Measurement of Soil Enzyme Activities

Soil enzyme activities have been often proposed as sensitive indicators of soil quality. As underlined by Schloter et al. (2003), "the ideal soil microbio-logicalandbiochemicalindicator todeter minesoil quality wouldbesimple to measure, should work equally well in all environments and reliably reveal which problems existed where". The authors considered enzymatic assays very helpful methodologies because they are short-term laboratory procedures, usually performed under standardised environmental conditions (use of sieved samples, optimal temperature and pH, and standardised water content), which prevent or minimise changes in the composition of soil biota and allow comparison of data from different soils obtained in different laboratories.

However, accurate and reliable methodologies are required to understand the cause-effect relationship between a considered parameter and an observed soil enzyme activity modification. Two different types of problems arise when measuring the enzyme activity of soil. The first concerns the selection of enzymes to be measured and mostly depends on the objectives of the research being conducted. The second problem relates to the intrinsic nature of the soil enzyme to be measured and concerns the selection of the proper method. However, it is also important to keep in mind that enzyme-like reactions occur in soil and may contribute to the measured enzyme activity (Ruggiero et al. 1996).

If the goal of the research is to characterise different soils for their microbial activity and/or to derive indices of soil microbial functional diversity, the activities of a great number of enzymes should be measured. Indeed, a single measure very unlikely could represent an indicator of the functional diversity, which reflects a multitude of biochemical pathways (Nannipieri et al. 1990; Schloter et al. 2003). Furthermore, attention should be paid to enzymatic activities of the main metabolic pathways, such as dehydrogenase or assimilatory nitrate reductase activities which are active in living cells (Dick 1994).

If the goal of the research is focused on process-level investigations such as response of soil to the application of fertilisers, amendments or pesticides, evaluation of land-use impacts, or monitoring soil pollution and recovery upon remediation actions, the measurement of enzyme activities involved in the transformation of specific substrates, without specific attention to their origin or location, may be suitable. Also in this case, the assay of a single enzyme activity is not sufficient for the reasons discussed above.

Enzyme assays are based on the quantitative evaluation of the product(s) released or of the substrate(s) consumed and thus they do not distinguish the contribution of the different enzyme categories to the overall enzyme activity (Burns 1982). As discussed above, the same enzyme can be intra-cellular in living cells, in dead cells or cell debris or extracellular, free or absorbed by soil colloids. By simplifying we can say that the enzyme can exist in soil, for example, in three forms: free in solution, inside a living cell and immobilised on an organo-mineral complex. It is obvious that these enzyme forms can behave in a different way. For example, the immobilised form can be very stable, and preserve its activity almost completely, whereas the activities of the other two forms can be inhibited by an inhibitor.

Nannipieri et al. (2002) have critically and thoroughly examined the different methodological approaches to distinguish among the different categories of soil enzymes. Particular attention has been paid to the identification and quantification of the intracellular and extracellular fractions contributing to the overall soil enzyme activity. This is definitely one of the most important problems in the interpretation of soil enzymatic activities. One method is based on the measurement of microbial growth and enzyme activity during incubation when soil is amended with C and N substrates. Each enzyme activity is plotted against microbial biomass, measured at the same incubation time; if a linear correlation exists, the extrapolation of the straight line to zero microbial biomass gives the intercept of the plot on the ordinate (enzyme activity) which represents the extracellular enzyme activity in case of a positive value. This approach assumes that enzyme assays detect both intracellular and extracellular enzyme activity. The second method assumes that enzyme measurements only detect extracellular enzyme activity (Klose and Tabatabai 2002). After CHCl3 fumigation or ultrasonic treatment it is possible to measure both intracellular and extracellular enzyme activity due to the lysis of microbial cells. This method presents the drawbacks that proteases are active during CHCl3 fumigation and partially hydrolyse enzymes (Renella et al. 2002) and released enzymes can be adsorbed by soil colloids with consequent denaturation (Fornasier 2002). These processes can obviously lead to an underestimation of the contribution of intracellular enzymes. Furthermore, the efficacy of CHCl3 fumigation on the lysis of microbial cells may change with changing soil structural properties, thus affecting the reliability of the method (Badalucco etal. 1997).

