BISC209/S11: Culture Media

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Wellesley College-BISC 209 Microbiology -Spring 2011

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

Perhaps we can add to this wonderful library of images of colonies of culturable microorganisms. [1]. The images can enhance your ability to select for desired Genera based on colony morphology.


The composition of medium is an important factor when attempting to culture microorganisms. The components and pH can be manipulated to favor the nutritional requirements of particular bacterial groups present if the desired microbes are unlikely to be successfully isolated in a general purpose medium. Keep in mind that the composition of selective enrichment media, particularly the difference between it and general purpose media, provides valuable information about the metabolic range of the microbes isolated on it.

The recipe for each enrichment medium described here includes 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 and agar:
Nutrient 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.

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 a nutrient 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 ingedient 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 on the formulations and types of media available in microbiology see: BD diagnostice Systems Difco catalog of media

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

Finding Denitrifying Methylotrophs (Hyphomicrobia) Bacteria :
Use Denitrifying Methylotrophs Medium with (DMMM) and without methanol(DMM)

  • Denitrifying Methylotrophs(Hyphomicrobium) Medium with methanol (DMMM) (Marine Biology Laboratory, Woods Hole, MA recipe)(DMMM medium): (1% Freshwater Base (FWB: 10% NaCl, 4% MgCl2*6H20; 1% CaCl2*2H2O; 2% KH2Po4 (acidic); 5% KCl); 0.02M 3-(N-morpholino)propanesulfonic acid (MOPS C7H15NO4S pH 7.2), 0.2mM Na2SO4,; 0.15mM K3PO4 pH 7.2; 5.0 mM NH4Cl, 0.5% KNO3; pH 7); 1% vitamin mix (Sigma product number M7150 Murashige and Skoog Vitamin Powder); 0.25% methanol
  • DMM medium: DMMM without 0.25% methanol.
  • DMM solid medium: DMM with 1.5% agar grown in a methanol gas enriched atmosphere chamber.

The bacteria in this group have two very distinct morphological forms (dimorphic): a motile, swarmer cell and a stalked, stationary, prosthecate cell. The motile cell is flagellated, typically with a polar or subpolar flagellum. The sessile cell attaches to the surface of some solid material using either an adhesive or a specialized outgrowth at one pole called a prostheca. Reproduction occurs by budding from the prosthecae and results in one motile and one sessile offspring.

Hyphomicrobia are facultative aerobes and chemoorganotrophs in the alpha-Proteobacteria group. They are common in aquatic environments and in nitrate rich environments. In nitrate-rich non-aquatic sites, such as soil, these organisms rely on other bacteria to degrade local organic molecules to a useable form. They use C1 carbons for nutrients and reduce nitrate to N2 for their energy needs. The enrichment protocol we will use relies on the ability of these bacteria to grow anaerobically on methanol and their ability to reduce nitrate to Nitrogen.

In pure culture, you should be able to find both morphologic forms in about equal ratio. Genera of this group we might find in soil include: Caulobacter, Asticcacaulis, Hyphomicrobium, Pedomicrobium, Hyphomonas, and Thiodendron (Find more reference information in The Prokayotes, on-line access available as directed in the Resources section of this wiki).

Isolation is based on their inability to grow on nutrient agar while they have the ability to grow anaerobically in the presence of methanol, while reducing nitrate to N2. (We will use screw cap tubes to reduce the oxygen gas exchange in the culture)

Primary Enrichment/Selection:
  • Weigh 0.5-1 g of soil (depending on the size of the tube) and add it to a screw cap tube containing Denitrifying Methylotroph (Hyphomicrobium) Medium with methanol(DMMM). Add more liquid media until the tube is completely full (so that you will create an anaerobic environment after the aerobic bacteria in the media use up all the oxygen.) Take precautions when handling the media as methanol is highly toxic. Always wear gloves.
  • Tighten the cap.
  • Incubate at 30°C for several days to 1 week.
  • Watch the culture for the development of turbidity.
  • When the solution begins to look turbid, slowly loosen the cap while watching for the appearance of bubbles rising in the liquid. The bubbles appear because of decreased pressure from loosening the cap and indicate the production of N2 gas by these denitrifying bacteria.

