BISC209/S12: Culture Media

American Society of Microbiology (ASM) Image library of bacterial colony morphologies

Perhaps we can add to this library of images of colonies of culturable microorganisms. Consult the microbe library maintained by the American Society of Microbiology at: | http://www.microbelibrary.org/ASMOnly/details.asp?id=2566&Lang=&ISkip=20. The images found there may help you recognize the desired Genera from colony morphology.

Background

The composition of medium is an important factor when attempting to culture microorganisms. The components and pH can be manipulated to favor the nutritional preferences of particular bacterial groups if the desired microbes are unlikely to be successfully isolated in a general purpose medium. Keep in mind that the composition of selective or enrichment media (particularly the differences between them and general purpose media) provides valuable information about the metabolic range of the microbes that grow there.

The recipe for each selective or enrichment medium described here includes the composition and concentration of all ingredients (in g/l or % [wt/vol or vol/vol]) and the desired final pH. Deionized (filtered) water is generally used unless a particular medium requires a purer type of water (distilled, salt-free, DNAase or RNAase free for example). Tap water is not used in media preparation because it may contain undesirable compounds such as chlorine, copper, lead, and detergents. Some of the media or reagents described here can be purchased from companies such as BBL Microbiology Systems or Difco Laboratories in a dehydrated, premixed form. If commercially purchased in dehydrated form, the manufacturer provides the instructions for preparation.

Pourite™ is an anti-foaming agent from American Scientific Products that is commonly added to non-commercial medium containing agar. One drop of Pourite™ is added to volumes up to 800 ml and 2 drops to volumes of more than 800 ml to prevent foaming.

General Purpose Media
Nutrient broth (NB) and Nutrient agar (NA):

NB: 0.3% Beef extract, 0.5% Peptone, Deionized water to 1 L at pH 6.6- 7.0 at 25°C

NA: 0.3% Beef extract, 0.5% Peptone, 1.5% Agar; Deionized water to 1 L at pH 6.6- 7.0 at 25°C

NA+Starch: NA (above) plus added 2.5% starch

Dilute NA: NA diluted 1:10 (0.03% beef extract, 0.05% Peptone, 1.5% agar)

Nutrient broth/agar is a moderately rich, general purpose, solid medium that meets the nutritional requirements of many culturable bacteria. It contains beef extract, soy digest, and enzymatically digested gelatin to support the growth of a wide variety of chemoheterotrophic organisms. Fungal growth is reduced in this media. In broth form, the solidifying agent (agar) is not included. Sometimes it is diluted to help slower growing microorganisms not be overshadowed by quicker growing or spreading microorganisms.

Defined vs. Undefined Media
These terms refer to media in which all ingredients and the concentrations of those ingredients are known (defined) or media that contains components that vary in concentration, molecular formula, or may or may not be uniform in different batches or brands of medium (undefined). A wider variety of microorganisms will grow on undefined media than on defined so most general purpose media, such as nutrient agar or tryptocase soy agar (TSA), are undefined. Some common ingredients in undefined media include tryptone (trypsin digested casein, a milk protein) or peptone (an enzymatic digest of varying proteins, often meat scraps). These are not pure substances. They contain left over enzymes, vitamins, salts, amino acids, peptides, and polypeptides. They are important sources of nitrogen. When we are interested in enrichment for particularly bacteria, it is more common to use defined media because we can control the sources of nitrogen or carbon. Many of the selective media are defined because your goal is to grow only one type of microorganism based on some known metabolic characteristic it shares with few others.

