0038-0717 Xl 04030745502.013 0 Copyrtphs 0 1981 Perpdmon Pies, Ltd
FLUOROMETRIC FOR SOIL LIPASE
A. B. C~CIPERand H. W. MORGAN Hamilton Science Centre, Ministry of Works and Development, Private Bag, Hamilton and Department of Biological Sciences, University of Waikato, Hamilton, New Zealand (Accepted20 December 1980) Summery-Optimum
conditions for the precise assay of soil lipases are described, using the fluorogenic substrate Cmethyl umbelliferone nonanoate (4MUN). The method utilizes a 3 h 0.1 M tetra-sodium pyrophosphate (pH 7.5) extraction, followed by a 10 min assay. Soil lipase is measured with average 95% confidence limits of 5.6%. Activity was optimal between 30-4o”C. Soil extracts characteristically possessed two pH-activity peaks, one each side of neutrality. Extracts were stable when stored at 4’C for at least 5 days.
INTRODUCIION The fate of lipids in soil, either those occurring naturally or as added wastes, is not well understood (Braids and Miller, 1975). A significant proportion of lipids entering soil will be in the form of triacylglycerols (or triglycerides), this being the primary storage fat in plant and animal tissue. The initial step in subsequent degradation will therefore involve the enzyme triacyiglycerol acylhydrolase (EC 184.108.40.206), more commonly termed lipase. Lipase activity in soil can be measured directly by incubation with an emulsified triglyceride substrate followed by analysis of the amount of fatty acid released (Pokorna, 1964). But this method requires incubation for 72 h to obtain detectable concentrations of fatty acid. Unless each soil is individually screened, errors due to fatty acid adsorption to soil colloids, can confuse interpretations of activity. Pancholy and Lynd (1972, 1973) described a sensitive ~uorometric technique for lipase assay using the butyrate ester of 4-methyl umbelliferone (4MUB) as substrate. The assay procedure involves a 1.5&y incubation of soil with olive oil, before extraction of lipase in phosphate buffer and subsequent release of the fluorescent Cmethyl umbelliferone (4MU) by hydrolysis of the non-fluorescent substrate. Such an incubation period is time consuming, and biased in favour of those soils capable of responding to lipid additions to the greatest extent. In addition, lipase measurements at the end of the experiment may bear no relationship to levels of lipase in unamended soil. Estimation of lipase activity in unamended soils is complicated by the possibility of the 4MUB substrate being hy~olysed by non-specific esterases (Pancholy and Lynd, 1971). We report the development of a rapid and specific method of lipase estimation, applicable to soils without prior addition of substrate. MATERIALSAND
Soils Surface samples (0-15cm) of four soils (Table 1) were collected and sieved (~2 mm). before storing at
field moisture in sealed plastic bags at 4°C for up to 2 yr.
The acetate, butyrate, octanoate and laurate esters of 4MU were obtained from ICN Laboratories, Cleveland, Ohio, U.S.A., the heptanoate, nonanoate, palmitate, and oleate esters from Koch-Light Laboratories, Colnbrook, England, and the 4MU from Eastman-Kodak, U.S.A. Stock solutions of esters and 4MU were prepared at 10 mM in 2~thoxyethanol and stored at 4°C. Working standards of 4MU at 10~~ and 100 fly were prepared daily by dilution with distilled water. Assay of soil lipase activity
To 1 g of field moist soil in 50 ml Erlenmeyer flasks, was added 10 ml of 0.1 M tetra-sodium pyrophosphate butler, pH 7.5. Flasks were incubated at 30°C with orbital shaking at 120 rev min-’ for 3 h, and the slurry centrifuged at 4000 g for 15 min. A 0.1 ml aliquot of the dark brown supernatant was added to 3 ml of 0.1 M Tris-HCl buffer pH 7.5 and equilibrated at 30°C for 5 min, then 0.1 ml of 10m~ 4-methyl. umbelliferone nonanoate substrate solution was added (giving a final concentration of 312~~) and vortex mixed. The resultant emulsion was placed in a Farrand Mk I spectrofluorometer and zeroed with excitation at 340nm and emission at 450 nm with 5 nm exit and entrance slits. The sample was incubated at 30°C for 10min and the increase in fluorescence determined. Rates of non-enzymic substrate hydrolysis were determined using the same procedure, except that boiled soil extracts were used. Standard curves to relate Auorescent response to 4MU concentration were constructed using the same matrix as sampies, again with boiled enzyme extract. The use of an identical matrix solution for standards is essential because excitation of the 4MU and therefore the degree of response, is altered by pH, the presence of substrate or the pyrophosphate extract. Pyrophosphate extracts caused considerable quenching of fluorescence of the 4MU standards. The addition of Ca2 +, a common cofactor in lipase assays (Desnuelle, 1972) did not increase some extracts additions mation of precipitates.
