Microb Ecol (1989) 17:159-170

MICROBIAL ECOLOGY 9 Springer-VerlagNew York Inc. 1989

The Characterization of Amidohydrolases in a Freshwater Lake Sediment Paul J. Sallis and Richard G. Burns Biological Laboratory, University of Kent, Canterbury, Kent CT2 7NJ, U.K.

Abstract.

T h e properties o f three amidohydrolases, i.e., urease (I) EC 3.5.1.5, L-asparaginase (II) EC 3.5.1.1, and L-glutaminase (III) EC 3.5.1.2, were studied in sediment samples taken from a shallow eutrophic freshwater lake. Sediment samples were air dried (ADS) and stored for at least 3 m o n t h s before being enzymically characterized. T h e p H o p t i m u m o f I, II, and III were p H 7.0, 8.4, and 6.5-7.0, respectively, while III in soluble extracts from ADS was most active between p H 8.0 and 9.0. T h e temperature response o f the three enzymes in ADS gave Ea values o f 38.9, 41.6, and 35.9 kJmol -~ for I, II, and III, respectively. K m and Vm~x values for ADS I, II, and III were 1.2 m M and 1.9 #mol NH3 g-Zh-~; 0.8 m M and 4.1 #mol NH3 g l h - l ; and 1.25 m M and 17.4 ~mol N H 3 g-~h -t. K m values for all three enzymes in ADS extracts were at least an order o f magnitude greater than those o f the ADS. The susceptability o f each e n z y m e to proteolysis was followed in ADS and fresh wet sediment and c o m p a r e d with that o f III in an ADS extract. All sediment enzymes were found to be m o r e resistant than the c o m m e r c i a l preparation o f bacterial L-glutaminase subjected to the same treatment. These results suggested that I, II, and III all exist to some extent as colloid-immobilized e n z y m e fractions in freshwater sediments and are analogous to the stable e n z y m e fractions in soils.

Introduction Freshwater sediments (FWS) have long been recognized as i m p o r t a n t sites o f mineralization in the aquatic e n v i r o n m e n t , and to have an equivalent role to that o f the soil in terrestrial ecosystems [21]. However, although soils have been the subject o f n u m e r o u s investigations and the enzymological basis o f nutrient mineralization has been extensively characterized for a n u m b e r o f key C, N, and S cycle enzymes [4], freshwater sediment enzymes have remained relatively unstudied. Enzymes in soil are known to exist in various forms [5], and there are a n u m b e r o f reports indicating that a similar distribution occurs in sediments [6, 16, 20, 28, 32]. For example, it is likely that some sediment enzymes are Present as stabilized forms associated with clay and h u m i c fractions, but this POssibility has been largely unexplored [1]. Instead, the study o f aquatic en-

160

P.J. Sallis and R. G. Burns

zymes has been directed towards the activities of proliferating microorganisms, e i t h e r in t h e w a t e r c o l u m n [29, 31] o r i n t h e u n d e r l y i n g s e d i m e n t . M o s t o f t h e s t u d i e s o f s e d i m e n t s h a v e b e e n c o n c e r n e d w i t h d i s s i m i l a t o r y C, N , a n d S c y c l i n g of CO2, NO;, and SO~ by anaerobic bacteria during the oxidation of organic m a t t e r [ 15, 37], r a t h e r t h a n w i t h t h e c o n t r i b u t i o n o f h y d r o l y t i c e n z y m e s t o t h e release s o l u b l e C, N , and S during the decomposition of sediment organic matter. I n t h i s p a p e r w e r e p o r t t h e c h a r a c t e r i z a t i o n o f t h r e e C/N c y c l e a m i d o h y d r o l a s e s , i.e., u r e a s e , L - a s p a r a g i n a s e , a n d L - g l u t a m i n a s e , i n t h e s e d i m e n t o f a shallow eutrophic freshwater lake and present evidence that these enzymes e x i s t in e x t r a c e l l u l a r f o r m s a n a l o g o u s t o t h e s t a b l e e n z y m e f r a c t i o n s o f soil.

