Short communication
Rapid assay for amidohydrolase (urease) activity in
environmental samples
R.L Sinsabaugh
a,*, H. Reynolds
b, T.M. Long
aaBiology Department, University of Toledo, Toledo, OH 43606, USA
bW. K. Kellogg Biological Station, Michigan State University, Hickory Corners, MI 49060, USA
Accepted 9 May 2000
Abstract
The use of microplate technology for enzyme assays has made it economical to measure a wide range of activities in environmental samples. Urease is one of the most widely measured soil enzyme activities, but current methods are cumbersome. We have developed a rapid, safe and sensitive assay that can be performed on microplates.q2000 Elsevier Science Ltd. All rights reserved.
Keywords: Amidohydrolase; Urease; Soil enzyme assay; Assay method; Microplate assay
Microbial ecology advances through technological improvements. For two decades, the commercial availabil-ity of chromogenic and ¯uorogenic substrates has facilitated research on enzyme activities in natural environments. Microplate technology has reduced the cost of enzyme assays, making it possible to assay up to 30 enzymes on a single plate (Foreman et al., 1998). Amidohydrolase (EC 3.5.1.5), commonly known as urease, is among the most commonly assayed soil enzymes. Interest in this enzyme stems from its role in the decomposition of urea fertilizer in agricultural systems (Bremner and Mulvaney, 1978; Xiaobin et al., 1995). More generally, it is considered an important agent for N mineralization in terrestrial and aqua-tic systems (Leftly and Syrett, 1973; Collier et al., 1999; Kandeler et al., 1999).
Urease decomposes urea into carbon dioxide and ammo-nium (Pettit et al., 1976). Activity is generally assayed by measuring the rate of NH4production (Tabatabai and
Brem-ner, 1972; Kandeler and Gerber, 1988). A number of assay procedures that variously require long incubations, large samples, specialized equipment, or hazardous chemicals have been published (Tabatabai and Bremner, 1972; Holland and Polacco, 1992; Klose and Tabatabai, 1999). We have developed a urease assay that is fast, sensitive, safe and can be performed on 96-well microplates.
A sample suspension is created by homogenizing 1.0 g soil (or 0.5 g litter) in 100 ml of 50 mM, pH 5.0 acetate
buffer, using a Brinkmann Polytron. More concentrated suspensions have more activity but they generate a higher absorbance background. We use pH 5 buffer to approximate the pH of bulk soil, but other buffers may be used to opti-mize sensitivity. Another option is to use puri®ed water and rely on the native buffering capacity of the soil. The sample suspension is poured into a wide mouth dish and placed on a magnetic stirrer. While stirring vigorously, 200ml aliquots are withdrawn and dispensed into the microplate wells using an 8-channel pipetter with wide ori®ce tips. We use three microplate columns (24 wells) per sample. Aliquots (10ml) of substrate solution (400 mM urea in deionized water), are dispensed into the wells of two columns (16 wells) yielding a ®nal urea concentration of,20 mM. While this concen-tration is well above reported Km values (Bremner and
Mulvaney, 1978), substrate saturation should be tested when applying this method to new soil types. The third column of wells acts as negative controls; each well receives 10ml of water. Each microplate also contains a substrate control: a column of wells that each contain 200ml of buffer 110ml of substrate solution.
Standards, prepared from ammonium chloride, can be added to the microplate. Standards ranging from 1±10mM (54±540mg NH4Cl l21, 18±80mg NH41l21) are good for
short incubations or samples with low activity. Wells in standard columns contain 200ml of standard and 10ml of water. The wells containing standards constitute the positive controls for the assay. Standards can also be prepared using sample suspensions which allows for the potential adsorp-tion of NH4 to particles. In this case, 10ml aliquots of Soil Biology & Biochemistry 32 (2000) 2095±2097
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standard, ranging from 20±200mM, are added to wells containing 200ml of sample suspension.
