Yeast Plate Count Lab
Making a Serial Dilution

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Robert Koch, Single-Colony Isolation, and Koch’s Postulates

Robert Koch, a German physician, is famous for determining the bacteria responsible for anthrax and tuberculosis. The chance observation of bacteria growing on the surface of a spoiling slice of boiled potato led Koch to realize that each spot, or colony, of bacteria had grown as a clone from a single contaminating bacterium that had previously landed on the potato. From this realization, Koch developed the technique called single colony isolation, in which a sample of bacteria from one colony may be used to inoculate a culture medium and from that, grow a pure culture of that species of bacterium.

In his work with bacteria, Koch also developed a series of methods for determining which species of bacterium is responsible for a given disease. In his honor, these four steps are collectively known as Koch’s Postulates. These say that to prove that a given bacterium causes a certain disease, the researcher must:

1. find the same bacterium species in all diseased individuals investigated,
2. isolate the bacterium from diseased individuals and grow it in a pure culture,
3. induce the disease in experimental animals by transferring bacteria from the culture, and
4. isolate the same bacterium from the animal after the disease develops.

Koch and other researchers at that time were also looking for a suitable medium upon which to grow bacteria for study. Gelatin was ruled out as a solidifying agent because (as any picnic-goer knows) a mixture of gelatin and water is not solid at body temperature (37° C). Also, because it is a polypeptide, it is digested by a number of species of bacteria, thereby losing its gelling ability. A housewife friend suggested that Koch try agar, noting that many cooks used it to solidify desserts, etc. in place of gelatin. Koch subsequently developed the use of agar, a sulfuric ester of a polysaccharide (poly = many, sacchar = sugar) complex derived from certain red algae (Japanese Isinglass, Gelidium spp., is the best, but also derived from other genera — obtained by boiling the seaweed for 6 hr in dilute H₂SO₄), to solidify nutrient liquid media. This allowed the growth on the surface of the medium of a specimen spread across it, thus isolating the various microorganisms (micro = small) which might be present. Agar was found to be ideal as a gelling agent — it melts/dissolves only when the media are heated to near boiling, will remain melted until cooled to around 40° C, and forms a gel that stays solid at body temperature. Because it is a complex polysaccharide, it is not degraded by the vast majority of bacteria, thus the medium remains solid and does not liquify as the bacteria grow in/on it, and therefore allows strict control of growth factors which may be limiting.

We Will Use Yeast

In this experiment, we will be using yeast (a fungus and something which is a lot safer for students who are just learning sterile techniques to handle) rather than bacteria to learn some of the techniques and methods used in microbiology. We will be making use of Koch’s conclusion that each viable cell (in this case yeast) that happens to land on the medium can/will grow into a small colony that we can see and count. We will be inoculating 4% glucose medium with an unknown number of yeast cells, and like Koch’s conclusion that each bacterial colony on the potato came from a single, live bacterium, we will assume that each yeast colony we find growing on our agar plates came from a single, live yeast (a colony-forming unit or CFU). We will use this information to calculate the average number of yeast cells in a packet of yeast.

However, there are too many yeast cells in a packet to count them all, so we must dilute them. If we keep track of how much they were diluted, we can use the number of colonies growing on our plates to “work backwards” to calculate the number of cells in a yeast packet. Because the yeast must be very dilute to get countable results, we need to do a serial dilution and to spend time discussing the math involved in figuring out the number of cells per packet.

Experiments by Dr. Fankhauser’s Microbiology students have shown that yeast, Saccharomyces cerevisiae (myce = fungus, Ceres = goddess of grain, visi = look or see), grows optimally on a nutrient agar supplemented with 4% glucose (gluco = sweet, ose = sugar or carbohydrate). Remember that glucose and dextrose (dextro = right) are just two names for the same thing. This medium contains nutrient broth which consists of 0.3% beef extract, and 0.5% of a pepsin digest of beef (peptone). It thus contains a broad variety of amino acids and vitamins providing a suitable medium for a wide variety of non-fastidious microorganisms (fastidious ones have very complex, specific nutritional requirements). The 4% glucose content especially encourages the growth of yeast with its ability to ferment at high rates. Thus, that is the medium we will be making for our yeast.