Materials and Methods

Sediment Sediment samples were collected from Westbere Lake (WBL), East Kent (National Grid Reference 190605, Sheet 179), a shallow eutrophic lake. Samples were taken from a littoral station covered by 2 m water using a hand-held sounding corer. Individual cores were extracted by a ramrod-type piston and the 0-3 cm horizons pooled and returned to the laboratory within 30 rain of collectiortThe slurry was then immediately passed through a sieve (.

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Fig. 1. pH/Activity response of(a) ADS urease, (b) ADS L-asparaginase, (e) ADS L-glutaminase, (d) ADS-extract L-glutaminase. Buffers: ([3) 0.1 M phosphate/citrate; (O) Tris/maleate (a) 0.1 M, (c) 0.2 M; (Q) 0.1 M Tris/HCl; ~1) phosphate (b) 0.5 M, (d) 0.1 M; (~,) diethanolamaine (h, c) 0.2 M, (d) 0.1 M; (•) 0.2 M bicarbonate; (*) 0.2 M acetate.

Results and Discussion

pH/Activity Response A D S urease was active o v e r the p H range 5.3-8.5 with an o p t i m u m p H near to neutrality (Fig. la). T h e overall shape o f the c u r v e resembles the classic bellshaped response shown by m o s t soluble e n z y m e s with extremes o f acidity or alkalinity causing extensive inactivation. T h e p H o p t i m a for urease in different soil types varies between p H 6.5 and 9.0 [2]. T h e p H o p t i m u m for A D S urease (pH 7,0) agrees closely with that r e p o r t e d for river s e d i m e n t urease [9] but is s o m e w h a t lower than the value o f p H 8.0 recorded for urease in fresh water [lll. In contrast to A D S urease, the shape o f the p H - a c t i v i t y curve for A D S

Arnidohydrolasesin Freshwater Sediment

163

L-asparaginase (optimum pH 8.4, Fig. 1b) has an extended alkaline response, with over 70% of the activity that was recorded at the pH o p t i m u m being expressed at pH 10.6. This is in general agreement with data published for soil [22] and marine sediment [30] L-asparaginase. The extreme alkaline tolerance Shown by ADS L-asparaginase has also been noted for other L-asparaginases [ l l , 17]. ADS L-glutaminase (optimum pH 6.5-7.0, Fig. lc) showed an almost conStant level of activity from pH 5.5 to pH 8.0, with over 95% o f the m a x i m u m activity being present throughout this range (Fig. l c). This broad plateau of high activity could either indicate that the activity comprises several distinct .L-glutaminases with widely ranging individual pH optima or that it is located m a microenvironment in which it is buffered against the pH value of the bulk 9aqueous phase (e.g., at a charged colloidal interface [18, 19]). The pH o p t i m u m is close to the pH 6.8-7.2 o f soil L-glutaminase [12, 23]. There was a sharp rise in activity between pI-I 4.5 and pH 5.5 which was also shown by both marine sediment [2] and freshwater [11] L-glutaminases. The response of ADS L-glutaminase to highly alkaline conditions more closely resembled that of ADS L-asparaginase than that of ADS urease, with enzyme activity at pH 10.0 being over 30% o f that recorded at the pI-I o p t i m u m . Although the pI-I/activity response of ADS extract L-glutaminase (optimum PH 8.0-9.0) has the same general appearance as that of the ADS L-glutaminase (Fig. l c, d), it is evident that the former is more active in highly alkaline COnditions, especially over the range pH 9.5-10.5. Furthermore, the sharp rise m activity between pI-I 4.5 and 5.5 recorded for the ADS enzyme occurred between pH 5.8 and 7.0 for the ADS extract enzyme (Fig. ld). This shift in PH profile towards the alkaline region suggests that the extracted L-glutaminase represents a fraction of ADS L-glutaminase which was intimately associated With a more acidic microenvironment (e.g., on humic colloids), than that of the bulk aqueous phase. Such responses have been artificially induced by enZYme immobilization on anionic surfaces [18, 19].

