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Interferences, limitations and an improvement in the extraction and assessment of cellulase activity in soil
Soil Biol. Biochem. Vol. 26, No. 1, pp. 65-73, 1994
Ekvier Science Ltd. Printed in Great Britain
Pergamon
0038-0717/94$6.00+ 0.00
INTERFERENCES, LIMITATIONS AND AN IMPROVEMENT IN THE EXTRACTION AND ASSESSMENT OF CELLULASE ACTIVITY IN SOIL* LISE K. GANDER,‘? CHARLESW. HENDRICKS* and JACK D. DOYLE’
‘ManTech Environmental Technology Inc. and *U.S. Environmental Protection Agency, Environmental Research Laboratory, 200 S.W. 35th Street, Corvallis, OR 97333, U.S.A. (Accepted
30 May 1993)
Summary-A practical modification of a reducing-sugar method to estimate soil (carboxymethyl) cellulase activity is described and used to compare three non-sterile soils. The original method involves the formation of a ferric-ferrocyanide complex in soil extracts prepared and exposed to substrate in 2M acetate buffer. Because high acetate concentrations compromise this reaction, such extracts require considerable (e.g. 20-30-fold) dilution prior to activity determinations. For soils with low organic matter, such as one used in this study, undetectable activity often results. We used 67 mu acetate buffer and were able to detect and compare cellulase activities in three diverse soils using undiluted extracts. The values obtained were more reproducible, but in a different range from those of the original method. Our modification yielded activities that appeared to be unchanged over the extraction buffer concentration range of 1t-140 mM acetate. The use of undiluted soil extracts prepared in less concentrated buffer simplifies and extends the practicality of the assay by: (a) reducing the importance of using matrix-matched standard curves; (b) requiring less sample manipulation and glassware, and (c) preventing the dilution to extinction of enzyme activity. It also improves the usefulness of the method as an indicator of ecological effects (biomass turnover) due to the introduction of nonindigenous microorganisms and chemicals of environmental concern. These factors make the method more competitive with other reducing-sugar assays used to measure cellulase activity in soil.
of soil cejlulase activity might be useful in environmental risk assessment. There are many calorimetric assays for soil cellulase activity (e.g. Benefield, 1971; Pancholy and Rice, 1973; Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990; Wirth and Wolf, 1992). Most involve reducing-sugar (or glucose) determinations, and use reagents such as dinitrosalicylic acid (DNS; Miller et al., 1960), alkaline copper (Somogyi, 1952) or glucose oxidase (Benefield, 1971). Carboxymethylcellulose is commonly used as a substrate, because of its high reactivity (Miller et al., 1960; Kanazawa and Miyashita, 1986). However, this substrate is less than ideal (Lindner et al., 1983; Breuil and Saddler, 1985). Differences in measurable cellulase activity can be method-specific, reflecting both the sensitivity of the detection method and the choice of substrate. Soil sources and extraction methods are also influential. Some researchers suggest calorimetric assays used to quantify released sugars are poor indicators of overall cellulase hydrolytic potential, particularly for samples with low B-glucosidase activities (Chan et al., 1989). Others indicate this finding is substrate-dependent (Rivers et al., 1984). In many cases, interferences are related to the degree of substrate hydrolysis (Rivers et al., 1984). Choosing an appropriate assay often means choosing appropriate compromises (Breuil and Saddler, 1985).
INTRODUCTION Cellulase [ 1,4-( 1,3; 1,4)-j?-D-glucan-4-glucanohydrolase] is responsible for the hydrolytic conversion of cellulose to glucose (saccharification). At least three classes of enzymes typically play roles in this process: endoglucanase (EC 3.2.1.4), exoglucanase (cellobiohydrolase; EC 3.2.1.91) and /?-glucosidase (cellobiase;
EC 3.2.1.21). The relative contributions of these three enzymes to overall cellulase (carboxymethylcellulase) activity are source-dependent (Coughlan and Ljungdahl, 1988) and have been investigated in a variety of plants, fungi, bacteria and soils. Through the action of cellulase, important energy sources for soil microorganisms are released. Hence, soil cellulase activity is an indicator of biomass turnover. Biomass and related measurements are useful in assessing soil stability, nutrient cycling and both agrochemical and microbial influence on soil metabolic processes. Increased interest in the ecological effects of toxic chemicals, hazardous wastes and genetically engineered microorganisms has prompted a need for methods to assess differences in perturbed vs non-perturbed soil. Because cellulase can influence biomass turnover and carbon cycling, determinations l@ Government of Canada. TAuthor for correspondence. 65
66
LISE K. GANDER et al.
Soil enzymes have lower activities than those from microbial and plant sources (Kanazawa and Miyashita, 1986). Also, even under optimized conditions of pH and temperature, extraction rates can be slow (Kanazawa and Miyashita, 1986). For this reason, combined soil extraction and incubation periods of 24-48 h are common for soil cellulase assays (Benefield, 1971; Pancholy and Rice, 1973; Schinner and von Mersi, 1990). Companion reducing-sugar or glucose determinations for cellulase activity estimates are relatively short (e.g. < 1 h), but vary with respect to sensitivity and reliability. Reactions involving DNS are convenient, but not very accurate, especially at low ( < 1 mM) reducing sugar concentrations (Lindner et al., 1983; Foroughi and Gunn, 1983). Also, results are influenced by dissolved oxygen, citrate buffer, divalent cations (Foroughi and Gunn, 1983) and the composition of the cellulase complex being measured (Breuil and Saddler, 1985). Because non-linear responses are generated in DNS assays, cellulase activities are dilution-specific (Breuil and Saddler, 1985). The Nelson-Somogyi alkaline copper assay (Somogyi, 1952) is less convenient, but more sensitive and reliable than that involving DNS (Lindner et al., 1983; Breuil and Saddler, 1985). The reaction is affected by incubation conditions, choice of substrate and other components in the cellulase complex; however, to a lesser extent than the DNS assay (Breuil and Saddler, 1985). Removal of trace metals from soil extracts improves Nelson-Somogyi detection limits (M. A. Tabatabai, pers. commun.). Unlike DNS, the alkaline copper reagent reacts stoichiometrically with the number of hemiacetal reducing groups present (Breuil and Saddler, 1985). However, biphasic responses with higher values at lower enzyme (sample) concentrations have been reported (Linder et al., 1983; Breuil and Saddler, 1985). These results are less reproducible and probably less representative of hydrolytic potential than those obtained for more concentrated samples (Breuil and Saddler, 1985). Values determined by the DNS method are often 45% higher than those by the Nelson-Somogyi method (Breuil and Saddler, 1985) which in turn are about 90% higher than those by the glucose oxidase-horseradish peroxidase assay (Kanazawa and Miyashita, 1985). This reflects the non-specific nature of reducing-sugar assays: i.e. substances other than glucose can be measured. The glucose oxidase-horseradish peroxidase method of Benefield (1971) is specific, reproducible and sensitive. This determination avoids the inaccuracies associated with co-measurement of non-glucose products of cellulolysis (e.g. cellobiose). Glucose oxidase reacts only with glucose. The reaction is compromised by pigments and ions released during soil extractions (Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990). Hence, soil extracts are often diluted or treated with phenol-binding polymers (Kanazawa and Miyashita, 1986) prior to
