 9.8 1.70 0.29 0 2 5.21 4.55 27.2 3.5 8 54 3.44 446 9.7 1.64 0.28 0 3 5.21 4.53 26.2 3.7 4 49 3.65 197 9.3 0.75 0.25 5.25 4.63 26.6 3.5 5 51 3.48 364 9.6 1.37 0.27 560 1 5.33 4.71 25.4 3.4 4 37 3.36 169 10.5 0.67 0.31 560 2 5.13 4.55 26.9 3.6 6 59 3.53 179 9.0 0.67 0.25 560 3 5,31 4.65 26,1 3.4 4 55 3.34 169 9.3 0.65 0.27 5.26 4.64 26.1 3.5 4 50 3.41 172 9.6 0.66 0.28 1120 1 5.38 5.13 27.6 3.5 3 51 3.45 170 4.8 0.62 0.14 1120 2 5.40 4.79 25.7 3.4 4 51 3.35 172 4.3 0.67 0.13 1120 3 5.24 4.65 26.7 3.9 4 60 3.84 201 3.2 0.75 0.08 5.34 4.86 26.7 3.6 4 54 3.54 181 4.1 0.68 0.11 2240 1 5.54 4.94 25.2 3.2 3 44 3.15 278 8.6 1.10 0.27 2240 2 5,49 4.88 25.8 3.2 5 53 3.14 318 9.3 1,23 0.29 2240 3 5.42 4.84 26.0 3.3 3 52 3.25 255 7.6 0.98 0.23 5.48 4.89 25.7 3.2 3 50 3.18 284 8.5 1.11 0.26 4480 1 5.60 5.04 26.1 3.4 4 46 3,35 127 6.1 0,49 0.18 4480 2 5.74 5.19 24.9 3,1 3 43 3.05 167 7.1 0.67 0.23 4480 3 5.53 4.94 25.2 3,5 4 60 3.44 201 6.4 0.80 0,18 5.62 5.06 25.4 3.3 4 50 3.28 165 6.5 0.65 0.20 6720 1 5.84 5.41 25.0 3.4 2 53 3.35 185 9.7 0.74 0.29 6720 2 5.84 5.37 24.1 3.2 3 54 3.14 177 8.8 0.73 0.28 6720 3 5.75 5.18 24.8 3.4 4 45 3.35 138 9.2 0.56 0.27 5.81 5.32 24.6 3.3 3 50 3.28 167 9.2 0.68 0.28 * Mean values are in bold • Table A6, Effect of liming on properties of soil under chisel till system (0-15 cm) at the Northwest Research Center, Sutherland, Iowa pH Inorganic N Microbial biomass Aglime Replication H20 CaCl2 Organic C Total N NH4-N N03-N Organic N C|,liC NMIC Cmic/org.C N,mi /org.N kg ECCE ha 1 g kg'1 soil mg kg1 soil- g kg1 —mg kg1 soil-- ...... 0 1 5,50 5.83 26.9 3.6 5 51 3.54 253 • 1.7 0,94 0.05 0 2 5,34 4,67 25,9 3.5 5 50 3.44 63 4.9 0,24 0,14 0 3 5,26 4,63 27.2 3.5 7 59 3.43 174 8.4 0.64 0.24 5.37 5.04 26.7 3.5 5 54 3.47 163 5.0 0.61 0.14 560 1 5.38 4.74 26.5 3.4 6 46 3.35 202 10,3 0,76 0.30 560 2 5,47 4.85 27.5 3.6 7 52 3.54 174 11.4 0.63 0.32 560 3 5.34 4.67 28,3 3.5 6 54 3.44 141 9.3 0,50 0.27 5.40 4.75 27.4 3.5 6 51 3.44 172 10.3 0.63 0.30 1120 1 5.57 4.92 29.4 3.6 7 49 3.54 154 11.2 0.52 0.31 1120 2 5.55 4.86 26.8 3.6 5 47 3.55 189 11.0 0.71 0.31 1120 3 5.47 4.78 27.3 3.6 5 45 3.55 163 10.3 0.60 0.29 5.53 4.85 27.8 3.6 5 47 3.55 169 10.8 0.61 0.30 2240 1 5.64 5.01 27.6 3.7 5 53 3.64 187 13.7 0.68 0.37 2240 2 5.59 4.92 26.4 3.6 5 48 3.55 177 13.7 0.67 0.38 2240 3 5.40 4.74 28.6 3.8 5 49 3.75 195 9,8 0.68 0.26 5.54 4.89 27.5 3.7 5 50 3.64 186 12.4 0.68 0.34 4480 1 5.70 5.06 27.1 3.3 5 44 3.25 149 7.0 0.55 0.21 4480 2 5.89 5.38 29.2 4.0 5 46 3.95 187 6.8 0,64 0,17 4480 3 5.57 4.92 27.5 3.9 5 46 3.85 197 3.6 0,72 0.09 5.72 5.12 27.9 3.7 5 45 3.68 178 5.8 0.64 0.16 6720 1 5,89 5.28 28.8 3.8 5 51 3.74 224 6.8 0.78 0.18 