Determination of Microbial Extracellular Enzyme Activity in Waters, Soils, and Sediments using High Throughput Microplate Assays

The overall goal of the following procedure is to rapidly assay microbial extracellular enzyme activity in a variety of natural environments. This is achieved by adding samples of water or soil slurries to microplate to yield adequate replication As a second step, artificial colormetric or fluoro metric substrate is added to the microplate wells, which allows for enzymatic reactions to occur. Next, either absorbence or fluorescence is measured in order to determine the amount of end product released during the reaction.

The results show differences in extracellular enzyme activity between samples based on the absorbence or fluorescence of the final reaction. Today we're gonna demonstrate how to measure extracellular enzyme activity in natural environments. This method can help answer key questions in microbial ecology, such as water rates of organic matter processing and nutrient mineralization in soils, sediments and waters Generally, individuals new to using this method will struggle because of difficulty in separating particles from the reaction solution, especially when assaying large numbers of samples.

A visual demonstration of this procedure is critical as a plate layout can be difficult to learn and taking measurements at multiple time steps can make this procedure more challenging. Chole metric analysis of extracellular enzyme activity for soils and sediments begins with preparation of substrate buffer and standard solutions as described in the text protocol for soil, prepare a slurry of each sample to be assayed in a sterile 15 milliliter centrifuge tube. Add a concentration of approximately one gram per milliliter using 50 millimolar acetate buffer for sediments.

Add enough acetate buffer to make the slurry easily pipee. The exact volume of slurry needed will vary according to the number of enzymes assayed, but a minimum of five milliliters is recommended. Vortex each slurry until all clumps of soil or sediment have dispersed.

And note the final volume. Clip the end of pipette tips with scissors prior to pipetting. As soil slurries tend to clog tips, re vortex each slurry and immediately pipette 150 microliters into each of six wells.

On a 96 well deep well block. It is important to vortex thoroughly and frequently. In order to keep soil particles in suspension, leave at least two wells per block empty to serve as a substrate control.

Prepare 1 96. Well block for each enzyme to be assay. Pour approximately five milliliters of acetate buffer into a pipette reservoir and use an eight channel pipetter to add 150 microliters of the buffer to the last two wells of each sample.

For the sample buffer controls and to the two substrate control wells. Pour approximately 10 milliliters of the appropriate p nitro fennel or PNP substrate solution into a pipette reservoir and use an eight channel pipetter to add 150 microliters of the substrate solution to the first four wells of each sample and to the two substrate control wells. Note the time as the reaction begins, as soon as the substrate solution is added.

Incubate plates at room temperature for 0.5 to four hours. Exact incubation time will vary depending on the activity level in samples and the enzyme to be assayed. While assays are incubating, prepare one clear 96 well microplate.

For each deep well block in order to read absorbance pipette, 190 microliters of distilled water and 10 microliters of one molar sodium hydroxide into each well of the microplate. Sodium hydroxide slows the enzymatic reaction and raises the pH, which enhances the color of the PNP released during the reaction after incubation. Centrifuge the 96 well blocks at 2000 to 5, 000 times G for five minutes to pellet soil particles.

Removal of the liquid without disturbing the pellet can be a challenge. One must avoid using too much soil sample initially. Be sure to centrifuge for long enough and use a careful but confident motion to withdraw the liquid.

Use a multi-channel pipette to withdraw 100 microliters from each. Well being careful to avoid the pellet. Transfer the liquid to the corresponding well on a prepared clear 96 well microplate turn on the Microplate reader and set up any necessary software record absorbance at 410 nanometers following determination of the dry mass of the samples as described in the text protocol, calculate final absorbance of each sample by subtracting the sample control absorbance from the sample assay absorbance.

If substrate controls have high absorbance of greater than 0.060, then subtract those. Also proceed to calculate enzyme activity in micromoles per hour per gram of dry mass using this equation. After organizing a black microplate for each enzyme following the example shown in the text protocol, pour approximately five milliliters of the first water sample into a pipette reservoir and use an eight channel pipetter to pipette 200 microliters into all of the wells.

In column one of the Microplate discard used pipette tips and repeat as needed for each water sample to fill columns one through nine. Set up controls to account for samples, standards, substrate and quenching on the same black microplate as the samples. Sample controls contain sample water and bicarbonate buffer and are not used in the activity calculations, but will demonstrate reading consistency throughout the course of the experiment.

The quench controls consist of sample water and a standard amount of the fluorescent tag and are used to measure the diffraction of fluorescence. In sample water substrate and standard controls are made up of either substrate linked or the standard fluorescent tag respectively and bicarbonate buffer. Pour approximately five milliliters of five millimolar bicarbonate buffer into a clean pipette reservoir.