assays for glucose. Only the latter permits low detection limits. The ferric-ferrocyanide reducing-sugar assay (Park and Johnson, 1949) has been applied by Schinner and von Mersi (1990) for soil cellulase activity measurements. The sensitivity of this assay is affected by the presence of aldehydes, phenols, catechols (Litwack, 1960), acids, high ionic strength (Cooper, 1977) and any compounds with significant oxidative or reductive potential (P. Turner, pers. commun.). Many of these interfering substances are found in soil extracts (Brady, 1974) originating from the soil itself or introduced with the extracting buffer. Hence, this cellulase determination is likely to generate matrixvariable responses of at least two types: those relating to the choice of soil, and those relating to the choice of extracting buffer. In our ‘research concerning the effect of environmental stressors on carbon cycling, we seek simple methods to assess differences in perturbed vs non-perturbed soil systems. Our early work with fertic-ferrocyanide approaches to determining cellulase activity suggested that results in soil systems might be dependent upon both extraction and dilution conditions (unpubl. observ.). We have evaluated the influence of soil extraction conditions and suggest not only a practical, more versatile modification of the Schinner and von Mersi (1990) method, but also potential limitations of the approach to assess soil cellulase activity. Additional information concerning soil properties, /I-glucosidase and dehydrogenase activities, and populations of indigenous microbes is presented and related to cellulase activity. MATERIALS AND METHODS
Soil characteristics and preparation
Four soils were collected as vegetation-free samples from the upper 5 cm of selected locations (meadow, agricultural, forest and riverbank) in Linn and Benton counties, Ore. (Tables 1 and 2). Meadow soil was used only in tests to establish appropriate concentrations of extracting buffer. Each soil was sieved (2 mm) and stored in a sealed plastic bag (room temperature, light-free) for not more than 2 weeks. Two days before use, soils were refrigerated (4°C) divided into two lots, and either adjusted to 60% of full water holding capacity (WHC; Pramer and Schmidt, 1965; Doyle et al., 1991), or left unadjusted (field-moist). Three subsamples were removed for replicate analyses of soil moisture (Pramer and Schmidt, 1965) selected chemical variables (Horneck et al., 1989) including pH (Stotzky et al., 1992), and the enumeration of indigenous microbes. All measurements were made in triplicate. Microbial enumeration
Indigenous bacterial colony forming units (cfu) and fungal propagules were enumerated from spreadplate distributions of serially diluted soil on selective
61
effects in assessing soil cellulase
Extraction
Table I. Chemical and microbioloeical characteristics of soils Agricultural (Malabon silty clay loam)
PH Organic matter (%) CEC (meq IOOg-‘) N&N (kg g-‘) NO,N (pg g-‘) P&g g-‘) KOlg g-‘) CaOlg g-‘) MgOlg g-‘) FeoCg g-‘) Total fungi (log propagules gg’) Total bacteria (log cfu g-‘)
60% WHC
Fieldmoist
60% WHC
Fieldmoist
8.48 ( f 0.25) 5.43 ( * 0.03) 3.76 26.12 8.04 6.37 34 437 II.3 3.5 130
36.89 ( f 0.54) 5.64 ( * 0.09) 3.87
34.65 ( f 0.81) 6.66 ( f 0.05) 13.83 40.92 3.06 8.62 I3 725 23.9 8.2 142
31.63 ( + 0.36) 6.60
I .26 ( k 0.08) 7.30 ( f 0.04) 0.37 14.09 6.35
5.83 ( + 0.04) 4.76 ( + 0.23)
5.80 ( + 0.06) 5.51 ( + 0.09)
5.47 (f0.16) 7.02 (kO.14)
WHC = water holding capacity; CEC = cation exchange capacity;
media To indicate the extent of microbial survival in soil extracts prepared for cellulase assays, aliquots of these preparations were plated also. Soil extract agar (Wollum, 1982) containing 100 fig cycloheximide ml-’ (Sigma) was used to enumerate total soil bacteria. Martin’s agar (Martin, 1950) was used to estimate fungal propagules. Dilutions were prepared (in sterile tap water) in triplicate per soil type. Two plates of each medium were inoculated per dilution or extract. Values were normalized to mean cfu (or propagules) g-r oven-dry soil equivalent prior to log,, transformations. Enzyme activity determinations
Enzyme activities are expressed as the mean ( f SEM) of three replicates, in units of product formed g-r oven-dry weight equivalent of (wet) soil and incubation time. The activities of soil /J-glucosidase and dehydrogenase were determined by standard methods (Tabatabai, 1982). Cellulase (endoand exocellulase) was extracted and assayed both using the Schinner and von Mersi (1990) variation of the Park and Johnson method (1949), and with the modifications described later. For these applications, a swollen suspension of carboxymethylcellulose sodium salt (medium viscosity; Sigma; 0.7% w/v, in acetate buffer) was used as the substrate. To indicate
Table 2. Cellulase activity in four concentrations of undiluted acetate extracting buffer Acetate Buffer (mM) II 53 I05 I40
Riverbank (Willamette silt loam)
Fieldmoist
Characteristic %H,O
Forest (Price-Ritner gravelly silty clay loam)
Endoglucanase activity (nmol glucose equiv. g-’ soil 24 h-‘) 467*52 439 * 53 453 f 93 398 + 77
Each activity is the mean + SEM of three replicates of fieldmoist, Willamette loam (meadow) soil (pH = 5.9, organic matter = 0.86%. Fe = 86 pg g-l).
( f ‘3.06) 13.99
5.40 (kO.10) 7.01 ( * 0.04)
3.07 (kO.18) 4.88 ( f 0.04)
60% WHC 19.80
( f 0.79) 7.12 ( + 0.02) 0.42
3.25 (50.19) 5.67 ( f 0.41)
+_values indicate the SEM of three replicates.
enzyme extraction efficiencies, acetate solutions containing 500 pg of crude endoglucanase from Trichoderma viride (Sigma) were extracted and assayed for cellulase activity in the presence or absence of soil. The combined cellulase assay and soil extraction procedure (Schinner and von Mersi, 1990) involves three reactions. (1) Soil (cellulase) is extracted under buffered, acidic conditions (pH 5.5, 5O”C, 24 h) and allowed to bind to carboxymethylcellulose substrate. This increases the number of glucose residues available from the modified cellulose. (2) Potassium ferricyanide is reduced (in the presence of excess potassium cyanide) under alkaline conditions (pH 10.5, lOO”C, 15min) to potassium ferrocyanide by glucose (and other reducing agents, constituting interferences). (3) Ferrocyanide reacts with excess ferricyanide (in the presence of ferric ammonium sulfate and a surfactant) under acidic conditions (pH < 2, room temp., 60min) to produce a ferric-ferrocyanide complex. This chromophore is measured spectrophotometrically (690 nm) and related to cellulase activity. The following modifications of the Schinner and von Mersi (1990) cellulase assay were used. (1) Controls: one soil-free control containing substrate and three replicate (soil-containing) substratefree controls were incubated with each reaction set. The former was used to zero the spectrophotometer and, when required, to dilute (post-reaction) offscalehigh samples. The technique of post-reaction dilutions was validated (agreement ranged from 97 to 115%) prior to being applied, by assaying 10 sets of three matched replicates. These matched replicates consisted of either three pre-reaction dilutions + three post-reaction dilutions (for extracts prepared
LISEK. GANDERet al.
