Peggy Campbell
1993 Woodrow Wilson Biology Institute


The purpose of this series of lab activities is to demonstrate both phenotypic and genotypic changes in bacteria that have been transformed with a gene that codes for antibiotic resistance.

In the first activity, students use a rapid method to render the bacterium Escherichia coli "competent" to uptake plasmid DNA. Escherichia coli cells are then transformed with the pAMP plasmid, which carries the gene coding for resistance to ampicillin. The recipient E. coli strain, MM294, lacks this gene; thus only transformed cells acquire the ability to grow on ampicillin. Ampicillin then is the selective agent in this experiment. E. coli cells are scraped from two large colony starter plates and suspended in two tubes containing a solution of calcium chloride. pAMP plasmid is added to ONE cell suspension, and both tubes are incubated at 0 deg C for 15 minutes. Following a brief heat shock at 42deg C, cooling, and the addition of LB broth, samples of the cell suspension are plated on two types of media: plain LB agar and LB agar with ampicillin. The plates are incubated for 12-24 hours at 37deg C then checked for bacterial growth. Only cells that have been transformed by taking up the plasmid DNA with the ampicillin-resistant gene will grow on the LB/amp agar plates. Students will also have growth on the plain LB plates by the untransformed cells (controls). This portion of the lab demonstrates the phenotypic changes associated with transformation.

In the second activity, students will perform a cell resuspension from the transformed and untransformed colonies, followed by a plasmid minipreparation on those cells, to enable them to demonstrate that only transformed cells have the plasmid present.

In the final activity, the students will complete a restriction digest and gel electrophoresis which will allow them to compare the untransformed with the transformed E. coli cells (containing the pAMP gene).


2nd year biology students


Prelab....1 or 2
Colony transformation....1 or 2
overnight suspension culture....1
plasmid miniprep....1
Gel electrophoresis....1
  • NOTE: If this is the first time you have attempted the rapid colony transformation, you may wish to use a kit, which contains not only the lab procedures and materials, but a complete teacher guide as well. When you have done it once with your students, you will be comfortable ordering the materials separately, and your cost will decrease dramatically. Good sources for kits are Carolina Biological Supply and Edvotek.

  • All materials are from Carolina Biological Supply unless otherwise noted.


    PART I: Rapid Colony Transformation

    A. Materials

    (for 24 students, divided into 8 groups of 3 students)
  • 2 E. coli strain MM294 starter cultures
  • 100 ul of pAMP (0.005 ug/ul) (#21-1438, $3.25)
  • 1 bottle LB agar (#21-6620, $17.25)
  • 1 bottle LB agar + amp (#21-6621, $18.50) (both these agars are sterile and ready to pour)
  • 32 plates (#21-4826, $6.00 for 20 plates)
  • 8 inoculating loops (#21-5826, 12 for $12.08)
  • 64 1ml sterile transfer pipettes (#73-6040, $76.34 for 200)
  • 24 15 ml culture tubes (#21-5078, $7.40 for 25)
  • 5 ml of 50 mM CaCl2
  • 8 beakers of ice 8 cell spreaders (#21-5820, $1.82 ea.)
  • parafilm or plastic wrap
  • 8 test tube racks
  • 8 lab markers
  • 8 Bunsen burners
  • masking tape
  • 37 degree incubator (or the top of a refrigerator)
  • 42 degree water bath (or hot water from the tap)
  • microwave oven (or boiling water bath) for melting sterile agar
  • disposal bag
  • 10% household bleach for cleanup

    Follow sterile techniques as outlined in this module. Student culture plates and station set up. (1 hour)
    1. Use presterilized, ready-to-pour agar as a convenience. It needs only to be melted in a microwave (carefully!) or boiling water bath, cooled to approximately 60deg C, and poured into sterile culture plates.

    2. Label at least 16 plates "LB" and 16 plates "LB/amp" on the bottom with a permanent marker and date them. Pour the LB broth into the LB plates to a thickness of about a quarter of an inch each; repeat for the LB/amp plates. Work as quickly as possible, using as sterile a technique as possible. Close each plate as soon as you have poured the agar, and do not disturb them until they have cooled and solidified. Once they have cooled, turn them upside down (to avoid condensation falling on the agar) and store until needed.

