Arizona State University College of Liberal Arts and Sciences

Experiment I
Recombinant Plasmid and Protein Expression


Week 1

1. Measures, techniques, and equipment.

2. Restriction digestion of pLIT-GFP and pET28-bgal.

3. Ligation of digested pLIT-GFP and pET28-bgal .

Week 2

1. Electroporation of E. coli to produce transformants.

2. Plating of cells on LB-agar medium with antibiotics to select for transformants.

3. Starting cultures of the transformants (one day after plating period) .

Week 3

1. PCR colony screen to identify recombinant clones.

2. Isolation of the recombinant plasmid from transformed E. coli cells.

Week 4

1. Restriction digestion of the recombinant plasmid.

2. Analysis of the digested recombinant plasmid and PCR reactions (week 3 lab) by gel electrophoresis.

3. Transformation of E. coli expression strain BL21(DE3) with recombinant plasmid.

4. Starting recombinant BL21(DE3) liquid cultures (day before lab).

Week 5

1. Collect BL21(DE3) cell samples.

3. Extract cell samples for native protein.

Week 6

1. Extract cell samples (denatured protein).

2. SDS-PAGE electrophoresis of proteins in cell samples.

Week 7

1. Metal affinity chromatography of GFP-6His.


Experiment I Overview

Recombinant DNA technology refers to the process of taking parts of two DNA molecules and putting them together to generate a new molecule that conveys new properties to the organism that receives this DNA. In this experiment, you will construct a recombinant plasmid designed to allow high-level expression of the inserted gene in E. coli cells.  The pET system (Novagen,, is depicted in Figure 1. (You may find the “pET System Tutorial” document as a pdf file on the Blackboard site.) The target gene is under control of the T7 promoter, which requires T7 polymerase for transcription (derived from bacteriophage T7). The trick is that the T7 polymerase gene has been inserted into the chromosome of the E. coli genome, under control of the lac promoter, which is inducible by galactose (in practice, by a galactose analog isopropyl -β-D-thiogalactoside, IPTG). Thus, cells can be grown in absence of IPTG to allow efficient growth without recombinant protein expression, and later addition of IPTG will cause expression of T7 polymerase, and consequently, the target gene. Moreover, in the pET28b vector that you will use, the target gene will be inserted in frame with a 6-histidine (6His) element, which will allow purification of the His-tagged protein by metal affinity chromatography.

You will make recombinant DNA by putting together two pieces of different plasmids. Plasmids are circular double-stranded DNA molecules that can range in size from less than 1 kbp (kilo base pairs, i.e., 1,000 base pairs) to more that 500 kbp. Plasmids are maintained in the bacterial cell as independent, autonomously replicating extrachromosomal entities. They contain an origin of replication that is recognized by DNA polymerase. They may also carry restriction endonuclease sites that are recognized by certain restriction enzymes (molecular scissors). Each of these enzymes cut the DNA at a specific DNA sequence. Moreover, plasmids used in the laboratory often carry one or more antibiotic resistance genes. These genes code for proteins that break down or modify the antibiotics and hence convey resistance to such antibiotics. The production of a recombinant DNA molecule is greatly facilitated by cleavage of the two DNA molecules that need to be recombined with restriction enzymes that leave compatible sticky ends on the DNA. These ends can then be ligated (enzymatically glued together).


Figure 1. Control elements of the pET expression system.

You will be given two different plasmids, pLIT-GFP and pET28-bgal (Figure 2). pLIT-GFP has an ampicillin resistance ( ampR) gene and pET28b has a kanamycin resistance ( kanR) gene. You will cleave both plasmids with restriction enzymes NcoI and XhoI, to release the fragment encoding green fluorescent protein (GFP) from pLIT-GFP, and ligate it with the vector pET28-bgal cleaved with the same enzymes. E. coli cells are transformed with the recombinant plasmid mixture. Selecting for cells that contain the desired recombinant plasmid, pET-GFP, is done by plating transformants on LB-agar that contains 50 µg/ml of kanamycin. As antibiotics are heat-sensitive, a sterile solution with the antibiotics is added to the agar after autoclaving, when the agar has cooled down to about 55°C.

plamid pET28-GFP


Figure 2. Construction of recombinant plasmid pET28-GFP.  pLIT-GFP (AmpR, ampicillin resistance marker) contains the GFP coding sequence between NcoI and XhoI sites. Ligation of the NcoI-XhoI digests of pLIT-GFP and pET28-bgal (KanR, kanamycin resistance marker) yields several different products, one of which is pET28-GFP, in which GFP gene is substituted for beta-galactosidase. The T7 promoter drives transcription of the adjacent gene when T7 RNA polymerase is present.

Experiment I

Week 1

Prelab Notes

To calculate the needed volume of a stock solution use the formula:

CiVi = CfVf ,

where Ci and Cf are the initial and final concentrations, and Vi and Vf are the initial and final volumes.


0.5 ml microcentrifuge tubes P200 and P20 micropipettors and disposable tips
Disposable gloves Microcentrifuge
ddH2O (dd = double-deionized) pLIT-GFP and pET28-bgal
NcoI (10 units/µl) and XhoI (20 units/µl) 10x BSA (10 mg/ml bovine serum albumin)
Ligase buffer (50 mM Tris-HCl, pH 7.5, 10 mM MgCl2 (magnesium chloride), 10 mM dithiothreitol, 1 mM ATP, 1 mg/ml bovine serum albumin) 10x NEB buffer 2 (50 mM NaCl, 10 mM
Tris-HCl, pH 7.5, 10 mM MgCl2, 1 mM dithiothreitol)
Ligase (400 units/µl)  
Glycogen (20 mg/ml)  
7.5 M ammonium acetate  
isopropanol (2-propanol)  


The typical order of adding reagents to the reaction vial is:

1. ddH2O

2. Buffer concentrates

3. DNA

Mix solution

4. Enzyme

Mix again

Restriction Digestion of Plasmids

1. Just prior to the lab, TAs prepared the enzyme mix, which contain (per 26 µl):

Reagent Volume, µl
ddH2O 19.25
NEB buffer 2 (10x) 3
BSA (10x) 3
NcoI 0.5
XhoI 0.25
Total 26 l

2. Label two microcentrifuge tubes using a permanent marker, with your initials and plasmid name (abbreviated “LIT” and “ET”).