activities of soil extracts. In of Ca2’ resulted in the forwhich interfered with the
A. B. COPPER and H. W. MORGAN
Table 1. Properties and lipase activities of four soils. Lipase activities were measured by the in situ and pyrophosphate extraction methods and are expressed as a mean + 2 SD of 10 separ-
ate assays Lipase activity (nmol CEC (m-equiv
Hamilton clay loam* Ruatangata clay* Horotiu silt loam? Waiteti sand*
5.7 5.3 5.5 5.5
4.6 6.5 12.1 9.7
at pH 7)
29.0 54.0 24.4 10.0
29.0 54.0 24.4 10.0
4 MU produced In situ
method 125 + 92+ 162 f 363 +
19 18 18 65
Pyrophosphate method 161 + 95 + 194 * 597 +
10 7 9 25
* New Zealand Soil Bureau Bulletin 26 (3). t Sarathchandra (1978). assay, therefore no Ca’+ was added. Variations on this standard method of assay are described in the appropriate results section. Organic matter and protein in pyrophosphate extracts were determined by the methods of Hayano (1977) and Lowry et al. (1951) respectively. Measurement of lipase activity in situ
To 0.1 g of field moist soil in a 50 ml Erlenmeyer flask was added 4.5 ml of 0.1 M Tris-HCl buffer pH 7.5. Flasks were equilibrated at 30°C and shaken for 10min at 120revmin-’ on an orbital shaker. After equilibration, 0.5 ml of 10 mM 4MUN substrate solution was added followed by shaking for a further 10min. Flasks were then cooled in an ice bath, the contents poured into pre-cooled centrifuge tubes and centrifuged at 4000g for 10min at 2°C. Supernatants were analysed fluorometrically for 4 MU concentration. To determine the extent of enzyme activity after 10min incubation, controls were run in which substrate was added to the soil suspension immediately before flasks were transferred to the ice bath. In addition, the extent of 4 MU adsorption onto each soil was determined by adding a range of 4 MU concentrations in 0.1 M TrisHCl buffer pH 7.5 to 0.1 g soil samples, incubating for lOmin, and measuring the fluorescence of the centrifuged supernatant. Enzyme activity by both procedures was expressed as nmo14 MU produced min- ’ g-i of dried soil. RESULTS
Comparison of in situ versus extracted enzyme activity
With the four soils tested, activity in the pyrophosphate extracts was equal to or greater than that determined when substrate was incubated directly with the soil (Table 1). Increased activity with the pyrophosphate extract may reflect the lack of diffusion barriers to the substrate which would exist in soils. The pyrophosphate extraction method was more precise than the in situ method (average 95% confidence limits of 5.6 and 16.0% respectively). The higher degree of variability with the in situ analysis arose principally from variations in lipase activity occurring after Cooling in ice following incubation. Attempts to terminate the enzyme activity more abruptly by adding acid or alkali were unsuccessful, either due to quenching of the fluorescence of 4MU or to non-enzymatic hydrolysis of the substrate. An additional complication with the in situ method was the necessity to correct
for adsorption to soil colloids of the 4 MU produced. Different adsorption capacities of each soil meant that this correction needed to be evaluated for each soil assayed, and coupled with the lower precision of the in situ procedure made the method less desirable when compared to the pyrophosphate extraction technique. Extraction of lipase activityporn soil