Selective / Differential / Enrichment media

Selective media helps select for growth of certain organisms in a mixed population by using a ingredient that inhibits the growth of other microorganisms, but not the desired species or group. Enrichment media can be considered a subgroup of selective media since its composition is usually designed to enhance the growth of certain microorganisms by including nutrients that the desired microorganism or group can use for an essential process while its competitors can not. Sometimes enrichment media also limits alternate sources of nutrition or contains an ingredient that inhibits the growth of competitors. Differential media does not select for any particular group by inhibiting or enhancing the growth of one group over competitors, but this type of medium is able to show a visible difference between or among groups of microorganisms. Media can be any permutation or combination of selective, differential, and/or enrichment, depending on its ingredients and its use.
For more information culture media you can access an e-book of the Difco manual of culture media at: | http://www.archive.org/details/difcomanualofdeh09dige

Media for isolation and identification of soil bacteria in a mixed population (Starting with either SOIL or SOIL EXTRACT)

Finding Nitrogen Cycling Bacteria & Citrate Carbon Source Users: Bacteria that can Extract Nitrogen from Ammonium Compounds

Ammonium as only nitrogen source & Citrate as only Carbon Source Users: Using Simmons Citrate Medium:
0.02% MgSO₄(Magnesium Sulfate), 0.1% NH₄H₂PO₄(monoammonium phosphate), 0.1% K₂HPO₄(dipotassium phosphate), 0.2% C₆H₅Na₃O₇(sodium citrate), 0.5% NaCl(sodium chloride), 2.5% agar, 0.008% C₂₇H2₇Br₂O₅SNa(bromothymol blue) at pH6.9

Simmons Citrate Medium is a defined, selective, medium that selects for microorganisms that can utilize citrate as their sole source of carbon in a medium containing inorganic ammonium salts as its only source of nitrogen. In 1923 an investigator named Koser invented a broth medium in which ammonium phosphate supplied the only source of nitrogen and each organic acid was added individually, allowing the introduction of carbon utilization as a diagnostic aid. Later, an investigator named Simmons converted Koser’s liquid medium to a solid by the addition of agar and added an indicator system by incorporating bromthymol blue. The exact nature of the alkaline reaction produced by the organisms that are able to use citrate as their sole source of carbon is poorly understood. It appears that the alkaline reaction that gives the color change characteristic of a positive Simmons-Citrate test most likely occurs when excess CO₂ is generated when citrate is cleaved by the enzyme citrase to form oxaloacetate. This by product is then decarboxylated to pyruvic acid and CO₂. The excess CO₂ combines with sodium and water to form sodium carbonate. Note that bacteria that utilize citrate in this way are able to extract nitrogen from ammonium phosphate in the medium, resulting in the production of ammonia which combines with water to form NH₄OH. These reactions in combination produce an alkaline pH (greater than 7.6), resulting in a color change in the indicator from green to blue. Some members of the genera Klebsiella in the Entrobacteriacea family are Simmons Citrate positive.

Isolation of Ammonium & Citrate Users from Simmons Citrate Selective Medium:

• Make a 1% soil extract as explained in the protocol in Lab1.
• Dilute the 1% soil extract serially (two 1:10 dilutions) to 10^-3 and 10^-4.
• Put 100μL of the 10^-2, 10^-3, and 10^-4 dilution in the center of separate Simmons Citrate medium culture plates and spread the diluted soil extract uniformly over the plates.
• Incubate the cultures at RT for a week or until visible colonies form.
• Count the colonies on one of the plates with 30-300 colonies and using a Sharpie number a few of the different appearing ones on the bottom of the plate or give them code names (your initial and a number or a more creative coding scheme).
• Some of the colonies on this plate may fluoresce, in order to see this, you will need to take a marker, safety goggles, the hand-held UV light on the instructors desk, and your plate to a completely dark location. Put on the glasses, turn on the UV light and shut off the white lights. If you see any colonies that "light up", try to mark them with a small dot or circle on the bottom side of the plate.
• Make observations in your lab notebook about the #, appearance, color, consistency &/or flouresence of any colonies growing on your Simmons Citrate plates.
• Each teammate should try to isolate a different looking colony from any of your Simmons Citrate plates (based on colony color, consistency, and/or fluorescence)
• Obtain a new sterile Nutrient Agar plate or for the fluorescent colonies use a new citrate plate (one plate for each student in your group).
• Each student should use her flame sterilized and cooled loop and touch it to the center of the colony on Simmons Citrate that you are attempting to isolate. DO NOT touch anything on this plate other than the desired colony.
• Follow the directions found in Streaking for Isolation in the Protocols section of this wiki carefully. Transfer bacteria from this colony to the O zone of a new NA plate.
• Flame and cool your loop before going back into sequential inoculated zones to transfer fewer and fewer bacteria as you streak each zone.
• Parafilm and save any plate that contains a colony you or your teammates are attempting to isolate. Store this plate(s) in the refrigerator. Discard unused dilutions.
• Check the plates frequently so you will know how fast your organism grows, and that their appearance or fluorescence is maintained.
• Your goal is to end with well isolated single colonies that grow from a single bacterium (all genetically identical) on nutrient agar (NA). It will probably take several transfers over the next few weeks to take a CFU from this original selective medium plate and grow bacteria from it in pure culture on nutrient agar.