6720 2 5.95 5.33 27.5 3.4 6 48 3.35 195 8.0 0.71 0.24 6720 3 6.00 5.47 27.7 3.6 6 51 3.54 175 8.5 0,63 0.24 5.95 5.36 28.0 3.6 5 50 3.54 198 7.8 0.71 0.22 * Mean values are in bold 274 Table Al. p-Glucosaminidase activity of soils under different croping systems at the NERC site in!996 and!997. P-Glucosaminidase activity Crop N -— 1996b 1997b- rotation" treatment Rt R2 R3 Mean Rt R2 R3 Mean kg ha1 .mrf111 g n .MifrnnVtpnnlii in upiiciiui If ry i suncnil 11Vi'L...—. ------C-c-c-c 0 37 30 45 37 45 62 42 50 180 36 43 27 35 54 39 65 53 C-sb-c-sb 0 24 25 26 25 36 41 37 38 180 33 33 26 31 43 38 34 38 c-Sb-c-sb 0 32 37 35 35 36 45 40 40 180 30 31 54 38 45 49 43 46 C-c-o-m 0 - - - - 46 49 50 48 180 - - - - 59 58 57 58 cCom 0 41 49 49 46 56 65 56 59 180 47 53 53 51 63 61 55 60 c-c-O-m 0 44 51 46 47 52 56 60 56 180 50 48 52 50 59 67 57 61 c-c-o-M 0 58 51 61 57 - - - - 180 65 52 62 60 - - - - Sb-sb-sb-sb 0 22 19 21 21 25 28 27 27 LSD P < 0.05° 7 7 LSD P < 0.05d 14 13 a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold letters indicate crop in which the sample was taken in 1996. In 1997, the samples wer taken in the crop following the capital bold letter b R = field replicate; subscript = replication number 0 Least significant difference due to crop rotation at 0 N d Least significant difference due to crop rotation atlSO N 275 Table A8. p-Glucosaminidase activity of soils under different cropping systems at the CWRC site in 1996 and 1997 P-Glucosaminidase activity Crop N 1996b 1997b 8 rotation treatment Ri RO Mean Ri R2 Mean kg ha1 -mg p -Nitrophenol kg1 soil h - C-c-c-c 0 34 32 33 26 32 29 180 47 58 53 37 52 45 C-sb-c-sb 0 38 30 34 41 29 35 180 44 28 36 42 38 40 c-Sb-c-sb 0 36 32 34 29 42 36 180 30 44 37 40 38 39 C-c-o-m 0 45 36 41 43 41 42 180 52 46 49 49 52 51 c-C-o-m 0 46 42 44 49 57 53 180 54 51 53 60 54 57 c-c-O-m 0 47 51 49 73 52 63 180 74 55 65 62 71 67 c-c-o-M 0 79 82 81 53 63 58 180 71 74 73 62 49 56 C-o-m-m 0 66 35 51 42 58 50 180 62 44 53 53 50 52 c-O-m-m 0 51 43 47 58 57 58 180 52 64 58 80 80 80 c-o-M-m 0 57 47 52 77 65 71 180 80 47 64 64 72 68 c-o-m-M 0 77 50 64 53 45 49 180 79 75 77 53 52 53 LSD P < 0.05e 17 19 LSD P < 0.05d 24 12 a C = corn, Sb = soybean, O = oats, M = meadow (alfalfa). Capital bold lett< indicate crop in which the sample was taken in 1996. In 1997, the samples weretaken in the crop following the capital bold letter b R = field replicate; subscript = replication number c Least significant difference due to crop rotation at 0 N d Least significant difference due to crop rotation at180 N Table A9. Regression equations and r values of the relationships between p-glucosaininidase activity and