Laboratory 6



Laboratory 6 Enumeration

Introduction

One of the central questions a researcher asks when studying any bacterial system is “how many bacteria are present in this sample?” This applies to environmental samples where we want to learn about a native population as well as to lab cultures where we are measuring physiological functions. There are many ways to count bacteria and each of them has advantages and disadvantages.

Measuring the optical density (O.D.) of a sample is an indirect method of determining the number of cells present. The amount of light of a specific wavelength that is absorbed by a culture is related to the number of cells. This is a very fast and easy way to count cells when possible. Unfortunately, in order to use this method, you first need to calculate a response factor (the ratio of the number of cells to the change in absorbance) for the specific cells and media that you are using. This requires use of an independent method for counting the cells.

Another common method for counting cells is to serially dilute the original sample and spread-plate the resulting dilutions. This is known as the plate-count method. The colonies that arise after incubation are counted to determine the original cell density. This method requires more time and supplies than measuring the optical density but it does result in an absolute number of cells rather than a relative value. The one important thing to remember when doing plate-counts is that it only counts the viable cells (those cells that are able to grow under the specific incubation conditions used).

If you want to count all of the cells present in a sample then a direct-counting method should be chosen. This generally involves using a microscope to visualize the cells. Special microscope slides that have surfaces calibrated into precise volumes can be used to determine exactly how many cells there are in a milliliter of sample. Sometimes stains are used to make the cells more visible. This method counts all cells whether they are alive or not. The decision as to whether viable or total cell counts are more relevant will depend on the experimental question being asked.

In this lab you will be counting the number of bacteria in an overnight culture of E. coli using three different methods: optical density, direct counts and plate counts. You will then be able to calculate a response factor for optical density and compare the direct count and viable count methods.

Materials

Equipment

- spectrophotometer

- microscopes

- Petroff-Hauser chamber

- bunsen burners

- spreaders

Cultures

- Escherichia coli (broth)

Supplies

- Tryptic Soy Agar (TSA) plates

- Liquid TSB media

- Pasteur pipettes and bulbs

- Crystal violet stain

- 95% ethanol

Procedures

Plate-Counts (Counting by Dilution Series)

Period 1

1. Make a dilution series of the E. coli culture in liquid TSB media.

a. Aseptically transfer 1 ml of the culture into a tube containing 9 ml of TSB medium.

b. Transfer 1 ml of the resulting tube into another tube with 9 ml of medium.

c. Repeat this process until you have produced 9 tubes. (10-1 – 10-9).

2. Spread 0.1 ml of each dilution tube onto a TSA plate.

3. Incubate at 25(C for two days.

Period 2

4. Count the number of colonies on plates that contain between 20 and 200 well-separated colonies.

5. Calculate the number of cells in the starting culture by multiplying the number of colonies times the inverse of the dilution.

Optical Density (Counting by Spectrophotometer)

1. Turn on the spectrophotometer and set it to 525 nm.

2. Make a dilution series of the original E. coli culture in liquid TSB media.

a. Transfer 2 ml of the culture into a tube containing 2 ml of TSB medium.

b. Transfer 2 ml of the resulting tube into another tube with 2 ml of medium.

c. Repeat this process until you have produced 4 new tubes. (1:2 – 1:16).

3. Set the transmittance to 0 with no sample in the chamber.

4. Set the absorbance to 0 using un-inoculated TSB medium as a blank.

5. Measure the absorbance of each of the dilution tubes used in the dilution series.

6. Plot the absorbance against the dilution factor.

Direct Counting (Petroff-Hauser Chamber)

1. Use a pipette to transfer 0.9 ml of the starting culture to a clean test-tube.

2. Add 0.1 ml of crystal violet stain to the culture.

3. Use a Pasteur pipette to transfer a drop of the stained culture into the Petroff-Hauser slide.

4. Carefully place the cover slip onto the calibrated surface of the counting chamber.

5. Place the chamber on the microscope stage and focus under the 20X objective to locate the large counting grids.

6. Switch to high power and count the number of bacteria in at least 20 of the small squares.

(If there are too many bacteria to count, make a 1:10 dilution of the culture and start over)

7. Average the number of cells and calculate the number per ml. (cubic centimeter).

Dilution Series

Lab #6 20 points

Name: Date:

Plate counts

Count the number of colonies on plates that contain between 20 and 200 well-separated colonies.

colonies dilution

Calculate the number of cells in the starting culture by multiplying the number of colonies times the inverse of the dilution.