3. Prepare the restriction digestion using the following table as a guide:

Reagents Tube 1 Tube 2
Enzyme mix 26 l 26 l
pLIT-GFP 4 l -----
pET28-bgal ----- 4 l
Total 30 l 30 l

4. Incubate the samples at 37 C for one hour for restriction digestion.

5. Place the samples at 70 C for 20 minutes to inactivate the restriction enzymes.


1. Just prior to the lab, TAs prepared the ligation mix, which contain (per 20 l):

Reagents Volume
ddH2O 16.5 l
Ligase buffer (10x) 3 l
Ligase 0.5 l
Total 20 l

Keep the ligation mix on ice at all times.

2. To the ligation mix add:

5 µl digested pLIT-GFP
5 µl digested pET28-bgal

3. Mix the sample by carefully pipetting half of the volume in and out several times. Avoid bubbles; if you do generate bubbles, microfuge the sample 5 seconds.

4. Incubate at room temperature 1 hour, or until 10 min before the end of lab period.

5. Precipitate the ligated DNA. To the ligation mix add:
      15 µl         7.5M ammonium acetate
        1 µl         20 mg/ml glycogen (carrier to enhance recovery of dilute DNA)
      46 µl         isopropanol

Mix well by flicking the tube with your finger several times, store at 4°C until next week.

6. Store unused restriction digests at –20 °C. You will need them again in Week 4.

Weekly Report Update

Make sure to address the following points:

1. The use of NcoI and Xho for generating a recombinant plasmid.

2. Possible ligation products, and selection procedure for pET28-GFP.

Experiment I

Week 2

Transformation is defined as the uptake and retention of foreign DNA. Transformation is one of the most commonly used methods of introducing vectors into bacterial cells for cloning purposes. As DNA cannot routinely enter a cell, cells are treated in a specific way to temporarily increase the cell membrane permeability. The goal of this lab is to introduce the recombinant plasmid prepared last week into E. coli cells, where it can be maintained and propagated.

Electroporation is the method that you will use today to transform E. coli cells. Electroporation means that you will use a short electric field pulse to induce microscopic pores (electropores) in the cell membrane. The idea here is to impose a short-lived electric field leading to formation of pores that live long enough to let in DNA molecules, but not so long as to introduce irreversible damage to the cells. If the electric field has the proper parameters, then the "electroporated" cells can recover by resealing the electropores spontaneously, and cells will continue to grow and express their genetic material--including the newly acquired vector. Transformants will be screened for resistance to kanamycin, as you wish to recover cells containing the desired recombinant plasmid (pET28-GFP transformants)..

In addition to transforming with the ligation mix, you will transform a different aliquot of E. coli cells with pET28-bgal as a positive control. Recall that pET28-bgal is the plasmid that you digested in Week 1 to provide the vector backbone for pET28-GFP.

It is important to run controls along with your sample to demonstrate the fidelity of your experiment. Electroporation of cells using a known concentration of a plasmid that has a particular marker gene can be used as a positive control.


Ice Microcentrifuge
Disposable gloves Electro-competent E. coli DH5α cells
Ligation mix precipitation from week 1 P1000, P200, P20, and disposable tips
pET28-bgal Electroporation cuvettes
LB-Agar plates containing 50 µg/ml kanamycin Glass spreader
YENB medium Sterile media tubes
Toothpicks Q-tips
Tape 95% ethanol


LB medium consists of,

per liter: 10 g bactotryptone
  5 g yeast extract
  10 g NaCl
  15 g bactoagar (for plates only)


YENB medium (pH 7.5) consists of,

per liter: Bacto tryptone 2.0% 20 g
  Bacto yeast extract 0.5% 5 g
  NaCl 10 mM 0.58 g
  KCl 2.5 mM 0.19 g
  MgCl2 10 mM 10 ml 1 M
  MgSO4 10 mM 10 ml 1 M
  Glucose 20 mM 20 ml 1 M

YENB medium is "richer" than LB, and helps to have as many transformants recover from the electroporation shock as possible.


1. Microcentrifuge the isopropanol precipitated ligation from Week 1 at full speed for 5 min. Orient the hinge of the tube toward the outside of the rotor, so that you can identify the side where the pellet will be.

2. Decant the supernatant to a waste container and invert the tube on a clean paper towel or kimwipe to drain, tapping gently.

3. Wash the pellet. Gently pipet 0.5 ml of 70% ethanol directed to the side of the tube away from the pellet, so that the pellet is not disturbed. Invert the tube once, microfuge for 1 min., and repeat step 2.

4. Dry the pellet in the Speed-vac for 5 min (heat setting low). If you see fluid remaining in the tube, dry for another 5 min. Since ammonium ions are volatile, any residual ammonium ions will evaporate, thus reducing ionic strength for electroporation.

5. Add 4 µl sterile ddH2O and pipet up and down 5 times to resuspend the pellet, and keep on ice.

6. (Done by TAs) E. coli DH5a cells were grown in YENB (low salt medium) to a cell density that corresponds to log phase growth. Cells were collected by centrifugation, and then washed several times with 10% glycerol, and flash-frozen. The low ionic strength of the solution prevents large currents that may permanently damage the cells during electroporation. The TAs will aliquot the cells in microfuge tubes at 40 µl each. Obtain 2 aliquot tubes and keep them on ice.

7. Use CLEAN, DRY and STERILE electroporation cuvettes. The cuvettes will be clean, dry and sterile before you use them, and make sure they are clean, dry, and sterile again before you leave!! After use, clean electroporation cuvettes by rinsing them with ddH2O and scrubbing the sides with a toothpick and a Q-tip. Sterilize the cuvettes and cuvette covers by soaking them in 70% ethanol for 10-20 minutes. Dry them thoroughly in a sterilized hood by putting them upside-down on an absorbent tissue (of course, you have not touched that part of the tissue!). It is important to maintain sterile conditions throughout this protocol. Use only sterilized cuvettes, microcentrifuge tubes, and pipet tips.

