3.5: Exercise 2 - observing yeast and bacterial cultures - Biology

3.5: Exercise 2 - observing yeast and bacterial cultures - Biology

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Students should work in groups of three. Each group will receive three cultures of S. cerevisiae (SC), S. pombe (SP), and E. coli (EC).

1. Label the three slides. Each slide will contain two samples for comparison. The slides are large enough to accommodate two samples—and two coverslips. Number the slides with a Sharpie (Use the frosted area, if the slide has one.) As you work, be sure to record which of the two samples is closer to the labeled end of the slide. Record your data in the spaces provided below or in your notebook.

Slide 1: compare cultures of S. pombe and S. cerevisiae.
Slide 2: compare cultures of S. cerevisiae and E. coli.
Slide 3: cultures of S. pombe and E. coli with the 100X objective (1000X).

2. Prepare concentrated suspensions of cultured cells. All of the cell cultures appear cloudy, because the cell densities are high. Even so, you will be observing a few microliters of cell suspension under the microscope, and cells may be few and far between. Concentrate
the cultures to ensure that there will be enough cells in the microscope’s field of view for meaningful observations.

  • Concentrate the cells by centrifuging the test tubes containing the cultures for a count
    of 10 in a microcentrifuge set at top speed. Hold down the Quick button on the Labnet microcentrifuges or the button between the two dials on the Eppendorf microcentrifuges.
  • Remove most of the culture medium with a P1000 micropipette, until the medium just covers the cell pellet.
  • Resuspend the cells with the vortex mixer.

3. Transfer and stain the cell samples.

• Transfer 2 μL of each cell suspension to the slide, using a P20 micropipette.
• Stain the cells by adding 6 μL of Gram’s Iodine to each cell suspension with a P20 micropipette. Cover each sample with a coverslip.

4. Observe samples of the three species using the 40X and 100X objectives. Each member of your group should observe the two species on the slide that he/she prepared. Work with another member of the group to make observations of the third species. Record your observations and answer the questions in the spaces below.


The oil immersion lens is required for the 100X objective, and this oil can damage the 40X objective. Be careful to follow the instructions below exactly.

The steps must be performed in the correct order to protect the lenses!

  1. Focus the microsope on EITHER an s. cerevisiae or s. pombe culture. Focus first with the 10X objective, then move to the 40X objective and refocus with the fine focus control. Record your observations.
  2. You are now ready to move to the 100X oil immersion objective! Rotate the nosepiece halfway between the 40X and 100X objectives. WITHOUT moving the stage, apply a drop of immersion oil on top of your coverslip where the light is shining through the slide. SLOWLYrotate the 100X objective into place, submerging it into the drop of oil. Use the FINE focus knob to bring the yeast into focus. Record your observations.
  3. Rotate the nosepiece halfway between the 40X and 100X objectives again. Do not attempt to move the stage with the 100X objective in place.
  4. Use the XY controls to move the stage to the second sample on the slide WITHOUT changing the focus.
  5. Rotate the 40X lens into position. Adjust the focus and record your observations.
  6. Repeat step 2 above and record your observations at 100X.
  7. Rotate the 100X objective out of position. Remove the slide and discard it in the glass waste. Clean the 100X lens with lens paper. Check that no oil is present on any of the other lenses.

S. cerevisiae observations
Sketch the cells that you see with the two different lenses in the boxes below.

What fraction of the cells show buds?
Can you detect any subcellular structures at 1000X magnification?

S. pombe observations
Sketch the cells that you see with the two different lenses in the boxes below.

What fraction of the cells show septa?
Can you detect any subcellular structures at 1000X magnification?

E. coli observations
Sketch the cells that you see with the two different lenses in the boxes below.

What fraction of the cells are motile?
How would you describe the structure of an E. coli?

Yeast Exercises

Microbiologists like to begin their experiments with a single colony, because the cells in a colony are the progeny of a single cell. A concern in all genetic experiments is unknown mutations that arise spontaneously and may affect the phenotype being studied. Spontaneous mutations arise constantly in all cells, with a rate of approximately 10-8/base/ generation. For S. cerevisiae, with a genome of 12 Mbp, most cells will have accumulated at least one mutation by the time that they have undergone 9-10 divisions. A colony, which has hundreds of millions of cells, is therefore a population of genetically very similar, but not completely identical, organisms.
Researchers commonly use streak plates to isolate single colonies. A streak plate is actually a serial dilution of an existing culture on solid media. Researchers begin a streak by picking up a small sample of yeast or another microorganism with a sterile instrument, which could be a platinum loop, a toothpick or micropipette tip. They then spread the culture by making a series of zig-zag strokes across the surface of the plate. The number of cells on the loop or toothpick decreases as the streak progresses. Consequently, streaks appear thickest at their starting points, and the streak thickness decreases until it is possible to detect well-isolated single colonies near the end of the streak. Because it may be difficult to resolve colonies from a single streak, many labs use a series of streaks on the same plate to separate colonies. Each new streak is done with a freshly sterilized loop that picks up cells by crossing over the tracks of the previous streak, before beginning a new series of zig-zags. In our experiments, we will use a multi-streak protocol, which allows us to culture multiple strains on a single plate of culture medium. (See the figure below.) The streak plates that you prepare will contain the stocks for future experiments. As you streak your strains, pay careful attention to detail to avoid cross-contamination or confusion about the identities of individual strains.