68 1.50
1.25
GLUCOSE (nmol ml- I) Fig. 1. Matrix-matched, soil-free, standard curves for (carboxymethyl) cellulase assay used to relate absorbance to product formation (enzyme activity). Each acetate concentration corresponds to the following dilutions of a 2 mM acetate stock: 64 nw = 3t-fold; 107 m&i= 21-fold; 129 mu = W-fold, and 320 mu = &fold. Correlation coefficients for the linear regressions are: 0.999 {no acetate), 0.995 (64 mbt), 0.996 (107 m@. 0.996 (I29m@, and 0.915 (320m~).
in 2 M acetate), or three undiluted extracts plus three post-reaction dilutions (for extracts prepared in 67 mM acetate). The mean absorbance of replicate, substrate-free controls was used for background subtractions. Thus, both soil and enzyme-free substrate contributions were taken into account. (2) Preparation of soil extract: rather than filtering the reaction mixture, ~nt~fugation (2 min at 12,000g; Beckman Microfuge 11) followed by removal of the supernatant was used to isolate extract for future reactions. (3) Assessment of microbial influence: soil extracts were assayed for cellulase activity both before and after filtering through 0.2pm membrane filters. (4) Extraction-incubation matrix: in addition to 2 M acetate, pH 5.5, diluted lO-30-fold for reducing sugar quantifications, various concentrations of pH 5.5 acetate buffer were examined. Because S&inner and von Mersi (1990) indicate that agricultural and forest soils extracted in 2 M acetate buffer must be diluted 20-30-fold prior to cellulase activity determinations, 67 m&racetate (equivalent to a JO-fold dilution of 2 M acetate) was most often selected. Mat~x-match~ standard curves of glucose [i.e. glucose standard (Sigma) + acetate] were used to compensate for acetate-induced signal quenching to establish linear relationships between absorbance and glucose concentration. (5) Activity units: cellulase activities are expressed on the basis of glucose equivalents g-r dry soil and total reaction exposure time. These units reflect the non-specificity of reducing-sugar assays,
which rely upon increased reducing potential (not necessarily free glucose residues) to measure the interaction between endo- and exocellulase (Sinsabaugh ef al., 1981). RESULTS
Soil-free standard curves constructed to assess the effect of pH 5.5 acetate (and other extracting buffers) on glucose quantification showed that signal quenching does occur, particularly at acetate concentrations of 129 and 320m~ (Fig. 1). The substitution of pH 5.5 citrate as an extracting buffer resulted in a 3-fold increase of the quench effect observed with acetate: 44 mhi citrate was equivalent to 129 ma4 acetate. No apparent differences in soil cellulase activity were found for a range of undiluted acetate extracting buffer concentrations (1 I-140 mM; equivalent to 190-14-fold dilutions of 2 M acetate) (Table 2). Soil-free, time-course assays of commerciallyavailable, crude endoglucanase from Trichina uiriak were examined both under extraction conditions using 2 mr+sacetate (S&inner and von Mersi, 1990), and those of our 67 mu acetate modification. Results indicated not only similar kinetics for the two approaches, but also that, in the absence of soil, our modification recovers at least as much cellulase activity as the original method (Figs 2 and 3). When soil spiked with standard enzyme was extracted and assayed in either 67m~ or 20- and 30-fold dilutions
Extraction effects in assessing soil cetlulase
69
24
Incubation Time (h) Fig. 2. Timecourse study of standard (T. vi&k) endoglucanase activity in soil-free assays using both 67 mhi acetate buffer (undiluted, except post-reaction, where required) and 30-fold (pre-reaction) dilutions of 2 M acetate buffer. Each value is the mean * SEM of three replicates. Correlation coefficients for the logarithmic curve fits are: 0.915 (67 mM) and 0.977 (2 M).
ABC 67&i
&et&e (1x)
2M Acewe wo
A
S
C
2M Amate (3mO
Fig. 3. Overall celhdase activity in soil-free extracts containing 500 pg endoglucanase from ?‘. viride (A), fieid-moist forest soil plus 5OO~g endoglucanase from T. viride (B), and field-moist forest soil (C). Measurements were in 67 mM acetate extracts (undiluted, except post-reaction, where required) and 2 mtu acetate extracts diluted @e-reaction) either 20- or 30-fold. Each value is the mean & SEM of three replicates.
LISEK. GANDERet al.
70
a
B
A
A
Agricultural tSoi1
B
A
B
Rivorbsnk sdl
Forest Soil
Fig. 4. Cellulase activity as measured in extracts prepared from field-moist soils (A) and those adjusted to 60% WHC (B), using undiluted 67 mM acetate buffer (PH 5.5). Extracts were assayed before and after filtering through a 0.2 pm membrane. Each value is the mean + SEM of three replicates.
of) 2rn~ acetate buffer, the cellulase activities obtained were far from equivalent to the combined activities of standard plus soil enzyme. Instead, enzyme recoveries were compromised to 27% (67 mM acetate extracts), 16% (20-fold dilutions of 2 M acetate extracts), and 78% (30-fold dilutions of 2 M acetate extracts) of soil-free, standard enzyme activities (Fig. 3). Cellulase activity was detectable for three diverse soils when extracts prepared in undiluted 67 mM acetate were analyzed (Fig. 4). The agricultural and forest soils were found to be 7-8 and 15-18 times higher, respectively, in cellulase activity than the riverbank soil. Filter-sterilization of these extracts gave no appreciable difference in activity (Fig. 4). Likewise, adjusting soil moisture from field-moist conditions to 60% WHC (Table 1) yielded no pronounced changes in measured activities of cellulase (Fig. 4), /J-glucosidase (Table 3), or dehydrogenase (Table 3). Large increases in water content (e.g.
Table 3. b-Glucosidase
and dehydrogenase
Agricultural (Malabon silty clay loam) Enzyme activity fi-Glucosidase @g PNP produced Dehydrogenase @g TPF produced
g-’ soil h-‘) g-’ soil 24 h-‘)
WHC = water holding three replicates.
adjustment of agricultural and riverbank soils to 60% WHC) also coincided with increases in bacterial populations, but left fungal numbers relatively unchanged (Table 1). There were no detectable fungi and, relative to unextracted soil, reduced bacterial (2.39 f 0.09 g-’ agricultural soil, numbers 3.70 + 0.03 g-’ forest soil, and 2.04 + 0.13 g-’ riverbank soil) in 67 mM acetate extracts. The numbers of recovered bacteria corresponded to about 0.1% of the original bacterial populations in these soils (Table 1). No fungi or bacteria were detected in corresponding 2 M acetate extracts. When 2M acetate extracts were diluted lo-, 20and 30-fold to determine cellulase activity (Schinner and von Mersi, 1990), the riverbank soil was found to be below detection limits (Fig. 5). Although raw absorbance values were lower, the activity units for other soils analyzed by this method were markedly higher (about 2-12 times; Fig. 5) than those attained with undiluted 67 mM extracts (Fig. 4). Background-
activities
Forest (Price-Ritner gravelly silty clay loam)
Riverbank (Willamette silt loam)
Fieldmoisl
60%WHC
Fieldmoist
6O%WHC
Fieldmoist
60% WHC
170+8
157*4
169 f 7
170&8
3*0
2&O
22Sk9
227 f 10
26 *
I
36 f 2
19*0
capacity; PNP =p-nitrophenol; TPF = triphenylfonnazan; + values indicate the
19+ SEM
I of
Extraction effects in assessing soil cellulase
Fig. 5. Cellulase activity as measured in extracts prepared from field-moist soils (A) and those adjusted to 60% WHC (B), using 2 M acetate buffer (PH 5.5) diluted lo-, 20-, and 30- fold (Schinner and von Mersi, 1990). Each value is the mean f SEM of three replicates. Cellulase activity was undetectable in riverbank soil (all dilutions) and in 30-fold dilutions of agricultural soil adjusted to 60% WHC.