    3. Set up the student work stations with the materials indicated above. You may choose to aliquot the amount of pAMP each group will use, or leave the solutions in the total amount indicated above to be shared by the students.

    4. After step 10 in the student instructions, you can have the students stop. Make sure to store their tubes in the cold (on ice in a refrigerator works fine) until the next step.

    5. Throughout these experiments, emphasize the importance of sterile technique.



    (for 24 students, 8 groups of 3) colony transformation plates and control plates from part I
  • 40 ml LB broth (#21-6660, $11.60 for 250ml)
  • 40 ml LB broth/amp (#21-6661, $12.25 for 250ml)
  • 10% bleach
  • 8 Bunsen burners
  • 8 pipette aids or bulbs w/10ml standard pipettes
  • Lysis buffer (sucrose, Triton X-100 (Sigma #X-100, 100ml @$11.00),
  • Tris-Cl, Na2 EDTA)
  • TE buffer
  • 8 50ml conical tubes (#21-5100, $12.19 for 25)
  • 8 permanent markers
  • 37deg C incubator
  • 16 1.5ml sterile/clear microfuge tubes (#21-5226, $31.10 for 1000) microcentrifuge (#21-4065, $450.00)

    Each student group should work with the following plates:
      i) LB agar plate with untransformed cells
      (positive control)

      ii) LB/amp agar plate with transformed cells



    (for 24 students; combine into a total of 1 to 8 groups, depending on supplies).

    Materials listed are per individual group

  • 1 gel box
  • 1 power supply
  • 20-30ml agarose (#21-7075, 5g for $7.00 )
  • 5 microfuge tubes
  • plasmid miniprep from untransformed colonies (no plasmid should be present)
  • plasmid miniprep from transformed colonies (plasmid should be present)
  • EcoRI restriction enzyme and buffer (New England Biolabs, 1-800-NEB-LABS, 10,000 units @ $44.00)
  • TAE buffer
  • loading dye (bromphenol blue)
  • 1kb marker
  • sterile transfer pipettes

    1. You may wish to have the students practice pipetting before they attempt to load their samples into the wells. Carolina Biological has kits for practicing. Having the students use two hands on the pipette will help steady it. The contents should be expelled slowly out, making sure the tip is not poking through the bottom of the well.

    2. In order to visualize the DNA bands on the gel, the gel needs to be stained in the buffer containing either ethidium bromide or methylene blue. Ethidium bromide gives better results, but it is a carcinogen, and should not be used by students. Also, the equipment used to see the bands is much more expensive than that for the methylene blue method (which does not give as good a resolution of bands). Both methods are described in great detail in DNA Science.

    3. At the end of the laboratory series, all bacterial cultures, tubes, pipettes, and other instruments that have come in contact with the cultures should be collected. Materials and glassware should be disinfected with a 10% bleach solution and placed in bio bag for disposal.



    1. Use a permanent marker to label one sterile 15ml tube +pAMP and the other tube -pAMP. Plasmid DNA will be added to the +pAMP tube; none will be added to the -pAMP tube. Put your name on the tubes as well.

    2. Use a sterile transfer pipette to add 250 ul of cold CaCl2 solution to each tube; place both tubes on ice.

    3. Use a sterile inoculating loop to transfer one or two large colonies (3mm) from the starter plate to the +pAMP tube: a. Sterilize loop (if metal) in Bunsen burner flame until it glows red hot. Then pass the lower half of the shaft through the flame.

      b. Stab the loop several times into the side of the agar to cool.

      c. Scrape up a visible cell mass, but be careful not to transfer any agar.

      d. Immerse loop tip in the CaCl2 solution and vigorously tap against wall of the tube to dislodge the cell mass. Hold tube up to light to observe the cell mass drop off into the CaCl2 solution. Make sure cell mass is not left on loop or on side of the tube.

      e. Reflame loop before placing on lab bench.

    4. Immediately resuspend cells in +pAMP tube by repeatedly pipetting in and out using a sterile transfer pipette. Be careful not to suction any cells up into the bulb!