Finding Nitrogen Fixing Bacteria:
Use Azotobacter Medium

Important nitrogen cycling bacteria, are able to aerobically use N₂ as their source of Nitrogen without a symbiotic partner. One group of these bacteria, Azotobacter, can be differentiated from other nitrogen fixers because the Azotobacters can use mannitol as their sole carbon source. We will not, initially, try to select for only members of the Azotobacter group of nitrogen fixers in our initial enrichment/selection but we could take away other sources of carbon other than mannitol later, in a secondary selection with a more selective medium, if we are interested in separating them from other nitrogen fixers. The Azotobacters are generally Gram-negative, large rods, or ovoid cells.

Medium: 0.08% K₂HPO₄; 0.02% KH₂PO₄, 0.02% MgSO₄ 0.01% CaSO_4; 0.0015% FeSO4/7H₂O; 0.00025% g NaMoO₃; 0.5% sucrose (or substitute mannitol for the sucrose if you want to select for Azotobacters---we will use sucrose not mannitol to be less selective).

Primary Enrichment/Selection:

• Measure 0.5 g of soil into 25 ml of liquid Azotobacter medium. (This is already aliquoted for you in small flasks with cotton plugs or a loose cover.)
• Mix well
• Place the flask in your closed bench cabinet so the culture will incubate in the dark at RT.

NEXT LAB: (7 days later) examine the air-liquid interface in your flask and look for a slimy growth. The slimy growth may only be on the sides of the flask or it may extend across the liquid surface (a pellicle).

Selection/Isolation:

• Take a loopful of the slime and place it in 1 ml of sterile water in a small tube. Cap the small tube and place the capped tube into an empty 16 mm tube.
• Vortex the tube to disperse the sample. (Vortexing in this way helps break up the other microbes that will be embedded in the slimy material. The other microbes are taking advantage of the by-products of Nitrogen compounds excreted by the N₂ fixers. )
• Isolation streak a sample of the diluted, vortexed slime suspension following the protocol in Streaking for Isolation onto Azotobacteria agar medium.
• Incubate at room temp or at 30 °C.

Isolation to Pure Culture:

Watch for the appearance of isolated, slimy colonies.

Continue to isolation streak until you think you have pure isolates.

To test for Azobacters by morphology:
Contaminants should be detectable on nutrient agar by differences in colony morphology. Although the Azotobacters may not appear characteristically slimy on nutrient agar, make a sub-culture onto NA by streaking out a well-isolated colony. If more than one type of colony appears on NA, Gram stain each different looking colony then restreak a colony with the typical Azotobacter bacterial morphology onto Azotobacter medium. Maintain the other non-Azotobacter looking bacteria on NA. What should Azotobacter bacteria look like in a Gram stain?

Characterization:
Once you have isolates in pure culture, begin to examine the cellular morphology, structure, and metabolism of your isolates as described in LAB 5.