First Day — Preparation of Medium

For this lab, work in groups. A batch/bottle of medium will make about 20 to 25 agar plates. Thus, making two batches/bottles per lab section of 15 to 20 people should suffice.

If not already present, assemble the necessary equipment and ingredients.

• 6 g nutrient broth powder
• 9 g agar-agar
• 24 g glucose (dextrose)
• (3 g NaCl, opt.)
• 600 mL dH₂O
• balance
• spoons or spatulas
• 1000 mL beaker
• 1-L bottle with cap
• funnel
• heat source (microwave or pot of boiling water on the stove)
• hot pads
• (spirit thermometer, 10 to 110° C, opt.)
• autoclave
• 16 to 20 sterile petri dishes
• incubator

Weigh out the dry ingredients.

1. Weigh the 1000-mL beaker, and record its apparent weight. If it’s too heavy for the balance, counterbalance it with an equal-sized beaker on the other balance pan.
2. In your lab notebook, add the required mass of the first reagent to the apparent weight of the beaker, and then set the balance to read that weight.
3. Add that dry reagent to the beaker with care until equal balance swings are achieved. Gently tap on the edge of the beaker with the spatula/spoon to better judge how “close” you are. If you go slightly over the required amount, do not remove any excess back to the original container. If/when you are weighing the nutrient broth powder, replace the cover immediately on the nutrient broth container because it is hygroscopic (hygro = moist or wet, scope = see or watch or look; it absorbs moisture from the air and “turns into a rock,” like brown sugar does in the summer). Move the balance weights to find the actual weight added and record that in your lab notebook.
4. For each of the other dry ingredients, again, as in the last two steps, add the new balance reading plus the required mass of the next reagent, then set the balance to read that weight. Weigh that ingredient into the same beaker along with the the previous ingredient(s), which does necessitate careful attention as you reach each “end point” because you cannot remove excess if you accidentally add too much.

Mix and autoclave the medium.

1. Add dH₂O to the beaker while stirring, and q.s. to 600 mL. Continue stirring until there are no lumps and the dry ingredients are thoroughly suspended/dissolved.
2. Use a funnel to transfer the medium to a 1-L bottle. Note that, at this point, the agar is not melted/dissolved in the medium and will settle out if given a chance. However, it is important to get it all as you transfer the medium to the bottle. Thus, it is necessary to swirl/stir the medium as it is being poured to help insure that all the agar gets poured into the bottle along with the other ingredients.
3. Heat the medium to melt/dissolve the agar. There are several ways of doing this – pick one. The “traditional” method is to place the bottle in a pot of boiling water on the stove, and while stirring with a thermometer (thermo = heat, meter = to measure), heat to boiling, but do not allow it to boil over, nor to burn on the bottom. A more “risky” method is to leave the mixture in the 1000-mL beaker, and set that directly on the stove or over a Bunsen burner to heat, while stirring with a thermometer (then, pour into the bottle after it’s heated). A “quick, modern” method is to microwave the medium: heat 2 min., swirl, heat 2 min, swirl, heat 1 min. No matter which method you use, have hot pads available, and be prepared to quickly turn off the heat source and then remove the bottle, if needed, to prevent it from boiling over.
4. Cap the bottle loosely, (if needed – if the medium will be stored in the bottle, label the bottle with the group name and date and/or apply autoclave tape to the lid), and place in the autoclave. When both groups have put their bottles in the autoclave, autoclave at 15 lbs. pressure for 15 min. (refer to autoclave protocol for how-to). Although set for 15 min, this process will take around 45 min to complete.
5. When the autoclave is done and the pressure is almost back down to zero, optionally use the stove to heat a pot of water to between 50 to 60° C. This will be used to cool the autoclaved bottles. If it’s cooler than 50° C, you risk having the bottles crack, and if it’s warmer than 60° C, it won’t cool the bottles as effectively. However, for the safety of the bottles, closer to 60° is better than closer to 50°.
6. When the autoclave pressure has returned to zero, carefully open the autoclave. Using hot pads, remove the bottle(s) and let them cool at room temperature for at least several minutes. After that, to speed the cooling process, the bottles may carefully be put into the warm water. Ideally, the bottles should be cooled enough that they feel “hot” but are just cool enough to be hand-held.