Concentration of cells in starting culture = cells/ml

Optical density

dilution tube absorbance

culture

1:2

1:4

1:8

1:16

Plot the absorbance against the dilution factor. (Attach graph)

Direct counts

Average the number of cells/square and calculate the number per ml. (cubic centimeter). Each small square is 1/20th X 1/20th of one millimeter square and 1/50th of one millimeter deep.

Average number of cells per square -

Cells per milliliter -

Cells per milliliter - (corrected for dilution by dye)

Response factor

Use the absorbance graph, the plate-count data and the direct-count data to calculate two separate response factors (# of cells / absorbance unit) for absorbance at 620nm for E. coli.

Plate count response factor cells/absorbance unit

Direct count response factor cells/absorbance unit

Questions:

1). Which response factor is more accurate for correlating absorbance to cell numbers? Why?

2). Give an example of an experiment in which you would want to use a direct count and an example of an experiment in which you would want to use a live count for quantifying bacteria.

Laboratory 7 Coliform Testing

Introduction

The determination of coliform bacteria in water is one of the oldest and most commonly used techniques in environmental microbiology. Coliform bacteria are used as indicators of potential health threats. The presence of bacteria in the coliform group in any sample is considered to be evidence of fecal contamination and therefore of the possibility that pathogenic organisms are present. The coliform group is functionally defined according to the way they are detected as “all aerobic and facultative anaerobic, gram-negative, nonspore-forming, rod-shaped bacteria that ferment lactose with gas and acid formation within 48hr at 35(C”. This definition was devised to detect E. coli but it also includes several other genera in the family Enterobacteriaceae including Klebsiella, Enterobacter and Citrobacter. All of these organisms are coliforms and will give a positive result in these tests but not all of them are necessarily the result of fecal contamination. In order to help clarify the results, and understand the implications of a coliform test, it is helpful to identify the number of “fecal coliforms” or to identify the species of bacteria detected. Fecal coliforms are defined as those coliforms that are capable of growth at 44(C. The increased temperature is meant to select only those organisms adapted to growth in the human body. Species typing is carried out by conducting several biochemical tests.

In this lab you will use two different methods two measure the number of coliforms present in surface water samples. The first method (membrane-filtration) is a one-step process where coliform colonies are counted directly from filters that have been incubated on m-Endo media. This method is relatively quick and easy but it is more prone to errors than the alternative “multiple tube fermentation” technique. Multiple tube fermentation is carried out in three separate steps. The “presumptive” step involves testing for gas production from lactose fermentation in selective media. The number of coliforms is determined by using the MPN method during this step. This is followed by the “confirmed” step where positive samples from the presumptive test are transferred to a more selective media and re-tested for gas production. Finally, the test is “completed” by isolating single colonies with a positive reaction on m-Endo agar.

You will further characterize the isolates from the completed step by using a commercial kit to determine the species.

Materials

Equipment

- incubator (35(C)

- bunsen burners

- inoculating loops

- pipettes and bulbs

- sterile graduated cylinders

- 100 ml dilution bottles

- sterile filter holders and vacuum flasks

Cultures

- environmental water samples

- Escherichia coli

- Enterobacter aerogenes

- Pseudomonas sp.

Media

- Durham tubes with lauryl tryptose broth (LTB) (2X and 1X)

- Durham tubes with brilliant green bile (BGB) broth

- m-Endo plates

- TSA plates and TSB media in tubes

- Tryptone broth

Supplies and Reagents

- membrane filters

- Enterotube II tests

- sterile water

- Kovac’s reagent

- 1% aqueous tetramethyl-p-phenylenediamine HCl

Procedures (Membrane Filtration Technique)

Period 1

1. Assemble a sterile filtration apparatus with a membrane filter

2. Aseptically transfer 25 ml of the sample to 225 ml of sterile distilled water.

3. Mixed diluted sample well.

4. Filter 10 ml of diluted water.

5. Aseptically transfer the membrane filter onto the surface of an m-Endo plate.

6. Replace the membrane filter and filter 50 ml of diluted water.

7. Aseptically transfer the membrane filter onto the surface of an m-Endo plate.

8. Replace the membrane filter and filter 100 ml of diluted water.

9. Aseptically transfer the membrane filter onto the surface of an m-Endo plate.

10. Incubate the plates upright at 35(C for 24 hrs.

Period 2

11. Count the number of dark red colonies with a metallic sheen.

12. Calculate the number of coliforms per 100 mL of the water sample.

Procedures (Multiple Fermentation Tube Technique)

