PROTEIN EXTRACTION FROM TRANSFORMED E. COLI CELLS:

Green fluorescent protein, originally isolated from a jelly fish, has become a powerful and widely used tool in biotechnology research. As the name suggests, GFP is fluorescent, meaning that when the protein is excited by an energy source like ultraviolet (UV) light, it will glow. In this case the fluorescence is a visible green (~500nm). We can use this property to see when and where the protein is expressed by the presence of green fluorescent light.

This lab will analyze the proteins present in your sample. Protein analysis can be done by migrating proteins through a gel and comparing protein sizes, we call this process SDS-PAGE. This stands for sodium dodecyl sulfate polyacrylamide gel electrophoresis. The gel is made of polyacrylamide, different from DNA electrophoresis gels, made of agarose. Proteins don’t have a uniform charge, like DNA (negatively charged) which is why it is important to coat the proteins with a buffer (SDS) so they will have a uniform mass to ratio charge. When SDS coats the proteins, it gives them a net negative charge, in a way so that the proportion of protein binding to SDS is relative to molecular mass.

This activity will allow you to see the difference in cellular proteins in E. coli regulated to express GFP and compare this to cells which do not express the protein. First, we extract all of the protein from the bacteria with Laemmli’s buffer, which contains detergents, to break open the bacterial cells and extract all of the protein from the cells. We then determine protein concentration to find out how much protein is in our samples. The proteins will be separated by size using SDS-PAGE. Once the gel has run, apply UV light to see if the band of green fluorescent protein reveals itself and then stain the separated proteins within the gel with a stain specific to proteins, Coomassie Blue. Though you should only see one green fluorescent band under UV light, the coomassie stained gel will allow you to identify differences between the protein profiles under different conditions.

MATERIALS/EQUIPMENTE NEEDED

For the class: Heat block Centrifuge Spectrophotometer - optional Vortex

For each student group: Four 1.7ml microcentrifuge tubes 10 ul loop 2 tubes of 500 μl LB broth Camiolo buffer

1. Label two 1.7ml microcentrifuge tubes, the one in LB+amp will be labeled White and the one

with LB+amp+arabinose will be labeled Green.2. Scoop ½ of glowing colony and ½ of non-glowing colony and add to 500 μl of LB broth

each.

Optional: Dilute sample 1/10 to measure OD. Add 900 μl of the broth used in the culture for blanking. Add 100 μl of sample and measure

OD. Multiply the spec reading number by 10 to get the real OD. OD must be in a range of 2.5-4.0 to get a good concentration of protein.

3. Centrifuge tubes for 5 min at 13000 rpm. 4. Dump the liquid out without dislodging the pellet. 5. Add 500 μl of Camiolo’s buffer to each sample. Vortex tubes for one full minute. 6. Label two microcentrifuge tubes

● White, + heat (W+)

● Green, + heat (G+)7. Transfer 250 μl of mixture in the Green tube to the G+ tube, do the same for the White tube. 8. Leave the two “no heat” tubes in a rack on your bench at room temperature and place the

two “+ heat” tubes in a hot block at 95C for 5 min. After heating, cool on the bench for 5 minutes and then centrifuge all four tubes for 3 mins to pellet cell debris and transfer the supernatant to a new labeled tube (1.7ml centrifuge tubes).

DETERMINING PROTEIN CONCENTRATION

Prepare Samples: You will be determining the protein concentration of your samples in groups of 4 students using protein standards of known concentrations

1. Label 1 cuvette identifying YOUR specific protein sample. For the protein standards, label 8 cuvettes with the known concentrations of the protein standards (see the photo below).

2. Add 1 ml of Bradford reagent into each appropriately labeled cuvette.

3. Pipette 20 μl of the known protein concentration standards, into each appropriately labeled cuvette. Place lid on to cuvette and invert several times.

4. Incubate at room temp at least 5 minutes, not longer than 1 hour, and measure absorbance at wavelength of 595 nm.

5. While the protein standards are incubating, prepare your protein samples by pipetting 20 l of your insect protein sample into the appropriately labeled cuvette. Place lid on to cuvette and invert several times. Incubate at room temp at least 5 minutes, not longer than 1 hour, and measure absorbance at wavelength of 595 nm.