Finding Spore Forming Bacteria
(Such as members of genera Bacillus and of Streptomyces in the Actinomycetes family
Use Glycerol Yeast Extract agar)

Spore forming bacteria are highly resistant to environmental stresses and to disinfection procedures. Among these hardy bacteria are members of the genus Bacillus or the Actinomycetes group of bacteria. All of these bacteria, particularly Streptomyces, are important sources of antibiotics. Actinomycetes often show a uniquely recognizable filamentous and/or leathery colonial morphology [1] that will help you find them.
The genus Bacillus, a member of the Firmicutes is one genus in a large group of Gram positive organisms. Bacillus spp. are known for the ability of the vegetative cell to produce a metabolically inactive state (a spore). It is likely you will find several subgroups of Bacillus growing on general purpose media or other of the media we use. Use web images to become familiar with the colony morphology of Bacillus so that you will recognize likely candidates and choose them for isolation.

We will use oven dried soil on selective media to enrich and select for spore forming bacteria that can survive oven drying and the high osmotic pressure in our selective medium. You should try to isolate and characterize several different appearing colonies from this medium. Do some research on the web to find images of macroscopic colonies and microscopic bacteria from these groups so you will recognize them if you find them.

Glycerol Yeast Extract Agar (GYEA): a selective medium to enrich for many of the spore forming, antibiotic producing bacteria in the Actinomycetes, Bacillus, and other groups of Gram positive spore formers.
Medium: 0.5% Glycerol ;, 0.2% yeast extract, 0.1% dipotassium phosphate, 1.5% agar.

Your instructors will heat dessicate (oven bake) your 3 one gram soil samples collected and weighed in LAB1 and return them to you in LAB2. You will use this heat shocked, dry soil sample to make a new soil extract for this protocol, which is based on spore resistance to dessication. Drying and heating the sample has encouraged spore generating bacteria to form a state that will allow them to survive harsh environmental conditions while killing off many of the microbes that can't make spores--- allowing survival in extreme heat and dehydration. Now we need to coax those spores back into their vegetative state.

Enrichment/Selection/Differentiation:

Your lab instructor will return (in LAB 2) the dried 1 g soil samples, that you weighed out in LAB 1.

• Record the dried weight from the 3 samples in your lab notebooks.
• Combine the 3 dried soil samples and then re-weigh to get 1 g of dried soil. Add it to 100 ml of dilute nutrient broth in a flask containing a magnetic stirrer.
• Agitate on a stir plate for 30 minutes.
• Allow the large particles to settle to the bottom of the flask for 30 minutes.
• Soak a sterile cotton swab in the broth.
• Swab section 1 of a labeled plate of glycerol yeast medium (GYEA) using your best isolation streak technique
• Allow the inoculum in section 1 to absorb into the agar before you,
• Follow the steps for Streaking for Isolation .
• Invert, and incubate the plate at RT.
• Check your plate for colonies daily.

Isolation to Pure Culture:

When well-isolated candidate colonies of the appropriate morphology appear, use the tip of a sterile toothpick to pick up a small but visible amount of growth, being careful not to touch anything but the tip of the colony. Isolation streak any interesting colonies (preferentially chose those that appear like "little volcanos" or "powdered sugar") onto new glycerol yeast plates (one colony/plate) using your flame sterilized inoculating loop after you have applied the growth from the toothpick to zone one of your streak plate. Note thatActinomycetes and Streptomycetes are often tough leathery colonies, so transfer of these colonies is sometimes difficult. The powdery area may indicate spore formation: take a sample from this area, if possible. In any case, try to "break off" a piece of the colony with your sterile loop or with a sterile toothpick and transfer that piece of a colony to zone one of the new medium and then use your loop for streaking out the other zones. The tiny spores on the surface of the colony are likely to transfer to the next plate or tube when you work with it. (That's a good thing this time.)

Actinomycetes colonies may be slow growing, so check your plates every few days for up to 2 weeks. Most of the genera in this Actinomycetes family produce a hyphal type of growth that will easily differentiate them from the large rod shaped Bacillus spp. Most members of both of these groups can form spores during their lifecycle.