  1. Your team will be assigned three different S. cerevisiae met strains to culture. The stock strains that you will use have been grown to a high density in liquid YPD medium. Gather the YPD cultures of the parent strains to be propagated, a platinum inoculation loop, and an agar plate with fresh YPD media. This plate will serve as your team’s stock plate for the semester.
  2. Divide your team’s stock plate into sectors by marking the bottom of the plate with a magic marker. CLEARLY label each sector with a code for the strain that will be streaked in it. As series of three streaks will be made within each sector, as described on the previous page. Keep the labels at the rim of the plate and use small letters. Note your initials and the date.
  3. Sterilize the inoculation loop by holding it in the flame of a Bunsen burner until it glows red. Cool the loop by briefly touching the surface of your agar plate before proceeding. (A hot loop will kill yeast cells that it contacts!)
  4. The cells in your stock cultures may have settled over time. Resuspend the cells using the vortex mixer. Working quickly, remove the cap the first culture tube. sterilized loop into the culture, remove the loop and place the cap back on the culture tube.
  5. Transfer cells to the appropriately labeled sector on your stock plate. First streak: make several zigzags across the outside edge of the sector with the loop. LIGHTLY touch the agar surface as you move the toothpick. Think of pushing a hockey puck across an ice rink, rather than digging a ditch. You do NOT want to put track marks in the agar! Replace the lid.
  6. Sterilize the loop in the flame and cool it on the agar surface before beginning the second streak. Make a second vertical streak from the rim of the plate toward the center, staying within the sector. The streak should cross the zigzags in the first streak. Close the lid and re-sterilize (and cool) the loop.
  7. Third streak: make a third series of zigzags that cross back and forth over the straight second streak, beginning at the outer edge of the plate and moving toward the center. Be careful to stay within the sector. Close the lid and sterilize the loop.
  8. Repeat steps 4-7 for each of your liquid cultures.
  9. Invert the plate and incubate it at 30 °C until individual colonies are visible, which is usually 24-48 hours.
Exercise 2 – Spot plates

Scientists use spot plates both to calculate the number of cells in cultures and to obtain information about the growth properties of strains on different media. The figure to the right shows an example of a typical spot plate. Each row represents a dilution series from a different yeast culture. The same volume of diluted culture is used for each spot. The dilution series is planned so that the most dilute spots contains a small number of individual colonies that can be distinguished from one another, typically less than ten.

Most commonly, investigators make a series of 1:10 dilutions in sterile water and then spot a few microliters of each dilution in a row. In this experiment, 5 μL aliquots were spotted from the serial dilutions. Note that it is possible to count individual colonies in the most dilute samples. This in turn enables you to calculate the number of viable cells in the original culture. In the top row, you can distinguish 4 colonies in the sample that has been diluted 100,000-fold. The original culture would have contained 400,000 cells in 5 μL, which corresponds to 80 million cells per mL (8 x 107 cells/mL).

In this experiment, you will use spot plates to estimate the cell densities of log phase and stationary phase cultures of S. cerevisiae. Each group will receive two cultures:

CL – S. cerevisiae log phase culture
CS – S. cerevisiae stationary phase culture
Spot the dilution series from each culture on a separate row of the plate. Be sure to LABEL the rows!

  1. Alignment grids are useful for preparing good-looking spot plates! Obtain an alignment grid (right) and mark the target positions for culture dilutions. Place an orientation mark at one point along the circumference.
  2. Label the plate with your initials and date with small letters around the BOTTOM rim of the dish. Put a hash mark on the edge of the plate to serve as an alignment marker.
  3. Prepare a series of five 1:10 dilutions from each culture using sterile water. (Diagrams in your lab notebook are often helpful in designing dilution series.) To prepare a serial dilution, first pipette 90 μL sterile water into five microcentrifuge tubes. Next, use a P20 to transfer 10 μL from the original culture into the first tube. Eject the tip into the appropriate waste container.
  4. Vortex the newly diluted culture to insure that the cells are uniformly distributed. With a fresh micropipette tip, transfer 10 μL from this tube to the second tube in the series. Repeat this step for tubes 3-5.
  5. Prepare the spot plate. Beginning with the last dilution in the series, spot 5 μL spots in a row. Vortex each dilution before spotting it, because cells may have settled. You will be able to use a single pipette tip for each dilution series, since you started with the most dilute cell suspension.
  6. Repeat step 3 for each culture that you are analyzing. Be careful to note in your lab notebook which culture has been spotted into each row on the plate!
  7. Leave the plate right side up for

Exercise 3 – Estimating cell densities with a spectrophotometer

The spectrophotometer provides a “quick and dirty” way to estimate the density of cells in a culture. In contrast to spot plates, which must be incubated for several days before colonies appear, spectrophotometer readings can be instantly converted into cell densities. On the other hand, the method does not discriminate between living and dead cells. The spectrophotometric method is based on light scattering by cells in the culture. When the light in a spectrophotometer hits a large particle such as a cell, light rays are deflected from a straight path and these light rays may not reach the detector. The greater the number of cells in a sample, the more light scattering occurs. The light scattering ability of a cell depends on its size and geometry, so a calibration curve is necessary to extrapolate optical density measurements to cell number. For example, the same number of yeast cells would scatter light more than the same number of bacterial cells, because the bacterial cells are much smaller.

