4-5 A dilution plating protocol

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Before beginning the dilution plating exercise, it is necessary for you to learn how to use micropipettors. We will be using them in this experiment and throughout the semester. Each micropipette costs $200 so please treat them with extreme care. Always adhere to the following rules when using micropipettors:

  • Never set the pipettor to above the upper limit or below the lower limit. Figure 4-19 lists the limits for the micropipettes that we use in this course
  • Figure 4-19 Limits of micropipettes

    MicropipetteVolume Range (µl)
    p200.5-20
    p20020-200
    p1000200-1000

    The limits of the micropipettes. Never go above or below these ranges as it may damage the micropipette.

  • Never point a pipettor up. This can cause liquid to run down into the pipettor and destroy it's parts.
  • When withdrawing liquids with the pipettor, always release the plunger slowly. This prevents liquid from rushing into the end of the pipette and clogging it up and is especially important with large volume pipettors (200-1000 µl).
  • Be sure you use the proper size tip for each pipettor.
  • Always use a new tip for each different liquid.
  • Use the correct pipettor for the volume that is to be dispensed. Never use the 200-1000 µl pipette to dispense volumes below 200 µl.

Protocol for Experiment 2

Period 1

Materials

Test tube containing solution to pipette (practice solution)

4 1.5-ml microcentrifuge tubes

20-200 µl micropipette

200-1000 µl micropipette

Yellow pipette tips

Blue pipette tips

Microcentrifuge

  1. Remove four 1.5-ml microcentrifuge tubes from the beaker. Label each tube 1 through 4 on the frosted labeling spot on the side. Place each in a microcentrifuge rack.
  2. Add the volumes of water listed in Figure 4-20 to tubes 1 through 4.
  3. Figure 4-20 Volumes to add

    TubeMeasurement 1Measurement 2Measurement 3Measurement 4
    125354570
    2306085-
    3125175450250
    4750200-50

    Add these volumed to tubes 1 through 4 as listed.

  4. To add the first volume (Measurement 1), take the 20-200 µl micropipette and set the dial to 20 µl. Follow the protocol below to dispense the volume into the tube:
  5. While holding the pipettor, open the box of yellow tips and firmly press the end of the pipettor into a tip. Remove the pipettor from the box, the tip should come along for the ride. Close the pipette tip box.
    • Press the plunger down to the first stop and place the end of the pipettor tip into the practice solution. The end of the pipette tip should be below the surface of the liquid, but not touching the bottom of the tube. Slowly release the plunger until it stops and then remove the tip from the liquid dragging it along the side of the test tube as you do. This will remove any excess liquid clinging to the side of the pipette tip.
    • Take the tube you labeled 1 and hold it at eye level. Place the pipette tip against the side of the microcentrifuge tube and expel the liquid by pushing the plunger down until the first stop. As you drag the tip along the side, push the plunger to the second stop to blow out any remaining liquid. Remove the micropipette from the tube.
    • To remove the pipette tip, hold the pipettor over the tip discard tray and press the white tip eject button located near your thumb. This will eject the tip.
    • Repeat step 3 for each measurement that you add to the tubes. Continue until both tube 1 and tube 2 have been completed. Take your tubes and place them opposite one another in the microcentrifuge. It is very important that tubes are balanced in the microcentrifuge, so make sure you place directly across from one another. If you have questions, please ask your instructor. Centrifuge the tubes for 2-3 seconds to force all liquid to the bottom of the tubes. Remove the tube and return to your lab bench.
    • Since you added a total of 170 µl to tubes 1 and 2, set the 20-200 µl micropipettor for 170 µl and withdraw the tubes contents. If the tube volume exactly fills the micropipette tip, it is time to celebrate! You did it right!.
    • Perform the additions for tubes 3 and 4 in a similar manner. Use the 200-1000 µl pipettor to dispense the larger volumes when necessary. Next, mix each tube briefly on a vortex mixer, and pulse 2-3 seconds in a microcentrifuge. Since a total of 1000 µl (or 1 ml) was added to tubes 3 and 4, set the 200-1000 µl pipettor to 1000 µl and remove the contents of each tube to check the accuracy of your pipetting.
    • Feel free to practice more until you are comfortable with the micropipettes.

Figure 4-7 A drawing of the dilution

A schematic for the dilution plating to be performed. Measure pipetting volumes carefully and mix tubes thoroughly. Eacg agar plate is inoculated with 0.1 ml.