Look for hard, white, ridged colonies (little "volcanoes") characteristic of Streptomyces or "powdered sugar" colonies with an indentation of agar around the colony. Bacillus spp. are also relatively easy to identify by colony morphology once you are familiar with their characteristic look.

Bacillus spp. are often able to spread across the surface of the agar, so isolation is sometimes difficult. If you are trying to isolate a Bacillus, shorten the incubation time so you can find the colonies when they are still small.

Once you think you have an isolate into pure culture, make a bacterial smear slides and Gram stain them (see Protocols for procedures) to allow you to examine the cellular morphology, arrangement, and cell wall structure of these bacteria. Look carefully for clear areas in the vegetative cells indicative of endospores. You will do an endospore stain in a later lab on any isolates that we expect to be spore formers, but look carefully for this preliminary indication of endospores.

Selective and Differential Media for Confirming Gram Stain Results

Selective Media for Gram positive Bacteria
Phenylethyl Alcohol Agar (PEA) PEA selects for the growth of gram positive organisms by inhibiting the growth of gram negative bacilli. The alcohol in the medium dissolves the gram negative lipid outer membrane that Gram positive bacteria lack. The thin layer of peptidoglycan, characteristic of gram negative bacteria allows entry of the phenylethyl alcohol into a gram negative bacterium, but not into a gram positive bacterium (gram positive bacteria have a much thicker, more impermeable layer of peptidoglycan in their cell wall). Alcohol interfers with DNA synthesis and kills bacteria with a gram negative cell wall structure. This medium is particularly useful at inhibiting the overgrowth of Gram negative Proteus species that tend to swarm (they are highly motile) all over an agar plate and, thus, using PEA medium makes isolating Gram positive organisms easier in a mixed population.
Recipe: 1.80% Bacto Agar, 1.50% Tryptone, 0.50% Phytone, 0.50% Sodium Chloride, 0.25% Phenylethyl Alcohol (PEA).

Postive control organism: Staphylococcus epidermidis

Selective and Differential Medium for Gram negative Bacteria
Eosin–Methylene Blue (EMB) Agar is a selective and differential medium used to enhance selection of gram negative bacteria. The medium contains peptone, lactose, sucrose, dipotassium phosphate, eosin and methylene blue dyes. Colonies produced by lactose non-fermentors are not dark blue or black because the dyes are not precipitated. The growth of gram positive bacteria is generally inhibited on EMB agar because of the toxicity of methlyene blue dye. In the low dye concentration found in this medium, the protective lipid outer membrane of gram negative bacteria prevents entry of this toxic water-soluble dye because the gram negative outer membrane is a phospholipid bilayer that repels many water-soluble substances. The cell wall structure of gram positive bacteria lacks the protective lipid based outer membrane and that makes them more sensitive to the toxicity of methyene blue.

Recipe:1% peptone, 1% Lactose, 0.2% dipotassium phosphate, 0.04% eosin Y, 0.0065% methylene blue 1.5% Agar. final pH 6.9-7.3

Table 2. Colonial appearance on EMB Agar after 18-24 hours at 35°C.

EMB is also a differential medium. Eosin and methylene blue act as indicators to differentiate between gram negative organisms that ferment lactose from those that do not ferment lactose. Most bacteria that ferment lactose form colonies on EMB agar that are dark blue to black due to precipitation of the dyes by the acid by-products of lactose fermentation. Lactose fermentors have dark colonies from dye precipitation while non-fermentors have light colored colonies. Additionally, E. coli often gives a characteristic green-metallic sheen on EMB, making this medium somewhat differential for E. coli to help distinguish it from other lactose fermenting gram negative bacteria.
Recipe: 0.04% Eosin Y, 0.0065% methylene blue, 1.0% peptone, 2.0% lactose, 0.2% K2HPO4, 1.5% agar, pH 7.1

Organism	Colonial Appearance
Escherichia coli	purple with black center/ green metallic sheen
Klebsiella pneumoniae	dark centered colonies/ sometimes a metallic sheen
Enterobacter aeorogenes	pink colonies/ no metallic sheen
Proteus mirabilis	colorless colonies
Salmonella typhimurium	colorless colonies

Reference: Dehydrated Culture Media and Reagents for Microbiology. DIFCO Laboratories, Detroit, MI. 1984.