Pipette 50 microliters of buffer into microplate wells one through nine in rows d and e to form two, replicate wells of sample controls per sample change pipette tips, then transfer 200 microliters of bicarbonate buffer to wells 10 through 12 in rows A and H.Although light is kept bright in this demonstration for filming purposes, ambient lighting should be reduced by dimming or turning off lights as the fluorescent standard is light sensitive. Pour approximately five milliliters of 10 micromolar, four methyl mone or MUB into a clean pipette reservoir, pipette 50 microliters into microplate wells one through 12 in row H.Then pipette 50 microliters into wells one through nine in rows G and F to form three replicates of quench controls per sample and overall standard controls. Either place the microplate in the dark or cover with an opaque lid to reduce light degradation of MUB, turn on the fluorimeter and set up any necessary software to be ready to read before adding the substrate.

For approximately five milliliters of the appropriate MUB lined substrate into a clean pipette reservoir, use a 12 channel pipetter to pipette 50 microliters into microplate wells one through 12 in row A and into wells one through nine in rows B and C to form three, replicate assays for each sample and three substrate controls immediately proceed to record fluorescence. Read the initial fluorescence immediately after substrate addition to the microplate after reading. Fluorescence incubate the microplate at room temperature either in the dark or covered with an opaque lid.

To reduce light degradation of MUB. The incubation time required to measure maximum potential enzyme activity will depend on the enzyme concentration within a sample. Since this is unknown, before the sample is assayed, a microplate will have to be read at multiple times steps.

For most enzymes, 10 to 15 minute intervals over the course of an hour is appropriate, although some samples with high concentrations will peak before 10 minutes. Continue reading fluorescence in the microplate at the designated intervals for at least an hour. Be sure to keep the microplate covered or in the dark between readings.

For each sample at each time interval, calculate the mean initial sample fluorescence, the mean final sample fluorescence, the mean standard fluorescence and the mean quench control fluorescence for each time interval. Calculate enzyme activity in m per hour per milliliter. Using the equation found in the text protocol.

Examine the activity values calculated for each time step, determine final potential activity from the time step with the highest activity. If activity values continue to increase, then later time steps may be required. If activity values fall throughout the course of the run, then run again with shorter time steps.

Soils and aquatic sediments typically have appreciable levels of extracellular en enzyme activity as a result of attached microbial communities growing on the surface of particles. This figure shows how this activity changes depending on the size of particles obtained from the surface sediment of a third order stream in northern Mississippi. USAA previous study has shown that the bacterial communities on sediment particles from this stream can be separated into three distinct groups based on molecular analysis of their community structure.

Those on 0.063 millimeter particles, those on 0.1 2 5, 0 0.25, and 0.5 millimeter particles and those on one millimeter particles. Analysis of patterns in extracellular enzyme activity supports this conclusion with phosphatase being similar on 0.1 2 5, 0 0.25, and 0.5 millimeter particles, but much higher on the one and 0.063 millimeter fractions when measured using the PNP linked substrate technique. Other enzymes such as beta glucco and n AGAs, show similar peaks on the finest particles, but are not elevated on one millimeter particles.

Highlighting the fact that different enzymes may show different environmental distributions and these can be elucidated using this assay. The relatively low error bars show that chole metric assays of enzyme activity on sediments are reproducible and thus amenable to statistical analysis. When comparing different and environmental samples, natural waters tend to have lower extracellular enzyme activity per milliliter than soils or sediments do per gram.

As such, they should be assayed flora metrically using MUB linked substrates. This figure shows how the activity of the enzymes, phosphatase, beta, glucco, and NGAs varies with depth. In a shallow lake in northern Mississippi, USA, the lake is known to be nutrient poor, which is also suggested by the relatively high activity of phosphatase and enzyme that microorganisms produce in order to acquire phosphate from organic compounds.

For all three of the enzyme's, assay samples collected at the water surface and at 50 centimeters depth showed similar activity. Whereas activity was elevated in the 100 centimeter sample, the sample was essentially taken from the water sediment interface and the presence of sediment particles in the sample likely accounts for the higher activity seen in the sample, especially for beta glucco and n AGAs. As with the PNP substrates, the low error bars show that even with just three replicate readings, the reproducibility of fluoro metric assays using MUB substrates is high.

Note that units for water sample enzyme activity are inal of substrate consumed, whereas those for enzyme activity on sediment particles are in micromoles of substrate consumed even though the per unit size is comparable. This highlights the fact that soils and sediments tend to have much higher extracellular activity than natural waters. It also demonstrates the increased sensitivity of the MUB linked substrate technique and the necessity of using this technique for assay extracellular enzyme activity in water samples Once mastered.

This technique can be used to assay nine water samples for up to enzymes in just over an hour While attempting this procedure for soils and sediments. It's important to use a known massive material so that activity can be expressed on a dry weight basis. After watching this video, you should have a good understanding of how to measure extracellular enzyme activity and a variety of natural environments using either color metric or lower metric techniques.