organic C and N, microbial biomass C and N in soils of the cropping systems Relationship Regression equation r Regression equation r NERC CWRC 1996 samples EGA vs Corg Y = -20.5 + 2.89X r = 0.70*** Y = -60.2 + 3.33X r = 0.76*** EGA vs Norg Y = -29.7 + 47.8X r = 0.75*** Y = -57.5 + 48.1X r = 0.76*** EGA vs Cmic Y = -2.53 + 0.20X r = 0.71*** Y = 1.86 + 0.13X r = 0.64*** EGA vs Nmic Y = 2.58 + 0.75X r = 0.76*** Y = 32.9 + 0.30X r = 0.33* 1997 samples EGA vs Corg Y = -24.9 + 3.45X r = 0.58*** Y = 1.88+ 1.41X r = 0.42** EGA vs Norg Y = 44.8 + 2.18X r = 0.03 Y = 17.4+ 15.7X r = 0.37* EGA vs Cmic Y = 23.2 + 0.12X r — 0.49*** Y = 30.4 + 0.07X r = 0.44** EGA vs Nlllic Y = 41.4 + 0.23X r = 0.21 Y = 44,2 + 0.28X r = 0.19 * P<0.05, ** P<0.01, P <0.001 EGA p-Glucosaminidase activity, Corg organic C, Norg organic N, Cmi(. microbial biomass C, Nmic microbial biomass N 277 Table AlO. Inhibition or activation of P-glucosaminidase activity in soils by trace elements Inhibition or activation of p-glucosaminidase Trace element in soils specified8 Element Oxidation state Clarion Nicollet Canisteo None 100 100 100 Ag I 80 27 50 Cu 95 67 68 Ba II 89 84 99 Cd 91 60 63 Co 137 156 168 Cu 94 66 67 Fe 93 92 93 Hg 65 45 28 Mg 115 120 106 Mn 107 122 105 Ni 88 86 82 Pb 93 84 95 Pb 94 82 99 Sn 81 88 79 Zn 92 82 68 A1 III 67 83 87 As 101 79 84 B 104 131 146 Cr 84 88 91 Fe 87 70 82 Se IV 94 67 82 Ti 88 83 69 V 88 84 91 As V 114 123 113 W VI 98 76 87 Mo 82 87 107 LSDb (P < 0.05) 1.1 0.8 1.9 * P-glucosaminidase activity of non-treated soils (mgp -nitrophenol kg"1 soil h"1) of Clarion soil = 90, Nicollet soil = 88 and of Canisteo soil = 36. b LSD = least significant difference. Table All. Effect of tillage and residue management on soil pHa Rep I Rep II Rep III Rep IV Mean LSD P< 0.05 Treatments water CaCl2 water CaCl2 water CaCl2 water CaCl2 water CaCl2 water CaCl2 No-till/Bare 7.2 6.3 7.1 6.4 6.5 5.8 5.1 4.3 6.5 5.7 0.03 0.05 No-till/Normal 6.9 6.2 7.2 6.5 6.6 5.8 5.0 4.2 6.4 5.7 0.07 0.02 No-till/2X Mulch 6.8 6.1 7.0 6.3 6.4 5.6 5.1 4.2 6.3 5.6 0.10 0.04 Chisel/Normal 7.1 6.4 7.1 6.5 7.0 6.3 4.9 4.2 6.5 5.9 0.02 0.03 Chisel/Mulch 7.1 6.5 7.1 6.5 7.1 6.5 5.2 4.3 6.6 6.0 0.02 0.06 Moldboard plow/Normal 6.9 6.2 6.9 6.2 7.0 6.2 4.9 4.1 6.4 5.7 0.02 0.05 Moldboard plow/Mulch 6.7 6.1 7.0 6.4 7.0 6.5 6.0 5.0 6.7 6.0 0.06 0.11 LSD P< 0.05 0.04 0.03 0.06 0.03 0.03 0.03 0.03 0.02 0.4 0.4 Depth of no-tiil/2X Mulch 0-5 cm 6.9 6.4 6.8 6.3 6.4 5.8 5.3 4.5 6.4 5.8 0.04 0.04 5-10 cm 6.9 6.3 6.9 6.3 6.4 5.7 5.1 4.2 6.3 5.6 0.06 0.05 10-15 cm 7.1 6.2 7.1 6.4 6.5 5.9 5.4 4.6 6.5 5.8 0.04 0.07 LSD P< 0.05 0.07 0.01 0.04 0.08 0.11 0.01 0.1 0.02 0.6 0.6 a soiliwater or soil:CaCl2 solution (0.01 M) = 1:2.5 ratio. 