Light scattering is measured with the spectrophotometer set to report absorbance. Because the principles used to measure light scattering and absorbance are different, the amount of light scattered by a solution is referred to as its “optical density” rather than its “absorbance.” The optical density of a sample analyzed at 600 nm is abbreviated OD600, with the subscript indicating the wavelength used for the measurement.

3.5: Exercise 2 - observing yeast and bacterial cultures - Biology

A simple dilution is one in which a unit volume of a liquid material of interest is combined with an appropriate volume of a solvent liquid to achieve the desired concentration. The dilution factor is the total number of unit volumes in which your material will be dissolved. The diluted material must then be thoroughly mixed to achieve the true dilution. For example, a 1:5 dilution (verbalize as "1 to 5" dilution) entails combining 1 unit volume of solute (the material to be diluted) + 4 unit volumes of the solvent medium (hence, 1 + 4 = 5 = dilution factor). The dilution factor is frequently expressed using exponents: 1:5 would be 5e-1 1:100 would be 10e-2, and so on.

Example 1: Frozen orange juice concentrate is usually diluted with 4 additional cans of cold water (the dilution solvent) giving a dilution factor of 5, i.e., the orange concentrate represents one unit volume to which you have added 4 more cans (same unit volumes) of water. So the orange concentrate is now distributed through 5 unit volumes. This would be called a 1:5 dilution, and the OJ is now 1/5 as concentrated as it was originally. So, in a simple dilution, add one less unit volume of solvent than the desired dilution factor value.

Example 2: Suppose you must prepare 400 ml of a disinfectant that requires 1:8 dilution from a concentrated stock solution with water. Divide the volume needed by the dilution factor (400 ml / 8 = 50 ml) to determine the unit volume. The dilution is then done as 50 ml concentrated disinfectant + 350 ml water.

2. Serial Dilution

A serial dilution is simply a series of simple dilutions which amplifies the dilution factor quickly beginning with a small initial quantity of material (i.e., bacterial culture, a chemical, orange juice, etc.). The source of dilution material (solute) for each step comes from the diluted material of the previous dilution step. In a serial dilution the total dilution factor at any point is the product of the individual dilution factors in each step leading up to it.

Final dilution factor (DF) = DF 1 * DF 2 * DF 3 etc.

Example: In a typical microbiology exercise the students perform a three step 1:100 serial dilution of a bacterial culture (see figure below) in the process of quantifying the number of viable bacteria in a culture (see figure below). Each step in this example uses a 1 ml total volume. The initial step combines 1 unit volume of bacterial culture (10 ul) with 99 unit volumes of broth (990 ul) = 1:100 dilution. In the second step, one unit volume of the 1:100 dilution is combined with 99 unit volumes of broth now yielding a total dilution of 1:100x100 = 1:10,000 dilution. Repeated again (the third step) the total dilution would be 1:100x10,000 = 1:1,000,000 total dilution. The concentration of bacteria is now one million times less than in the original sample.

3. Making fixed volumes of specific concentrations from liquid reagents:

V 1 C 1 =V 2 C 2 Method

Very often you will need to make a specific volume of known concentration from stock solutions, or perhaps due to limited availability of liquid materials (some chemicals are very expensive and are only sold and used in small quantities, e.g., micrograms), or to limit the amount of chemical waste. The formula below is a quick approach to calculating such dilutions where:

V = volume , C = concentration in whatever units you are working.

(stock solution attributes) V 1 C 1 =V 2 C 2 (new solution attributes)

Example: Suppose you have 3 ml of a stock solution of 100 mg/ml ampicillin (= C 1 ) and you want to make 200 ul (= V 2 ) of solution having 25 mg/ ml (= C 2 ). You need to know what volume ( V 1 ) of the stock to use as part of the 200 ul total volume needed.

V 1 = the volume of stock you will start with. This is your unknown.
C 1 = 100 mg/ ml in the stock solution
V 2 = total volume needed at the new concentration = 200 ul = 0.2 ml
C 2 = the new concentration = 25 mg/ ml

By algebraic rearrangement:

V 1 = ( V 2 x C 2 ) / C 1

V 1 = (0.2 ml x 25 mg/ml) / 100 mg/ml

and after cancelling the units,

V 1 = 0.05 ml, or 50 ul

So, you would take 0.05 ml = 50 ul of stock solution and dilute it with 150 ul of solvent to get the 200 ul of
25 mg/ ml solution needed. Remember that the amount of solvent used is based upon the final volume needed, so you have to subtract the starting volume form the final to calculate it.

4. Moles and Molar solutions (unit = M = moles/L)

Sometimes it may be more efficient to use molarity when expressing chemical concentrations. A mole is defined as exactly 6.023 x 1023 atoms, or molecules, of a substance (this is called Avagadro's number, N). The mass of one mole of an element is its atomic mass (g) and is noted for each element in the periodic table. Molecular weight is the mass (g) of a substance based on the summed atomic masses of the elements in the chemical formula. Formula weight refers to chemicals for which no discrete molecules exist for example, NaCl in solid form is made up of Na+ and Cl- ions, but there are no true molecules of NaCl. The formula weight of 1 mole NaCl would therefore be the sum of 1 atomic mass of each ion. The molecular weight (or FW) is provided as part of the information on the label of a chemical bottle. The number of moles in an arbitrary mass of an element or compound can be calculated as:

number of moles = weight (g)/ atomic (or molecular) weight (g)

Molarity (M) is the unit used to describe the number of moles of an element or compound in one liter (L) of solution (M = moles/L) and is thus a unit of concentration. By this definition, a 1.0 M solution is equivalent to one molecular weight (g/mole) of a compound brought up to 1 liter (1.0 L) volume with solvent (e.g., water) at a fixed temperature (liquids expand and contract with temperature and thus can change molarity).