A range of extractants was tested for efficiency of recovery of lipase activity from soil (Table 2). Highest lipase activities were obtained in extracts of 0.1 M tetra-sodium pyrophosphate pH 7.5. Most of the extractable lipase activity was recovered by a single 3 h extraction with pyrophosphate. When the soil residue, of other extractants used, was re-extracted with tetrasodium pyrophosphate, the combined activities of both extractants approached that present in a single pyrophosphate extraction. Exceptions were soils initially extracted with either EDTA or 0.1 M NaOH which caused irreversible inhibition of lipase activity. The extraction procedure of Pancholy and Lynd (1972) recovers only 14% of the lipase activity extracted by pyrophosphate. Tetra-sodium pyrophosphate is a general soil organic matter extractant which has been used to extract soil urease (Nannipieri et al., 1974). Although the amount of urease extracted did not correlate with the amount of extracted soil organic matter, lipase activity extracted from the Hamilton clay ioam was more closely correlated to extractable soil organic matter than extractable protein (Table 3). The effect of extractant to soil ratio on the efficiency of extraction was determined for a range of soils using tetrasodium pyrophosphate as extractant. In all cases the amount of activity recovered obeyed a Langmuir curve, with greater than 90% of the theoretical maximum activity (as determined from regression analysis) being extracted at solution to soil ratios of 1O:l. Results for Hamilton clay loam are presented in Fig. 1. Solution to soil ratios of 1O:l were routinely used because at higher dilutions efficient extraction was compromised by excessive dilution. Choice of substrate
A range of commercially-available fatty-acyl-esters of 4 MU were tested for their suitability as substrates for extracted soil lipase determination. Kinetic values, K, (substrate cont. at 0.5 V,,,,,) and V,,,,, (maximum velocity of enzyme activity), are presented for these
Fluorometric assay of soil lipase
Table 2. Comparison of the efficiency of extraction of lipase from Hamilton clay loam by 0.1 M tetra-sodium pyrophosphate and a range of other extractants all used at soil to extractant ratios of 10 to 1 for 3 h Lipase activity (nmo14 MU produced g- 1min- i)
In extract Distilled water Tris-HCl 0.1 M (pH 7.5) Phosphate buffer 0.1 M (pH 7.5) Phosphate buffer 0.1 M (pH 7.5)* Phosphate buffer 0.1 M + 2 M urea Phosphate buffer 0.1 M + 1 M KC1 Phosphate buffer 0.1 M + 1 M KC1 + 0.01 M EDTA Acetate buffer 0.5 M pH 5.8 Tetra-Na pyrophosphate 0.1 M (pH 7.5) Tetra-Na pyrophosphate 0.1 M + 10% ethanol Tetra-Na pyrophosphate 0.1 M + 2 M urea NaOH 0.1 M
16 38 50 23 104 54 6
In pyrophosphate extract of residue 142 120 110 141 45 100 5
14 160 156 155 4
140 5 6 8 4
* Extracted according to the method of Pancholy and Lynd (1972) using a soil to extractant ratio of 2: 1 for 30 min. Table 3. Effect of extraction time on the recovery of lipase activity, organic matter and total protein from Hamilton clay loam extracted with 0.1 M tetra-sodium pyrophosphate
Lipase activity (nmol 4 MU produced g- ’ mini)
Organic matter (absorbance at 280nm g-i)
Protein (mg g- i)
1 2 3 6 15
122 144 160 156 156
152 175 190 218 262
1.5 1.7 1.9 2.4 4.8
substrates and a Hamilton clay loam pyrophosphate extract in Table 4. Acetate and butyrate esters are poor substrates for lipase estimations due to the possibility of non-specific esterase action, high rates of non-enzymatic hydrolysis under the conditions employed, and low rates of enzymatic hydrolysis. The medium chain length esters (Cs, Ca and Crz) were
attacked most rapidly and possessed suitably low K, values (high affinities). Essentially similar results were obtained when pyrophosphate extracts from 10 New Zealand topsoils were assayed against the same range of substrates. The nonanoate ester was selected as the most appropriate substrate and was routinely used in assays at a concentration of 312~~, well above the highest K, value obtained for all soil extracts assayed with this ester. The extent of 4MUN hydrolysis by soil extracts was linear with time for at least the first 15 min, even Table 4. Michaelis-Menten
constants for the hydrolysis of fatty-acyl-esters of 4 MU by soil lipase extracted from a Hamilton clay loam Substrate
Esters of 4 MU
Fig. 1. Effect of soil to extract ratio on the recovery of lipase activity from Hamilton clay loam.