Period 1

Most Probable Number (MPN) Determination (Presumptive Test)

1. Inoculate 5 tubes of 2X lauryl tryptose broth with 10 ml of your water sample and 5 tubes of 1X broth with 1 ml.

2. Dilute your water sample 1:10 by adding 10 ml to 90 ml of sterile water.

3. Inoculate 5 tubes of 1X broth with 1 ml of the diluted sample and 5 tubes of 1X broth with 0.1 ml.

4. Incubate the tubes at 35(C for two days.

Period 2

5. Examine each tube for gas production.

6. Record the results for each tube and use the MPN Table to calculate the number of coliforms per 100 mL of the water samples.

Confirmed Test

1. Inoculate BGB tubes with a loop full of culture from each of the most dilute positive tubes from the presumptive test.

2. Incubate the tubes at 35(C for two days.

Period 3

3. Examine each tube for gas production.

Completed Test

1. Streak two m-Endo plates from each positive BGB tube.

2. Incubate one plate at 35(C and the other at 44(C for 24 hrs.

Period 4

3. Examine the plates for the presence of green, metallic colonies.

Species Typing

1. Select a positive E. coli colony from the m-Endo plates.

2. Streak a TSA plate with that colony from the m-Endo plate.

3. Incubate at 35(C for 24 hrs.

Procedures (Enterotube II kit)

Period 5

Oxidase Test

1. Pick a colony from the TSA plate using a wooden applicator stick and smear it onto a filter soaked in tetramethyl-p-phenylenediamine.

2. Development of a purple color indicates a positive reaction.

3. Record the result on the Data sheet.

Inoculating the Tube

1. Remove both caps from the Enterotube without touching the wire inside.

2. Use the tip of the inoculating wire to pick one large (2-3mm) colony from your TSA plate (there should be a visible amount of cells on the wire).

3. Inoculate the tube by first twisting the wire then withdrawing it through all 12 compartments while slowly turning it.

4. Reinsert the wire back through all 12 compartments until the notch in the wire is even with the opening of the tube.

5. Break off the handle end of the wire by bending it.

6. Use the handle of the wire to punch holes through the foil covering the air inlets of the last eight compartments (adonitol, lactose, arabinose, sorbitol, Voges-Proskauer, dulcitol/PA, urea and citrate).

7. Replace both caps.

8. Incubate at 37(C for 24 hrs.

Period 6 Reading the Results

1. First read the color reactions for all of the tests except indole and Voges-Proskauer and record the results on the ID form.

2. Use a syringe and needle to pierce the plastic film and add one or two drops of Kovac’s reagent to the H2S / indole compartment.

3. Development of a red color in the compartment indicates a positive result.

4. Record the result on the ID form

5. Calculate the biotype number and determine the isolate’s identity from the code book.

6. If the code book indicates that it is necessary, perform the Voges-Proskauer test for confirmation.

a. Add two drops of 20% KOH and three drops of alpha naphthol to the chamber.

b. Development of a red color within 20 min. indicates a positive reaction.

Lab #7 20 points

Name: Date:

Water sample name:

Membrane filter test

# of coliform colonies:

10 ml 50 ml 100 ml

# of coliform cells per 100 mL

MPN test

Presumptive test

# of positive tubes

10 ml 1.0 ml 0.1 ml 0.01 ml

Most probable number

Confirmed test

How many of the BGB tubes are positive? /

How does this change the estimate of how many coliforms were in the original sample?

Completed test

How many of the EMB plates showed typical coliform colonies at 35˚C?

How many of the EMB plates showed typical coliform colonies at 44˚C?

Species typing (Attach Enterotube ID form)

Indole test Species identity

Laboratory 8 Chlorination

Introduction

The final step in the treatment of sewage and other wastewaters often involves disinfecting the treated water before discharging it into a river or lake. This is done to kill any pathogenic organisms that may have survived through the initial stages in treatment. Chlorination is the most common method used for the disinfecting stage of wastewater treatment. Chlorine gas is bubbled through the water where it hydrolyses to form hypochlorous acid (HOCl) and hypochlorite (OCl-).