8. Label and prechill electroporation cuvettes by putting them on ice for at least 5 minutes.

9. Pre-warm a bottle of YENB at 37 °C.

10. Add 2 µl of the appropriate DNA to labeled microcentrifuge tubes with 40 µl E. coli cells and mix thoroughly by gently flicking the tube with your finger 5 times. Avoid pipetting the cells to mix, as cells can be damaged by shearing. Make sure to use a fresh pipette tip for each sample.

Tube 1           2 µl precipitated ligation mix
Tube 2           2 µl pET28-bgal positive control, 10 pg/µl DNA

Incubate the samples on ice for 5 minutes.

11. Carefully (avoid shearing cells!) pipette the contents from one tube into a dry and prechilled electroporation cuvette and tap the bottom gently on the bench top to propel the fluid to the bottom of the cuvette. Put the cuvette back on ice. Make sure that there are no air bubbles in the sample in the cuvette as they may explode upon electroporation!

Do one sample at a time for steps 12-14!!

12. Put the cuvette into the electroporator, pulse at 2.5 kV and record the time constant. Any time constant above 4 ms is OK, while a shorter time constant is indicative of rapid capacitor discharge and therefore high electric current flow that can cause irreversible damage to the cells.

13. Immediately after the pulse, use a P-1000 Pipetman to transfer 1 ml of YENB (prewarmed to 37 °C) medium into the cuvette, and immediately decant into to a glass culture tube, and apply the cap.

14. Place the capped tube in the 37 °C rotary incubator before you start your next electroporation and incubate the tube for 45 minutes. During this time, the antibiotic resistance marker gene(s) will be expressed, and appropriate transformants will be able to form colonies on plates in the presence of an antibiotic.

15. Clearly label the bottom (not the lid) of the LB-Kan agar plates with a permanent marker. Prepare 2 plates for the ligation, and one for the positive control.

16. For ligation transformation, gently pipette 0.25 ml cells on one plate, and the remaining volume on the other. For the positive control pET28-bgal, use 0.1 ml. Spread the cells with a sterilized glass spreader. Do the ligation cells first, both plates without sterilizing the glass spreader. Then sterilize the spreader before doing the positive control.

Sterilize the glass spreader by dipping it in 70% ethanol and briefly passing it through a burner flame to ignite the ethanol. Place the 70% ethanol beaker away from the burner so as not to ignite the ethanol in the beaker. Cool the glass spreader for 15 seconds in air, and then placing it on the periphery of the agar plate before spreading the cells. Repeat glass spreader sterilization between samples.

17. Place the plates inverted in the 37°C incubator, and make sure you are careful when moving or removing plates… gravity tends to send unsupported lids to the floor. Colonies should be visible within 16 hours. You need to come to the lab tomorrow to remove your plates from the incubator and record your results (estimate number of colonies per plate).

18. The same day that you remove your plates from the incubator, use a sterile P-200 pipette tip with a P-200 pipettor to pick 3 single colonies on the ligation transformation plates. Transfer the cells to labeled microcentrifuge tubes containing 50 µl of sterile ddH2O, pipetting up and down a few times to disperse the cells. With 10 µl each colony suspension, inoculate a 2 ml culture in a labeled glass tube of liquid LB medium containing 50 µg/ml kanamycin, and place the culture in the rotating wheel at 37 °C. The cultures will be used to isolate the recombinant plasmid in the next lab period. TAs will remove the cultures after about a day of growth, and will place them in the refrigerator.

19. Place your plates and picked colony suspensions in the refrigerator. The colony suspensions will be used for colony PCR screening in Week 2.

Prelab Questions

1. What is the purpose of electroporation?

2. Discuss the procedure for the selection of transformed cells carrying the recombinant plasmid.

3. What proportion of recombinant kanamycin-resistant colonies are expected to be the desired pET38-GFP? What other recombinant ligation products are possible?

Weekly Report Update

1. Discuss the electroporation procedure.

2. What are your transformation results? (Number of colonies on the various plates).

Experiment I

Week 3

This week you will prepare and digest plasmid DNA from the 3 liquid cultures that were inoculated 6 days ago, and also perform PCR on the cell suspensions used to inoculate the cultures. Next week, samples will be analyzed by gel electrophoresis to determine fragment sizes. The method for plasmid DNA preparation used in this lab is the alkaline lysis miniprep method, which is a simple and efficient technique to isolate small amounts (1-10 µg) of plasmid DNA from 1-3 ml of transformed E. coli culture. The DNA isolated from this procedure is pure enough for most of the testing done in a molecular biology laboratory such as restriction digestion, agarose gel electrophoresis, and E. coli transformation. An overnight culture originating from a single colony of E. coli that may contain the plasmid of interest is harvested, and the cells are lysed. Plasmid DNA is then separated from cellular proteins, lipids, and chromosomal DNA.

To fully understand this procedure, you will need to become familiar with the molecular and biochemical effects of the reagents used in the protocol:

Materials for plasmid preparation

2 ml overnight culture of plasmid-containing E. coli grown in LB + 50 l/ml ampicillin 10% SDS
TE solution (10 mM Tris, 1 mM EDTA, pH 8.0)  
TER (TE + 15 µg/ml RNase A, DNase-free)  
7.5 M ammonium acetate  
100% isopropanol 70% ethanol
2 M NaOH ddH2O
P20, P200, P1000 and pipette tips 1.5 ml microcentifuge tubes
Ice bucket Microcentrifuge
Paper towels  

RNase preparation for TER (NOTE: typical preparations of bovine pancreatic RNase are contaminated with DNase.)  10 mg/ml RNase A, DNase free- 3 ml:

    30 mg RNase A
    3 ml ddH2O

    Mix by vortexing

Place the tube in boiling water for 10 minutes, allow to slowly cool to room temp, then store at -20 °C. Heat denatures both RNase and DNase, but  RNase renatures on slow cooling, while DNase does not.