Secondary Enrichment/Selection:

  • a. Transfer 2 ml of the culture liquid to a fresh screw cap tube of the same DMMM medium with methanol and incubate at 30°C. Incubate for several days. (This is a secondary enrichment.)
  • b. Check for Nitrogen bubbles, as described previously, and then isolation streak a plate of DMM (hypomicrobia) solid medium without methanol. Incubate in a closed jar containing an open tube of methanol, ask your instructor if you are not sure where this jar is located and work with it in the hood. Wear gloves and remember that methanol is toxic! Incubate at 30°C. Your instructor will change the tube of methanol every few days.

Isolation to Pure Culture:
After 3 days or so, examine the streaked plates using a magnifying glass or the colony counter. Look for tiny, "drop-of-water" colonies, possibly volcano shaped. Select a few likely colonies and streak for isolation on DMM solid medium plates that have NO methanol. After the bacteria are transferred to zone one of your new plate using a sterile toothpick or inoculating needle, you can use your loop to streak out the other zones. Keep streaking for isolation until you have pure colony isolates. (Other organisms can overgrow the Hyphomicorbia and mix in so that it is difficult to separate them in the streak plates.) Note that Hyphomicrobia tend to attach to other bacteria. Reduce the chance of contamination by transferring newly arising colonies as soon as you see them and using the inoculating needle or sterile toothpick rather than a loop to pick up the bacteria from the center of your colony. Transfer to a new plate of DMM and put the plate into the jar with the tube of methanol.

Finally, when you feel confident you have a pure isolate of a denitrifying methylotroph based on colony appearance, perform an isolation streak onto nutrient agar. If anything grows, you do not have a pure colony of Hyphomicrobia, as these bacteria can not grow on nutrient agar; however, you may have bacteria of different group of methylotrophic bacteria (Pseudomonas spp, Lolium spp, Methylobacterium spp etc. so you can continue your characterization of the isolate.

Maintenance in lab atmospheric conditions (without methanol)
Once your isolate is in pure culture on DMM, isolation streak onto Peptone Yeast Calcium Maintenance for Hyphomicrobium (PYCM: 0.25% peptone; 0.05% yeast extract; 1 mM CaCl2; 2 mM MgSO4; 1% agar). Also streak out a new subculture onto DMM. If the isolate grows successfully on PCYM, you can maintain your isolate exclusively on PYCM medium at RT in room air. If it does not grow well on PYCM medium, continue to maintain on DMM solid medium in the jar with liquid methanol.

Finding Nitrogen Cyclying Bacteria: Azotobacter
Use Azotobacter Medium

Azotobacter, important nitrogen cycling bacteria, are able to aerobically use N2 as their source of Nitrogen without a symbiotic partner. They can use mannitol as their sole carbon source, although we are not using that capability for our initial selection. They are generally Gram-negative, large rods, or ovoid cells.

Medium: 0.08% K2HPO4; 0.02% KH2PO4, 0.02% MgSO4 0.01% CaSO4; 0.0015% FeSO4/7H2O; 0.00025% g MoO3; 0.5% sucrose (2010,2011).

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).

  • 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 purity from contaminants:
    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 the colony with the correct bacterial morphology onto Azotobacter medium. What should Azotobacter bacteria look like in a Gram stain?
  • Characterization:
    Once you have an appropriate isolate 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 [2] 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 or survive the heat or lack of moisture. Now we need to coax those spores back into their vegetative state. The medium uses cycloheximide to inhibit fungal growth since many fungi make spores, too. Cycloheximide is highly toxic to eukaryotic cells (yours!) so use caution and wear gloves when working with it or with media containing cycloheximide.


    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 and the thin layer of peptidoglycan allows entry of the phenylethyl alcohol into the cell which then interfers with DNA synthesis. This medium is particularly useful at inhibiting the overgrowth of Gram negative Proteus species that tend to swarm (they are highly motile) and, thus, make isolation of Gram positive organisms difficult in a mixed population .
    Recipe: 1.80% Bacto Agar, 1.50% Tryptone, 0.50% Phytone, 0.50% Sodium Chloride, 0.25% Phenylethyl Alcohol (PEA)..