    5. Return +pAMP to ice.

    6. Transfer a second mass of cells to the -pAMP tube as described in steps 3 through 5 above. Use a new transfer pipet to resuspend cells.

    7. While both tubes are on ice, use a new sterile transfer pipette to add 10.0 ul of 0.005 ug/ul pAMP solution directly into cell suspension in the +pAMP tube. Tap tube with finger to mix. Avoid making bubbles in the suspension or splashing suspension up the sides of the tubes.

    8. Return +pAMP tube to ice. Incubate both tubes on ice for an additional 15 minutes.

    9. While your tubes are incubating, use a permanent marker to label two LB plates and two LB/amp plates with your name and the date. Take the time to predict what your results will be.

      a. Label one LB/amp plate +. This is the experimental plate.

      b. Label the other LB/amp plate -. This is the negative control.

      c. Label one LB plate +. This is a positive control.

      d. Label the other LB plate -. This is a positive control.

    10. Following a 15 minute incubation, heat shock the cells in both tubes:

      a. Carry the ice beaker to the water bath. Remove the tubes from the ice and IMMEDIATELY immerse them in the 42deg C water for 90 seconds.

      b. Immediately return both tubes to ice for at least 1 additional minute.

      NOTE: At this point, you may need to give your tubes to your teacher for overnight storage. Make sure your tubes are clearly labelled.

    11. Place +pAMP and -pAMP tubes in test tube rack at room temperature.

    12. Use a sterile pipette to add 250 ul of LB broth to each tube. Gently tap tubes with finger to mix.

    13. Use the matrix below as a checklist for spreading +pAMP and -pAMP cells on the four different plates:
      Agar Used         +pAMP          -pAMP
         *                *              *   
       LB/amp           100 ul         100 ul
        LB              100 ul         100 ul

    14. Spread cells over the surface of each plate.

    15. Let plates set for several minutes to allow suspension to become absorbed into the agar. Wrap the plates together with tape, place them upside down in the incubator, and incubate for 12-24 hours.

    16. After initial incubation, store plates at 4deg C to arrest E. coli growth.

    17. Save all plates with colonies on them for the next step.

    18. Take time for responsible cleanup:

    a. place all items that have come in contact with E. coli into the "bio-bag" provided by your teacher after disinfecting with 10% bleach or disinfectant.

    b. wipe down lab bench with disinfectant.

    c. wash hands before leaving lab.


    1. Observe the plates, and count the number of colonies on each. Observe colonies through the bottom of the culture plate, using a marker to mark each colony as it is counted. If cells are too dense, record as "lawn". Were results as expected? Explain.

    2. Compare and contrast the growth on each of the following pairs of plates. What does each pair of results tell you about the experiment?

      a. +LB and -LB (i.e., LB agar with +pAMP compared to LB agar without pAMP)

      b. -LB/amp and -LB

      c. +LB/amp and -LB/amp

      d. +LB/amp and +LB



    1. Label 2 sterile 50 ml tubes with name, date, and which colony you will be suspending (i.e., tube #1 = untransformed, tube #2 = transformed).

    2. Use 10 ml pipette to sterilely transfer 5 ml of LB broth into tube #1. Use a second pipette to transfer 5 ml of LB/amp broth into tube #2.

    3. Locate a well-defined colony on your LB plate.

    4. Sterilize inoculating loop and scrape up a cell mass from your plate.

    5. Sterilely transfer colony into culture tube #1.

    6. Repeat steps 4 & 5 for the LB/amp plate.

    7. Reflame loop before placing it on the lab bench.

    8. Loosely replace caps to allow for air flow.

    9. Incubate 1 or more days at 37deg C until ready for miniprep.

    10. Shake culture tubes to resuspend E. coli cells.

    11. Label two 1.5 ml tubes with your initials and indicate "untransformed" (#1) and "transformed" (#2). Use sterile transfer pipette to transfer 1 ml of your overnight LB suspension into the tube labeled "untransformed". Use a second pipette to transfer 1 ml of your overnight LB/amp broth into the tube labeled "transformed".

    12. Close caps, and place tubes in microcentrifuge rotor. Make sure you place the tubes opposite for balance. Spin for 1 minute to pellet cells.