Method for plasmid preparation

1. If cells have settled, shake the culture tubes gently to resuspend the E. coli cells.

2. Use a P-1000 to transfer 1.5 ml of each of the 3 cultures into appropriately labeled 1.5 ml microcentrifuge tubes. Use a separate pipet tip for each culture.

3. Close the caps well and centrifuge the tubes for 1 minute in a properly balanced microcentrifuge rotor at ~3/4 speed (~12,000xg). For some microfuges this may be full speed.

4. While cells are spinning, make NS solution (0.5% SDS/0.1 M NaOH) using the 10% SDS and 2 M NaOH stocks (this solution should be made fresh daily):

                                Stock          1 ml
0.1 N NaOH          2 M          0.05 ml
0.5% SDS             10%        0.05 ml
Water (sterile)                          0.9 ml

5. Remove the supernatant from the tubes with a P-1000 being careful not to disturb the pellet. Quick-spin 5 sec in the microfuge to pull down residual supernatant, and remove it with another P-1000 tip, being careful not to disturb the pellet.

6. Resuspend each pellet in 50 µl of TE, by gentle digging and stirring with a P-200 pipet tip, followed by moderate vortexing. Be sure to resuspend well, as cells in clumps will not lyse and yield will suffer.

7. Add 300 µl of NS solution to each tube. Mix gently by inverting tubes four  times. Do not vortex or mix vigorously because this can cause shearing of the chromosomal DNA, which prevents it from pelleting in the next step.

8. Incubate at room temperature 1-2 min (longer times may cause irreversible denaturation of plasmid.)

9. Add 200 µl of ice-cold 7.5 M ammonium acetate solution to each tube. Mix by rapidly inverting the tubes several times. A white precipitate will appear immediately.
NOTE: Ammonium acetate solution is filter sterilized; DO NOT autoclave.

10. Incubate tubes on ice for 5 minutes.

11. Centrifuge for 5 minutes in a balanced microcentrifuge to pellet chromosomal DNA, cell debris, etc.

12. While tubes are spinning, label 3 clean, sterile 1.5 ml tubes and add 0.33 ml isopropanol to each tube.

13. Transfer the supernatants to the tubes containing isopropanol, cap, and vortex to mix well. Avoid taking the pellet and floating debris. Discard old tubes containing precipitate in biohazard waste.

14. Let stand at room temperature for 2 minutes, then centrifuge for 5 minutes in a balanced microcentrifuge at full speed to pellet the nucleic acids. Align the tubes in the rotor so that the cap hinges point outward. Nucleic acid pellets are not always visible, but by orienting the tubes in the rotor in this manner the pellet will form at the bottom of the tube under the hinge during centrifugation.

15. Decant the supernatant to a waste container and invert the tube on a clean paper towel or kimwipe to drain, tapping gently. The pellet should stick firmly, but can be loose. NOTE: Plasmid DNA may be in a pellet at the bottom or streaked on the side of the tube and may be only slightly visible or translucent.

16. Wash the pellet. Gently pipet 0.5 ml of 70% ethanol directed to the side of the tube away from the pellet, so that the pellet is not disturbed. Invert the tube once, observing the pellet to remain attached, and decant the wash to a waste container and invert the tube on a clean paper towel or kimwipe to drain, tapping gently. If you see that the pellet became detached, centrifuge the tube for 1 min at full speed before decanting.

17. Dry the pellet in the Speed-vac for 5 min (heat setting low). If you see fluid remaining in the tube, dry for another 5 min. If not required immediately, pellets can be air-dried on the bench for 15-20 min. Observe closely by holding the tubes up to the light to verify that all fluid is evaporated.

18. Add 30 µl of TER (TE + 15 µg/ml RNase A) to the tube to resuspend the DNA pellet by gently flicking with a finger or vortexing. Store the tubes at -20 °C for use next week.

Materials for colony screen PCR

E. coli colony suspensions from last week
pET28-GFP positive control plasmid (10 pg/µl)
ddH2O, sterile
Taq polymerase (5 units/µl)
Taq polymerase buffer (10X, with Mg2S04 at 2 mM for 1X)
dNTP solution (5 mM each dATP, dCTP, dGTP, dTTP)
oligonucleotide primers, 100 µM each
            GFP-Nco (forward): 5’-ATACCATGgtgagcaagg
         pET26-F (reverse): 5’-agccaactcagcttcct
P20, P200, P1000 and pipette tips         
0.2 ml PCR tubes
Ice bucket
PCR thermal cycler

Method for colony screen PCR

1. 1. Just prior to the lab, TAs prepared the PCR reaction mix, which contains (per 15 µl):
Reagent                    Volume, µl
ddH2O                          11.45 µl
10X Reaction buffer   2.0 µl
5 mM dNTPs                0.8 µl
Primer 1                        0.25 µl
Primer 2                        0.25 µl
Taq polymerase          0.25 µl
Total                              15 µl

NOTE: Keep on ice until reactions are started.

2.  Label 5 PCR tubes (one for each of 3 colony  suspensions, and one for positive control, one for negative control), and add 15 µl of the reaction mix to each tube.

3.  Vortex the colony suspensions briefly to resuspend bacterial cells, and add 5 ml bacterial suspension to its numbered tube, and mix by pipetting up and down 3 times. NOTE: The buffer contains Triton X-100, a detergent, and pipetting air into the mix will generate bubbles; thus be careful not to introduce air when mixing. If bubbles are excessive, spin the tube 5 sec in a microcentrifuge using a 1.5 ml as adaptor.

4. For negative control add 5 µl sterile ddH2O, and for positive control, add 5 µl of pET28-GFP (10 pg/µl).
NOTE: Pipet the positive control last, in order to minimize the chance of contaminating the other reactions. It is extremely easy to contaminate your pipettor with plasmid DNA; thus use care and avoid splashing when pipetting.