Butyrate (C,) Heptanoate (C,) Octanoate (Cs) Nonanoate (C,) Laurate (C, J Palmitate (C,,) Oleate (Cis:i)
MichaelkMenten constants I’,,,,, (nmol4 MU produced g-’ mu-‘) K, (FM) 35 17 12 12 13 19 38 18
30 36 74 142 162 120 32 47
A. 3. COOPERand H. W. MORGAN
Fig. 3. Effect of pH on rate of hydrolysis of 4MUN by
Fig. 2. Hydrolysis of 4MUN with time by pyrophosphate extracts of four soil types. O----O. Hamilton clay loam; 0-G. Ruatangata clay: A---A, Horotiu silt loam: m---a, Waiteti sand.
with the high activity of the Waiteti sand extract (Fig. 2). Incubations for 10 min were used routinely.
Maximal rates of hydrolysis of 4 MUN substrate by extracts from the four soils occurred between 3t34O”C. with relatively littfe change in activity up to 5O’C. Because of an increased rate of non-enzymatic substrate hydrolysis at higher temperatures, 3o’C was selected for routine assay. The extracted lipase proved to be extremely stable and extracts could be conveniently stored with little loss of activity. With the soil extracts tested no activity was lost when stored at -20°C or 4’C for up to 5 days (Table 5). Air drying of soils before analysis decreased measured activity by between lO-SOY/,depending upon the soil. No detectable difference in extractable lipase activity was apparent between soil when collected fresh or after storage at 4’C for up to 2 yr.
pyrophosphate extracts of Hamilton clay loam, Q--O: Ruatangata clay, C----D: Horotiu silt loam. n---a: and Waiteti sand, D----m. Buffers used were citric acid-Na citrate (pH 4.c6.2). potassium phosphate (pH .5.2-&O),and Tris-HCI (pH 7.C-lO.O),all at 0.1 M.
Effect of pW
The excitation of 4 MU varies with pH (Fink and Koehler, 1970). In establishing pH-profiles excitation at 340 nm was used in preference to 365 nm, as at the latter wavelength 4MU exhibits a sharp increase in fluorescence over the pH-range 6.0-8.0. No quenching of 4 MU or enzyme activity due to the buffer systems used was detected. Pyrophosphate extracts from the four soils characteristically displayed two distinct pH optima; one at pH 6.S7.0, the other at pH W-8.5 (Fig. 3). The proportion of activity present in each peak varied depending on the soil. The choice of pH for routine assay is therefore somewhat arbitrary and may be dependent on soil type. Ideally the assay would be run at two pHs. As a compromise, a pH of 7.5 avoids the problems of increased non-enzymatic substrate hydrolysis at alkaline pH, and depression of fluorescence of 4 MU standards (prepared in a matrix of buffer, extract and 4 MUN) at acid pH.
Table 5. Effect of storage at different temperatures on relative activity of lipase extracted from soils by 0.1 w tetra-sodium pyrophosphate -20 c 5 days 10 days Soil Hamilton clay loam Ruatangata clay Horotiu silt loam Waiteti sand
I 10 0
100 100 100 100
95 100 100 100
Temperature of storage 4c 20 c 5 days 10 days 5 days 10 days (“, activity) 100 100 100 100
84 .88 96 100
88 90 95 100
73 77 90 97
Fluorometric assay of soil lipase DI!XU!WON
An assay for lipase activity using fluorescent substrates allows for rapid and sensitive determinations of enzyme concentrations. The initial application of this technique to soil enzymes (Pancholy and Lynd, 1972, 1973), did not fully exploit the inherent potential of this new approach. The use of the butyrate ester of 4 MU has two major disadvantages, the more serious being the hydrolysis of the substrate by nonspecific esterases as well as lipases. Furthermore, the rate of hydrolysis of 4 MUB by the lipases is not as great as some of the longer chain-esters, e.g. nonanoate (C,) and laurate (C,,). Also, the nonaquatic hydrolysis of the shorter chain-length esters is greater than that of the longer chain-Iength esters. A~thou~ this can be compensated for by the use of adequate controls, the overall sensitivity of the method is diminished. Pancholy and Lynd (1972) used phosphate buffer to extract soil lipase, thus overcoming adsorption of the 4 MU product to soil colloids, and diffusion barriers presented by the soil aggregates to an applied substrate. Both of these