Collecting Data

6. After incubation, clean off the clear non-ribbed side of the cuvette labeled 0.00 mg/ml using a Kimwipe, and place the cuvette in the Spectrophotometer. Line up the clear sides with the arrow. Push the Zero button and remove cuvette when you see 0.00. You should only have to Zero one time. Preforming this step “blanks” the instrument, making all the subsequent measurements relative to this “blank.”

7. With the other cuvettes, wipe off the sides using a Kimwipe and place them in the Spectrophotometer. Line up the clear sides with the arrow. Push the Test button and record the absorbance values.

8. Test your tissue samples in the same manner as described in step 7.

DETERMINING PROTEIN CONCENTRATION OF SAMPLES

Standards:Protein concentration

(mg/ml)Absorption

1.0 (blank) 00.1250.250.500.751.001.502.00

Samples: Raw data Sample name Absorption

Use Excel to find out the equation for the linear best fit line from the absorption data of the protein standards (these instructions are for Excel in Windows 7, yours may differ):

1. Open an Excel spread sheet containing a column for protein standard concentrations, and an additional column with their corresponding absorbances (as shown above).

2. Highlight the values, click on insert, and choose the scatter plot with no line function.

3. Go to chart layouts, choose trendline. In order to see the equation, go into trendline options and click “display equation on chart”.

4. Determine protein concentrations of your insect samples using displayed equation, y=mx+b where y is your Absorbance value of your insect protein sample, and you are solving for x. Therefore x is your protein concentration in mg/ml.

Samples: Processed data

Sample name Absorption Protein concentration (mg/ml)

PROTEIN STANDARDS AND PROTEIN SAMPLE ANALYSIS

If the absorbance value of your sample falls beyond the maximum absorbance value of the protein standard of the highest concentration, it will be necessary to add less of your protein sample for the protein assay.

For protein samples that are greater than >0.5 A from the protein standard of highest concentration,

dilute your protein sample by ¼ for the Bradford protein concentration analysis, by adding 5 μl of

your protein sample and 15 μl of DI water into your cuvette with 1ml of Bradford reagent. Then measure the Absorbance (after a 5 min incubation), determine the concentration, and multiple the concentration by 4 in order to calculate the concentration of protein in your sample.

For protein samples with values between 0.2 and 0.5 A higher than the Absorbance of the highest

concentration protein standard, dilute your protein sample by ½ by adding 10 μl of your protein

sample and 10 μl of DI water into your cuvette with 1ml of Bradford reagent. Then measure the Absorbance (after a 5 min incubation), determine the concentration, and multiple the concentration by 2 in order to calculate the concentration of protein in your sample.

Only the protein samples with absorbance values that fall beyond the absorbance of the protein standard of highest concentration need to be diluted. In other words, if your protein sample absorbance falls within the range of the protein standard absorbance values, it will not be necessary to dilute your sample.

ANALYZING DATA AND CALCULATING DILUTIONS FOR SDS-PAGE

You will want to load an equal amount of protein for your protein gel. This is most easily achieved if you start with all of your samples at the same concentration. The final concentration of all of your samples will be determined by the sample that had the lowest concentration unless it is less than 0.25 mg/ml

Calculation for sample dilution Dilute samples in Laemmli’s Buffer instead of Camiolo because Laemmli’s Buffer has

bromophenol blue which dyes the sample making it helpful to visualize when loading the gel. Expect protein concentrations to be in a range of 0.8 to 1.5 mg/ml. Make the necessary

calculations to have an end concentration of 0.5 mg/ml to load the gel.