The measurement of the enzyme activities of soil before and after ul-trasonication has also been proposed as a reliable method to distinguish between intra- and extracellular enzyme activities in soil (Badalucco, pers. comm.). Apart from the same problems of the CHCl3 fumigation method, the ultrasonic treatment of soil can facilitate the disaggregation and frac-tionation of mineral particles and organic matter, thus favouring the possible release of enzymes associated to soil colloids and whose activity could not be determined before the ultrasonic treatments. Indeed the consistent increase in acid phosphatase activity (up to 156% greater than the untreated sample) has been hypothesised to be due to ultrasonication with release of a large portion of acid phosphatase activity, undetected when associated to soil colloids (De Cesare et al. 2000). However, it was concluded that the release of intracellular acid phosphatase after lysis of cells upon sonication could not be excluded.

Different chemical agents have also been added to the enzyme assays during the incubation to inhibit the microbial growth and the consequent increases in enzyme activity due to new synthesis of proteins. Toluene is one of the most used antiseptic agents in enzyme assays, especially in the hydrolases assays, but it presents some disadvantages such as direct inhibitory or stimulatory effects on the activity of the tested enzymes (Acosta-Martinez and Tabatabai 2002) or the increase of microbial cell permeability with consequent overestimation of the intracellular enzyme activity (Skujins 1978; Nannipieri et al. 2002). However, Klose and Tabatabai (1999a,b) demonstrated that both intracellular arylsulfatase or urease activities were not completely detected upon toluene treatment and that the enzymes were not inhibited by toluene.

A reasonable approach to determine the so-called stabilised extracellular enzyme activity could involve the efficient sterilisation of soil prior to the assay. Numerous methods are available to sterilise soil (Trevors 1996). However, the approach presents the following drawbacks: the method cannot be efficient in eliminating soil microbial populations; the chemical and physical properties of soil may be altered, with consequent changes in the relative fractions of immobilised enzymes; intracellular enzymes could be released upon sterilisation. Thus either an under- or overestimation of the extracellular enzyme activity of the investigated soil is possible.

The contribution of the different enzyme categories to the overall enzyme activity in soil might indirectly be evaluated by comparing properties and behaviour of enzyme activities of soils with those of synthetic enzyme model systems. This approach was followed to study the effect of three herbicides (atrazine, paraquat, glyphosate) and one insecticide (carbaryl) on invertase, urease and phosphatase activities present as: (1) free purified enzymes, which should simulate the fractions of free enzymes in soil solution; (2) synthetic clay-, organo- and organo-clay-enzyme complexes, which should simulate enzyme-soil colloid associations; and (3) soil enzymes (Gianfreda et al. 1993, 1994, 1995, 2002; Sannino and Gianfreda 2001). Activation, inhibition or no influence depended on the pesticide and the enzyme preparation thus suggesting that the "state" of the enzyme in soil is important and no generalisations can be made. Complex responses were also obtained when the effects of the four pesticides on soil enzyme activities were studied (Gianfreda et al. 1994,1995; Sannino and Gianfreda 2001). Increases, decreases and no effects of four pesticides on the activity of invertase, urease and phosphatase were obtained in 22 soils sampled from different sites of Italy and characterised by different physical-chemical properties (see Sect. 12.7). Furthermore, the response of some soil enzyme activities to four pesticides was similar to that of one of the model systems, probably because the model was prevalent in the relative soil. In addition, the clays of soil protected phosphatase activity against the inhibitory effect of carbaryl and atrazine. This was confirmed by the fact that phosphatase immobilised on montmorillonite was less affected by the pesticides than the free and organo- and organo-mineral-complexed phosphatase.

The conclusion of all the findings reported so far is that unfortunately methodologies capable of accurately and unambiguously measuring each of the soil enzyme components are not available to date.