Enzyme Kinetics ADS urease did not obey Michaelis-Menten kinetics at low urea concentrations. This was evident for urea concentrations below 5 m M when Eadie-Hofstee plots were convex to the origin. However, enzyme kinetics within the range 550 rnM urea were typical, and Eadie-Hofstee plots gave an apparent Km of 1.2 rnM (Table 1). In contrast, soil urease has generally been found to obey Michaelis-Menten kinetics [26, 33], and therefore ADS urease more closely resembled the kinetic response of river sediment urease [9]. The anomalous kinetic response of this enzyme below 5 m M urea could be due to one or more of the following: (1) ADS enzyme is composed of two or more kinetically distinct fractions (e.g., both low and high affinity ureases), which could occur either if the urease was produced by different microbial species or if the identical enzyme became located in different microenvironments within the sediment colloidal matrix; (2) urea was adsorbed at surfaces giving locally elevated concentrations in the vicinity of the enzyme at low bulk-phase concentrations ofsubstrate; (3)

164 Table 1.

P+ J. Sallis and R. G. Burns Kinetic constants of ADS amidohydrolases

Enzyme ADS urease ADS L-asparaginase ADS L-glutaminase ADS-extract urease ADS-extract L-asparaginase ADS-extract L-glutaminase

Km (mM) 1.2 0.8 1.25 15.5 5.5 15.4

Correlation V ..... coefficient (#mol g-~ (r) h ~) 0.957 0.970 0.983 0.970 0.972 0.965

1.9 4.1 17.4 0.047 0.089 3.1

the ability o f urea to act as a hydrogen-bond disrupting agent allowed it to progressively alter the structure of sediment organic colloids, resulting in the unmasking o f enzyme active sites as urea concentrations were increased. However, at higher substrate concentrations, the Km value for ADS urease o f 1.2 mM (Table l) agrees with those reported for soil [33] and freshwater [10] ureases, which were 1.8-3.3 mM and 1.1 mM, respectively. In contrast to urease, ADS L-asparaginase was found to obey MichaelisMenten kinetics over the substrate range 0.8-4.0 mM. The Eadie-Hofstee plot was linear and indicated an apparent Km value of 0.8 m M L-asparagine (Table 1). This value is considerably higher than the Km values o f many bacterial L-asparaginases [34] which show a particularly high affinity for their substrate (Kin 0.01 mM). However, a similar disparity between Michaelis constants of a sediment L-asparaginase and those of bacterial L-asparaginases from which it was assumed to have been derived has been recorded for marine sediment L-asparaginase [30], suggesting that the L-asparaginases in these sediments are located at diffusionally restricted sites within the sediment matrix. ADS L-glutaminase also obeyed Michaelis-Menten kinetics over the range 1-10 mM L-glutamine, giving an apparent K~ of 1.25 m M (Table 1). This value contrasts with the figure o f 0.1 mM recorded for marine sediment L-glutaminase [8]. As for L-asparaginase, Km values differ by orders of magnitude from those o f bacteria which have Km values as low as 0.01-0.001 mM L-glutamine [34]_. Although Eadie-Hofstee plots of ADS L-glutaminase show the enzyme to be almost kinetically homogeneous over the observed substrate range, there was evidence from assays at L-glutamine concentrations above 60 mM that ADS L-glutaminase also consisted of a heterogeneous population of one or more kinetically distinct enzymes (the fraction having a Km value of 1.25 mM being by far the largest component). Thus, elevated L-glutamine levels (50 x Kin) were found to produce rates of hydrolysis some 15% higher than expected from the kinetically determined Vm,x value of 17.4 #tool g-~ h -~ (Table 1). The possible origins of such a low substrate-affinity fraction have been described above. The substrate dependence of urease, L-asparaginase, and L-glutaminase activity in ADS extract showed no major deviations from Michaelis-Menten kinetics over the substrate ranges 5-200 mM urea, 2-100 mM L-asparagine, and 10-100 mM L-glutamine. The Km values for each o f the extracted enzymes