corrected absorbance values ranged from 0.070 to 0.366 for 30-lo-fold dilutions of 2mM acetate extracts of forest soil. Undiluted 67 mM acetate extracts had typical absorbance values of 0.418-0.535. Extreme dilution-specific variabilities in cellulase activity were observed for the same 2rn~ acetate soil extracts prepared and analyzed by the Schinner and von Mersi (1990) method (Figs 3 and 5), even in the presence of high amounts of added standard enzyme (Fig. 3). Pronounced differences associated with water adjustment were also noted in the cellulase activities for these extracts. This is in contrast to all the other enzyme activity measurements conducted, including cellulase activities for 67 mM extracts. Both /I-glucosidase and overall cellulase activities were lowest for riverbank soil (Table 3, Figs 4 and 5). However, the profile of relative /I-glucosidase activity for all three soils differed from that observed for cellulase dete~inations with 67m~ extracts. Mean /I-glucosidase activities for both treatments (with and without water adjustment) of agricultural, forest and riverbank soils were 164 : 170 : 2.4, respectively (Table 3; equivalent to 68: 71: 1). Cellulase activities for the same three soils showed relative ratios of 362:780:49 (Fig. 4; equivalent to 7: 16: 1). Contrary to reports of inverse relationships between soil dehydrogenase and cellulase activities (Schinner and von Mersi, 1990), the profile for relative dehydrogenase activity (Table 3) was similar to that seen for cellulase activity (Fig. 4) for all three
soils. Forest soil had the highest dehydrogenase and cellulase activities, while those for agricultural and riverbank soils were considerably lower. DISCUSSION
Our modification of using an undiluted soil extract prepared in a less concentrated buffer was effective in assessing overall cellulase activity in a variety of soils, including two containing < 1% organic matter. The original method of Schinner and von Mersi (1990) seems impractical for soils with low organic matter, probably because cellulase activity is diluted to extinction. In fact, Schinner and von Mersi (1990) recommend determining xylanase rather than cellulase activity in such soils. Sparingly small amounts of soil are involved in cellulase activity dete~inations of 20-30-fold dilute extracts. This could promote high sample-to-sample variability, and might account for both the differences in cellulase activity observed for the same 2 M acetate extract diluted over a range of values and those between field-moist and 60% WHC soils (Fig. 5). That cellulase activities for extracts prepared in 2 rnM acetate were lower with decreased soil amounts (produced by increased soil extract dilutions, or higher water content in soils) suggests the original method is easily biased when solutions representing different amounts of soil are used for ferric-ferrocyanide-based determinations. This problem
12
LISEK. GANDER et al.
is avoided by determining glucose equivalents in undiluted soil extracts prepared in less concentrated buffer. Such a modification not only increases the reproducibility and the sensitivity of the application, but also permits the post-reaction dilution of off-scale high samples. The use of undiluted, less concentrated, extracting buffer eliminates some of the difficulties associated with the Schinner and von Mersi (1990) approach to estimating soil cellulase activity. However, compromised recoveries of combined standard plus soil enzyme activities (Fig. 3) suggest that results are matrix-dependent, regardless of whether or not soil extracts are diluted. Of all the samples prepared and analyzed in this manner, the 30-fold dilution of soil extract spiked with standard enzyme came closest to approaching the activity of soil-free standard enzyme alone (Fig. 3). This suggests the exogenous enzyme may be binding to the soil or the presence of soil-contributed interferences. Presumably, interferences are diluted to undetectable concentrations in highly diluted soil extracts. Such factors might account for the lower cellulase activities (relative to l&30-fold dilute 2 M acetate extracts; Fig. 5) obtained for undiluted 67 mM acetate extracts (Fig. 4), which involve more amounts of soil. Thus, the ferric-ferrocyanide cellulase assay appears to suffer from similar drawbacks as that involving DNS (Breuil and Saddler, 1985). Both generate dilution-specific, matrix-variable responses of at least two types: those relating to the choice of extracting buffer, and those relating to the properties of the soil. In both the Schinner and von Mersi method (1990) and our modified cellulase assay, enzyme extraction and substrate exposure occur in one 24 h step. This promotes saturation kinetics and fails to distinguish between enzyme extractability and true activity. Also, cellulase activity units are expressed as a function of dry soil (rather than protein concentrations) for the entire 24 h reaction. Combined, these facts suggest that activities determined by these methods must be relative. However, other soil enzyme assays involve combined 24 h enzyme extraction and substrate exposure, and use similar activity units (Pancholy and Rice, 1973; Tabatabai, 1982). These assays provide relative values which are commonly used as indicators of perturbed vs non-perturbed systems (Malkomes, 1991). In soil applications, an extracting buffer of high ionic strength would be more likely to kill indigenous microorganisms and mask differences in ionic character of the soils used, providing a more universal testing system. However, high ionic strength is a major interference in the ferric-ferrocyanide reducing-sugar assay (Cooper, 1977). Thus, we investigated the rationale and practice of using acetate (pH 5.5) and other choices of extraction-reaction buffers. Acetate buffers are commonly used for soil cellulase extractions (Pancholy and Rice, 1973; Kanazawa and Miyashita, 1986; Wirth and Wolf, 1992). Cellulase
activity has a pH optimum of ca 5 (Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990). The enzyme should form its strongest ionic attachments to carboxymethylcellulose (pK,-4) at pH -3-5. Because the only stoichiometric reaction involved in ferric-ferrocyanide-based reducing sugar determinations is compromised by formic acid (Litwack, 1960), the use of acetate is somewhat questionable. Yet, in comparing acetate (two carbon) and citrate (six carbon) buffers, we found that although both cause signal quenching, acetate does so to a lesser extent, and thus remains a preferable choice. Also, soil extracts prepared in acetate buffers are less pigmented (Kanazawa and Miyashita, 1986). For greater versatility in assessing various soil types for relative cellulase activity (especially those expected to have low values), the use of an undiluted, less concentrated (e.g. 67 mM) acetate solution seems advisable. Our modification of using a less concentrated extraction buffer for cellulase determinations permitted the analysis of undiluted soil extracts, including those of low activity. This broadens the versatility of the method and makes it more competitive with other reducing-sugar assays used to determine cellulase activity. However, for maximum confidence in soil cellulase activities estimated by this approach, we recommend the following: (1) identical preparation of replicate soil extracts and controls; (2) inclusion of an internal standard, such as commercially-available cellulase enzyme, assayed in the presence and absence of each soil type, as an indicator of both enzyme extraction efficiency and the extent of soil-contributed interferences; (3) recognition that activity values are relative, and probably should not be compared across soil types. The determination may be most useful for assessing the effects of different treatments within one soil type. We believe our modification and recommendations increase the practical application of this cellulase assay as an indicator of biomass turnover and ecological effects due to the introduction of non-indigenous microorganisms and chemicals of environmental concern. thank Marcia Bollman and Drs Thomas L. Bott. Robert Sinsabauah and Paul M. Turner for their review and comments on the manuscript. We thank Lynn Bucao, Bonnie Hugley (ManTech Environmental) and Central Analytical Laboratory (Oregon State University, Corvalhs, Ore.) for technical support. This research was funded by the U.S. Environmental Protection Agency and conducted in accordance with an approved quality assurance plan, and has been subjected to Agency review. Acknowledgements-We
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Lindner W. A., Dennison C. and Quicke G. V. (1983) Pitfalls in the assay of carboxymethylcellulase activity. Biotechnology
and Bioengineering
Litwack G. (1960) Carbohvdrates.
25,377-385. In Exoerimental
_ Bio-
chemistry: .A Loboratory kanual, pp. c88. Wiley, New York. Malkomes H.-P. (1991) Existing alternative tests to measure side-effects of pesticides on soil microorganisms: dehydrogenase activity. Toxicological and Environmental Chemistry 30, 167-176. Martin J. P. (1950) Use of acid, rose Bengal and streptomycin in the plate method for estimating soil fungi. Soil Science 69, 215-232.
73
Miller G. L., Blum R., Glennon W. E. and Burton A. L. (1960) Measurement of carboxymethylcellulase activity. Analytical
Biochemistry
2, 127-l 32.
Pancholy S. K. and Rice E. L. (1973) Soil enzymes in relation to old field succession: amylase, cellulase, invertase, dehydrogenase, and unease. Soil Science Society of America Proceedings
37, 47-50.
Park J. T. and Johnson M. J. (1949) A submicro determination of glucose. Journal of Biological Chemistry 181, 149-151.
Pramer D. and Schmidt E. L. (1965) Experimental Soil Microbiology. Burgess, Minneapolis. Rivers D. B., Gracheck S. J., Woodford L. C. and Emert G. E. (1984) Limitations of the DNS assay for reducing sugars from saccharified lignocellulosics. Biotechnology and Bioengineering
26, 800-802.