Sterilize a work area and pour the plates.

1. On one of the lab tables, attach Bunsen burners to the desk stopcocks. Obtain a striker, a squirt bottle of 70% ethanol, a scissors and a sealed bag of plastic petri dishes for each bottle of medium that has been made. Use the alcohol and a paper towel to “sterilize” an area of the desktop. Then, working on the cleaned area of the table, cut the top off the bag of petri dishes, invert the bagged dishes, and slowly and carefully, slide the bag up, off the petri dishes (save the bag for future use). After the petri dishes are out of the bag, they may be re-stacked in convenient-height stacks of about 3 or 4 dishes each, cover-side up. Remember, these petri dishes are sterile inside, and need to remain sterile inside. Do not open them up.
2. Working on the sterile field, each student should pour a stack of two to three plates using sterile technique as demonstrated by your instructor (your group/class should use up all the medium in your bottle — a total of 20 to 25 plates). To make it easier to pour the plates, stacks of 3 or 4 sterile plates should be positioned near the edge of the desk. Remove the cap from the bottle and hold it with the little finger of the hand that’s holding the bottle (do NOT set it down on the desk). Flame the lip of bottle to sterilize it.
3. Starting with the bottom plate in your stack, use your other hand to lift that plate’s lid and the rest of the stack of plates above it straight up. Do not set these down on the table, and do not hold them to the side of the plate that’s being filled. Rather, hold them straight above the plate being filled as a “shield” to help protect that plate from airborne bacteria falling into it. Fill each plate about half full.
4. After all the plates in that stack have been poured, flame the lip of the bottle, and loosely replace the cap (until the next person pours his/her plates). Carefully slide your stack of plates to an open area of the desktop and to help them cool more quickly, carefully unstack them so each plate is resting directly on the desktop. If all the plates in your bag have been used up, and there is still some medium in the bottle, look for plates that are a bit “low” — that contain a bit less medium than the others — and top them off.
5. When all the plates have been poured and all the medium in the bottle has been used up, extinguish the Bunsen burner and immediately rinse the bottle with hot water before any left-over medium in it has a chance to gel. Any sterile, unused plates may carefully be returned to their plastic bag.
6. When the plates have cooled and solidified, invert them, place them in a dish tub (or other, similar-sized container), tape a label on the tub/container with the lab section, instructor’s name, and date, then place into the 37° C incubator. Incubate the plates for 48 hr to check for sterility.

Second Day — Serial Dilution and Inoculation of Plates

If not already present, assemble the necessary equipment and ingredients.

• per student:
  • test tube rack to hold 16×150-mm (large) test tubes (from under the sink)
• on first pair of lab tables (repipet stations):
  • rack with capped, sterile, 16×150-mm (large) test tubes (at least 3 tubes per student)
  • 2 wax pencils
  • 2 repipets with sterile dH₂O, set for 9.9 mL
  • 2 Bunsen burners
  • at least 1 striker
  • squirt bottle of 70% EtOH
• on second pair of lab tables (serial dilution stations):
  • 250-mL beaker containing 100 mL dH₂O, on magnetic stirrer (note: need one 100 mL of water per class, so the other may, temporarily, be empty)
  • unopened packet of fresh bakers’ yeast (1 per class)
  • canisters of sterile 0.1-mL pipets (at least 3 pipets per student)
  • at least 2 pipet bulbs or pipet fillers
  • 2 Bunsen burners
  • at least 1 striker
  • used pipet container
  • vortex
  • squirt bottle of 70% EtOH
• on third pair of lab tables (spectrophotometer stations):
  • spectrophotometer set at 660 nm
  • lens paper
  • set of 2 clean, matched cuvettes (one containing dH₂O) in plastic test tube rack (“small” test-tube size)
• on fourth pair of lab tables (plating-out stations):
  • 2 turntables
  • 2 spreaders, each in a beaker with 95% EtOH
  • canisters of sterile 1.0-mL pipets (at least 1 per student)
  • at least 2 pipet bulbs or pipet fillers
  • 2 Bunsen burners
  • at least 1 striker
  • used pipet container
  • 2 wax pencils
  • squirt bottle of 70% EtOH
• on fifth pair of lab tables:
  • sterile 4%-glucose nutrient agar plates made last lab period
  • plastic “dish” tub into which to put inoculated plates
• other:
• incubator set for 37° C