In enzyme measurements, as for other microbiological and biochemical measurements, soil samples should be representative of the natural situation of studied soils (representative sampling) and the sampling strategy for biochemical and microbiological soil properties should consider their temporal and spatial dynamics (Dick et al. 1996a). Therefore, the sampling time, the number of samples and their horizontal and vertical distribution as well as the sampling procedure and device must be defined. Simple rules may be suggested in carrying out an accurate sampling procedure. To prevent temporal heterogeneity and variability, sampling should take place repeatedly during a given period of time and possibly at the same time intervals. Composite or well-mixed samples can be used and in this case a mean of the investigated soil property is obtained, provided the sampling area has uniform properties (Dick et al. 1996b). Moreover, if the aim is to obtain any information on the variability of the studied enzyme at the microsite levels, a great number of samples should be collected and geostatistical analyses should be carried out to adequately define spatial variability of the studied enzyme (Bonmati et al. 1991).

If soil pretreatment and storage are not important for measuring physical and chemical properties of soil, they are crucial in the determination of biochemical activities (Brohon et al. 1999). A short storage (maximum 10-15 days or less) of field-moist soils at 4 °C is suggested for a reliable determination of the majority of soil enzyme activities if the measurements cannot be done immediately after sampling (Alef and Nannipieri 1995; Gianfreda and Bollag 1996; Tabatabai and Dick 2002). Changes in enzyme activities during storage depend on the period and temperature of soil storage, and the physical-chemical characteristics of the investigated soils (Brohon et al. 1999). Storage of air-dried soils has been proposed for handling soils before enzyme assay (Burns 1978a; Alef and Nannipieri 1995; Gianfreda and Bollag 1996; Tabatabai and Dick 2002), but measurements with fresh samples are more reliable (Rao et al. 2003). The response of enzyme activities to air-drying of soil as well as storage of moist soil may be enzyme-specific (Luo et al. 1996). Probably, the activities of enzymes stabilised by soil colloids are less affected than those associated to living cells by air-drying (Ladd 1985; Gianfreda and Bollag 1996). The response of enzyme activity of air-dried soils can be different in respect to that of the same enzyme activity of moist soil, thus indicating that they are different. The inhibition of several trace elements on aspartase activity (Senwo and Tabatabai 1999) was greater in air-dried than in field-moist soils whereas opposite behaviours have been observed for cellulase (Deng and Tabatabai 1995) and arylamidase activities (Acosta-Martinez and Tabatabai 2001).

Assay conditions such as presence or absence of a buffer, pH, substrates and their concentration, temperature, shaking of soil, inhibitors of microbial proliferance, etc., markedly affect the measured activity, as previously discussed (Burns 1978a; Alef and Nannipieri 1995). Usually, a soil enzyme assay is based on the use of a buffered solution of a synthetic, artificial substrate at a concentration high enough (saturation concentration) to be assumed constant throughout the time course of the enzymatic reaction and assuming a zero-order kinetics. Moreover, substrate concentration should be very much higher than that of the enzyme to allow a reaction rate proportional to the enzyme concentration.

Miller et al. (2001) suggested the use of low concentrations of substrate, so as to simulate environmental conditions. They measured two types of for-mamide hydrolase activity in a barley soil: a low (Km values of 30-60 mM) and a high (Km values of 0.5-1.0 mM) affinity enzyme activity with the latter representing the enzyme activity of active microorganisms. A similar approach was followed to measure high- and low-affinity l-glutamine synthetase (Sallis and Burns 1989) and l-histidine ammonia-lyase (Burton and McGill 1989, 1991) activities. Probably, under natural conditions, microorganisms should use high-affinity systems operating at low substrate concentration (Unanue et al. 1999).

The use of different substrates may be helpful to better understand the role of an enzyme activity in nutrient transformations in soil. For instance, Tabatabai et al. (2002) studied the activity of acyl arylamidase (an enzyme involved in one of the limiting steps of N-mineralisation) of several Iowa soils towards eight substrates, differing in their amino acid moiety. Kinetic (Km and Vmax) and thermodynamic (AHa and Ea) parameters depended on soil and the amino acid moiety, thus suggesting that enzyme isoforms were present in soils (Tabatabai et al. 2002).