Example 1: To prepare a liter of a molar solution from a dry reagent

Multiply the molecular weight (or FW) by the desired molarity to determine how many grams of reagent to use:

Suppose a compound’s MW = 194.3 g/mole

to make 0.15 M solution use 194.3 g/mole * 0.15 moles/L = 29.145 g/L

You would dissolve the specified mass of reagent in a fraction of the total volume of solvent (at STP) and then raise the volume to exactly one liter by adding additional solvent and mixing thoroughly.

Example 2: To prepare a specific volume of a specific molar solution from a dry reagent

A chemical has a FW of 180 g/mole and you need 25 ml (0.025 L) of 0.15 M (M = moles/L) solution. How many grams of the chemical are needed to make this solution?

#grams/desired volume (L) = desired molarity (mole/L) * FW (g/mole)

by algrebraic rearrangement,

#grams = desired volume (L) * desired molarity (mole/L) * FW (g/mole) #grams = 0.025 L * 0.15 mole/L * 180 g/mole

after cancelling the units,

#grams = 0.675 g

Solutions Containing Multiple Reagents

Complex solutions such as buffers, salines, fixatives, etc., may be comprised of multiple chemical reagents. In preparation of these solutions, each reagent is dealt with separately in determining how much to use to make the final solution. For each, the volume used in the calculations is the final volume of solution needed.

More examples of worked problems: Chemistry

5. Percent Solutions (% = parts per hundred or grams/100 ml)

Many reagents are mixed as percent solutions either as weight per volume (w/v) when starting with dry reagents OR volume per volume (v/v) when starting with liquid reagents. When preparing solutions from dry reagents, the same mass of any reagent is used to make a given percent concentration although the molar concentrations would be different.

Weight percent (w/v) = [mass of solute (g)/ volume of solution (ml)] x 100, and,

Volume percent (v/v) = [volume of solute (ml)/ volume of solution (ml)] x 100

For example, a 100 ml of 10% solution of any dry reagent would contain 10 g dry reagent in a final volume of 100 ml. A 10% (v/v) solution would contain 10 ml solute/ 100 ml solution volume.

Example 1: If you want to make 200 ml of 3 % NaCl you would need 0.03 g/ml x 200 ml = 6.0 g NaCl in 200 ml water.

When using liquid reagents the percent concentration is based upon volume per volume , and is similarly calculated as % concentration x volume needed = volume of reagent to use .

Example 2: If you want to make 2 L of 70% ethanol from 100% ethanol you would mix 0.70 ml/ml x 2000 ml = 1400 ml ethanol with 600 ml water.

To convert from % solution to molarity , multiply the % solution by 10 to express the percent solution grams/L, then divide by the formula weight.

Molarity = (grams reagent/100 ml) * 10
xxxxxxxxxx FW

Example 1 : Convert a 6.5 % solution of a chemical with FW = 325.6 to molarity,

[(6.5 g/100 ml) * 10] / 325.6 g/mole = [65 g/L] / 325.6g/mole = 0.1996 M

To convert from molarity to percent solution , multiply the molarity by the FW and divide by 10:

% solution = molarity * FW
xxxxxxxxxx 10

Example 2: Convert a 0.0045 M solution of a chemical having FW 178.7 to percent solution:

[0.0045 moles/L * 178.7 g/mole] / 10 = 0.08 % solution

6. Concentrated stock solutions - using "X" units

Stock solutions of stable compounds are routinely maintained in labs as more concentrated solutions that can be diluted to working strength when used in typical applications. The usual working concentration is denoted as 1x. A solution 20 times more concentrated would be denoted as 20x and would require a 1:20 dilution to restore the typical working concentration.

Example : A 1x solution of a compound has a molar concentration of 0.05 M for its typical use in a lab procedure. A 20x stock would be prepared at a concentration of 20*0.05 M = 1.0 M. A 30X stock would be 30*0.05 M = 1.5 M.

7. Normality (N): Conversion to Molarity

Normality = n*M where n = number of protons (H+) in a molecule of the acid.

Example : In the formula for concentrated sulfuric (36 N H2SO4), there are two protons, so, its molarity= N/2. So, 36N H2SO4 = 36/2 = 18 M.

Describing Colonial Morphology of Bacteria

By looking closely at the colonial growth on the surface of a solid medium, characteristics such as surface texture, transparency, and the color or hue of the growth can be described. The following three characteristics are readily apparent whether you’re looking at a single bacterial colony or more dense growth, without the aid of any type of magnifying device.

Texture—describes how the surface of the colony appears. Common terms used to describe texture may include smooth, glistening, mucoid, slimy, dry, powdery, flaky etc.