Get ready to do the lab.

1. At the second station, your instructor will suspend the contents of a package of yeast in 100 mL of water. This willbe mixed thoroughly (using a magnetic stirrer) for at least 5 to 10 min. If stations are set up on both sides of the room, your instructor may divide the yeast suspension between two beakers and place one on each magnetic stirrer.
2. If you haven’t already done so, obtain a “big” test tube rack from under the sink.

At the first table (repipet station):

1. Obtain three, sterile, capped, 16×150-mm test tubes and use the wax pencil to label them “2,” “4,” and “6” to represent the three serial dilutions (10^–2, 10^–4, and 10^–6) you will be making.
2. Check to insure that the repipet is set for 9.9 mL.
3. Obtain or share one paper towel. Squirt some of the 70% EtOH onto the table top, and wipe to sterilize it.
4. As demonstrated by your instructor, remove the lid from a test tube and hold it with your little finger — keep the cap off the tube the minimum time necessary and do not set it down. Flame the lip of the tube. Raise the plunger on a repipet to fill it, then push down to deliver 9.9 mL of sterile dH₂O into that tube using a repipet.
5. Again, flame the lip of the test tube, and replace the lid.
6. Repeat for your other two tubes.

At the second table (serial dilution station):

1. Begin with a sterile field by using a paper towel (obtain or share) to wipe the desktop with 70% EtOH (95% will dehydrate the bacteria rather than being absorbed to kill them). It is necessary to do this once at the beginning of the lab, but use your judgement if you think it needs to be done again (if someone leans on the table). Squirt some of the 70% EtOH onto the table top, and wipe to sterilize it.
2. Make sure you understand how the pipet filler works and how to use it before you start working with sterile equipment.
3. Perform a 10⁶ serial dilution of the yeast suspension as follows. So that you get “good” results and your plates don’t get contaminated, it is very important to not rush through this, but rather, to concentrate on what you are doing. This procedure is extremely important in microbiological labs, and is one of the crucial techniques in aseptic (a = not or without, septi = rotten or putrid) technique. While the steps may seem overly detailed in the following narrative, care in learning proper technique at the beginning establishes good technique for the rest of your life. Compare these detailed steps with the demonstration given by your instructor. Patience pays off. Go slowly at the beginning, and verbally (not physically) assist your fellow students as they work through the steps.
4. Do the first dilution:
   1. So that your dilutions don’t become “contaminated” by droplets of more-concentrated yeast solution, you must use a fresh 0.1-mL pipet each time.
   2. Out of the 100 mL of yeast suspension that was made, you will use 0.10 mL to do the serial dilution. Note that this 0.10 mL contains 1/1000 or 10³ of the yeast in the original packet (0.10/100 = 1/1000).
   3. With your rack of sterile test tubes right there and “ready-to-go,” obtain a sterile, 0.10-mL pipet from the canister. Only touch the pipet you are withdrawing, and only touch it by the top end — it is sterile, and touching it will transfer bacteria from your hands and make it non-sterile.
   5. Pass the tip end of the pipet through the Bunsen-burner flame to make sure it is sterile. Fit the pipet into the end of a pipet filler.
   6. Using your writing hand, use the pipet and pipet filler to obtain 0.10 mL of the yeast suspension from the beaker on the magnetic stirrer. Use caution because the 0.01-mL pipets are “tiny” and will fill up a lot more quickly than the 5.0-mL and 1.0-mL pipets you’ve used before. Then, tilt the pipet slightly horizontally so that fluid moves up slightly into the pipet and doesn’t drip during transfer.