We incubate the microplates at 208C for up to 18 h. Higher temperatures will increase sensitivity. Ammonium concentration in the wells is quanti®ed using reagent pack-ets from Hach (Loveland, CO 80539, USA) which are marketed for water testing kits (salicylate reagent Ð cata-log no. 23952-66; cyanurate reagent Ð catacata-log no. 23954-66). The contents of the salicylate reagent packets are dissolved in deionized water (one packet per ml) and 40ml of the salicylate reagent is added to each well, includ-ing controls and standards. After a 3 min reaction period, the contents of the cyanurate reagent packets are dissolved in deionized water (one packet per ml) and 40ml of cyanurate reagent is dispensed into each well. Color development is complete in 20 min. The absorbance (ABS) of each well is then read at 610 nm using a spectrophotometric plate reader.
Urease activity is expressed as nmol NH41released g dry
soil21h21.
Net ABS(mean ABS of assay wells)2(mean ABS of sample control wells)2(mean ABS of substrate control wells).
Activity (nmol NH4g21h21) Net ABS £V=E£T£
M£A:
whereVis the volume of the sample suspension (100 ml),E the extinction coef®cient, DABS/nmol NH4, the slope of
ABS vs. [NH4] (for our machineE is about 25 pmol2 1
),T the incubation time, h,M the mass of soil or litter used to make the sample suspension, A the volume of the sample aliquot assayed (0.2 ml).
It is worth noting that analytical variation among repli-cate wells can be considerable. In our work, coef®cients of variation range from 5 to 20% depending on site and sample type. Well-to-well variation can be minimized by adequate homogenization, vigorous mixing of samples while dispen-sing, and good pipetting technique. By using 16 replicate wells, the standard error of the mean is,5%.
We have used this assay to measure urease activities at the sites of three long-term research experiments. The Duke Forest site, near Durham, NC, is a Pinus taeda (loblolly pine) plantation, planted in 1983, which is currently subject to a Free Air Carbon Enrichment (FACE) experiment (DeLucia et al., 1999). The soil at Duke has a high clay content and is classi®ed as an Ultic Al®sol, pH 5, with an organic matter content of 0.5% (Allen et al., 2000). The Oak Ridge, TN, site, also used for a FACE experiment, is a Liquidamber styraci¯ua(sweet gum) plantation, planted in 1989. Soil at the Oak Ridge site is classi®ed as a Typic Dystrocrept with a shaly silt loam A horizon. The third site near Pellston, MI, is a 90 y old Acer saccharum (sugar maple) stand with soil classi®ed as sandy, mixed, Frigid Typic and Al®c Haplorthods and an organic matter content of 3.5%. Since 1987, parts of it have been amended with 30 kg N ha21y21.
The Duke and Oak Ridge sites were sampled in May, July and September 1999. On each date, twelve 2£15 cm2cores were collected in each of six plots. At the Duke site, litter samples were also collected from several locations in each plot. Composite samples from each plot were assayed for urease activity on the day of collection. The Pellston site was sampled on a single date in September 1999. Composite soil samples from three control and three N-amended plots were assayed.
At the Duke Forest, activity varied from undetectable to 0.73mmol g21h21 mean0:32;n18in soil and from 0.11 to 5.37mmol g21h21 mean1:43;n18in litter. Activity was lower in Oak Ridge soil, ranging from unde-tectable to 0.71mmol g21h21 mean0:21; n18: At the Pellston site, activity in control plots averaged 1.63mmol g21h21 compared to 0.98mmol g21h21 in the N-treated plots.
These values fall in the range of published reports for forest systems. The availability of a microplate assay for urease facilitates long term monitoring and makes it easier to include urease in pro®les of extracellular enzyme activity.
Acknowledgements
We thank William Schlesinger and George Hendrey for access to the Duke Forest FACE site, Rich Norby and Gerry O'Neill for access to the Oak Ridge FACE site, and Don Zak for access to the Michigan gradient sites. The enzyme assays were done by Tammy Long and Kellyanne Driscoll.
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