Microbial, plant and animal cells can be present in soil and release enzymes upon cell death and lysis or for physiological reasons such as hydrolysis
Liliana Gianfreda: Dipartimento di Scienze del Suolo, della Pianta e dell'Ambiente, Universita di Napoli Federico II, Via Universita 100, 80055 Portici, Napoli, Italy, E-mail: [email protected]
Pacifico Ruggiero: Dipartimento di Biologia e Chimica Agroforestale ed Ambientale, Universita di Bari, Via Amendola 165/A, 70126 Bari, Italy, E-mail: [email protected]
Soil Biology, Volume 8 Nucleic Acids and Proteins in Soil P. Nannipieri, K. Smalla (Eds.) © Springer-Verlag Berlin Heidelberg 2006
of polymers in oligomers or monomers to be taken up by cells. Therefore, soil is rich in enzymatic proteins which catalyse reactions involved in energy transfer, nutrient cycling, environmental quality and crop productivity (Dick 1994; Tabatabai 1994). Thus, the number of enzymes in soil is expected to be very high and at least 500 enzymes can have a critical role in both C and N cycles. However, the number of enzyme activities so far measured in soil is much more limited (Dick et al. 1996a; Gianfreda and Bollag 1996; Nannipieri et al. 2002). For instance, the enzymatic activities involved in the N cycle, and identified in soil, are summarised in Table 12.1. This number is by no means exhaustive, considering all N transformations occurring in soil.
Some of the enzyme activities listed in Table 12.1 were detected many years ago (Skujins 1967, 1976; Burns 1978a), whereas others have been measured only recently (Acosta-Martinez and Tabatabai 2000,2001). Most of them were tested by using synthetic rather than natural substrates. This along with other methodological aspects may lead to an erroneous evaluation of the meaning of enzyme measurements in soil, as discussed later.
The majority of the examples reported earlier (Gianfreda and Bollag 1996; Nannipieri et al. 2002) and those summarised in Table 12.1 refer to enzyme-like activities of soil rather than to activities of purified enzyme proteins. Unlike what occurs with other living systems, very few enzymes or enzyme-like activities have been extracted from soil and their purification has been generally unsuccessful (Mayaudon 1986; Tabatabai and Fu 1992; Nannipieri et al. 1996; Chap. 4). Generally, these enzymes have been extracted as organo-enzyme complexes, i.e. as enzyme activities associated with organic material (see Chap. 4).
Various intracellular and extracellular enzymatic forms contribute quantitatively and qualitatively to the overall enzymatic activity of soil (Burns 1978b, 1982). The same enzyme can have a different origin (i.e. from bacteria, fungi, plants, and a range of macroinvertebrates) and occupy different locations not only in the producing cell, but also in the extracellular soil matrix (Burns 1978b, 1982). The same enzyme can be intracellular or associated to the external surface of the originating cell (cells can be alive, dead, proliferating or non-proliferating; active enzymes can be also associated to cell fragments), and, if secreted outside the cell, they can be free in soil solution, adsorbed by clays or entrapped by humic molecules. Thus it may have different properties, and show different behaviours, in response to a given factor.