Hydrolysis CL2 ( HOCl + H+ + CL-

Ionization HOCl ( H+ + OCl-

Hypochlorous acid is a strong oxidizing agent that acts by reacting with cell components to cause irreparable damage. Hypoclorite is less effective as an oxidizer. Together these two compounds are referred to as the free residual chlorine. Hypoclorous acid can also be formed by adding sodium hypochlorite (NaOCl) to the wastewater. Sodium hypochlorite is the active ingredient in Clorox( bleach.

NaOCl + H2O ( HOCl + NaOH

In addition to killing cells, the free residual chlorine also oxidizes non-living organic matter and several inorganic components of wastewater such as ammonia, iron and sulfides. When hypochlorous acid reacts with ammonia, monochloramine and dichloramine are formed. These compounds are also oxidizers and contribute to the disinfection process. They, along with some chloro-organic compounds, are referred to as combined residual chlorine. When enough chlorine has been added to completely oxidize the combined residual chlorine then free residual chlorine will begin to accumulate. This process is known as breakpoint chlorination.

Although chlorination is performed in order to kill all of the remaining pathogens in treated wastewater it is not always completely effective. Some organisms will only be injured by the chlorine and some will escape treatment altogether. When this happens, there is potential for re-growth of the pathogens after they have been discharged into clean water. This is why it is so important to be sure that the breakpoint has been reached. In this lab we will be carrying out a simple chlorination series with different concentrations of sodium hypochlorite and measuring the free residual chlorine and the total chlorine with a colorimetric assay. Then we will evaluate the survival of coliforms in the treated wastewater.

Materials

Equipment

- spectrophotometer

- incubator

Cultures

- sewage samples

Supplies

- Clorox bleach

- Hach Test Kit

- m-Endo plates

- beakers and flasks

- 16 X 150mm test tubes with 9.9ml of phosphate buffer

- pipettes and bulbs

Procedures

Chlorination

1. Dilute a sample of Clorox bleach by adding 1ml of bleach to 499ml of distilled water.

2. Dispense 60ml portions of “sewage” into seven 125 ml Erlenmeyer flasks.

3. Add 0, 0.5, 1, 2, 3, 5 or 8 ml of diluted bleach (1:500) to each of the flasks.

4. Mix each sample well and allow it to sit for 20min.

5. Follow the directions to measure both TOTAL and FREE Chlorine for each flask.

6. Select three flasks: one that has no free or total chlorine; one that has no free chlorine but does have measurable total chlorine; and one with a measurable free chlorine concentration.

7. Test each of these flasks for coliforms as described below.

Chlorine, Free [Method 10069 (0 – 5.0 mg/L)]

1. Press [PRGM] key. The display will show “PRGM?”.

2. Press [8] [ENTER]. The display will show “mg/L, CL2” and the ZERO icon.

3. Fill two sample cells with 10 ml of the sample.

4. Add the contents of one “DPD Free Chlorine” packet to one sample cell. Cap the cell and swirl vigorously to dissolve the powder.

5. Within one minute of reagent addition, add deionized water to each cell in order to fill to the 25 ml mark. Cap and mix both cells.

6. Place the diluted sample without reagent into the cell holder and tightly cover with the instrument cap.

7. Press [ZERO]. The cursor will move to the right and the display will read “0.00 mg/L CL2”. Remove the blank.

8. Place the sample with reagent into the cell holder and tightly cover with the instrument cap.

9. Press [READ]. The cursor will move to the right and the result will be displayed. Record this value.

Chlorine, Total [Method 10070 (0 – 5.0 mg/L)]

1. Press [PRGM] key. The display will show “PRGM?”.

2. Press [8] [ENTER]. The display will show “mg/L, CL2” and the ZERO icon.

3. Fill two sample cells with 10 ml of the sample.

4. Add the contents of one “DPD Total Chlorine” packet to one sample cell. Cap the cell and swirl vigorously to dissolve the powder.