5.  Load tubes in the thermal cycler and start, cycle parameters:

            1 cycle:  95 °C for 3 min to lyse the cells and denature the DNA

            25 cycles:      93°C x 30 sec
                                    52°C x 30 sec (temp depends on primer length and G/C content)
                                    72°C x 30 sec (~30 sec per kb of amplicon; expect 791 bp)

6. Store the tubes at -20 °C for gel analysis next week.

Prelab Questions

1. What are the roles of NaOH and SDS in the plasmid miniprep procedure?

2. Why is it necessary to avoid vigorous mixing (e.g. vortexing) of the solution after adding NaOH/SDS?

3. Note that in the PCR reaction, one of the primers binds to GFP (GFP-Nco) and the other primer (pET26-R) binds to the vector DNA. How does this choice of primers help to distinguish recombinant pET28-GFP from other possible recombinant ligation products (refer to the plasmid map in Fig. 3 below)? Can you suggest other primers that might also work as well?

map of pET28-GFP

Figure 3. Map of pET28-GFP. Primers for PCR are indicated in black type, with arrows indicating the orientation. T7 promoter drives transcription to produce GFP mRNA. A primer also exists that binds the T7 promoter. RBS, ribosome binding site, where translation initiation occurs.

Experiment I

Week 4

This week you will perform restriction enzyme digestion of your (hopefully) recombinant plasmids prepared last week, and you will perform agarose gel electrophoresis to analyze both the restriction digests and the PCR reactions from last week. Then you will select one of the three plasmid preparations to use for transformation of an E. coli protein expression strain, BL21(DE3), by electroporation. In the (unlikely) event that you fail to obtain any recombinant clones, you may use a recombinant plasmid obtained by one of your successful classmates. In the (extremely unlikely) event that none of you obtained a recombinant clone, you will be provided a sample of pET28-GFP by your TA.

Remember that the recombinant plasmid pET28-GFP was obtained by ligating pLIT-GFP and pET29-bgal fragments that were obtained using NcoI and XhoI restriction enzymes. This indicates that the pET28-GFP plasmid should have both NcoI and XhoI restriction sites (refer to the diagram of plasmids in lab 1). Note that the NcoI site (CCATGG) contains the start codon (underlined) for the GFP coding sequence, and thus it is very important that this site is confirmed in your recombinant clone, in order to expect translation initiation to occur. Furthermore, the XhoI site (CTCGAG) lies within the coding sequence just 5’ of and in frame with the 6His coding sequence. Thus, it is also important to verify the XhoI site in your recombinant clone. You will digest plasmid DNA with NcoI and XhoI restriction enzymes, and analyze the size of the fragments by gel electrophoresis.

Restriction Digestion Materials

disposable gloves  
pET28-GFP plasmid preparations
enzyme mixture      
37 °C incubator
70 °C water bath       


1. DNA restriction enzymes should always be added last! The typical order for reagents in restriction digests is:
          * Water
          * Buffer, containing a specific salt (NaCl, MgCl2) concentration.
          * DNA
          * Mix the sample by pipetting up and down 3 times.
          * Enzyme

 2. Restriction enzyme buffers usually are provided as concentrated stocks, and need to be added in a volume so that the final concentration is 1x. Using inappropriate concentrations of buffer may result in the inactivation of the restriction enzyme.

 3. Mix the sample after DNA addition by pipetting the total volume into a pipet tip; then dispense the total volume into the restriction digestion tube, and gently mix the sample with the enzyme. This is to prevent the exposure of the enzyme to unfavorable salt concentrations or pH conditions.


Restriction Digestion of Cloned Plasmid

1. Use a permanent marker to clearly label 3 tubes, one for each of your 3 plasmid preparations.

2. Just prior to the lab, TAs prepared the enzyme mix, which contain (per 22 µl):

Reagent                       Volume, µl
ddH2O                              15.25
NEB buffer 2 (10x)            3
BSA (10x)                           3
NcoI (10 units/µl)               0.5
XhoI (20 units/µl)               0.25
Total                                  22

3. Prepare the digest reactions by adding 22 µl of the enzyme mix to each of your 3 tubes; then add 8 µl of each plasmid prep to its correctly labeled tube, and mix by pipetting up and down 3 times.

4. Incubate the samples at 37 °C for one hour.

Agarose gel electrophoresis

After treating the DNA molecules with restriction enzymes, the fragments are separated according to size (number of base pairs) using a 0.8% (w/v) agarose gel. Agarose is a form of agar, which is a polysaccharide isolated from seaweed. An electric field runs through the gel causing the DNA that is negatively charged (due to phosphate groups) to migrate toward the positive pole; the larger the fragment the slower it moves, due to the molecular sieving effect of the gel matrix. To provide electrolytes that conduct electricity, agarose gels are submerged in a Tris/boric acid/EDTA or Tris/acetic acid/EDTA buffer.

DNA is colorless, and therefore migration of DNA molecules is hard to visualize on the agarose gel. Therefore, dyes are added to the DNA samples before loading them on the agarose gel. The loading dye (tracking agent) is made of two different chemicals- bromophenol blue and xylene cyanol. The faster-moving, purplish band is bromophenol blue, and it migrates along the agarose gel at the same rate as DNA of approximately 600 base pairs (bp). The slower-moving, bluish band is xylene cyanol and it migrates at the same rate as DNA of approximately 4000 bp.

To visualize DNA, DNA molecules are stained using ethidium bromide. Ethidium bromide is an intercalating agent, which binds between the two strands of double-stranded DNA. Ethidium bromide bound to DNA is very fluorescent, and emits orange light when excited with ultraviolet (UV) light. Therefore, after staining, DNA fragments can be visualized using an ultraviolet lamp (260-360 nm).


      Usually, it is important to estimate the size of double-stranded, linear DNA on gels. For this purpose, a small sample of a "1 kb ladder" generally is run on a gel, next to samples. The 1 kb ladder (Figure 4) is a commercially available mixture of linear DNA of known sizes, ranging from a few hundred bp to 12 kb. By comparing the mobility of the ladder and samples, one can determine the approximate size of DNA fragments. The ladder can also be used to estimate the concentration of DNA fragments by comparing the brightness of the bands.