   7. Without delay, and without touching anything with the pipet, pick up the first (“2”) test tube with your non-writing hand, grip the cap with the little finger of the pipet hand and gently remove with twisting-pulling motion. Hold the cap in that little finger and do not lay it down.
   8. Flame the lip of the test tube, then if needed, set the test tube back in the rack. Deliver the 0.10 mL of yeast suspension into the test tube, remembering to puff out the last drop.
   9. If needed, pick up the test tube. Reflame the lip of the test tube. Replace the cap and return the test tube to the rack. Do this, first, as soon as possible, before dealing with the pipet, etc.
   10. Remove the pipet from the pipet filler, and place it in one of the used-pipet containers.
   11. Mix the contents of the test tube well with a vortex. Note that this means you have used that 0.1 mL of solution to make 10 mL of solution — 100 times as much, thus the concentration is 1/100 (10^–2) as strong as it was before.
5. Do the second dilution:
   1. Obtain a new, sterile 0.10-mL pipet. As before, pass the tip end of the pipet through the Bunsen-burner flame to make sure it is sterile. Fit the pipet into the end of a pipet filler.
   2. Without delay, and without touching anything with the pipet, pick up the first (“2”) test tube with your non-writing hand, grip the cap with the little finger of the pipet hand and gently remove with twisting-pulling motion. Hold the cap in that little finger and do not lay it down.
   3. Flame the lip of the test tube, then if needed, set the test tube back in the rack. Use the pipet and pipet filler to obtain 0.10 mL of the yeast suspension from the first test tube. Use caution because the this solution is more clear than the previous one, and is very difficult to see as it rises (quickly!) in the pipet.
   4. If needed, pick up the test tube. Reflame the lip of the test tube. Replace the cap and return the test tube to the rack.
   5. Without delay, and without touching anything with the pipet, pick up the second (“4”) test tube, remove the cap, and flame the lip of that test tube, then if needed, set the test tube back in the rack. Deliver the 0.10 mL of yeast suspension into the test tube.
   6. Reflame the lip of the second test tube. Replace the cap and return the test tube to the rack. Then, remove the pipet from the pipet filler, place it in one of the used-pipet containers, and mix the second test tube with the vortex.
   7. You now have a solution that is 100 × 100 = 10,000 (10⁴) times as dilute. Note that the 0.10 mL used to make this dilution contains 10^–3 × 10² = 10⁵ of the yeast in the original packet.
6. Do the third dilution:
   1. Again using sterile technique and a new, sterile pipet, use this same procedure to transfer 0.10 mL from the second test tube (“4”) to the third (“6”) one. Remember to mix this tube with the vortex, too.
   2. Notice that you now have a solution that is 100 × 10,000 = 1,000,000 (10² × 10⁴ = 10⁶) times as dilute. Also, the 0.10 mL used to make this dilution contains 10^–5 × 10^–2 = 10^–7 of the yeast in the original packet.

At the third table (spectrophotometer station):

1. Check to make sure the wavelength of the spectrophotometer is set to 660 nm. Use the cuvette of dH₂O to blank the spectrophotometer.
2. In the other cuvette, pour some of the contents of your first (“2”) tube only (assuming you did your dilutions correctly, you’re done with that tube, now). Note that this solution is no longer sterile, so make sure you do not mistakenly use the solution in your third tube!
3. Use the spectrophotometer to determine the A₆₆₀ of the 10² dilution and record that number in your lab notebook. Since everyone is working from the same initial yeast solution and everyone is doing the same dilutions, that means everyone’s absorbance readings should be about the same, so this step is just to double-check.