Transparency—colonies may be transparent (you can see through them), translucent (light passes through them), or opaque (solid-appearing).

Color or Pigmentation—many bacteria produce intracellular pigments which cause their colonies to appear a distinct color, such as yellow, pink, purple or red. Many bacteria do not produce any pigment and appear white or gray.

Figure 3. Bacteriological descriptions of colonial morphology

As the bacterial population increases in number, the colonies get larger and begin to take on a shape or form. These can be quite distinctive and provide a good way to tell colonies apart when they are similar in color or texture. The following three characteristics can be described for bacteria when a single, separate colony can be observed. It may be helpful to use a magnifying tool, such as a colony counter or dissecting microscope, to enable a close-up view of the colonies. Colonies should be described as to their overall size, their shape or form, what a close-up of the edges of the colony looks like (edge or margin of the colony), and how the colony appears when you observe it from the side (elevation).

Figure 4 shows a close-up of colonies growing on the surface of an agar plate. In this example, the differences between the two bacteria are obvious, because each has a distinctive colonial morphology.

Figure 4. Two different types of bacterial colonies on an agar plate.

Using microbiology terms, describe fully the colonial morphology of the two colonies shown above. A full description will include texture, transparency, color, and form (size, overall shape, margin, and elevation).

Form (shape, margin, elevation): ____________________________________________

The number of available media to grow bacteria is considerable. Some media are considered general all-purpose media and support growth of a large variety of organisms. A prime example of an all-purpose medium is tryptic soy broth (TSB). Specialized media are used in the identification of bacteria and are supplemented with dyes, pH indicators, or antibiotics. One type, enriched media, contains growth factors, vitamins, and other essential nutrients to promote the growth of fastidious organisms, organisms that cannot make certain nutrients and require them to be added to the medium. When the complete chemical composition of a medium is known, it is called a chemically defined medium. For example, in EZ medium, all individual chemical components are identified and the exact amounts of each is known. In complex media, which contain extracts and digests of yeasts, meat, or plants, the precise chemical composition of the medium is not known. Amounts of individual components are undetermined and variable. Nutrient broth, tryptic soy broth, and brain heart infusion, are all examples of complex media.

Figure 1. On this MacConkey agar plate, the lactose-fermenter E. coli colonies are bright pink. Serratia marcescens, which does not ferment lactose, forms a cream-colored streak on the tan medium. (credit: American Society for Microbiology)

Media that inhibit the growth of unwanted microorganisms and support the growth of the organism of interest by supplying nutrients and reducing competition are called selective media. An example of a selective medium is MacConkey agar. It contains bile salts and crystal violet, which interfere with the growth of many gram-positive bacteria and favor the growth of gram-negative bacteria, particularly the Enterobacteriaceae. These species are commonly named enterics, reside in the intestine, and are adapted to the presence of bile salts. The enrichment cultures foster the preferential growth of a desired microorganism that represents a fraction of the organisms present in an inoculum. For example, if we want to isolate bacteria that break down crude oil, hydrocarbonoclastic bacteria, sequential subculturing in a medium that supplies carbon only in the form of crude oil will enrich the cultures with oil-eating bacteria. The differential media make it easy to distinguish colonies of different bacteria by a change in the color of the colonies or the color of the medium. Color changes are the result of end products created by interaction of bacterial enzymes with differential substrates in the medium or, in the case of hemolytic reactions, the lysis of red blood cells in the medium. In Figure 1, the differential fermentation of lactose can be observed on MacConkey agar. The lactose fermenters produce acid, which turns the medium and the colonies of strong fermenters hot pink. The medium is supplemented with the pH indicator neutral red, which turns to hot pink at low pH. Selective and differential media can be combined and play an important role in the identification of bacteria by biochemical methods.

Think about It

  • Distinguish complex and chemically defined media.
  • Distinguish selective and enrichment media.

The End-of-Year Picnic

The microbiology department is celebrating the end of the school year in May by holding its traditional picnic on the green. The speeches drag on for a couple of hours, but finally all the faculty and students can dig into the food: chicken salad, tomatoes, onions, salad, and custard pie. By evening, the whole department, except for two vegetarian students who did not eat the chicken salad, is stricken with nausea, vomiting, retching, and abdominal cramping. Several individuals complain of diarrhea. One patient shows signs of shock (low blood pressure). Blood and stool samples are collected from patients, and an analysis of all foods served at the meal is conducted.

Bacteria can cause gastroenteritis (inflammation of the stomach and intestinal tract) either by colonizing and replicating in the host, which is considered an infection, or by secreting toxins, which is considered intoxication. Signs and symptoms of infections are typically delayed, whereas intoxication manifests within hours, as happened after the picnic.

Blood samples from the patients showed no signs of bacterial infection, which further suggests that this was a case of intoxication. Since intoxication is due to secreted toxins, bacteria are not usually detected in blood or stool samples. MacConkey agar and sorbitol-MacConkey agar plates and xylose-lysine-deoxycholate (XLD) plates were inoculated with stool samples and did not reveal any unusually colored colonies, and no black colonies or white colonies were observed on XLD. All lactose fermenters on MacConkey agar also ferment sorbitol. These results ruled out common agents of food-borne illnesses: E. coli, Salmonella spp., and Shigella spp.