Schinner F. and von Mersi W. (1990) Xylanase-, CMtellulase and invertase activity in soil: an improved method. Soil Biology & Biochemistry
22, 51 l-515.
Sinsabaunh R. L.. Benefield E. F. and Linkins A. E. (1981) -Cellulase ‘activity associated with the decomposition of leaf litter in a woodland stream. Oikos 36, 184-190. Somogyi M. (1952) Notes on sugar determination. Journal of Biological Chemistry 195, 19-23. Stotzky G., Broder M. W., Doyle J. D. and Jones R. A. (1992) Detection and assessment of ecological effects resulting from the release of genetically engineered microorganisms to terrestrial environments. Advances in Applied Microbiology
38, l-99.
Tabatabai M. A. (1982) Soil enzymes. In Methoak of Soil Analysis, Part 2 (A. L. Page, R. H. Miller and D. R. Keeney, Eds), pp. 903-947. American Society of Agronomy, Madison. Wirth S. J. and Wolf G. A. (1992) Micro-plate colourimetric assay for endo-acting cellulase, xylanase, chitinase, 1,3-/Jglucanase and amylase extracted from forest soil. Soil Biology & Biochemistry
24, 511-519.
Wollum A. G. II (1982) Cultural methods for soil microorganisms. In Metho&of Soil Analysis, Part 2 (A. L. Page, R. H. Miller and D. R. Keeney, Eds), pp. 781-802. American Society of Agronomy, Madison.
Ekvier Science Ltd. Printed in Great Britain
Pergamon
0038-0717/94$6.00+ 0.00
INTERFERENCES, LIMITATIONS AND AN IMPROVEMENT IN THE EXTRACTION AND ASSESSMENT OF CELLULASE ACTIVITY IN SOIL* LISE K. GANDER,‘? CHARLESW. HENDRICKS* and JACK D. DOYLE’
‘ManTech Environmental Technology Inc. and *U.S. Environmental Protection Agency, Environmental Research Laboratory, 200 S.W. 35th Street, Corvallis, OR 97333, U.S.A. (Accepted
30 May 1993)
Summary-A practical modification of a reducing-sugar method to estimate soil (carboxymethyl) cellulase activity is described and used to compare three non-sterile soils. The original method involves the formation of a ferric-ferrocyanide complex in soil extracts prepared and exposed to substrate in 2M acetate buffer. Because high acetate concentrations compromise this reaction, such extracts require considerable (e.g. 20-30-fold) dilution prior to activity determinations. For soils with low organic matter, such as one used in this study, undetectable activity often results. We used 67 mu acetate buffer and were able to detect and compare cellulase activities in three diverse soils using undiluted extracts. The values obtained were more reproducible, but in a different range from those of the original method. Our modification yielded activities that appeared to be unchanged over the extraction buffer concentration range of 1t-140 mM acetate. The use of undiluted soil extracts prepared in less concentrated buffer simplifies and extends the practicality of the assay by: (a) reducing the importance of using matrix-matched standard curves; (b) requiring less sample manipulation and glassware, and (c) preventing the dilution to extinction of enzyme activity. It also improves the usefulness of the method as an indicator of ecological effects (biomass turnover) due to the introduction of nonindigenous microorganisms and chemicals of environmental concern. These factors make the method more competitive with other reducing-sugar assays used to measure cellulase activity in soil.
of soil cejlulase activity might be useful in environmental risk assessment. There are many calorimetric assays for soil cellulase activity (e.g. Benefield, 1971; Pancholy and Rice, 1973; Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990; Wirth and Wolf, 1992). Most involve reducing-sugar (or glucose) determinations, and use reagents such as dinitrosalicylic acid (DNS; Miller et al., 1960), alkaline copper (Somogyi, 1952) or glucose oxidase (Benefield, 1971). Carboxymethylcellulose is commonly used as a substrate, because of its high reactivity (Miller et al., 1960; Kanazawa and Miyashita, 1986). However, this substrate is less than ideal (Lindner et al., 1983; Breuil and Saddler, 1985). Differences in measurable cellulase activity can be method-specific, reflecting both the sensitivity of the detection method and the choice of substrate. Soil sources and extraction methods are also influential. Some researchers suggest calorimetric assays used to quantify released sugars are poor indicators of overall cellulase hydrolytic potential, particularly for samples with low B-glucosidase activities (Chan et al., 1989). Others indicate this finding is substrate-dependent (Rivers et al., 1984). In many cases, interferences are related to the degree of substrate hydrolysis (Rivers et al., 1984). Choosing an appropriate assay often means choosing appropriate compromises (Breuil and Saddler, 1985).
INTRODUCTION Cellulase [ 1,4-( 1,3; 1,4)-j?-D-glucan-4-glucanohydrolase] is responsible for the hydrolytic conversion of cellulose to glucose (saccharification). At least three classes of enzymes typically play roles in this process: endoglucanase (EC 3.2.1.4), exoglucanase (cellobiohydrolase; EC 3.2.1.91) and /?-glucosidase (cellobiase;
EC 3.2.1.21). The relative contributions of these three enzymes to overall cellulase (carboxymethylcellulase) activity are source-dependent (Coughlan and Ljungdahl, 1988) and have been investigated in a variety of plants, fungi, bacteria and soils. Through the action of cellulase, important energy sources for soil microorganisms are released. Hence, soil cellulase activity is an indicator of biomass turnover. Biomass and related measurements are useful in assessing soil stability, nutrient cycling and both agrochemical and microbial influence on soil metabolic processes. Increased interest in the ecological effects of toxic chemicals, hazardous wastes and genetically engineered microorganisms has prompted a need for methods to assess differences in perturbed vs non-perturbed soil. Because cellulase can influence biomass turnover and carbon cycling, determinations l@ Government of Canada. TAuthor for correspondence. 65
66
LISE K. GANDER et al.