Figure 4. Fermentas GeneRuler™ 1 Kb ladder.

It is important to note that approximate size determinations using a 1 kb ladder are valid for linear, double-stranded DNA only. It does not work for circular DNA, as the supercoiled DNA runs faster, and nicked-circular DNA runs a bit slower than linear DNA of the same size.


dd H2O Micropipettors + disposable tips
Disposable gloves Microcentrifuge
10x BSA Agarose gels with EtdBr (made by TA)
1x Tris-Borate-EDTA (TBE) buffer 1 kb DNA ladder
6x Loading dye 10 g/ml ethidium bromide
Disposable gloves Gel tray and comb
Electrophoresis box 60 C water bath (for agarose)
60 °C water bath (for agarose)  
37 °C water bath  

Method: Gel electrophoresis

The TAs will perform steps 1-7 before lab, so that the gels are ready to run when you arrive.

1. Weigh 0.25 g of agarose in a 125-250 ml flask.

2. Add 30 ml TAE.

3. Microwave for 1 minute and 30 seconds at 30% power level to dissolve the agarose, and verify that agarose granules have disappeared. Using a mitt or several paper towels, grasp the flask and swirl gently—being careful to avoid touching the hot flask!

4. Place the flask in a 60 °C water bath.

5. Place rubber stoppers on both sides of the gel tray. Be sure to secure the rubber stoppers to prevent gel leakage.

6. Place the gel comb in the gel tray and adjust the teeth of the comb to be 2 mm above the tray surface.

7. Pour the gel slowly into the tray that is placed on a flat and horizontal surface. The gel should cover only one third of the height of the comb teeth. Use a large pipet tip to remove large bubbles. Let sit for 10-20 minutes to let the agarose solidify.

8. When the agarose has solidified, cover the gel on the tray with TAE buffer.

9. Gently pull the comb by holding it with both hands, without wiggling or rocking, to prevent deformation of the wells (do not pull comb without submerging the gel in the buffer).

10. Pour the buffer in the electrophoresis box.

11. Remove the rubber stoppers and place the gel (with tray) in the electrophoresis box.

12. Immediately immerse the gel with 1x TAE buffer.

13. Arrange your sample tubes:

1           Digested pLIT-GFP (week 1)
2           Digested pET28-bgal (week 1)
3           Digested pET28-GFP #1
4           Digested pET28-GFP #2
5           PCR product #1 (week 3)
6           PCR product #2 (week 3)
7          1 kb ladder (provided by TA)

14. For each sample, place 2 µl of loading dye in a well on a microtiter plate or on a small piece of parafilm and add 10 µl of each sample (be careful to remember which sample is where if you use parafilm).

15. Mix by pipetting the full volume of sample (12 µl) in and out. Load each sample in a separate well in the gel. Use a fresh tip for each sample. Load samples using the following method:

    * Position the pipet tip over the well using one hand and support the Pipetman with the other hand.
    * Center the tip over the well and place it under the TAE buffer covering the well. Dip the tip gently in the well, but do not dip deeply to prevent puncturing the gel and losing your sample. If there is air in the pipet tip, carefully press the plunger of the pipettor to eject the air bubble while being careful not to lose your sample. An air bubble that is depressed in the well can cap the well. This will cause the sample to flow in the buffer outside the well.
    * Cover the electrophoresis box, turn on the power supply, and adjust the voltage to 70 V. Have the gel run for 40-50 minutes, periodically observing the migration of the marker dyes.
    * Turn of the power supply, and carefully remove your gel from the gel electrophoresis tray into a container for transport to the AlphaImager
    * Using the AlphaImager, view the gel on a UV transilluminator, and photograph it.

Method: Transformation of E. coli strain BL21(DE3)

Now that you have identified a recombinant clone (or perhaps 3 of them!) you will use plasmid DNA from a selected clone to transform E. coli BL21(DE3), which is a specially constructed genotype that will allow inducible expression of GFP protein using the pET28 vector system. Although it is possible to transform BL21(DE3) directly with the ligation mix used to transform strain DH5a in week 2, it is preferable to characterize the recombinant DNA obtained from DH5a first. DH5a will maintain greater stability of DNA inserts, and moreover will not express the inserted gene in the pET28 vector, since it cannot express the T7 polymerase needed to transcribe the gene to make mRNA. Although this is not a problem with GFP, other proteins may be toxic to bacterial cells and inhibit growth.

1. The electroporation method is the same as that used in week 2 for DH5a cells, only this time you will be provided with electro competent BL21(DE3) cells. Your TA will provide you with an aliquot of electro competent cells, to which you will add 1 µl of your plasmid.

2. After incubation of DNA with competent cells, follow the protocol as described for the week 2 lab, except:


Prelab Questions

1. Discuss the use of a ladder in gel electrophoresis.

2. What size bands do you expect to see in the pLIT-GFP, pET28-bgal, and pET28-GFP restriction digests? (Refer to the maps in Figure 2, week 1).

3. What is the role of IPTG for inducing expression of GFP in BL21(DE3) cells transformed with pET28-GFP?

Weekly Report Update

1. What is the purpose of restricting the pET28-GFP recombinant plasmid?

2. Discuss agarose gel analysis of samples and controls. Include the gel photograph.

3. Note whether GFP fluorescence is observed in BL21(DE3)/pET28-GFP cells +/- IPTG.

Reagents for agarose gel electrophoresis

6x Loading dye
0.25% (w/v) bromophenol blue
0.25% (w/v) xylene cyanol FF
30% (w/v) glycerol

1x TAE: 40 mM Tris-acetate pH 8.0, 1 mM EDTA

Experiment I

Week 5


Your cultures of E. coli BL21(DE3)/pET28-GFP, which you inoculated 6 days ago into Staby™Switch auto-inducible medium, should have undergone induction of T7 polymerase as the culture matured. See the web site for a full explanation. As the bacterial cells metabolize proprietary medium components during growth, the resulting compounds will automatically induce protein expression at high cell density. Thus, it is not necessary to add IPTG (isopropyl -β-D-thiogalactoside) to induce expression. The production of T7 polymerase in these bacterial cells allows transcription of the GFP gene inserted downstream of the T7 promoter.