Differential Medium For Assessment of Soil Exoenzymes: Amylase, Cellulase, Phosphatase

Nutrient Agar (NA) General Purpose Medium is used to determine comparative number of total culturable bacteria: and for growth of non fastidious organisms once in pure culture. For plate counts use your P200 micropipet and sterile tips, dispense 100µl of a soil extract dilution (choose a dilution that should give you between 30-300 CFUs) onto a pre-labeled Nutrient agar plate. Use a sterile, disposable spreader to evenly distribute the diluted soil extract all over the culture plate. Repeat for two other dilutions (one 10fold more and one l0 fold less dilute).
Nutrient Agar General Purpose Medium:
0.3% Beef extract, 0.5% Peptone, 1.5% Agar at pH 6.6- 7.0 at 25°C.

Starch Medium is used to determine the % of amylase producing (starch digesting) culturable microbes when compared to the total number counted on NA: Using your P200 micropipet and sterile tips, dispense 100µl of a soil extract dilution (choose a dilution that should give you between 30-300 CFUs) into the center of a pre-labeled Nutrient agar plate. Use a sterile, disposable spreader to evenly distribute the diluted soil extract all over the culture plate. Repeat for two other dilutions (one 10 fold more and one l0 fold less dilute).
Starch medium :
2.5% (wt/vol) soluble starch in Nutrient Agar

Reference: Beishir, Lois. 1996. Microbiology in Practice 6th ed. HarperCollins Publishers Inc. New York. Module 33: 301-306.

Cellulose Medium is used to determine the % of cellulolytic microbes (those producing cellulase) when compared to the total number counted in NA : Using your P200 micropipet and sterile tips, dispense 100µl of a soil extract dilution (choose a dilution that should give you between 30-300 CFUs) onto a pre-labeled plate of Cellulose medium. Use a sterile, disposable spreader to evenly distribute the diluted soil extract all over the culture plate.
Cellulose Congo Red Agar:
0.05% K2HPO4; 0.025% MgSo4; 0.188% ashed, acid washed cellulose powder; 0.02% Congo red, 0.5% Noble Agar, 0.2% gelatin, 10%(vol/vol) sterile soil extract (Soil extract prepared as follows:105 g of air-dried sieved soil and 660 ml of deionized water are placed in a 1 litre bottle and autoclaved once at 15 psi for 15 minutes, then again after 24 hours. The contents of the bottle are left to settle for at least a week and then the supernatant is decanted and filtered. The final pH should be 7.0 - 8.0.)
Reference: Hendricks, Charles W., Doyle, J.D., Hugley, B. (1995) A New Solid Medium for Enumerating Cellulose-Utilizing Bacteria in Soil. Applied and Environmental Microbiology. May: 2016-2010.

Phosphate Medium (Pidovskaya medium) is used to determine the % phosphate solubilizing microbes (those producing phosphatases) in a soil community: Using your P200 micropipet and sterile tips, dispense 100µl of a soil extract dilution (choose a dilution that should give you between 30-300 CFUs) onto a labeled Pidovskaya medium plate.
Pidovskaya Medium:
1.0% glucose, 0.05% yeast extract, 0.01% Calcium Chloride (CaCl₂), 0.025% Magnesium Sulfate (MgSO⁴.7H₂0), 0.251% Calcium Phosphate [Ca(PO₄)], 2.0% agar.
References: Pikovskaya, R.I. 1948. Mobilization of phosphorus in soil in connection with the vital activity of some microbial species. Mikrobiologiya 17, 362-370, modified by Pranjal Baruah (2007) Isolation of phosphate solubilizing bacteria from soil and its activity. Biotechindia.files.wordpress.com/2007/12/isolation.pdf.