Generally, the activity of free enzymes is considered very low and thus its contribution to the measured enzyme activity can be neglected also considering their short life as free proteins in an inhospitable environment such as soil (Nannipieri and Gianfreda 1998). The fact that free extracellular
Table 12.1. Soil enzyme activities involved in the N cycle
Enzyme (trivial name) Function Natural substrates
Amidase (EC 220.127.116.11) Hydrolysis of C-N bonds other Peptides, amides than peptide bonds in linear amides
Arylamidase Hydrolysis of a N-terminal Arylamides
(EC 18.104.22.168) amino acid from peptides, amides or arylamides
Peptidases Peptides hydrolysis Peptides
Denitrification activity Denitrification of nitric Nitric compounds
(DEA) compounds to N2O and N2
l-Asparaginase Hydrolysis of asparagine L-Asparagine
Formamide Miller et al. (2001); Dodor and
l-Leucine Acosta-Martinez and Tabatabai
(2001,2002); Dodor and Tabatabai (2002); Tabatabai et al. (2002)
/?-Naphthylamide Brown (1985); Burket and Dick hydrochloride (1998); Marx et al. (2001);
Leucine-4-nitroaniline Vepsalainen et al. (2001);
l-Leucine 7-AMC* l-Arginine 7-AMC l-Tyrosine 7-AMC Lys-ala 7-AMC
Luo et al. (1996); Simek and Hopkins (1999); Simek et al. (2002)
Frankenberger and Tabatabai (1991a); Dodor and Tabatabai (2003)
Enzyme (trivial name) Function l-Aspartase (EC 22.214.171.124) l-Glutaminase (EC 126.96.36.199)
l-Clutamine synthetase (EC 188.8.131.52) l-Histidase (EC184.108.40.206)
Nitrogen as e
Hydrolysis of L-aspartate to fumarate and NH3 Hydrolysis of glutamine to glutamate and NH3
Reaction between NHJ-N and l-glutamic acid Hydrolysis of hystidine to urocanate and NH3 Reduction of NOJ to NOj
Natural substrates L-Aspartic acid L-Glutamine
L-glutamic acid and NHj-N L-Histidine
Natural and added proteins l-Aspartic acid l-Glutamine
NH+-salts and nitrates l-Histidine
Casein, N-benzoyl-l-argininamide (BAA), N-benzyloxy-carbonyl-l-phenylalanyl l-leucine (ZPL)
Senwo and Tabatabai (1999); Dodor and Tabatabai (2003) Frankenberger and Tabatabai (1991b); Dodor and Tabatabai (2003) McCarty (1995); Landiet al. (1999) Frankenberger (1983); Burton and McGill (1989,1991)
Abdelmagid and Tabatabai (1987); Fu and Tabatabai (1989)
Tann and Skujins (1985); Martinez-Toledo et al. (1988); Gajendiran and Mahadevan (1989); Halsall and Gibson (1991) Bonmati etal. (1998); Gianfreda and Bollag (1996); Nannipieri et al. (2002); Tabatabai and Dick (2002)
Enzyme (trivial name) Function Natural substrates Substrates used in the References assay
Enzyme (trivial name) Function Natural substrates Substrates used in the References assay
Reaction between cyanide and
Tabatabai and Singh (1976, 1979);
thiosulfate to produce thyocianate
of sulfur organically bound compounds
Singh and Tabatabai (1978); Ray et al. (1985); Szadjach (1996)
Hydrolysis of urea into ammonium and carbon oxide
Bremner and Mulvaney (1978); O'Toole (1991); Kandeler et al. (1999a); Klose and Tabatabai (1999a); Sinsabaugh et al. (2000); Taylor et al. (2002)
*7-AMC = 7-amino-4-methylcoumarin
*7-AMC = 7-amino-4-methylcoumarin enzymes are not extracted from soil seems to prove the low importance of this enzymatic category in soil.
Another aspect of the complexity of enzyme activities in soil is their in situ distribution, i.e. their spatial variability in a soil profile and their localisation in soil structural fractions of different nature and size. As reviewed by Gianfreda and Bollag (1996), earlier studies on this matter were performed by Ross and Speir and their co-workers and by Perez-Mateos and his collaborators.
Bergstrom and Monreal (1998) and Bergstrom et al. (1998b, 2000) found that arylsulfatase activity was spatially decreased with depth and water content along a slope of the Ap horizon in a Gray Brown Luvisol (Hapludalf), sampled on 74 positions following harvest of soybean (Glycine max L. Merr.) and fall tillage. Phosphatase activity spatially depended on organic C and inorganic P contents whereas dehydrogenase, urease, glutaminase and^3-glucosidase activities showed little or no spatial dependence. The six enzyme activities were also measured in a Rego Humic Gleysol (Aquoll) sample (0-23 cm depth) strip-cropped to corn (Zea mays L.) and soybean (Glycine max L. Merr.) and these samples were collected at 20 cm spacing along transects across adjacent plant rows (Bergstrom and Monreal 1998). Enzymatic activities of soil samples collected from within crop rows were compared with those of samples collected between crop rows. The spatial pattern of all six soil enzyme activities was sometimes influenced by the crop and they behaved similarly as general indices of microbial activity at the scale of measurement.
Generally, enzyme activities decline with depth and are related to mi-crobial activities and C and N organic contents of soil (see Sect. 12.7).