Figure 2. Gram-positive cocci in clusters. (credit: Centers for Disease Control and Prevention)

Analysis of the chicken salad revealed an abnormal number of gram-positive cocci arranged in clusters (Figure 2). A culture of the gram-positive cocci releases bubbles when mixed with hydrogen peroxide. The culture turned mannitol salt agar yellow after a 24-hour incubation.

All the tests point to Staphylococcus aureus as the organism that secreted the toxin. Samples from the salad showed the presence of gram-positive cocci bacteria in clusters. The colonies were positive for catalase. The bacteria grew on mannitol salt agar fermenting mannitol, as shown by the change to yellow of the medium. The pH indicator in mannitol salt agar is phenol red, which turns to yellow when the medium is acidified by the products of fermentation.

The toxin secreted by S. aureus is known to cause severe gastroenteritis. The organism was probably introduced into the salad during preparation by the food handler and multiplied while the salad was kept in the warm ambient temperature during the speeches.

  • What are some other factors that might have contributed to rapid growth of S. aureus in the chicken salad?
  • Why would S. aureus not be inhibited by the presence of salt in the chicken salad?

Key Concepts and Summary

  • Chemically defined media contain only chemically known components.
  • Selective media favor the growth of some microorganisms while inhibiting others.
  • Enriched media contain added essential nutrients a specific organism needs to grow
  • Differential media help distinguish bacteria by the color of the colonies or the change in the medium.

Multiple Choice

EMB agar is a medium used in the identification and isolation of pathogenic bacteria. It contains digested meat proteins as a source of organic nutrients. Two indicator dyes, eosin and methylene blue, inhibit the growth of gram-positive bacteria and distinguish between lactose fermenting and nonlactose fermenting organisms. Lactose fermenters form metallic green or deep purple colonies, whereas the nonlactose fermenters form completely colorless colonies. EMB agar is an example of which of the following?

  1. a selective medium only
  2. a differential medium only
  3. a selective medium and a chemically defined medium
  4. a selective medium, a differential medium, and a complex medium

Haemophilus influenzae must be grown on chocolate agar, which is blood agar treated with heat to release growth factors in the medium. H. influenzae is described as ________.

Fill in the Blank

Blood agar contains many unspecified nutrients, supports the growth of a large number of bacteria, and allows differentiation of bacteria according to hemolysis (breakdown of blood). The medium is ________ and ________.

Rogosa agar contains yeast extract. The pH is adjusted to 5.2 and discourages the growth of many microorganisms however, all the colonies look similar. The medium is ________ and ________.

Think about It

What is the major difference between an enrichment culture and a selective culture?

Critical Thinking

Haemophilus, influenzae grows best at 35–37 °C with

5% CO2 (or in a candle-jar) and requires hemin (X factor) and nicotinamide-adenine-dinucleotide (NAD, also known as V factor) for growth. [1] Using the vocabulary learned in this chapter, describe H. influenzae.

Methods Summary

The A. annua cytochrome b5 cDNA sequence was identified from a trichome expressed sequence tag library (NCBI accession 35608) by searching for sequence similarity to Crepis alpina cytochrome b5 type 11 (ref. 23). A cDNA encoding A. annua ADH1 was identified and the encoded protein characterized essentially as described for ALDH1 (ref. 11). Growth of strains, general genetic methodology and construction of synthetic genes were as described 3 . Yeast strains were derived from Y337 (ref. 3). Salient features of DNA constructs are as follows: (1) for expression of ERG9 from the CTR3 promoter the MET3 promoter was replaced with nucleotides −1 to −734 of the CTR3 promoter, integration being selected by d -serine 24 (erg9Δ::dsdA_PCTR3-ERG9) or nourseothricin 25 (erg9Δ::natA_PCTR3-ERG9) resistance (2) for reduced expression of cytochrome P450 reductase (CPR1), CPR1 was removed from plasmid pAM322 (ref. 3) by digestion and recircularization to generate pAM552, which expresses only ADS and CYP71AV1. A single copy of CPR1 was expressed from the GAL3 promoter (PGAL3) (nucleotides −1 to −660) integrated between GAL1 and GAL7 (gal1/10/7Δ::natA_PGAL3-CPR1-TCYC1, in which TCYC1denotes the CYC1 terminator) (3) a single integrated copy of A. annua cytochrome b5 was expressed from the GAL7 promoter (nucleotides −1 to −725 leu2::hisMXΔ::kanA_PGAL7-CYB5-TCYC1) (4) a single integrated copy of A. annua aldehyde dehydrogenase (ALDH1) was expressed from the GAL7 promoter, selecting for hygromycin B (ref. 25) (ndt80::hphA_PGAL7-ALDH1-TTDH1 and his3::hphA_PGAL7-ALDH1-TTDH1) (5) a single integrated copy of A. annua alcohol dehydrogenase (ADH1) was expressed from the GAL7 promoter, selecting for uracil prototrophy (natAΔ::URA3_PGAL7-ADH1-TTDH1 and gal80Δ::URA3_PGAL7-ADH1-TGAL80). Flask and fermentor culture conditions were essentially as described 3 . Fermentations requiring IPM contained 400 ml IPM added to 800 ml fermentor volume before inoculation. Artemisinic acid was purified from IPM by aqueous extraction at pH 10.7, followed by precipitation at pH 5.0. Assays for amorph-4,11-diene and artemisinic acid are essentially as described 3 . Artemisinic alcohol and artemisinic aldehyde were monitored by gas chromatography with flame-ionization detection.