Amidohydrolasesin FreshwaterSediment

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was approximately tenfold higher than that of the corresponding ADS enzyme (Table 1). The extraction of urease from soil also brought about a concomitant rise in K~n [24], with some authors attributing the elevated constant to the selection of an "adsorbed" urease from the various types of urease fractions in the parent soil [24]. These findings would support the view that in ADS, arnidohydrolases exist to some extent as adsorbed colloidal enzyme fractions. An anomalous kinetic response was, however, observed for ADS extract urease which showed a continuing increase in enzyme activity with increasing urea COncentrations above 200 mM urea. Consequently, the observed activity sometimes exceeded the kinetically determined value of Vma~ shown in Table 1 in routine assays containing high concentrations of urea (1.5 M) as substrate. This effect was thought to result from the hydrogen bond disruption by urea as described above.

Temperature Response In general, all three ADS enzymes showed an exponential increase in activity with increasing assay temperature before the onset of thermal inactivation (Fig. 2a, c, e). However, above the inactivation temperature, differences between the enzymes were apparent. Urease had the highest temperature optimum at 45~ and also continued to show high activity above the temperature optimum With over 90% of the maximum activity being expressed at 60~ In contrast, both L-asparaginase (optimum 38~ and L-glutaminase (optimum 40~ rapidly declined in activity at temperatures above their optima (Fig. 2c, e). However, as their substrates are unstable above this temperature, it would be surprising to find an enzyme that had evolved to catalyse the reaction above 40~ Arrhenius plots for each ADS enzyme were linear for data measured below the temperature optima (Fig. 2b, d, f). ADS urease had an activation energy (Ea) of 38.9 kJ mol-'. This is very close to the Ea for both soil [7, 13, 25] and freshwater [10] ureases. However, Ea values for ADS L-asparaginase (41.6 kJ rnol-~) and ADS L-glutaminase (35.9 kJ mol -~) compare less favorably with the equivalent values for the enzymes in fresh water (35.2 and 22.1 kJ mol -~, respectively [ 10]).

Resistance to Proteolysis All three enzymes in ADS were highly susceptible to inactivation by pronase (Fig. 3a, b, c) with less than 20% of the original activity surviving the 20 hours Preincubation. In the case of L-asparaginase and L-glutaminase, preincubation With buffer or BSA did not produce the same reduction in activity as preinCUbation with pronase, and much of the original activity remained even after the 20 hours preincubation (Fig. 3, b, c). In contrast, the urease activity of ADS declined by over 80% in response to all three preincubation conditions, and the rewetting process in itself was enough to induce a considerable inactivation of the enzyme (Fig. 3a). A similar but less complete loss of soil urease activity, which followed rewetting, was considered to result from desorption

166

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of urease from adsorbed (and therefore previously protected) sites [36]. Alternatively, the preincubation of ADS may have reduced the redox potential of the sediment sufficiently to solubilize inhibitory metal ions, a process which is thought to account for the inhibition of soil urease following the waterlogging of soil [271. There was a less marked effect of preincubation on the activity of FWS enzymes. FWS urease activity declined by approximately 20% for each preixacubation condition over the 20 hour period. FWS L-asparaginase and FWS L-glutaminase showed similar resistances to pronase, with more than 90% of