At the fourth table (plating-out station):

1. As before, obtain or share one paper towel. Squirt some of the 70% EtOH onto the table top, and wipe to sterilize it.
2. Obtain two, sterile, 4%-glucose nutrient-agar plates from the fifth table. Pay attention to what you’re doing, and make sure you take your plates from the stack of sterile plates, and not some that someone else has already inoculated. Hold your chosen plates up to the light and examine them to verify that they are sterile and nothing is growing on them.
3. Label the bottom of each of your plates in small letters (you’ll need to be able to see through the plate) with the date, your initials, and aliquot size (0.2 or 0.1 mL).
4. Use a 1.0-mL pipet to plate out 0.10 and 0.20 mL of the 10⁶ dilution onto the corresponding plates.
1. Obtain a sterile 1.0-mL pipet from the canister. As above, flame the pipet and fit it into a pipet filler.
2. Again using your little finger to hold the lid, and without touching the pipet to anything, remove the lid of your third (“6”) test tube, and flame the mouth of the test tube.
3. Draw at least 0.3 mL of liquid into the pipet, and adjust the level to a convenient 0.10-mL marking.
4. Reflame the mouth of the test tube and replace its cap.
5. Working quickly and accurately (so all of your sample doesn’t end up in one spot on the plates), from that pipetful of liquid, release 0.1 mL into the “0.1 mL” plate and 0.2 mL into the “0.2 mL” plate.
5. Place the pipet into the used-pipet receptacle. Place the 0.1 mL plate onto the turntable. You want to do that one first since it contains less liquid, and therefore is slightly more likely to have all the liquid absorb into one spot unless you work quickly.
6. Obtain a spreader and shake off the excess EtOH. When it is no longer dripping, pass the spreader briefly through the flame of a Bunsen burner to ignite the rest. DO NOT HOLD A FLAMING SPREADER OVER THE BEAKER OF EtOH!
7. When the flaming has stopped, slightly lift the lid of the 0.1-mL petri dish and touch the spreader to the inside of the top of the petri dish to cool it. Hold the lid of the petri dish over the dish to shield the dish from possible airborne contamination, and with that same hand, slowly turn the turntable. With your other hand, hold the spreader so it skims across the surface of the agar to spread your sample. Do not press down on the spreader or you might damage agar surface. Again, during this process, hold the petri dish lid above the plate to reduce airborne fallout contamination. When the fluid has been absorbed, the spreader will drag with a bit more difficulty, so you know that sample is “done.”
8. Place the spreader back into the 95% EtOH.
9. Remove the 0.1-mL plate from the turntable and place the 0.2-mL plate on the turntable.
10. As above, flame the spreader, then spread the second plate.
11. Once both of your plates are “done,” invert both plates (agar side up), and place in the designated container to be incubated at 37° C for 48 hr (2 days). The plates should be stored agar-side-up in the incubator so any condensation that is formed will be absorbed back into the agar rather than dripping onto the plate and “messing up” your colonies.
12. Note that since you are using only 0.1 mL (or 0.2 mL) out of the 10 mL in the last dilution, the 0.1 mL spread on the first plate contains 10^–7 × 10^–2 = 10^–9 of the yeast in the original packet (and the 0.2-mL sample, thus, contains twice as much, or 2 × 10^–9 of the yeast from the packet).

Third Day — Counting Yeast Colonies

If not already present, assemble the necessary equipment and ingredients.

• petri plates from last period
• wax pencil and/or clicking counter
• opt. colony counter or other light source

Count the number of colonies on each of your plates. In theory, the 0.20-mL plate should have approximately twice the colonies as the 0.10-mL plate. Record these numbers in your lab notebook.

Assuming that one yeast cell can grow to form a colony of yeast as it grows and divides (it is a colony-forming unit), calculate the number of colony-forming units (CFU) in the original yeast package Note that the 0.1-mL plate is 10^–9 as concentrated as the original package. The 0.2-mL plate is 2 × 10^–9 as concentrated as the original package. Thus, the package of yeast would contain 10⁹ × [number of colonies on the 0.1-mL plate] or 5 × 10⁸ × [number of colonies on the 0.2-mL plate] CFUs (actual yeast cells).

Once you have gathered all your data and completed all your calculations, submit your data online. Once everyone has submitted their data, you may print out a copy of the class data for your lab notebook.