Period 2

Materials

A 1/10 dilution of hamburger

4 saline or 0.85% saline dilution blanks (9 ml)

4 plates of Plate Count Agar (PCA)

4 plates of MacConkey Agar (MAC)

Pipettors and sterile tips

  1. Label one plate of PCA and one plate of MAC for each of the following plated dilutions (as we defined them on page 121, q.v.): 10-3, 10-4, 10-5 and 10-6.
  2. 3. Label the four 9 ml dilution blanks with the dilutions to make of the hamburger as follows: 10-2, 10-3, 10-4 and 10-5.
  3. With the P1000 pipettor and a blue pipettor tip, aseptically transfer 1 ml of the 1/10 hamburger dilution to the dilution blank labeled 10-2; discard the tip into the disinfectant. (Alternatively, if the hamburger dilution is provided in a 1 ml amount, you can dump the contents of the dilu-tion blank into the hamburger tube.)
  4. Mix this dilution well by holding the tube on the Vortex mixer* for about 5-10 seconds as demonstrated by the instructor; an actual vortex must be achieved for proper mixing. Note that this tube is a 1/100 (i.e., 10-2) dilution of the original, undiluted hamburger.
  5. Referring to the diagram on the next page, continue making serial dilutions of the hamburger, using a new pipettor tip for each new dilution. (Be sure you are discarding the tips into disinfec-tant!) Thus, by making 1 ml inoculations into the 9 ml dilution blanks, we will achieve the ham-burger dilutions which you have labeled on the tubes.
  6. With the P200 pipettor and a yellow tip, aseptically transfer 0.1 ml of the 10-2 dilution onto each of the plates marked 10-3. (Why are these plates marked 10-3 and not 10-2?) With new tips, continue in like manner to inoculate the remaining plates.
  7. If such a device is not available, hold the tube between thumb and forefinger in one hand and flick the bottom of the tube with the fore-finger of the other hand in order to achieve a vortex. (Do not simply roll the tube between the hands or lightly tap the tube. Don't be too gentle, yet don't shake the tubes end-to-end!)

  8. Perform the following with care! (See page 118 including the safety precaution.)
    • Sterilize a hockey stick according to the directions near the bottom of page 118, making sure that the glass hockey stick has cooled sufficiently before immersing it into the ethanol.
    • Spread the inoculum over the entire surface of the 10-6 plates. Proceed to do the same with the remaining plates, moving from the more dilute to the more concentrated inocula. During the course of spreading the plates, the hockey stick need not be re-flamed, as long as you proceed from the more dilute to the more concentrated inocula. When all plates have been spread, resterilize the hockey stick and let it cool before returning it to the drawer.
  9. Incubate the plates (inverted!) and incubate at 30°C for 2 days. If the next period is 3 or more days away, bring the plates to the tray on the stage for 2-day incubation (followed by refrigera-tion which will arrest growth and development of the colonies, preventing overgrowth).

Figure 4-21 depicts a movie showing the dilution plating technique.

Period 3

Figure 4-8 Dilution plating results

Colonies growing on a plate of PCA agarColonies growing on a plate of MAC agar

Colonies growing on PCA agarColonies growing on MAC agar

Example results from a dilution plating. One gram of fresh hamburger hours was diluted as described in the procedure for this expeirment and then plated on MAC agar and PCA Row 1, MacConkey Agar; Row 2 PCA. If you are doing this virtually, you can count the colonies on the plate and determine the CFU per gram of hamburger. Record your values in your notebook.

  1. . Total Aerobic Plate Count: Choose one PCA plate which appears to have between 30 and 300 colonies on it and count the colonies on the plate. As in the example above, determine the number of colony-forming units per gram of the hamburger. Note that this method is technically neither total nor aerobic with regard to microorganisms. Not all microorganisms are able to grow on this medium even though it is termed an all-purpose medium. Aerobic refers to the incubation conditions, not the oxygen relationship of the organisms. Figure 4-8 shows a picture of typical results. The PCA plate has been diluted to 10-4. This is the total dilution. This would be an appropriate plate to count.
  2. . Total Gram-Negative Plate Count: (Note the selective and differential aspects of this medium as discussed on page 126.) Choose one MAC plate which appears to have between 30 and 300 colonies on it and count the colonies on the plate. Determine the number of gram-negative colony-forming units per gram of the hamburger. If distinctly red or pink colonies can be differentiated clearly, what appears to be the relative proportion of lactose fermenters? (E. coli is among the gram-negative organisms which can ferment lactose and thus produce red colonies on MacConkey Agar.) The MAC plate has been diluted to 10-3. This is the total dilution.
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