5. Press [TIMER][ENTER]. A two-minute reaction period will begin.

6. After the timer beeps, add deionized water to each cell in order to fill to the 25 ml mark. Cap and mix both cells.

7. Place the diluted sample without reagent into the cell holder and tightly cover with the instrument cap.

8. Press [ZERO]. The cursor will move to the right and the display will read “0.00 mg/L CL2”. Remove the blank.

9. Place the sample with reagent into the cell holder and tightly cover with the instrument cap.

10. Press [READ]. The cursor will move to the right and the result will be displayed. Record this value.

Counting Coliforms

Period 1

1. Make a dilution series of each sewage sample into buffered water.

a. Aseptically transfer 0.1 ml of the sample into a tube containing 9.9 ml of buffered water.

b. Transfer 0.1 ml of the resulting tube into another tube with 9.9 ml of water.

c. Repeat this process until you have produced 5 tubes. (10-2 – 10-10).

2. Spread 0.1 ml of each dilution tube onto an m-Endo plate.

3. Incubate the plates upside-down at 37(C for 24 hrs.

Period 2

4. Count the number of dark red colonies with a metallic sheen.

5. Calculate the number of coliforms per milliliter of the water sample.

Dilution Series

Lab #8 10 points

Name: Date:

Chlorination:

|Chlorine |0 |0.5 |1 |2 |3 |5 |8 |

| | | | | | | | |

|Total | | | | | | | |

| | | | | | | | |

|Free | | | | | | | |

Coliform counts:

10-3 10-5 10-7 10-9 10-11 cells/ml

Flask 1

(no chlorine)

Flask 2

(no free chlorine)

Flask 3

(free chlorine)

Question:

1). How would you expect the data to change if the contact time with chlorine was increased from 20 min. to 1 hour? Why?

Laboratory 9 Rossi-Cholodny Slides

Introduction

The distribution of microorganisms in soil is heterogeneous. Microbes need nutrients and water to survive and these resources are not evenly distributed in soil. The structure of soils is composed of particles of inorganic and organic matter and the pores in between these particles. The pore spaces may be filled with water or air. Bacteria are mostly found attached to particles growing in small micro-colonies wherever nutrients can be found. Filamentous organisms such as actinomycetes and fungi make up much of the biomass. When they grow around soil particles they help to cement them into aggregates. These aggregates have their own internal pore spaces. The filaments also stretch between aggregates.

Often when soils are examined microbiologically, the native structure is destroyed and it is impossible to determine the original distribution of the microbes in their natural habitat. One way to visualize how microbes are distributed in soil is to use the soil contact slide method developed by Rossi and Cholodny (1938). In this technique, glass slides are buried in soil and allowed to incubate. Bacteria and fungi attach to the glass as though it was a mineral particle and grow on the surface. When the slide is carefully uncovered and stained, the microbes remain in their original positions and their associations with particles and each other can be seen.

In this lab we will be staining and examining slides that have been buried in soils that have received the following amendments: 1.6g of yeast extract, 0.2g of inorganic fertilizer, 1.6g of yeast extract and 0.2g of inorganic fertilizer, or no addition. All of the slides were incubated for ~2 weeks.

Materials

Supplies

- slides and coverslips

- staining racks and trays

- Pasteur pipettes and bulbs

Reagents

- crystal violet

Equipment

- Phase contrast microscopes

Activities

- Soil slide staining

Procedures

Soil Slide Stain

1. Start with a microscope slide that has had ¾ of its length buried in a beaker of garden soil and incubated for over one week.

2. Remove the slide by tilting it backwards so that the top of the slide is not scraped off.

3. Shake off any large soil particles and clean the bottom of the slide with a moist paper towel.

4. Allow the slide to thoroughly dry.

5. Fix the cells by passing the slide through a bunsen burner flame.

6. Gently tap the slide to remove as much soil as possible.

7. Flood the smear with crystal violet stain and let it sit for 60 seconds.

8. Gently rinse the slide with water. Try to remove as much of the stain as possible without disturbing the cells.

9. Blot with a paper towel and allow the slide to dry.

10. View under the microscope.

11. Look for fungal filaments, actinomycetes and bacterial micro-colonies.

12. Record all observations.

Lab #9 10 points

Name: Date:

Observations:

[pic]

Laboratory 10 Soil Enzyme Activity Part I (Polymer Degradation)

Inroduction

Complex polymers such as starch, cellulose, proteins and nucleic acids make up an important nutrient source for bacteria living in the soil. These polymers are components of the cells of other organisms (plants, animals, fungi and bacteria) in the soil. They become available after those cells die. Because of their large size, most polymers are insoluble in water. This means that in their native state, they cannot be taken up into bacterial cells for metabolism. Some bacteria can only use the polymers after they have been broken down into soluble monomers such as glucose, amino acids or nucleotides. They depend on other organisms to degrade the polymer structure. Other bacteria are able to cleave the polymer bonds by hydrolysis (breaking by the addition of water). This process requires the bacteria to produce enzymes that travel outside of the cell to attack the insoluble polymers. Although these enzymes allow the organism to break down a plentiful nutrient source in the soil, the cells need to expend a lot of energy to make the enzymes. In addition, many of the solubilized monomers will diffuse away from the cell and be taken up by other bacteria.