Dr. Fankhauser’s Dilution Practice Problems

In your lab notebook, do the dilution practice problems which follow.

Because solutions in science are often much more concentrated than are desired or can be managed for a given protocol, it is frequently necessary to dilute these solutions. This requires a working knowledge of the principles of diluting, dilution factors, concentration factors and the calculations involved. High dilutions are usually expressed exponentially.

First, some definitions:

Aliquot:

a measured sub-volume of sample

Diluent:

material with which the sample is diluted

Dilution factor:

ratio of final volume (aliquot plus diluent volume) divided by the aliquot volume

Concentration factor:

ratio of aliquot volume divided by the final volume

Example: You make a dilution by adding 0.1 mL specimen to 9.9 mL of diluent which gives a final volume of 10 mL:

• Dilution Factor = final volume/aliquot volume = (0.1 + 9.9)/0.1 = 1 to 100, 1:100 or 10²
• Concentration Factor = aliquot volume/final volume = 0.1/(0.1 + 9.9) = 0.01 or 10^–2

To prepare a desired volume of solution of a given dilution:

1. Calculate the volume of the aliquot:
   

   aliquot volume = concentration factor × final volume

2. Calculate the volume of the diluent:
   

   volume of diluent = (final volume - sample aliquot volume)

3. Measure out the correct volume of diluent, add the correct volume of aliquot to it, mix.

SAMPLE PROBLEMS:

1. How much sample is required to prepare 10 mL of a 1 to 10 dilution, and how much diluent would you need?
2. What is the dilution factor when 0.2 mL is added to 3.8 mL diluent? What is the concentration factor?
3. What should the aliquot and diluent volumes be to prepare 5 mL of a 10² dilution?
4. You have 0.6 mL of sample, and want to dilute it to a fiftieth of its present concentration. How much diluent will you add, and what will the final volume be?
5. How would you prepare 20 mL of a 1:400 dilution?
6. What is the dilution factor when you add 2 mL sample to 8 mL diluent?
7. You want 1 L of 0.1 M NaCl, and you have 4 M stock solution. How much of the 4 M solution and how much dH₂O will you measure out for this dilution?
8. You add a pint of STP gas treatment to a 12-gal. fuel tank, and fill it up with gas. What is the dilution factor?
9. You diluted a bacterial culture 10⁶, and plated out 0.2 mL and got 45 colonies on the plate. What was the concentration of bacteria in the original undiluted culture?
10. A hard one: You have 100.0 mL of dH₂O. How much glycerine would you have to add in order to make a 2.00% v/v (volume per volume) solution of the glycerine? (Hint: it requires a little algebra.)
11. Here’s another “English system” one (for people who aren’t interested in cars and STP?): if you are making homemade ice cream, and put 1’tsp of vanilla in a 1-gal. batch of ice cream mix, what is the dilution factor?

(The answers are on the data-submission Web page.)

Other Things to Include in Your Notebook

Make sure you have all of the following in your lab notebook:

• all handout pages, including the dilution practice problems (in notebook or separate protocol book)
• all notes you take during the introductory mini-lecture
• all notes and data you gather as you perform the experiment
• all requested calculations based on those data
• print-out of class data (available online)
• drawings (yours!) of all “new” equipment used, including:
  • autoclave (also cross-referenced to autoclave protocol)
  • bottle used to autoclave the agar
  • petri dish
  • repipet (detail of scale set for 9.9 mL)
  • magnetic stirrer/hot plate
  • stir bar
  • 16×150-mm test tube(s) with cap(s)
  • 0.1-mL pipet
  • turntable
  • spreader
  • clicking counter
  • colony counter (detail of grid)
  • incubator (including controls)
• drawing or xerox of your “finished” plate the day we count the colonies
• optional sample of yeast packet
• answers to all discussion questions, a summary/conclusion in your own words, and any suggestions you may have
• evidence that you have at least tried to work the practice problems (not just answers, but actually showing your work)
• any returned, graded pop quiz