Kandeler and co-workers have emphasised the importance of studies on the micro-scale distribution of enzyme activities and soil microorganisms in particle-size soil fractions and their variations at a small-scale level in response to different agricultural practices (i.e. organic amendments, tillage; Stemmer et al. 1998, 1999; Kandeler et al. 1999a,b,d; Gerzabek et al. 2002; Poll et al. 2003). These studies can be useful: (1) to study the importance of different soil fractions on the protection of microorganisms and enzyme activities associated with organo-mineral particles (Kandeler et al. 1999d); (2) to elucidate if and how the metabolic activities are distributed in different particle size fractions (Gerzabek et al. 2002); and (3) to evaluate the contribution of different particle size fractions to the turnover of organic C, by following the response of microbial processes to a range of long- and short-term organic C inputs (Stemmer et al. 1999; Gerzabek et al. 2002; Poll et al. 2003).
The poor knowledge about the location of enzymes in differently sized particle fractions may have been due to the procedures used for physical fractionation in the past, which greatly reduced both soil microbial biomass and enzyme activities (Ruggiero et al. 1996). It was concluded that soil microorganisms were mainly associated with the silt and clay fraction (Ladd et al. 1996). In recent investigations, a more preservative fractionation method preceded by low-energy sonication of soil (because it minimised destruction of labile particulate organic matter) was adopted (Stemmer et al. 1998, 1999; Kandeler et al. 1999a,b,c; Gerzabek et al. 2002; Poll et al. 2003).
The distribution of enzyme activities between bulk and rhizosphere soil and the accumulation of enzymes at soil-plant root interfaces have been studied (Tarafdar and Jungk 1987; Badalucco and Kuikman 2001; Naseby and Lynch 2002). A high activity of arylsulfatase per unit of microbial biomass-S was observed in rape rhizosphere soil, and organic acids were considered the most efficient substrates in promoting the production of the enzyme by the soil microflora (Vong et al. 2003). The acid phosphatase and dehydrogenase activities of rhizosphere soils from 13 nodulated legume species sampled at two savanna sites with very different chemical properties were significantly higher than the respective enzyme activities of bulk soils (Izaguirre-Mayoral et al. 2002). Differences among species and between sites were recorded.
Although root-associated microorganisms often are assumed to be the main producers of enzymes in the rhizosphere soil, an important contribution of enzymes can, however, derive from enzymatic proteins produced by plant roots and released in their surrounding rhizosphere soil. These enzymes are usually wall-associated enzymes and catalyse the formation of products which are taken up by plant roots or rhizosphere microorganisms (Gramss et al. 1999b; Chroma et al. 2002; Harvey et al. 2002).
The enzymes typically more abundant in the rhizosphere are phos-phatases whose activities are related to the depletion of organic P (Tarafdar and Jungk 1987). Furthermore, a strict correlation between phosphatases originating from plant roots and P nutrition of plants was demonstrated, particularly in rhizosphere soils of mycorrhiza-infected plants (Halstead and McKercher 1975; Dick et al. 1983; Chhonkar and Tarafdar 1984; Reddy et al. 1987; Tarafdar and Jungk 1987; Tarafdar and Claassen 1988; Haus-sling and Marschner 1989; Tadano and Sakai 1991; Fox and Comerford 1992; Tarafdar and Marschner 1994). The localisation of phosphatase activity in rhizosphere soil was carried out by microscopic observations of soil sections, after their treatment with suitable compounds, which allowed enzymes in soil structure to be localised, or reaction with substrates giving rise to products easily visualisable as coloured compounds (Ladd et al. 1996; Joner et al. 2000).
Grierson and Adams (2000) concluded that seasonal and spatial heterogeneity of acid phosphatase activity of rhizosphere soil depended on plant species composition and root type (cluster roots or ectomycorrhizal roots), in a study comparing the enzyme activity, ergosterol (as indicator of fungal biomass) and microbial P in soils from a forest of southwestern Australia dominated by Jarrah (Eucalyptus marginata Donn ex SM.) with and without Banksia grandis Wiild. Jarrah is characterised by an extensive surface system of fine lateral roots with ectomycorrhizal associations, while Banksia grandis is the producer of a mat of cluster (proteoid) roots and is capable of growing if fire is excluded from these forests. Increases in acid phosphatase activity were associated to increases in microbial P and ergosterol concentrations of soils and, thus, it was concluded that most of the measured enzyme activity was due to fungal activity.