    13. Pour off supernatant from tubes into waste beaker for later disinfection. Do not disturb cell pellets. Invert tubes, and tap gently on surface of clean paper towel to drain thoroughly.

    14. Add 0.35 ml of lysis buffer solution to tubes. Resuspend pellets by pipetting solution in and out several times. Check to make sure the solution is homogeneous before continuing.

    15. Close the caps and place in 42deg C water bath for 3 minutes. Immediately cool on ice for 5 minutes.

    16. Spin for 5 minutes with tubes BALANCED in the microfuge.

    17. Pour the supernatant into two clean 1.5 ml tubes.

    18. Add 400 ul of isopropanol to supernatant. Close caps and mix vigorously by rapidly inverting several times. Stand at room temperature for 2 minutes.

    19. Place tubes in a balanced configuration in microfuge and spin for 5 minutes.

    20. Pour off supernatant.

    21. Add 1.0ml iced 70% ethanol to the tubes and invert the tubes gently a few times. Do NOT resuspend! This is a cleaning step only. Centrifuge again in the microfuge for 2 minutes.

    22. Again, carefully pour off the supernatant. Stand the tubes upside down on a paper towel to allow all of the liquid to drain off, making sure the pellets don't slide down the tube.

    23. Let dry 5-10 minutes until all the alcohol smell is gone. Resuspend the pellets by vortexing (vigorous mixing) in 50 ul of TE buffer. Give to your teacher in labeled tubes to be stored in the cold.



    1. Set up the following mixtures in different microfuge tubes and incubate for 30 min at 37deg C. Make sure to add the solutions in the order listed:

      a. 2 ul l buffer, 3 ul pAMP plasmid DNA ('transformed' prepared in part II)

      b. 2 ul buffer, 10 ul control DNA ("untransformed" prepared in part II)

      c. 10 ul marker ladder

      d. 1 ul buffer, 10 ul control DNA ("untransformed"), 1 ul EcoRI, 2 ul EcoRI buffer

      e. 1 ul, 3 ul pAMP plasmid DNA ("transformed"), 1 ul EcoRI, 2 ul EcoRI buffer

    2. While the DNA is being digested by the EcoRI restriction enzyme, pour the gel. (This can also be done ahead of time, just make sure the gel is completely covered by buffer so it doesn't dry out).

    3. Allow the gel to solidify for approximately 20 minutes.

    4. When the gel has solidified, pour approximately 150 ml TAE electrophoresis buffer into the chamber to completely cover the gel. Wait a couple of minutes, then carefully and slowly pull the comb straight up so the gel will not break.

    5. After the digest incubation has ended, heat the restriction enzymes by putting the digests at 65deg C for 5 minutes. Then remove 5 ul of each digest to a new microtube (don't forget to label them)and add 3 ul of loading dye solution to your samples.

    6. Load the samples into the wells in the order below. If you mix up the order, record your actual order in your notes.

      Lane 1. 10 ul 1 KB Ladder MW Marker, dye included (letter "c" above)

      Lane 2. 1.0 ul undigested pAMP plasmid DNA (letter "a" above)

      Lane 3. 5.0 ul EcoRI-digested pAMP DNA (letter "e" above)

      Lane 4. 5.0 ul EcoRI-digested control DNA (letter "d" above)

      Lane 5. 10 ul undigested control DNA (letter "b" above)

    7. When all the samples are loaded, close the lid on the gel box and attach the electrical leads, taking care not to jostle the box. Connect the red (positive) leads to the red jack, and the black (negative) leads to the black jack.

    8. Turn on the power and adjust to the appropriate voltage as indicated by your power supply (in general, lower voltages result in a longer running time). Electrophorese your samples until the dye has reached the middle of the gel.

    9. Follow the staining procedures as given to you by your teacher.


    Remember that the migration of DNA molecules in agarose gels is roughly proportional to the inverse of the log of their molecular weights (sizes). Sketch the banding patterns present on the gel and compare the sizes to those found on the 1 KB Ladder Marker. What do the results indicate? Plot their sizes (y-axis) against the distances migrated (mm, x-axis) onto semi-log paper or using computer graphics.

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