You will collect cell pellets, save a sample for SDS-PAGE in Week 6, and extract cells in a non-denaturing buffer to obtain a mixture of cellular proteins that includes the induced GFP-6His. The extracts will be stored at -20 °C until Week 7, when you will perform metal affinity purification of the His-tagged GFP.


BL21(DE3)/pET28-GFP cultures (induced and uninduced)
1.5 ml microcentrifuge tubes
P-20, P-200 pipettors and tips
Extraction buffer (10 mM Tris, pH 8.0; 50 mM NaCl)
Lysozyme (10 mg/ml in water)
Dry ice (solid CO2)


Method: Bacterial cell extraction

1. Transfer the 4 ml liquid cultures to labeled 1.5 ml microcentrifuge tubes (2 each for induced and uninduced cultures).

2. Centrifuge to sediment cells 1 min in a microcentrifuge and discard supernatants. Drain pellets thoroughly and place tubes on ice.

3. Examine the cell pellets under long-wave UV light and record the fluorescent appearance of the induced vs. uninduced samples.

4. Resuspend each pellet in 75 µl of ice-cold extraction buffer by vortexing, and combine the 2 induced samples to make a single 150 µl sample; do the same for the 2 uninduced pellets.

5. Transfer 20 µl of each sample of resuspended cells into labeled tubes (for later use in SDS-PAGE analysis), and store the tubes at –20 °C.


Cell lysis
For larger-scale cultures, sonication is recommended to lyse cells. However, the process is only efficient for cell suspensions of greater than 2 ml, representing culture volumes of at least 40 ml. For lysing smaller cultures (≤ 12 ml), the freeze/thaw method below is recommended and can be performed in standard 1.5 ml microcentrifuge tubes.
Note: Except where noted, keep all samples and tubes on ice.

6. Add 1 µl of lysozyme solution to each of the 2 tubes containing 130 µl cell suspensions. Vortex tubes gently or finger-flick the tubes to disperse lysozyme, which digests the bacterial cell walls. Incubate tubes at room temperature for 5 minutes.

7. In a fume hood, prepare a dry ice bath in an ice bucket by adding dry ice and isopropanol until the consistency becomes slushy.

8. Transfer the lysozyme-treated cell suspension sample tubes to a flotation carrier, and place tubes containing in the dry ice bath until cells are frozen solid (~ 20 seconds).

9. Transfer the tubes to 37 °C bath until the suspension becomes fully liquid, ~ 1 minute.

10. Repeat freeze/thaw cycle 9 times.

11. Spin at full speed in a microcentrifuge for 10 minutes to remove insoluble material.

12. Transfer the supernatants to fresh tubes and store at -20 °C.
Note: If the lysate is too viscous for pipetting, DNase I may be added to a final concentration of 10 µg/ml in order to degrade the high molecular weight DNA.

Prelab Question

1. Why is the T7 polymerase gene in E. coli strain BL21(DE3) under the control of an inducible promoter?

2. What is the role of lysozyme in the extraction of bacterial cells?

Experiment I

Week 6: Denaturing Polyacrylamide Gel Electrophoresis (SDS-PAGE)


Experiment I

In this lab, you will extract a small aliquot of BL21(DE3)/pET28-GFP cells in SDS-PAGE sample buffer and then resolve the proteins by electrophoresis on a SDS-PAGE gel. SDS (sodium dodecyl sulfate, aka sodium lauryl sulfate, SLS) is an ionic detergent that binds to proteins, denaturing them (unfolding the polypeptide chain), and resulting in nominally linear molecules that sieve uniformly in the polyacrylamide gel.  Moreover, the negative charges of SDS impart a uniform charge to mass ratio to all proteins, thus causing resolution solely by length of the polypeptide chain. This works for most proteins that have a pI in the range of 5 to 9, but proteins that are extremely acidic or basic may often migrate at unexpected positions, because in addition to their size, their charge contributes to their mobility behavior in the electric field.

Fully denaturing conditions require that disulfide bonds (covalent links between 2 cysteine residues in the polypeptide chain) be broken. A reducing agent such as β-mercaptoethanol is included in the sample buffer for this purpose.



BL21(DE3)/pET28-GFP cultures (induced and uninduced)
0.5 ml microcentrifuge tubes
P-20, P-200 pipettors and tips
Narrow gel-loading tips
Electrophoresis cells
Latex gloves
SDS-PAGE sample buffer (2X)
SDS-PAGE electrode buffer
Protein size standard ladder solution
Gel fixation solution (50% methanol, 7% acetic acid)
PageBlue protein staining solution
ddH20 for destaining


1.  Obtain the 20 µl samples of induced and uninduced cells from Week 5, and to each tube add 20 µl of 2X SDS sample buffer, vortexing gently to mix, and keep on ice.

2. Prepare the electrophoresis cells with gels. Open the pre-made gel package and carefully remove the gel, which is sandwiched between 2 glass plates. Apply 2 gels to the buffer dam (one on each side) as instructed by your TA, and place it inside the buffer vessel.

3. Fill the central buffer chamber of the gel unit with electrode buffer until the top of the gel is covered. Add more electrode buffer to the bottom of the container vessel until the bottom of the gel is covered. Look for air bubbles that may become trapped under the gels, and blow them out with a Pasteur pipette that has a curved end (produced by bending after slightly softening in a flame).

4. Close the lids of the sample tubes firmly, and apply closures if available. Place the tubes in a heat-block or water bath at 85 °C for 10 min. The SDS detergent action will lyse the cells and denature proteins.

5. Microcentrifuge the samples at full speed for 2 min to pellet insoluble material.

6. Using narrow-bore pipet tips, load 15 µl of each sample into a well, and load one well with the prescribed volume of protein size standard ladder. Be careful not to allow spill-over of the samples into adjacent wells.