    Mannitol Salt Agar (an alternative to PEA, not used in 2010)

    Postive control organism: Staphylococcus epidermidis

    Selective and Differential Medium for Gram negative Bacteria
    Eosin–Methylene Blue (EMB) Agar is a differential medium for the detection of Gram negative enteric bacteria. The medium contains peptone, lactose, sucrose, dipotassium phosphate, eosin and methylene blue dyes. Eosin and methylene blue act as indicators to differentiate between Gram negative organisms that ferment lactose and those that do not ferment lactose. Most bacteria that ferment lactose form colonies on EMB agar that are dark blue to black with a metallic sheen due to precipitation of the dyes by the acid by-products of fermentation. Colonies produced by lactose non-fermentors are not dark blue or black. The growth of Gram positive bacteria is generally inhibited on EMB agar because of the toxicity of methlyene blue dye. In low concentration, the protective lipid outer membrane of Gram negative bacteria prevents entry of the toxic water soluble dye while the more porous cell wall of Gram positive bacteria without the protective outer membrane 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, in that it can be used to visually differentiate Gram negative lactose fermenting bacteria from non-fermenters. Lactose fermentors have dark pigmented colonies while non-fermentors have light colored colonies. E. coli often gives a green-metallic sheen on EMB, making this medium somewhat differential for E. coli.
    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 amalyase 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 :
    0.3% (wt/vol) soluble starch in Nutrient Agar or 3g/liter.
    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.

    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). Record as number of starch digesting organisms in that 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 plated*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.

    Differential Medium For Assessment of ammonia, nitrate/nitrite

    Grow organisms in the appropriate medium so you have freshly grown isolation streak plates. (1-3 days old).

    You will inoculate tubes of peptone meat extract soft agar (PM) medium today.
    Recipe: 0.5%(5 g/L) peptone; 0.3% (3g/L) meat extract; 1 L water; pH 7.

    1. Use a sterile 5mL pipet to pipet 5ml of sterile DMM broth into sterile 16x100mm culture tubes (one for each isolate to be tested).
    2. Use a sterile loop or toothpick to add a small amount of growth from a well isolated colony growing in pure culture on a recent (1-3 days old) DMM agar isolation streak plate. Mix well.
    3. Compare the turbidity to a #5 McFarland turbidity standard available at the instructor’s bench. Add more medium or more culture until the turbidity appears to match the standard.
    4. Label 6 prepared tubes (3 screw capped and 3 covered loosely, preferably 16X100mm) containing 8ml peptone meat extract medium (PM) for each isolate.
    5. Transfer 200μl from one of the diluted (DMM)isolate cultures to each of 3 sterile screw cap tubes containing peptone meat extract medium (PM).
    6. Using your P1000 and sterile tips, add more liquid peptone broth (PM) medium until the tubes are completely full.
    7. Tighten the cap leaving no air interface. The goal is to create an anaerobic environment to facilitate the growth of anaerobic Hyphomicrobia bacteria.
    8. Add 200μl of the diluted DMM culture to the each of 3 PM tubes that are loosely covered (not screw cap).
    9. Test one of the loosely capped tubes immediately for the presence of ammonia, and nitrate/nitrite using the appropriate test strips. Directions for using the test strips are detailed below. 10. Incubate all the cultures in your team’s rack at 30°C.
    11. Prepare a data sheet for your team’s isolates. Arrange a schedule with your teammates for someone to come in to test all the cultures each day and to record the results on the data sheet. Look for evidence of growth (turbidity). Test all 6 cultures each day for the presence of ammonia, nitrate and nitrite using the test strips. Have a column or row on your data sheet to record personal observations (smell, color, etc). PROTOCOL for TESTING: Ammonia, and Nitrite/Nitrate

    The detection limit of these strips is: for Ammonia (6ppm - mg/L)), nitrite (10ppp-mg/L), and nitrate (200ppm-mg/L).

    1. Avoid contaminating your cultures by aseptically removing about 200-500ul of culture solution and placing it in a small test-tube. Discard the tips in an orange biohazard bag.
    2. You can use the same tube for both strips today and you will take a new sample each day that you perform the test. Be sure you have organized a good system for recording the data daily so all teammates will be provided with the data.
    3. Carefully remove one ammonia test strip at a time and reseal the container.
    4. Dip the ammonia test strip in the solution for 10 seconds, remove slowly so the strip will not drip.
    5. Use the color chart on the bottle to determine ppm (mg/L).
    6. Discard the strip in an orange biohazard bag immediately.

    7. Carefully remove a nitrate/nitrite test strip from the container and reseal the container.
    8. Dip into the 1 ml tube solution and remove the strip 2 times. Remove the strip so the pads face up.
    9. Do not shake the strip.
    10. Wait 60 seconds.
    11. Determine the concentration (ppm = mg/L) using the color chart on the Nitrate/Nitrite container.
    12. Discard properly.

    Links to Labs

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