Soil enzymes have lower activities than those from microbial and plant sources (Kanazawa and Miyashita, 1986). Also, even under optimized conditions of pH and temperature, extraction rates can be slow (Kanazawa and Miyashita, 1986). For this reason, combined soil extraction and incubation periods of 24-48 h are common for soil cellulase assays (Benefield, 1971; Pancholy and Rice, 1973; Schinner and von Mersi, 1990). Companion reducing-sugar or glucose determinations for cellulase activity estimates are relatively short (e.g. < 1 h), but vary with respect to sensitivity and reliability. Reactions involving DNS are convenient, but not very accurate, especially at low ( < 1 mM) reducing sugar concentrations (Lindner et al., 1983; Foroughi and Gunn, 1983). Also, results are influenced by dissolved oxygen, citrate buffer, divalent cations (Foroughi and Gunn, 1983) and the composition of the cellulase complex being measured (Breuil and Saddler, 1985). Because non-linear responses are generated in DNS assays, cellulase activities are dilution-specific (Breuil and Saddler, 1985). The Nelson-Somogyi alkaline copper assay (Somogyi, 1952) is less convenient, but more sensitive and reliable than that involving DNS (Lindner et al., 1983; Breuil and Saddler, 1985). The reaction is affected by incubation conditions, choice of substrate and other components in the cellulase complex; however, to a lesser extent than the DNS assay (Breuil and Saddler, 1985). Removal of trace metals from soil extracts improves Nelson-Somogyi detection limits (M. A. Tabatabai, pers. commun.). Unlike DNS, the alkaline copper reagent reacts stoichiometrically with the number of hemiacetal reducing groups present (Breuil and Saddler, 1985). However, biphasic responses with higher values at lower enzyme (sample) concentrations have been reported (Linder et al., 1983; Breuil and Saddler, 1985). These results are less reproducible and probably less representative of hydrolytic potential than those obtained for more concentrated samples (Breuil and Saddler, 1985). Values determined by the DNS method are often 45% higher than those by the Nelson-Somogyi method (Breuil and Saddler, 1985) which in turn are about 90% higher than those by the glucose oxidase-horseradish peroxidase assay (Kanazawa and Miyashita, 1985). This reflects the non-specific nature of reducing-sugar assays: i.e. substances other than glucose can be measured. The glucose oxidase-horseradish peroxidase method of Benefield (1971) is specific, reproducible and sensitive. This determination avoids the inaccuracies associated with co-measurement of non-glucose products of cellulolysis (e.g. cellobiose). Glucose oxidase reacts only with glucose. The reaction is compromised by pigments and ions released during soil extractions (Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990). Hence, soil extracts are often diluted or treated with phenol-binding polymers (Kanazawa and Miyashita, 1986) prior to
assays for glucose. Only the latter permits low detection limits. The ferric-ferrocyanide reducing-sugar assay (Park and Johnson, 1949) has been applied by Schinner and von Mersi (1990) for soil cellulase activity measurements. The sensitivity of this assay is affected by the presence of aldehydes, phenols, catechols (Litwack, 1960), acids, high ionic strength (Cooper, 1977) and any compounds with significant oxidative or reductive potential (P. Turner, pers. commun.). Many of these interfering substances are found in soil extracts (Brady, 1974) originating from the soil itself or introduced with the extracting buffer. Hence, this cellulase determination is likely to generate matrixvariable responses of at least two types: those relating to the choice of soil, and those relating to the choice of extracting buffer. In our ‘research concerning the effect of environmental stressors on carbon cycling, we seek simple methods to assess differences in perturbed vs non-perturbed soil systems. Our early work with fertic-ferrocyanide approaches to determining cellulase activity suggested that results in soil systems might be dependent upon both extraction and dilution conditions (unpubl. observ.). We have evaluated the influence of soil extraction conditions and suggest not only a practical, more versatile modification of the Schinner and von Mersi (1990) method, but also potential limitations of the approach to assess soil cellulase activity. Additional information concerning soil properties, /I-glucosidase and dehydrogenase activities, and populations of indigenous microbes is presented and related to cellulase activity. MATERIALS AND METHODS
Soil characteristics and preparation
Four soils were collected as vegetation-free samples from the upper 5 cm of selected locations (meadow, agricultural, forest and riverbank) in Linn and Benton counties, Ore. (Tables 1 and 2). Meadow soil was used only in tests to establish appropriate concentrations of extracting buffer. Each soil was sieved (2 mm) and stored in a sealed plastic bag (room temperature, light-free) for not more than 2 weeks. Two days before use, soils were refrigerated (4°C) divided into two lots, and either adjusted to 60% of full water holding capacity (WHC; Pramer and Schmidt, 1965; Doyle et al., 1991), or left unadjusted (field-moist). Three subsamples were removed for replicate analyses of soil moisture (Pramer and Schmidt, 1965) selected chemical variables (Horneck et al., 1989) including pH (Stotzky et al., 1992), and the enumeration of indigenous microbes. All measurements were made in triplicate. Microbial enumeration
Indigenous bacterial colony forming units (cfu) and fungal propagules were enumerated from spreadplate distributions of serially diluted soil on selective
61
effects in assessing soil cellulase
Extraction
Table I. Chemical and microbioloeical characteristics of soils Agricultural (Malabon silty clay loam)
PH Organic matter (%) CEC (meq IOOg-‘) N&N (kg g-‘) NO,N (pg g-‘) P&g g-‘) KOlg g-‘) CaOlg g-‘) MgOlg g-‘) FeoCg g-‘) Total fungi (log propagules gg’) Total bacteria (log cfu g-‘)
60% WHC
Fieldmoist
60% WHC
Fieldmoist
8.48 ( f 0.25) 5.43 ( * 0.03) 3.76 26.12 8.04 6.37 34 437 II.3 3.5 130
36.89 ( f 0.54) 5.64 ( * 0.09) 3.87
34.65 ( f 0.81) 6.66 ( f 0.05) 13.83 40.92 3.06 8.62 I3 725 23.9 8.2 142
31.63 ( + 0.36) 6.60
I .26 ( k 0.08) 7.30 ( f 0.04) 0.37 14.09 6.35
5.83 ( + 0.04) 4.76 ( + 0.23)
5.80 ( + 0.06) 5.51 ( + 0.09)
5.47 (f0.16) 7.02 (kO.14)
WHC = water holding capacity; CEC = cation exchange capacity;
media To indicate the extent of microbial survival in soil extracts prepared for cellulase assays, aliquots of these preparations were plated also. Soil extract agar (Wollum, 1982) containing 100 fig cycloheximide ml-’ (Sigma) was used to enumerate total soil bacteria. Martin’s agar (Martin, 1950) was used to estimate fungal propagules. Dilutions were prepared (in sterile tap water) in triplicate per soil type. Two plates of each medium were inoculated per dilution or extract. Values were normalized to mean cfu (or propagules) g-r oven-dry soil equivalent prior to log,, transformations. Enzyme activity determinations
Enzyme activities are expressed as the mean ( f SEM) of three replicates, in units of product formed g-r oven-dry weight equivalent of (wet) soil and incubation time. The activities of soil /J-glucosidase and dehydrogenase were determined by standard methods (Tabatabai, 1982). Cellulase (endoand exocellulase) was extracted and assayed both using the Schinner and von Mersi (1990) variation of the Park and Johnson method (1949), and with the modifications described later. For these applications, a swollen suspension of carboxymethylcellulose sodium salt (medium viscosity; Sigma; 0.7% w/v, in acetate buffer) was used as the substrate. To indicate
Table 2. Cellulase activity in four concentrations of undiluted acetate extracting buffer Acetate Buffer (mM) II 53 I05 I40
Riverbank (Willamette silt loam)
Fieldmoist
Characteristic %H,O
Forest (Price-Ritner gravelly silty clay loam)
Endoglucanase activity (nmol glucose equiv. g-’ soil 24 h-‘) 467*52 439 * 53 453 f 93 398 + 77
Each activity is the mean + SEM of three replicates of fieldmoist, Willamette loam (meadow) soil (pH = 5.9, organic matter = 0.86%. Fe = 86 pg g-l).
( f ‘3.06) 13.99
5.40 (kO.10) 7.01 ( * 0.04)
3.07 (kO.18) 4.88 ( f 0.04)
60% WHC 19.80
( f 0.79) 7.12 ( + 0.02) 0.42
3.25 (50.19) 5.67 ( f 0.41)
+_values indicate the SEM of three replicates.
enzyme extraction efficiencies, acetate solutions containing 500 pg of crude endoglucanase from Trichoderma viride (Sigma) were extracted and assayed for cellulase activity in the presence or absence of soil. The combined cellulase assay and soil extraction procedure (Schinner and von Mersi, 1990) involves three reactions. (1) Soil (cellulase) is extracted under buffered, acidic conditions (pH 5.5, 5O”C, 24 h) and allowed to bind to carboxymethylcellulose substrate. This increases the number of glucose residues available from the modified cellulose. (2) Potassium ferricyanide is reduced (in the presence of excess potassium cyanide) under alkaline conditions (pH 10.5, lOO”C, 15min) to potassium ferrocyanide by glucose (and other reducing agents, constituting interferences). (3) Ferrocyanide reacts with excess ferricyanide (in the presence of ferric ammonium sulfate and a surfactant) under acidic conditions (pH < 2, room temp., 60min) to produce a ferric-ferrocyanide complex. This chromophore is measured spectrophotometrically (690 nm) and related to cellulase activity. The following modifications of the Schinner and von Mersi (1990) cellulase assay were used. (1) Controls: one soil-free control containing substrate and three replicate (soil-containing) substratefree controls were incubated with each reaction set. The former was used to zero the spectrophotometer and, when required, to dilute (post-reaction) offscalehigh samples. The technique of post-reaction dilutions was validated (agreement ranged from 97 to 115%) prior to being applied, by assaying 10 sets of three matched replicates. These matched replicates consisted of either three pre-reaction dilutions + three post-reaction dilutions (for extracts prepared
LISEK. GANDERet al.