A buffered vs. an unbuffered condition is usually chosen when the disappearance of substrate and/or the formation of reaction products may change the pH of the soil slurry, since an optimal pH value is required during the assay. Acid and alkaline soil phosphatase activities prevail in acid and alkaline soils, respectively (Eivazi and Tabatabai 1977), and thus Dick et al. (2000) proposed that acid (AcdP) and alkaline phosphatase (AlkP) activity should also be used as pH indicators.

Foranassayofsoilenzymeactivitytobereliableandapplicable, itshould be tested and validated with various soils with different properties. Indeed, major problems may arise from both the adsorption of substrates and/or productson soil particlesandfrom thepossibleinterferencebyelementsor compounds present in soil. For example, p-nitrophenyl derivatives which are common substrates in the determination of several soil enzyme (acid and alkaline phosphatases, glucosidases, galactosidases, and arylsulfatases) activities are hydrolysed to p-nitrophenol (pNP) which maybe adsorbed by soil colloids. To avoid an incorrect evaluation of pNP-substrate-based activity, the adsorption of p-nitrophenol should be quantified. Thus the calibration curve should be prepared by using different amounts ofp-nitrophenol in the presence of the soil under investigation (in the amount used in the assay) and under the experimental conditions (temperature, incubation time, addition of reagents) of the enzyme assay. Similar problems may occur when NH+ is detected as a measure of urease activity in soils with different NH+-fixing capabilities. Urease activity of NH+-fixing and non-fixing soils was affected by the used buffer (Tris-, borate- or non-buffered; O'Toole 1991). Since small changes occur in substrate concentration under saturating substrate conditions, product appearance rather than substrate consumption is usually monitored so as to achieve higher analytical sensitivity.

Some compounds or elements of soils under investigation may react with the reaction products thus affecting the measured activity. For example, Cu reacts abiotically with triphenylformazan, the end product of dehydrogenase activity measured by using tetrazolium salt (TTC) as acceptor of electrons (Chander and Brookes 1991). Unfortunately, Cu also reacts with 2-p-iodophenyl-3-p-nitrophenyl-5-phenyltetrazolium formazan (INTF), the endproductof2-p-iodophenyl-3-p-nitrophenyl-5-phenyltetrazolium chloride (INT), the alternative substrate used to measure the dehydrogenase activity of soil (Obbard 2001). Paraquat affects the p-nitrophenol determination at 400/405 nm in alkaline environment (Gianfreda et al. 1993). Coloured compounds extracted from soil with some buffers can affect the colorimetric determination of substrates or reaction products.

The use of substrate derivatives, with enzymatic production of fluorescent end products, has been proposed as a valid alternative to increase the sensitivity of the enzymatic assays and to avoid some of the problems cited above (Marx et al. 2001). These assays are based on the use of fluorophore-labelled artificial substrates, derived from water-soluble flu-orophores (e.g. hydroxy- and amino-substituted coumarins, fluorescein, and rhodamine), which are enzymatically transformed into highly fluorescent, water-soluble products with optical properties different from those of the parent compound. The enzyme assay is rapid, can be used to measure several enzyme activities of soil with the microplate approach, and it allows the measurement of several soil samples (Wirth 1992; Wirth and Wolf 1992; Sinsabaugh et al. 2000; Marx et al. 2001; Vepsalainen et al. 2001; Stemmer 2004; Wittmann et al. 2004). The advantages and disadvantages of the microplate fluorimetric assay and thep-nitrophenol method are summarised in Table 12.2.

Although the multiple-substrate approach provided results closely correlated to those obtained with the classical enzyme assays, it presented some drawbacks that may limit its application to different soils (Stemmer 2004). Indeed, the substrate recovery was incomplete because of its adsorption onto the soil matrix, soils with different physical and chemical properties behaved differently, competitive inhibition occurred due to chemically similar substrates, and data from soils with contrasting pH values could not be compared because the multiple-substrate assay was conducted at soil pH (Stemmer 2004).