279 ACKNOWLEDGEMENTS Many people have made this study possible. First and foremost, I would like to express my sincere appreciation and gratitude to my major professor, Dr. M. A. Tabatabai, for his guidance and encouragement through the period of my studies and research at Iowa State University. His enthusiasm and commitment towards research was inspirational. This accomplishment would not be possible without his guidance. I would like to express my appreciation to Dr. R.M. Cruse, Dr. T.E. Fenton, Dr. A.L. Pometto III, and Dr. L B. Tabatabai for taking the time to serve on my committee, along with Dr. S.T. Henning for soil sampling. Also it is necessary to extend my special thanks to Dwi Rovita Lee, Dr. Veronica Acosta-Martinez and Dr. Susann Klose, for supporting me not only academically but also for being the closest friends I had during my graduate study. Many thanks to my research colleagues, Dr. Fahad Alromian, Dr. Daniel Dodor, and Marcia LaCorbiniere for their friendship. I wish there was a much better way to describe my appreciation and love to my parents, Gullii and Muzaffer Ekenler. From my heart, thank you for being such wonderful and supportive parents. Without your sacrifice and encouragement, I would not be where I am. Thanks one more time for your unconditional love and support in all the moments of my life. My sisters, Gulay, Rukiye, and Eylem and my brother Mahir, I am deeply grateful for your love and 280 encouragement. Although you live thousands of miles away from me, you are always close to my heart and in my thoughts. My lovely nephews, Berdan and Adar, you have also been the angels and one of my happy sources during the difficult time of my life in the US. I love both of you and wish the best of everything in your life. My warmest thanks go to the Borges family. Thank you for your love and prayers. Dear Josie, I appreciate your support and coming here to help me during the last part of my Ph.D journey. You are one of the special persons who made it possible to complete my degree this semester. My deepest and warm appreciation goes to my little boy Deniz Jason. Thank you for being such a good baby during my pregnancy and after you were born. Your easy-going and well-behaved nature have allowed me to focus on my work and have made this degree possible. Finally, I am deeply grateful to my dear husband Rogerio Borges. He was and is my everything; my love and my best friend. He was the one to keep my spirit up during my most difficult times in the US. He made many sacrifices for my happiness, not only to complete this program, but also to make me successful in my career.