68 1.50
1.25
GLUCOSE (nmol ml- I) Fig. 1. Matrix-matched, soil-free, standard curves for (carboxymethyl) cellulase assay used to relate absorbance to product formation (enzyme activity). Each acetate concentration corresponds to the following dilutions of a 2 mM acetate stock: 64 nw = 3t-fold; 107 m&i= 21-fold; 129 mu = W-fold, and 320 mu = &fold. Correlation coefficients for the linear regressions are: 0.999 {no acetate), 0.995 (64 mbt), 0.996 (107 m@. 0.996 (I29m@, and 0.915 (320m~).
in 2 M acetate), or three undiluted extracts plus three post-reaction dilutions (for extracts prepared in 67 mM acetate). The mean absorbance of replicate, substrate-free controls was used for background subtractions. Thus, both soil and enzyme-free substrate contributions were taken into account. (2) Preparation of soil extract: rather than filtering the reaction mixture, ~nt~fugation (2 min at 12,000g; Beckman Microfuge 11) followed by removal of the supernatant was used to isolate extract for future reactions. (3) Assessment of microbial influence: soil extracts were assayed for cellulase activity both before and after filtering through 0.2pm membrane filters. (4) Extraction-incubation matrix: in addition to 2 M acetate, pH 5.5, diluted lO-30-fold for reducing sugar quantifications, various concentrations of pH 5.5 acetate buffer were examined. Because S&inner and von Mersi (1990) indicate that agricultural and forest soils extracted in 2 M acetate buffer must be diluted 20-30-fold prior to cellulase activity determinations, 67 m&racetate (equivalent to a JO-fold dilution of 2 M acetate) was most often selected. Mat~x-match~ standard curves of glucose [i.e. glucose standard (Sigma) + acetate] were used to compensate for acetate-induced signal quenching to establish linear relationships between absorbance and glucose concentration. (5) Activity units: cellulase activities are expressed on the basis of glucose equivalents g-r dry soil and total reaction exposure time. These units reflect the non-specificity of reducing-sugar assays,
which rely upon increased reducing potential (not necessarily free glucose residues) to measure the interaction between endo- and exocellulase (Sinsabaugh ef al., 1981). RESULTS
Soil-free standard curves constructed to assess the effect of pH 5.5 acetate (and other extracting buffers) on glucose quantification showed that signal quenching does occur, particularly at acetate concentrations of 129 and 320m~ (Fig. 1). The substitution of pH 5.5 citrate as an extracting buffer resulted in a 3-fold increase of the quench effect observed with acetate: 44 mhi citrate was equivalent to 129 ma4 acetate. No apparent differences in soil cellulase activity were found for a range of undiluted acetate extracting buffer concentrations (1 I-140 mM; equivalent to 190-14-fold dilutions of 2 M acetate) (Table 2). Soil-free, time-course assays of commerciallyavailable, crude endoglucanase from Trichina uiriak were examined both under extraction conditions using 2 mr+sacetate (S&inner and von Mersi, 1990), and those of our 67 mu acetate modification. Results indicated not only similar kinetics for the two approaches, but also that, in the absence of soil, our modification recovers at least as much cellulase activity as the original method (Figs 2 and 3). When soil spiked with standard enzyme was extracted and assayed in either 67m~ or 20- and 30-fold dilutions
Extraction effects in assessing soil cetlulase
69
24
Incubation Time (h) Fig. 2. Timecourse study of standard (T. vi&k) endoglucanase activity in soil-free assays using both 67 mhi acetate buffer (undiluted, except post-reaction, where required) and 30-fold (pre-reaction) dilutions of 2 M acetate buffer. Each value is the mean * SEM of three replicates. Correlation coefficients for the logarithmic curve fits are: 0.915 (67 mM) and 0.977 (2 M).
ABC 67&i
&et&e (1x)
2M Acewe wo
A
S
C
2M Amate (3mO
Fig. 3. Overall celhdase activity in soil-free extracts containing 500 pg endoglucanase from ?‘. viride (A), fieid-moist forest soil plus 5OO~g endoglucanase from T. viride (B), and field-moist forest soil (C). Measurements were in 67 mM acetate extracts (undiluted, except post-reaction, where required) and 2 mtu acetate extracts diluted @e-reaction) either 20- or 30-fold. Each value is the mean & SEM of three replicates.
LISEK. GANDERet al.
70
a
B
A
A
Agricultural tSoi1
B
A
B
Rivorbsnk sdl
Forest Soil
Fig. 4. Cellulase activity as measured in extracts prepared from field-moist soils (A) and those adjusted to 60% WHC (B), using undiluted 67 mM acetate buffer (PH 5.5). Extracts were assayed before and after filtering through a 0.2 pm membrane. Each value is the mean + SEM of three replicates.
of) 2rn~ acetate buffer, the cellulase activities obtained were far from equivalent to the combined activities of standard plus soil enzyme. Instead, enzyme recoveries were compromised to 27% (67 mM acetate extracts), 16% (20-fold dilutions of 2 M acetate extracts), and 78% (30-fold dilutions of 2 M acetate extracts) of soil-free, standard enzyme activities (Fig. 3). Cellulase activity was detectable for three diverse soils when extracts prepared in undiluted 67 mM acetate were analyzed (Fig. 4). The agricultural and forest soils were found to be 7-8 and 15-18 times higher, respectively, in cellulase activity than the riverbank soil. Filter-sterilization of these extracts gave no appreciable difference in activity (Fig. 4). Likewise, adjusting soil moisture from field-moist conditions to 60% WHC (Table 1) yielded no pronounced changes in measured activities of cellulase (Fig. 4), /J-glucosidase (Table 3), or dehydrogenase (Table 3). Large increases in water content (e.g.
Table 3. b-Glucosidase
and dehydrogenase
Agricultural (Malabon silty clay loam) Enzyme activity fi-Glucosidase @g PNP produced Dehydrogenase @g TPF produced
g-’ soil h-‘) g-’ soil 24 h-‘)
WHC = water holding three replicates.
adjustment of agricultural and riverbank soils to 60% WHC) also coincided with increases in bacterial populations, but left fungal numbers relatively unchanged (Table 1). There were no detectable fungi and, relative to unextracted soil, reduced bacterial (2.39 f 0.09 g-’ agricultural soil, numbers 3.70 + 0.03 g-’ forest soil, and 2.04 + 0.13 g-’ riverbank soil) in 67 mM acetate extracts. The numbers of recovered bacteria corresponded to about 0.1% of the original bacterial populations in these soils (Table 1). No fungi or bacteria were detected in corresponding 2 M acetate extracts. When 2M acetate extracts were diluted lo-, 20and 30-fold to determine cellulase activity (Schinner and von Mersi, 1990), the riverbank soil was found to be below detection limits (Fig. 5). Although raw absorbance values were lower, the activity units for other soils analyzed by this method were markedly higher (about 2-12 times; Fig. 5) than those attained with undiluted 67 mM extracts (Fig. 4). Background-
activities
Forest (Price-Ritner gravelly silty clay loam)
Riverbank (Willamette silt loam)
Fieldmoisl
60%WHC
Fieldmoist
6O%WHC
Fieldmoist
60% WHC
170+8
157*4
169 f 7
170&8
3*0
2&O
22Sk9
227 f 10
26 *
I
36 f 2
19*0
capacity; PNP =p-nitrophenol; TPF = triphenylfonnazan; + values indicate the
19+ SEM
I of
Extraction effects in assessing soil cellulase
Fig. 5. Cellulase activity as measured in extracts prepared from field-moist soils (A) and those adjusted to 60% WHC (B), using 2 M acetate buffer (PH 5.5) diluted lo-, 20-, and 30- fold (Schinner and von Mersi, 1990). Each value is the mean f SEM of three replicates. Cellulase activity was undetectable in riverbank soil (all dilutions) and in 30-fold dilutions of agricultural soil adjusted to 60% WHC.