Further evidence on the effect of plant species on enzyme activities of rhizosphere soil were provided by Gramss et al. (1999a) who demonstrated that several members of Fabaceae, Gramineae and Solanaceae released considerable amounts of peroxidase, laccase, monophenol monoxygenase, and proteinase-lipase-esterase activity, expressed as fluorescein diacetate hydrolase, in the rhizosphere of both sterile and non-sterile soils. Such enzyme activities were lower in non-sterile than sterile soils probably because they were inactivated by proteases of soil microorganisms. The protein preparations from planted soils contained plant peroxidase as evidenced by sodium dodecyl sulfate/polyacrylamide gel electrophoresis (SDS-PAGE) analyses and isolectrofocusing (Gramss et al. 1999a). The production of cell-wall-associated peroxidase was also demonstrated in vitro in some cultures of plants of various species and morphology (Chroma et al. 2002; Harvey et al. 2002).
The capability of a soil to show a catalytic and enzymatic activity, even in the absence of any living cells, led to the conclusion that active enzymatic proteins must act in soil as stable complexes with inorganic and organic soil colloids. As defined by Gianfreda and Bollag (1996), these enzymes can be regarded as "naturally immobilised enzymes".
Two main features differentiate a soil enzyme from its pure preparation: the kinetic behaviour and the sensitivity to various denaturing agents. Soil enzymes usually have lower Vmax values and higher Km constants, indicating a lower catalytic efficiency and a reduced substrate affinity, than the respective purified enzymes from different sources (Nannipieri and Gianfreda 1998). Similarly, the extensive bibliography shows different activity-pH or activity-temperature profiles (Nannipieri and Gianfreda 1998). These peculiar behaviours can partially be explained by assuming that soil enzymes are immobilised on soil supports and, consequently, work in a heterogeneous system (McLaren and Packer 1970; McLaren 1975, 1978; Nannipieri and Gianfreda 1998). However, the exact mechanism by which enzymes are immobilised and consequently stabilised in soils is not completely clarified.
Numerous studies have examined the adsorption of proteins on clay minerals and the properties of the resulting clay-protein complexes, because the clay fraction can play an important role in the immobilisation of soil enzymes (Kiss et al. 1975; Theng 1979; Boyd and Mortland 1990). This topic is discussed in Chap. 7. On the other hand, experimental evidence obtained on the interactions between proteins and organic substances indicates that soil organic matter can also have an important role in the immobilisation of enzymes in soil (Ladd and Butler 1975). Lahdesmaki and Piispapen (1992) concluded that the extraction, the fractionation by gel filtration and the dilution damaged or destabilised the protective mechanism (against environmental stresses in soil) by soil colloids on protease, cellulose and amylase.
The main strategy used for understanding the relationships between immobilised enzymes and their clay or humic supports was to study properties and behaviours of enzyme complexes, obtained in vitro by the interaction between enzymes and clay minerals, humic substances or organo-mineral complexes (Ladd and Butler 1975; Burns 1978a; Boyd and Mortland 1990; Gianfreda et al. 2002). The support, the enzyme, the bond between enzyme and support, the chemical or physical mechanism implicated in the immobilisation process are all important in the formation of the complex and markedly affect the properties of the resulting complex. Cation-exchange adsorption mechanisms, van der Waals type forces, ionic or hydrophobic bonds, can hold the enzymes to clay surfaces. Ion exchange, entrapment within organic networks, ionic or covalent bonding, may account for the stable association between enzymes and humic materials (Ladd and Butler 1975; McLaren 1975; Burns 1978a, 1982, 1986; Theng 1979; Boyd and Mortland 1990).
Enzyme activities in soil are resistant to proteolysis, thermal and chemical denaturation (Nannipieri et al. 1996). To achieve an increased stability, enzymatic proteins must preserve not only their three-dimensional structure or conformation, but also their plasticity to explicate their catalytic action. Several mechanisms may contribute to enhance enzyme stability upon immobilisation on solid supports: the tightening of the protein structure by covalent (mono or multipoint) bindings to an insoluble support or by "cage effects" inside a polymeric gel; the change of the enzyme microenvironment; the steric hindrance created by the surrounding matrices with reduced molecular mobility and capacity of unfolding; an increased physical protection towards proteases. The presence of special molecules such as sugars, polyalcohols and organic polymers or of an organic phase can also lead to increased enzyme stability (Gian-freda and Scarfi 1991). These mechanisms and conditions can also be valid for soil enzymes. The physical and chemical conditions surrounding enzymes immobilised by clay minerals, entrapped in the humic matrix, biological membranes or by particular subcellular structures, are different from those of aqueous solutions. Thus the kinetic characterisation of these immobilised enzymes has to take into consideration the heterogeneous phase in which they carry out their activity (Nannipieri and Gianfreda 1998).
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