In this lab we will be examining four pure cultures for their ability to hydrolyze starch casein and gelatin. We will also plate soil dilutions on these same substrates to identify native soil organisms that are effective polymer degraders.

Materials

Equipment and Supplies

- incubator (30(C)

- bunsen burners

- spreaders and inoculating loops

- 99ml bottles with sterile water

- 1ml pipettes and pumps

Cultures

- soil samples

- Serratia marcesens

- Escherichia coli

- Pseudomonas fluorescens

- Bacillus megaterium

Media and Reagents

- starch plates

- gelatin plates

- skim milk plates

- iodine solution

Procedures

Period 1

1. Divide one of each type of polymer plate into four sections.

2. Inoculate each plate with all four of the pure cultures (one in each section).

3. Set up a soil dilution series as shown in the diagram.

4. Use the dilutions to spread plate onto all three types of polymer plate (12 plates in all).

Period 2 (after 1 week)

Casein Hydrolysis

1. Examine each of the skim milk plates for colonies that show a zone of clearing around them.

Starch Hydrolysis

1. Flood each of the starch plates with an iodine solution.

2. Look for clearing zones around the colonies.

Gelatin Hydrolysis

1. Look for clearing zones around the colonies.

Laboratory 10 Soil Enzyme Activity Part II (Alkaline Phosphatase Assay)

Materials

Equipment

- incubator (37(C)

- pH strips

- clinical centrifuge

- 5 ml pipettes and pumps

- screw-top tubes (wide-mouth)

- 16 X 100 mm test tubes

- balance

- spectrophotometer (460nm)

- drying oven (100(C)

- aluminum weighing dishes

Samples

- one soil sample treated with organic fertilizer, inorganic fertilizer, both or untreated

Media and Reagents

- buffer (pH 10)

- 2 mM p-nitrophenol

- 0.5 M CaCl2

- PNPP test solution

(para-nitrophenol phosphate in buffer)

Preparations

Reagents

1. Make a 2mM solution of p-nitrophenol (100ml)

2. Make pH=10 buffer (100ml)

3. Make 0.5 M CaCl2 solution (100ml)

4. Set up a standard curve of p-nitrophenol in 16 X 100mm test tubes diluted in 0.5M CaCl2 (2.0, 1.0, 0.5, 0.25, 0.125 and 0.0625mM; 3ml per tube)

Laboratory 10 Soil Enzyme Activity Part II (Alkaline Phosphatase Assay)

Introduction

Bacteria and fungi that break down insoluble nutrient sources in the soil produce extracellular enzymes. These are proteins that are produced inside the cell and exported out into the soil solution. The enzymes are active outside the cell where they catalyze reactions to break down the structure of the nutrient source to make it more accessible. The amount of an extracellular enzyme in the soil depends on the metabolic abilities of the soil organisms, the number of organisms present, the presence of substrate and the environment of the soil (pH, temp., ionic strength etc.). Because enzymes are costly for the cells to make, they are tightly regulated. Enzymes will only be made when they are needed.

One example of a common extracellular enzyme in soil is alkaline phosphatase. This enzyme is produced by many organisms in the soil. Its purpose is to remove the phosphate molecule from organic compounds such as phospholipids and nucleic acids. Once the phosphate is cleaved it becomes soluble and can be taken up by the cell. This is a very important activity because phosphate is often the limiting nutrient for microbial growth in soil.

In this lab you will be measuring the amount of active enzyme in soil samples by using a chromogenic substrate assay. In the presence of alkaline phosphatase, the colorless chemical para-nitrophenol phosphate is converted to para-nitrophenol, which is bright yellow. The amount of product formed can be measured using a spectrophotometer and the amount of enzyme activity can be calculated. You will also calculate the dry weight of the soil in order to standardize the results. The soils that you will be analyzing have been kept moist and incubated for ~2 weeks with the following amendments: 1.6g of yeast extract, 0.2g of inorganic fertilizer, 1.6g of yeast extract and 0.2g of inorganic fertilizer, or no addition.