corrected absorbance values ranged from 0.070 to 0.366 for 30-lo-fold dilutions of 2mM acetate extracts of forest soil. Undiluted 67 mM acetate extracts had typical absorbance values of 0.418-0.535. Extreme dilution-specific variabilities in cellulase activity were observed for the same 2rn~ acetate soil extracts prepared and analyzed by the Schinner and von Mersi (1990) method (Figs 3 and 5), even in the presence of high amounts of added standard enzyme (Fig. 3). Pronounced differences associated with water adjustment were also noted in the cellulase activities for these extracts. This is in contrast to all the other enzyme activity measurements conducted, including cellulase activities for 67 mM extracts. Both /I-glucosidase and overall cellulase activities were lowest for riverbank soil (Table 3, Figs 4 and 5). However, the profile of relative /I-glucosidase activity for all three soils differed from that observed for cellulase dete~inations with 67m~ extracts. Mean /I-glucosidase activities for both treatments (with and without water adjustment) of agricultural, forest and riverbank soils were 164 : 170 : 2.4, respectively (Table 3; equivalent to 68: 71: 1). Cellulase activities for the same three soils showed relative ratios of 362:780:49 (Fig. 4; equivalent to 7: 16: 1). Contrary to reports of inverse relationships between soil dehydrogenase and cellulase activities (Schinner and von Mersi, 1990), the profile for relative dehydrogenase activity (Table 3) was similar to that seen for cellulase activity (Fig. 4) for all three
soils. Forest soil had the highest dehydrogenase and cellulase activities, while those for agricultural and riverbank soils were considerably lower. DISCUSSION
Our modification of using an undiluted soil extract prepared in a less concentrated buffer was effective in assessing overall cellulase activity in a variety of soils, including two containing < 1% organic matter. The original method of Schinner and von Mersi (1990) seems impractical for soils with low organic matter, probably because cellulase activity is diluted to extinction. In fact, Schinner and von Mersi (1990) recommend determining xylanase rather than cellulase activity in such soils. Sparingly small amounts of soil are involved in cellulase activity dete~inations of 20-30-fold dilute extracts. This could promote high sample-to-sample variability, and might account for both the differences in cellulase activity observed for the same 2 M acetate extract diluted over a range of values and those between field-moist and 60% WHC soils (Fig. 5). That cellulase activities for extracts prepared in 2 rnM acetate were lower with decreased soil amounts (produced by increased soil extract dilutions, or higher water content in soils) suggests the original method is easily biased when solutions representing different amounts of soil are used for ferric-ferrocyanide-based determinations. This problem
12
LISEK. GANDER et al.
is avoided by determining glucose equivalents in undiluted soil extracts prepared in less concentrated buffer. Such a modification not only increases the reproducibility and the sensitivity of the application, but also permits the post-reaction dilution of off-scale high samples. The use of undiluted, less concentrated, extracting buffer eliminates some of the difficulties associated with the Schinner and von Mersi (1990) approach to estimating soil cellulase activity. However, compromised recoveries of combined standard plus soil enzyme activities (Fig. 3) suggest that results are matrix-dependent, regardless of whether or not soil extracts are diluted. Of all the samples prepared and analyzed in this manner, the 30-fold dilution of soil extract spiked with standard enzyme came closest to approaching the activity of soil-free standard enzyme alone (Fig. 3). This suggests the exogenous enzyme may be binding to the soil or the presence of soil-contributed interferences. Presumably, interferences are diluted to undetectable concentrations in highly diluted soil extracts. Such factors might account for the lower cellulase activities (relative to l&30-fold dilute 2 M acetate extracts; Fig. 5) obtained for undiluted 67 mM acetate extracts (Fig. 4), which involve more amounts of soil. Thus, the ferric-ferrocyanide cellulase assay appears to suffer from similar drawbacks as that involving DNS (Breuil and Saddler, 1985). Both generate dilution-specific, matrix-variable responses of at least two types: those relating to the choice of extracting buffer, and those relating to the properties of the soil. In both the Schinner and von Mersi method (1990) and our modified cellulase assay, enzyme extraction and substrate exposure occur in one 24 h step. This promotes saturation kinetics and fails to distinguish between enzyme extractability and true activity. Also, cellulase activity units are expressed as a function of dry soil (rather than protein concentrations) for the entire 24 h reaction. Combined, these facts suggest that activities determined by these methods must be relative. However, other soil enzyme assays involve combined 24 h enzyme extraction and substrate exposure, and use similar activity units (Pancholy and Rice, 1973; Tabatabai, 1982). These assays provide relative values which are commonly used as indicators of perturbed vs non-perturbed systems (Malkomes, 1991). In soil applications, an extracting buffer of high ionic strength would be more likely to kill indigenous microorganisms and mask differences in ionic character of the soils used, providing a more universal testing system. However, high ionic strength is a major interference in the ferric-ferrocyanide reducing-sugar assay (Cooper, 1977). Thus, we investigated the rationale and practice of using acetate (pH 5.5) and other choices of extraction-reaction buffers. Acetate buffers are commonly used for soil cellulase extractions (Pancholy and Rice, 1973; Kanazawa and Miyashita, 1986; Wirth and Wolf, 1992). Cellulase
activity has a pH optimum of ca 5 (Kanazawa and Miyashita, 1986; Schinner and von Mersi, 1990). The enzyme should form its strongest ionic attachments to carboxymethylcellulose (pK,-4) at pH -3-5. Because the only stoichiometric reaction involved in ferric-ferrocyanide-based reducing sugar determinations is compromised by formic acid (Litwack, 1960), the use of acetate is somewhat questionable. Yet, in comparing acetate (two carbon) and citrate (six carbon) buffers, we found that although both cause signal quenching, acetate does so to a lesser extent, and thus remains a preferable choice. Also, soil extracts prepared in acetate buffers are less pigmented (Kanazawa and Miyashita, 1986). For greater versatility in assessing various soil types for relative cellulase activity (especially those expected to have low values), the use of an undiluted, less concentrated (e.g. 67 mM) acetate solution seems advisable. Our modification of using a less concentrated extraction buffer for cellulase determinations permitted the analysis of undiluted soil extracts, including those of low activity. This broadens the versatility of the method and makes it more competitive with other reducing-sugar assays used to determine cellulase activity. However, for maximum confidence in soil cellulase activities estimated by this approach, we recommend the following: (1) identical preparation of replicate soil extracts and controls; (2) inclusion of an internal standard, such as commercially-available cellulase enzyme, assayed in the presence and absence of each soil type, as an indicator of both enzyme extraction efficiency and the extent of soil-contributed interferences; (3) recognition that activity values are relative, and probably should not be compared across soil types. The determination may be most useful for assessing the effects of different treatments within one soil type. We believe our modification and recommendations increase the practical application of this cellulase assay as an indicator of biomass turnover and ecological effects due to the introduction of non-indigenous microorganisms and chemicals of environmental concern. thank Marcia Bollman and Drs Thomas L. Bott. Robert Sinsabauah and Paul M. Turner for their review and comments on the manuscript. We thank Lynn Bucao, Bonnie Hugley (ManTech Environmental) and Central Analytical Laboratory (Oregon State University, Corvalhs, Ore.) for technical support. This research was funded by the U.S. Environmental Protection Agency and conducted in accordance with an approved quality assurance plan, and has been subjected to Agency review. Acknowledgements-We
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