Practical Work for Learning

Class practical

This activity involves skills for safe handling of microbial material. Students compare the activity of an enzyme extract with that of two enzyme-producing microbes and can consider the industrial importance of enzymes and microbes.

This practical is based on an investigation called Breakdown of protein by microbes (319 KB) published in Practical Microbiology for Secondary Schools © Society for General Microbiology.

Lesson organisation

Give each working group (1-2 students) a milk-agar plate. They will have to use sterile paper discs to collect samples of two microbes and one enzyme extract, all using sterile technique. Organise the work so that not all the groups want bottle A at the same time!

Written information about industrial processes using microbes and enzymes (particularly proteases) would be a useful resource for this lesson. See links at end.

There is scope here for developing skills of sterile technique to become the focus as well as exploring the enzyme activity.

Apparatus and Chemicals

For each group of students:

Marker pens to label the plates

Tape to close up the plates (Note 2)

For the class – set up by technician/ teacher:

Bacillus subtilis nutrient broth culture, 2 cm 3 in a bottle/ test tube

Saccharomyces cerevisiae malt extract broth culture, 2 cm 3 in a bottle/ test tube

Milk-agar plates, 2 per group (Note 1)

Virkon solution 1% w/v (see manufacturer’s instructions)

Bottles of sterile distilled water, 1 per 4 groups

Bottles of 0.1% trypsin solution, 1 per 4 groups

Health & Safety and Technical notes

Carry out a full risk assessment before planning any work in microbiology (Note 1 for more details).
Check the standard procedures for more details of Maintaining and preparing cultures, Aseptic techniques, Making up nutrient agars, Pouring an agar plate and Incubating and viewing plates safely.

1 Before embarking on any practical microbiological investigation carry out a full risk assessment. For detailed safety information on the use of micro-organisms in schools and colleges, refer to Basic Practical Microbiology – A Manual (BPM) which is available, free, from the Society for General Microbiology (email This email address is being protected from spambots. You need JavaScript enabled to view it. ) or go to the safety area of the SGM website ( or refer to the CLEAPSS Laboratory Handbook.

2 To make milk-agar, make up and sterilise nutrient agar. Allow to cool to 40-50 ºC and add pasteurised milk (10% by volume) aseptically and mix carefully. The milk should be pasteurised and freshly bought but can be skimmed, semi-skimmed or full-cream. See Standard procedures: Making up nutrient agars for more details.

3 Make up 0.1% trypsin solution. Refer to CLEAPSS Hazcard and Recipe card. Powdered enzyme is harmful, but solutions less than 1% are considered LOW HAZARD. Make up fresh for each day of use as it degrades over time.

4 Make your own assay discs by punching discs around 6mm diameter from filter paper, or chromatography paper using a cork borer or a hole punch. Wrap in aluminium foil, in sets of four, and sterilise by autoclaving.


SAFETY: Use sterile technique when handling microbial cultures. Incubate and view plates safely. Sterilise and/ or dispose of all contaminated material using appropriate methods.


a Inoculate nutrient broth using sterile technique from a clean culture of Bacillus subtilis at least 48 hours before use. Inoculate malt extract broth with Saccharomyces cerevisiae at the same time. (See Standard procedures: Maintaining and preparing cultures of bacteria and yeasts)

b Calculate the quantity required and prepare just enough milk-agar (Note 2) for the investigation (12-15 ml for normal depth in a 90 mm Petri dish).

c Prepare a suitable solution to disinfect the work area for the investigation and afterwards. Suitable disinfectants include sodium chlorate(I) (hypochlorite) at concentrations providing 1000 ppm available chlorine for general surface cleaning, or 2 500 ppm chlorine for discard pots, or VirKon (follow manufacturer’s instructions).

d Prepare 0.1% trypsin solution (Note 3).

e Sterilise distilled water in enough McCartney/ Universal bottles.

f Paper discs can be purchased (Whatman Antibiotic Assay discs) or you can make your own (Note 4).


a Keep one milk-agar plate unopened as a control.

b Provide each working group with one milk-agar plate.

c Students use a marker pen to mark four sections on the base. Label the sections A, B, C, and D. Write a key to record the treatment of each disc, for example


In response to the COVID-19 pandemic, the University of KwaZulu-Natal (South Africa) switched to on-line classes. I developed a series of computational exercises using the Jupyter notebook ( as a substitute for the molecular biology labs normally run during my course. Jupyter notebooks are becoming the de facto standard for scientific data analyses because they are widely available, open-source and can be annotated with explanatory text and images. 1, 2 Furthermore, as the code within the notebook is run as a series of “cells,” any errors in the code can be readily identified and corrected.

Demonstrating Cellular Respiration and Fermentation

Cellular respiration and fermentation are 2 of the most challenging concepts for introductory biology students, who may become so consumed by memorizing steps of the Krebs cycle and glycolysis that they lose sight of the big picture. The following demonstrations place aerobic cell respiration and fermentation firmly in grasp. First, students observe respiration in germinating seeds by detecting the carbon dioxide produced. Next, they observe the carbon dioxide gas produced by yeast fermentation.