7. When all samples are loaded, apply the lid of the unit, connect to a power supply, and set the voltage to 125 V.

8. During the run, observe the progress of the bromophenol blue dye in the sample buffer, which runs near the front ahead of the proteins. If your protein size standard sample is pre-stained, you will be able to observe the separation of the different size proteins during the run.

9. When the run is nearing the end, prepare a dish of 100 ml gel fixation solution (a P-1000 pipet tip box works well for this purpose).

10. When they dye has reached ~1 cm from the bottom of the gel (or 1 hr), turn off the power and remove the lid.

11. Remove the gel sandwiches from the unit, and carefully pry apart the glass plates as demonstrated by your TA. Wear gloves and be careful to avoid breaking the glass! The gel will remain adhering to one glass plate.

12. Carefully pick up the gel by a corner and peel it off the glass plate, and place it in the dish of gel fixation solution, place on a shaker (~30-40 rpm) for 10 minutes.

13. While holding the gel(s) with a gloved finger, pour off the fixation solution to a disposal container, and add 100 ml ddH2O to the gel dish.

14. Microwave with no cover on high power for 20 seconds, and shake for 3 minutes. Discard the wash. Repeat twice for a total of 3 washes.

15. Add 20ml (or enough to completely cover the gel) of PageBlue Protein Staining Solution, microwave for 15 seconds on high power (do not boil). Place on the shaker (30-40 rpm) for 20 min.

16. Decant and save the stain solution (it can be reused 3 times), and wash the gel in ddH2O.

17. Observe stained bands and photograph on the AlphaImager with transmitted white light. If the gel is not destained stained well enough to photograph, cover and seal the gel dish with saran wrap and keep at room temperature until later in the day, or the following day.



Sample Buffer (SDS Reducing Buffer)
3.55 ml deionized water
1.25 ml 0.5 M Tris-HCl, pH 6.8
2.5 ml glycerol
2.0 ml 10% (w/v) SDS
0.2 ml 0.5% (w/v) Bromophenol Blue
9.5 ml Total volume
Store at room temperature.
Use: Add 50 µl β-mercaptoethanol to 950 µl sample buffer prior to use.

10x Electrode (Running) Buffer, pH 8.3 (makes 1 L)
30.3 g Tris base
144.0 g glycine
10.0 g SDS
Dissolve and bring total volume up to 1,000 ml with deionized water. Do not adjust pH with acid or base. Store at 4°C. If precipitation occurs, warm to room temperature before use.
Use: Dilute 50 ml of 10x stock with 450 ml deionized water.


Prelab Questions

1. The GFP-6His recombinant protein expressed in this study is a polypeptide chain of 247 amino acids. What is the approximate molecular weight that you expect for this protein?

2. What is the role of SDS in denaturing proteins for electrophoresis?

Experiment I

Week 7:  Metal Affinity Purification of His-tagged GFP

You constructed recombinant plasmid pET28-GFP in a way that adds a C-terminal 6-histidine (6His) peptide to GFP. Due to the ability of some divalent cations like nickel and cobalt to coordinately bind multiple His residues, poly-His containing proteins can be conveniently purified by chromatography on media containing immobilized metal ions. Normally, this process is performed using a column to contain the immobilized metal media. The crude protein extract is added to the top of the column and the proteins percolate through the matrix, where poly-His proteins bind to the metal ions. After washing the column by running buffer through the column to remove non-specifically bound proteins, the His-tagged protein is eluted by washing the column with a buffer containing a high concentration of imidazole, a compound that competes with His for binding to metal ions. Figure 5 depicts the His-cobalt interactions in the TALON® metal affinity system (Clontech, You can find the a pdf file on the Blackboard site that contains the user manual for this system.

metal affinity system


Figure 5. Diagram of TALON® metal affinity system. A, TALON resin; a Sepharose bead bearing the tetradentate chelator of the Co2+ metal ion. B, Polyhistidine-tagged recombinant protein binds to the resin.

In order to save time and expense, you will instead use a mini-batch procedure with small amounts of TALON resin in 1.5 ml microcentrifuge tubes. The crude bacterial cell extracts will be mixed with the resin to allow binding of GFP-6His, and washing will be performed by sedimenting the resin in a microcentrifuge and resuspending it in washing buffer.


BL21(DE3)/pET28-GFP cell extracts from Week 5

TALON® metal affinity resin equilibrated in binding buffer
1.5 ml microcentrifuge tubes
P-200 and P-1000 pipettors and tips
Disposable gloves
Washing buffer
Elution buffer

Method: TALON metal affinity chromatography

1. Obtain your induced bacterial cell extract from Week 5, stored at –20 °C. If they are too viscous to pipet with a P-200, they will need to be treated with DNase I (10 µg/ml).

2. Obtain a 50 µl aliquot of TALON metal affinity resin in 1.5 ml microcentrifuge tube from your TA.

3. Transfer the cell extract into the tube with the TALON metal affinity resin, and invert to mix.

4. Keep at room temperature for 10 min, inverting to mix every 30 sec.

5. Allow the resin to settle to the bottom of the tube, illuminate the tube with long-wave UV lamp, and record your observations of relative fluorescence of the resin vs. the supernatant.

6. Centrifuge the tube for 10 sec., and remove the supernatant with a P-1000.

7. Add 1 ml of wash buffer and resuspend the resin by inversion of the tube, or vortexing gently. Let the resin settle, and again observe the relative amounts of fluorescence in the resin vs. supernatant.

8. Repeat step 6.

9. Add 0.5 ml of elution buffer and resuspend the resin. Let the resin settle, and again observe the relative amounts of fluorescence in the resin vs. supernatant.



Binding buffer:  10 mM Tris-HCl, pH 8.0, 50 mM NaCl

Washing buffer:  Binding buffer + 10 mM imidazole

Elution buffer:  Binding buffer + 150 mM imidazole

Prelab questions


1. Describe the principle of metal affinity purification of His-tagged proteins.

2. What is the role of imidazole in the process?



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