Arnidohydrolases in Freshwater Sediment

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their original activities surviving the 20 hours preincubation. The relatively high susceptibility o f ADS enzymes to proteolytic inactivation could have resulted from either an increased permeability o f microbial cells containing the enzymes, or from a significant change in the structure o f the colloidal support in air drying. The response o f ADS extract and c o m m e r c i a l bacterial L-glutaminase to Preincubations with buffer or pronase solutions is shown in Figure 4. ADS extract L-glutaminase was stable for 6 hours in buffered solution but was rapidly Inactivated by pronase. However, at least 20% o f the activity remained after the 6 hours preincubation. Bacterial L-glutaminase (Sigma, type II) was stable

168

P. J, Sallis and R. G. Burns

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tO preincubation with buffer, but was totally inactivated by pronase after approximately 2 hours preincubation (Fig. 4). In order to confirm that pronase a m e n d m e n t s were o f c o m p a r a b l e efficacy in both ADS a n d FWS, the proteolytic activity o f a m e n d e d and control sediments were measured over the same 20 hour time course. The results indicated that although there was a decline in pronase activity in FWS it was at or above the level o f pronase in A D S t h r o u g h o u t the preincubation. Furthermore, the a m e n d e d level o f pronase greatly exceeded the native proteolytic activity o f both FWS and A D S at all times. The proteolytic resistance o f A D S extract L-glutaminase was found to be no higher than that o f A D S L-glutaminase. However, the kinetic and p H / a c t i v i t y data o f this e n z y m e fraction suggests that it was largely c o m p o s e d o f L-glumminase in close association with (immobilized by) sediment organic material. Thus, it would appear that unlike soil organic matter, which affords enzymes some protection against proteolytic attack [26], an apparently similar association does not impede the access o f soluble proteolytic enzymes to the sites o f complexed enzymes in ADS.

Acknowledgment. This work was supported by the Natural Environment Research Council. References 1. Billen G (1982) Modelling the processes of organic matter degradation and nutrients recycling in sedimentary systems. In: Nedwell DB, Brown CM (eds) Sediment microbiology.Acaderaic Press. London, pp 15-52 2. Bremner JM, Mulvaney RL (1978) Urease activity in soils. In: Burns RG (ed) Soil enzymesAcademic Press, London, pp 149-196 3. Bullock P, Loveland PJ (1974) Mineralogical analyses. In: Avery BW, Bascomb CL (eds) soil survey laboratory methods. Harpenden Press, Rothamsted, pp 57--69 4. Burns RG (1978) Enzyme activity in soil: Some theoretical and practical considerations. In: Burns RG (ed) Soil enzymes. Academic Press, London, pp 295-340