Materials

Equipment

- incubator (37(C)

- pH strips

- clinical centrifuge

- 5 ml pipettes and pumps

- screw-top tubes (wide-mouth)

- 16 X 100 mm test tubes

- balance

- spectrophotometer (460nm)

- drying oven (100(C)

- aluminum weighing dishes

Samples

- one soil sample treated with organic fertilizer, inorganic fertilizer, both or untreated

Media and Reagents

- buffer (pH 10)

- 2 mM p-nitrophenol

- 0.5 M CaCl2

- PNPP test solution

(para-nitrophenol phosphate in buffer)

Procedures

Phosphatase Assay

1. Weigh out two 2-gram portions of each soil sample and pour them into labeled screw-cap tubes.

2. Pipette 5ml of 0.5 M CaCl2 solution into each tube and shake well.

3. Pipette 1ml of PNPP solution into one tube from each soil sample.

4. Pipette 1ml of phosphate buffer into the other tube to serve as a control.

5. Incubate all tubes at 37(C for 1 hour.

6. Centrifuge the screw-cap tubes in the clinical centrifuge for 5 min.

7. Transfer 4ml of the supernatant into labeled 16 X 100mm test tubes.

8. Centrifuge the test tubes in the clinical centrifuge for 5 min.

9. Transfer 3ml of the supernatant into clean test tubes (filter if needed).

10. Set the wavelength on the spectrophotometer to 460nm.

11. Set the absorbance to zero with a blank tube containing 3 ml of CaCl2.

12. Read and record the absorbance for each of your samples.

13. Use pH strips to check the pH of each sample.

14. Read and record the absorbance of the prepared standards.

15. Plot the absorbance vs. concentration to make a standard curve.

Water Content Analysis

1. Weigh four aluminum dishes (one for each soil) and record the weights.

2. Weigh out ~10g of each soil sample in an aluminum dish. Record the exact weight.

3. Put the samples in a 100(C oven overnight and let them cool in a desicator.

4. Weigh each of the dried samples and record the weight.

Lab #10 25 points

Name: Date:

Polymer Degradation

| |Starch |Casein |Gelatin |

| |Hydrolysis |Hydrolysis |Hydrolysis |

|Ps. fluorescens | | | |

|S. marcesens | | | |

|B. megaterium | | | |

|E. coli | | | |

Soil Dilution Observations:

How many colonies? What percent have clearing zones? How much diversity?

Casein Plates

Gelatin plates

Starch plates

Alkaline Phosphatase Assay

Spectrophotometer Readings

| | |Net Absorbance |Concentration of |

| |Absorbance |(absorbance – control) |p-Nitrophenol |

|Organic fertilizer | | | |

|control (organic) | | | |

| | | | |

|Inorganic fertilizer | | | |

|control (inorganic) | | | |

| | | | |

|Combined ferilizer | | | |

|Control (combined) | | | |

| | | | |

|Unamended | | | |

|control (unamended) | | | |

Standard Curve

|Concentration |2.0 mM |1.0 mM |0.5 mM |0.25 mM |0.125 mM |0.063 mM |

|absorbance | | | | | | |

Water Content Analysis

|Sample |organic |inorganic |combined |unamended |

|Dish weight | | | | |

|Wet weight with dish | | | | |

|Wet weight – dish weight | | | | |

|Dry weight with dish | | | | |

|Dry weight – dish weight | | | | |

|Water content | | | | |

Calculations

Water Content of Soil

Water content = (wet weight – dry weight) / dry weight

Dry weight = (wet weight / water content + 1)

Enzyme Activity

One unit of enzyme activity (U) is defined as the amount of enzyme that is able to convert 1 (mole of substrate to product in one minute. For soil assays, activity is reported as U per gram of dry soil

1. Calculate the amount of p-nitrophenol that was produced using the standard curve (remember that the total volume of liquid was 6ml but you only measured 3ml).

_______ µmoles in 6 mls

2. Divide the amount of product by the number of minutes that the samples were incubated to find the value of U

_______ µmoles / minute

3. Calculate the dry weight of the soil sample that was used in the incubation.

_______ grams

4. Calculate the activity per gram of dry soil.

_______ U / gram of dry soil

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