As has been extensively discussed, the present enzyme assays give potential rather than real enzyme activities due to experimental conditions adopted in the determination (Burns 1978a, 1982; Nannipieri et al. 1990, 2002; Gianfreda and Bollag 1996). For this reason, Schloter et al. (2003) have suggested that the present soil enzyme assays give a measure of the enzyme activity "encoded in the soil genotype" and not a "phenotypic activity". This obviously restricts the usefulness of enzymatic activity data and limits their interpretation.

Another problem is the lack of standardisation because different protocols are used to determine an enzyme activity in soil, as indicated by Burns (1978b) for assaying urease activity. Unfortunately, this variability still continues, as shown in Table 12.3, which reports the experimental conditions used for assaying soil urease in the last 5 years. It is worth noting that different conditions have been adopted by the same research groups in different studies and no reasonable explanation of such variability is provided.

Another problem is the presence of enzyme-like reactions which cannot be distinguished by enzyme reactions with the present assays (Huang 1995; Ruggiero et al. 1996; Nannipieri et al. 2002). Clays, metal oxides and hydroxides can catalyse a large number of reactions; e.g. organic

Table 12.2. Comparison between the 4-methylumbelliferone (MUB) micro-plate fluorimetric assay and the p-nitrophenol assay for measuring soil enzyme activity (from Marxetal. 2001)

4-Methylumbelliferone The products

Measured fluorimetrically

Excitation at 365 and emission at 460 nm

MUB fluorescence is pH-dependent

Highly sensitive especially at low concentrations

No known side effects

Quenching of fluorescence by soil particles and phenolic compounds

MUB molecule is highly mobile

The assay

Soil added volumetrically Small quantity of substrate needed (|l) Short incubation time (35 min) Substrate difficult to dissolve Continuous monitoring of product release over time (1 read cycle min-1) Measurement directly carried out in the reaction medium

High numbers of samples and replicates processed at the same time (96-well plate-1)

Plate set-up and measurement in less than 1h

Disposable materials (micro-plates, universal bottles, pipette tips) Results

Automatic calculation of activity rates (relative units of fluorescence min-1) Miscellaneous p-Nitrophenol

Measured colorimetrically Absorbance at 405 nm p-Nitrophenol absorbance is pH-dependent Low sensitivity p-Nitrophenol interferes with organic material p-Nitrophenol is adsorbed

Soil added gravimetrically Large amounts of substrate required (ml) Medium to long incubation time (1-24 h) Substrates dissolve instantaneously A single reading at the end of the incubation

Need to stop the reaction and extract the product prior to measurement (measurement must not be delayed, otherwise there is formation of Na2CO3) Maximum of 50 assays per run

Assay preparation, incubation, product extraction and measurement >3.5 h

Use of glass equipment with time-consuming preparation

Withdrawal of background absorbance

Expensive analytical equipment Analysis at low cost

[fluorimetric plate-reader, multichannel (digital) pipettes, micro-plates]

A limited number of potential A large number of substrates available substrates available compounds, usually of phenolic nature, are transformed in their oxidised products, and these abiotic reactions are involved in the humus formation (Ruggiero et al. 1996).

Table 12.3. Experimental conditions used in soil urease assays


Klose and


Dilly and Nannip-

Taylor et al. (2002)

Caravaca and


Tabatabai (1999a)

et al. (2000)

ieri (2001)

Roldân (2003)

et al. (2004)

Urea concentration

0.20 M

0.02 M

0.08 M

8.9 mM


Not specified




Not specified



Not specified



50 mM acetate (or deionised H20)

Not specified

75 mM borate buffer

0.1 M phosphate

0.1 M phosphate

Incubation temperature

37 C

20 C

37 C

20 C

30 C

37 C

Incubation time

2 h


2 h


1.5 h


Soil/aqueous solution

Not specified






ratio (w/v)


Not specified




Flow injection





analyser coupled

reagent kit

reagent kit

reagent kit

with a spectrophotometer

Soil status, treatment

Field-moist soil

Soil homogen-

Fresh soil

Field-moist soil

Field-moist soil

Soil moistening

or pre-treatment

isation in acetate buffer pH 5

stored at 4 C

stored at 4 C

to 50% WHC and preincubation for 7 days at 25 : C

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