factors would make the determination of lipase activity in vivo more difficult and less accurate (see Table 1). However, we have shown that phosphate buffer is a poor extractant, recovering less than 30”,/,of total detectable lipase activity in Hamilton clay loam (Table 2). Furthermore, these results were obtained after a 3 h extraction and a 10: 1 extractant to soil ratio as against a 30min extraction and 2: 1 extractant to soil ratio described by Pancholy and Lynd (1972). The standard assay we propose using 0.1 M tetrasodium pyrophosphate as an extractant gives a 7-fold increase in the recovery of lipase activity over the method of Pancholy and Lynd (1972). Practically all extractable lipase was recovered after a single pyrophosphate extraction whereas with methods using phosphate buffer only a small proportion of the lipase present is extracted. Because the natural substrates for lipases are insoluble, the enzyme must operate at a solution-substrate interface and partition between these phases. Thus extractants such as phosphate buffer are unlikely to be effective in removing adsorbed enzyme in a single extraction. One possibility is to use surfactants to dissipate the interface effect but this may lead to some enzyme inactivation. The addition of urea, which disrupts hydrophobic bonds, improves the efficiency of phosphate buffer as an extractant, but 0.1 M tetra-sodium pyrophosphate proved to be superior to all other extractants (Table 2). Enzyme activity in pyrophosphate extracts was closely correlated with extractable organic matter but not extractable protein, possibly indicating that the bulk of the lipase is bound to organic colloids. Similar suggestions have been made for unease (Burns er al., 1972) and protease (Ladd, 1972). Properties of the extracted soil lipase bear similari-
ties to those of purified hpases from plant and animal
sources. Thus the soil enzyme extract preferentially degrades the medium chain length (Cs-CIZ) esters of 4 MU as does pancreatic and plant lipases (Jacks and Kircher, 1967). This is probably a compromise between an affinity of the lipase for the hydrophobic regions of long chain esters and spatial considerations which would tend to favour the short chain esters. Lipases are generally regarded as highly-stable enzymes (Desnuelle, 1972) and the soil enzyme proves to be no exception. Enzyme activity can be preserved by cold storage and was not diminished after 2 h at 70°C. ~eknowledge~ent-This research was partially funded by a University Grants Committee Scholarship to ABC. REFERENCES BRAIDS 0.
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HAYANOK. (1977) Extraction and properties of phosphodiesterase from a forest soil. Soil Bioiow “. cf Biochemistry
JACKST. J. and KIRCHERH. W. (1967) Fluorometric assay for the hydrolytic activity of lipase using fatty acyl esters of 4-methylumbelliferone. Analytical Biochemistry 21,
279-285. LADD J. N. (1972) Properties of proteolytic enzymes extracted from soil. Soil Biology & Biochemistry 4, 227-237.
LOWRY0. B., RO~E~URGH N. J., FARR A. L. and RANDALL R. V. J. (1951) Protein measurement with the Fohn phenol reagent. Journal of Biological Chemistry 193, 265-275.
NANN~PIERIP., CECCANTIB., CERVELLIS. and SEQUI P. (1974) Use of 0.1 M nvronhosnhate to extract urease from a podzol. Soif Biol&y d Bio&mistry 6,359362. PANCHOLYS. K. and LYNDJ. Q. (1971) Microbial esterase detection with ultraviolet fluorescence. Applied Microbiology 22, 939-941. PANCHOLY S. K. and LYNDJ. Q. (1972) Quantitative fluorescence analysis of soil lipase activity. Soil Biology & Biochemistry 4, 257-259. PANCHOLYS..K. and LYNDJ. Q. (1973) Interactions with soil lipase activation and inhibition. Proceedings Soil Science Society of America 37, 51-52. POKORNAV. (1964) Method of determining the lipolytic activity of upland and lowland peats and muds. Soviet Soil Science 1, 85-87. SARATHCHANDRA S. U. (1978) Nitrification activities of some New Zealand Soils and the effect of some clay types on nitrification. New Zealand Journal of Agricuirural Research 21, 615-621.