Incubate all cultures at room temp until mature colonies have formed and then refrigerate before the bacteria overgrow.

Count the total number of colonies on the Nutrient Agar plate that has between 30-300 colonies and record the dilution. Assess total culturable CFUs for that dilution.

Find the starch plate with between 30-300 total colonies and note the dilution. Flood the starch plate with a thin layer of iodine and count the number of colonies that show starch digestion activity as a clear zone or non-blue halo around the colony. Iodine turns from reddish brown to blue-black in the presence of starch; therefore, bacteria in those colonies that do NOT turn black have produced and secreted amalyase enzyme that digested the starch in the area around the colony into simpler sugar. Those bacteria that do not produce amalyase are unable to break down starch, revealed by the color change to blue-black in the medium surrounding those colonies. Record the number of starch digesting organisms and the dilution.

Find the cellulose plate showing between 30-300 total colonies. Count the number of colonies that show cellulose digestion activity by looking for positive digestion as a clear zone or halo around the colony. Record as number of cellulose digesting organisms in that dilution.

Do the same for the assessment of number of phosphate digestion microbes in a particular dilution. The positive colonies will be red that show phosphate solubilizing activity.

Calculating the % of digestion positive microbes in the total culturable population

Use the soil extract dilution on the plates counted to normalize all the calculations to CFUs/gram of soil (wet weight) for each assessment medium. If you divide the number of colonies counted by the volume of inoculum plated, times the dilution factor of that inoculum, you will obtain the number of that type of bacteria per gram of soil.

For example, if you counted 150 colonies on the 10^-3 plate the calculation is:
150/(0.1ml vol. of inoculum*1X10^-3dilution)= 150X10⁴ which in scientific notation is written as 1.5X10⁶ CFU/gram

Once you calculate the total number of aerobically culturable bacteria (cfu/g) on the general purpose media, you can determine the % of the total number able to solubilize phosphate by dividing the number of phosphatase positive colonies by the total number of culturable colonies---if the colonies counted are compared from the same plate dilutions.

This calculation of the % of cultured bacteria that are positive for each tested enzymatic activity: (# positive colonies/total count on nutrient agar X 100) gives you a sense of the prevalence and variety of soil organisms in a community with particular substrate utilizing potential.

Medium For Assessment of motility and ability to reduce nitrate

Motility and nitrate reduction medium Mannitol Nitrate Motility Medium

1% Casein Peptone, 0.75% Mannitol, 0.1% Potassium Nitrate, 0.004% Phenol Red, 0.35% Bacteriological Agar. pH 7.6 at 25°C

Mannitol Nitrate Motility Medium is a semisolid general purpose medium that provides nitrogen, minerals, amino acids and nutrients from the Casein peptone. The fermentable carbohydrate, Mannitol, provides an energy source. Potassium Nitrate provides additional nutrients and organisms capable of reducing nitrate show increased motility. Phenol red is a pH indicator. Agar is included as a solidifying agent.

The inoculating needle is used to introduce a single deep inoculum to the center of the soft agar pour with stab technique. Motile bacteria diffuse away from the line of inoculation clouding the otherwise transparent medium. Non motile organism only grow along the stab line. If mannitol is fermented the medium changes color from red to yellow. After incubation, Gries reagent (2 drops of solution A and 3 drops of solution B, are added to the surface of the medium to test for Nitrate reduction. Nitrate-negative organisms are unable to reduce nitrates and no color change is observed after adding the reagent. If the organism is nitrate positive a pink or red color indicate that nitrates have been reduced to nitrites.

Gries Reagent: Solution A: 0.8% Sulfanilic Acid in 5N Acetic Acid. Solution B: 0.001% Alpha-Naphthylamine in 5N Acetic Acid.

Links to Labs

Lab 1
Lab 2
Lab 3
Lab 4
Lab 5
Lab 6
Lab 7
Lab 8
Lab 9
Lab 10
Lab11
Lab 12