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5. Burns RG (1982) Enzyme activity in soil: Location and a possible role in microbial ecology. Soil Biol Biochem 14:423-427 6. Chandramohan D, Devandran K, Natarajan R (1974) Arylsulphatase activity in marine sediments. Mar Biol 27:89-92 7. Dalai RC (1975) Effect of toluene on the energy barriers of urease activity of soils. Soil Sci 120:256-260 8. Dharmaraj K, Selvakumar N, Chandramohan D, Natarajan R (1977) L-Glutaminase activity in marine sediments. Indian J Mar Sci 6:168-170 9. Duddridge JE, Wainwright M (1982) Enzyme activity and kinetics in substrate-amended river sediments. Water Res 16:329-334 10, Frankenberger WT, Johanson JB (1983) Amidohydrolase activities in natural waters. Pol Arch Flydrobiol 30:319-329 11. Frankenberger WT, Page AL (1983) The effects of acidity and alkalinity on the stability of amidohydrolases in freshwater. J Envir Qual 12:459-462 12. Galstyan ASh, Saakyan EG (1973) Determination of soil L-glutaminase activity. Dolk Akad Nauk SSSR 209:1201-1202 13. Gould WD, Cook FD, Webster GR (1973) Factors affecting urea hydrolysis in several Alberta soils. PI Soil 38:393-401 14. Jones JG (1979) Microbial nitrate reduction in aquatic sediments. J Gen Microbiol 115:27-35 15. Jones JG (1982) Activities of aerobic and anaerobic bacteria in lake sediments and their effect on the water column. In: Nedwell DB, Brown CM (eds) Sediment microbiology. Academic Press, London, pp 107-145 16. Kiss S, Dragan-Bularda M, Radulescu D ( 1975) Biological significance ofenzymes accumulated in soils. Adv Agron 27:25-87 17. Lea pJ, Fowden L, Miflin BJ (1978) Purification and properties of Asparaginase from Lupinus sp. Phytochem 17:217-222 18. Levin y, Pecht M, Goldstein L, Katchalski E (1964) A water-insoluble polyanionic derivative of trypsin. 1. Preparation and properties. Biochem 3:1905-1913 19. McLaren AD, Babcock KL (1959) Some characteristics of enzyme reactions at surfaces. In: Hayashi T (ed) Subcellular particles. Ronald Press, New York, pp 23-35 20. Meyer-Reil LA (1981) Enzymatic decomposition of proteins and carbohydrates in marine sediments: Methodology and field observations during sampling. Kieler Meeresforsch Sonderh 5:311-317 21. Mortimer CH (1941) The exchange of dissolved substances between mud and water in lakes (parts I and II). J Ecol 29:280--329 22. Mouraret M (1965) L-Asparaginase activity in soil. Memoires, Orstom, Paris 23. Omura H, Sato F, Hayano K (1983) A method for estimating L-glutaminase activity in soils. Soil Sci P1 Nutr (Tokyo) 29:295-303 24. Paulson KN, Kurtz LT (1970) Michaelis constant of soil urease. Soil Sci Soc Am J 34:70-72 25. Perucci p, Guisquiani PL, Scarponi L (1982) Nitrogen losses from added urea and urease activity of a clay-loam soil amended with crop residues. PI Soil 69:457--463 26. Pettit NM, Smith ARJ, Freedman RB, Burns RG (1976) Soil urease: Activity, stability and kinetic properties. Soil Biol Biochem 8:479--484 27. Pulford ID, Tabatabai MA (1988) Effect of waterlogging on enzyme activities in soils. Soil Biol Biochem 20:215-219 28. Radulescu D, Kiss S (1971) Enzyme activities in the sediments of the Zanoaga and Zanoguta lakes (Retezat mountain mass). In: Progress in Roumanian palinology. Ed Acad Rep Soc Rom, Bucharest, pp 243-248 29. Satoh y (1981) Decomposition of urea by a size-fractionated planktonic community in a eutrophic reservoir in Japan. Hydrobiologia 83:153-160 30. Selvakumar N, Chandramohan D, Natarajan R (1977) L-Asparaginase activity in marine sediments. Curr Sci 46:287-291 31. Sepers BJ (1981) Diversity of ammonifying bacteria. Hydrobiologia 83:343-350 32. Skiba U, Wainwright M (1983) Assay and properties of some sulphur enzymes in coastal Sands. PI Soil 70:125-132

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33. Tabatabai MA (1973) Michaelis constant ofurease in soils and soil fractions. Soil Sci Soc Am Proc 37:707-710 34. Wade HE (1980) Synthesis and functions of microbial asparaginases and glutaminases. In: Payne JW (ed) Microorganisms and nitrogen sources: Transport and utilization of amino acids, peptides, proteins and related substances. Wiley Interscience, Chichester, pp 563-575 35. Zantua MI, Bremner JM (1975) Comparison of methods of assaying urease activity in soils. Soil Biol Biochem 7:291-295 36. Zantua MI, Bremner JM (1977) Stability of urease in soils. Soil Biol Biochem 9:135-140 37. Zeikus JG (1983) Metabolic communication between biodegradative populations in nature. In: Slater JH, Wittenbury R, Wimpenny JW (eds) Microbes in their natural environments, Cambridge University Press, Cambridge, pp 423--462