Demonstrating Aerobic Cellular Respiration with Germinating Seeds

Have students observe the respiration of germinating seeds, using bromthymol blue as a pH indicator. You can keep this demonstration simple by observing pH changes in solution as seeds respire. Alternatively, you might add variables to the demonstration (e.g., compare the respiration rates of various plant seeds with the same total mass). Additionally, you can test whether photosynthesis is occurring by placing duplicate samples in the dark.


  • Hydrated Seeds
  • Sealable Bag
  • Bromthymol Blue, 0.04%
  • Sodium Hydroxide, 1%
  • Test Tubes
  • Rubber Stoppers
  • 250-mL Flask
  • Test Tube Rack
  • Distilled Water or Springwater
  • Paper Towel

Safety tip

Note: Wear gloves and safety goggles while handling the bromthymol blue solution and sodium hydroxide.


  1. Two days before the demonstration, rehydrate the seeds (we recommend wheat, barley, peas, or mung beans). Place the seeds in a cup or beaker. Use dechlorinated water to cover the seeds to a depth at least 3 times their height in the container to compensate for the expansion of the seeds as they swell. Allow the seeds to soak overnight.
  2. Pour off the remaining water and fold the seeds into a wet paper towel. Place the towel in a sealable bag. Close the bag and store the seeds in a dark place over a second night.
  3. Prepare the bromthymol blue solution by adding 1.5 mL (about 30 drops) of 0.04% bromthymol blue to 80 mL of springwater or distilled water. The prepared bromthymol blue solution should be green, but the shade of green may vary depending on the pH of your water source. A color change will easily be observed if the solution given to students is slightly basic. To create a slightly basic solution, add the sodium hydroxide dropwise to the bromthymol blue until the color changes from green to a deep blue (10 to 20 drops).
  4. Explain the use of bromthymol blue as a pH indicator to the class. The chemical bromthymol blue is an indicator that appears blue in an alkaline (base) solution and yellow in an acidic solution. Carbon dioxide that is added to the solution surrounding the seeds combines with water to form carbonic acid (H2CO3), turning the bromthymol blue to a yellow-green color. Remove the carbon dioxide from solution and the bromthymol blue will turn a deeper blue.
  5. Fill a test tube ¾ full with rehydrated seeds. Pour the bromthymol blue solution over the seeds until the tube is completely full. Seal the tube with a rubber stopper.
  6. Prepare a control tube with only bromthymol blue solution. Fill the tube until it is completely full, and seal it with a rubber stopper.
  7. Store the samples until the end of the class period or overnight and have students make observations regarding the color of the solution or any other changes they observe. (You may set up duplicate samples in the dark to rule out any possible effects of photosynthesis on the experiment.)

Class discussion

Have students consider the seeds&apos respiration. Oxygen is present in the test tube, and the carbon dioxide gas is a product of the process of aerobic cellular respiration. (To a much lesser extent and for a limited time, germinating seeds may also perform anaerobic respiration, which also produces carbon dioxide.) Discuss the purpose of the control tube with only bromthymol blue solution.

Students may think that the seeds are photosynthesizing. If chlorophyll and light are present, the seeds may be photosynthesizing. Students should know that plants&apos photosynthesis uses carbon dioxide and produces oxygen. While additional oxygen would not significantly change the pH, removal of carbon dioxide from solution would make the solution more basic and turn the bromthymol blue a deeper blue color.

Demonstrating Fermentation with Yeast

Students observe the carbon dioxide gas produced as yeast ferments sugar. You can simply demonstrate yeast fermentation of a 10% glucose solution, or you can introduce variables and have students compare the degree of fermentation (as indicated by the volume of carbon dioxide generated) occurring under each condition. Suggested conditions to vary include the concentration of glucose, the sugar type, the temperature of incubation, and the amount of light.


  • Sugar(s): Glucose, Sucrose, Lactose, and/or Sucralose (artificial sweetener)
  • Packet of Activated Dry Yeast
  • Water (distilled or spring)
  • Fermentation Tube(s) (a plastic graduated test tube and a larger glass test tube)
  • Beaker or Test Tube Rack
  • 2 100-mL Flasks
  • 100-mL Graduated Cylinder
  • Balance
  • Hot Plate


  1. Prepare the 10% sugar solution(s) in distilled water. Yeast can ferment glucose and sucrose but not lactose or sucralose (artificial sweetener).
  2. Prepare the yeast suspension immediately before class.
    1. Warm 70 mL springwater or distilled water to approximately 37° C and add 7 g of yeast (1 packet). If you are using a 15-mL conical tube for the small tube, this volume will be sufficient for 8 setups. Scale up or down depending on the desired volume.
    2. Activate the yeast by swirling it for 1 to 2 min. Make sure the yeast distributes evenly in suspension. The presence of clumps leads to inconsistent results. Start a fresh yeast suspension for each class period as the rate of fermentation diminishes over time.

    Class discussion

    The volume of carbon dioxide released is proportional to the quantity of sugar fermented and is therefore a measure of fermentation. Fermentation results from the action of enzymes. In yeast cells, enzymes convert sugars into carbon dioxide and ethanol. A yeast can ferment a particular type of sugar only if the yeast cell contains the proper enzyme(s) to break down that molecule. Baker&aposs yeast contains the enzymes to break down glucose and sucrose but not lactose or sucralose.

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