For each sample, calculate how much of the sample to add to Laemmli’s Buffer to make

0.1 ml (= 100 μl) final volume. For example: if the lowest measured concentration sample was 0.5 mg/ml and you had another sample that was 1.2 mg/ml, use the following formula to make a final

volume of 0.1 ml (100 μl) of 0.5 mg/ml protein:

(final concentration) X (final volume) = (sample concentration ) X (sample volume)

(final concentration) X (final volume) --------------------------------------------- = sample volume

Sample concentration

(0.5 mg/ml) (0.1 ml) = (1.2 mg/ml) (x)

(0.5 mg/ml) (0.1 ml) ----------------------- = (x) (1.2 mg/ml)

x = 0.041 ml = 41 μl

Volume of Laemmli’s Buffer needed = (100 μl – sample volume)

Volume of Laemmli’s Buffer needed = (100 μl – 41 μl) = 59 μl Laemmli’s Buffer for a final concentration of 0.5 mg/ml for your more concentrated sample.

Sample name Final protein concentration

Volume of sample needed

Volume of Laemmli’s

needed

Why is it important to load the same amount of protein in your samples?

Protein Separation and visualization using SDS-PAGE:

1. Gather the gel running apparatus which is the clear plastic tank, the beige gel holder, and lid with wires on top. Two to four gels can share a power supply. 

2. Prepare the gel by removing the plastic cassette from the package. There will be a little buffer in the package which can be eliminated. Place the cassette containing the gel into the beige gel holder with the taller plastic plate of the cassette facing the outside of the holder. If the numbers written on the top of the gel cassette are backwards then the gel is in backwards and the apparatus won’t work. If you are unsure if the gel is in correctly, PLEASE ASK. 

On one side of the gel holder is a buffer dam.  If you are running one gel, leave the dam in place.  If you are running two gels, remove the dam.  (DO NOT THROW THE DAM AWAY!) Also make sure the buffer dam is in correctly otherwise the buffer will leak out and the apparatus won’t work. In this case the words “Buffer Dam” SHOULD be written backwards. If you are unsure if the buffer dam is in correctly, PLEASE ASK. 

3. Clamp the gel and buffer dam in the gel holder. Fill the inner chamber with Teo-Tricine SDS running buffer to the top and check to see if you have any leaks (gels sometimes leak from the bottom if they are not seated correctly).  If a leak is detected, empty the buffer into the tank, as this will be your outer buffer, and refill the inner chamber and look for leaks.

4. Place the gel holder into the clear plastic tank. Make sure that the red side of the gel holder matches with the red sticker of the gel tank. Then fill the outside of the chamber with Teo-Tricine SDS to the “2 gel” line on the clear plastic gel box. 

5. These gels are stored in a viscous solution that will make loading your samples into the wells difficult. Rinse out the wells with the buffer in the chamber with a p1000 and 1 ml of buffer releasing the solution above the wells such that it will displace the solution in the well.  Then run a small amount of 

6. Two groups will share a gel. Load the gel with 5 μl of molecular weight marker and 30 μl of your samples of extracted protein in the below order.  You will need to vortex your samples again before pipetting, even then the mixture may be extremely viscous.  Take care to get 30 μl in your tip which should look like this: 

Lane 1: EMPTYLane 2: Group 1 Green, +heatLane 3: Group 1 Green, no heatLane 4: Group 1 White, + heatLane 5: Group 1 White, no heatLane 6: Group 1 MarkerLane 7: Group 2 MarkerLane 8: Group 2 White, no heatLane 9: Group 2 White, + heatLane 10: Group 2 Green, no heatLane 11: Group 2 Green, + heatLane 12: EMPTY

7. Place the lid on the tank. Plug the wires from the lid into the power supply pay attention to match the colors.  Set the power supply to 100 volts and start the run.  Let the gels run approximately 15-20 min and turn the power supply up to 130 volts.  Turn up the power to 150 volts in another 10 min. Let gel run until smallest fragment in the MW Marker lane is a few cm from the bottom. Total run time will be 40-50 min.

8. Perform the following steps to retrieve your gel from the gel apparatus:a. To stop running the gel, turn off the power supply and remove the wires from the power

supply. b. Remove the lid from the tank and remove the gel holder from the tank. Carefully pour the

buffer on the inside of the gel holder down the drain. c. Release the gel holder clamps and remove the gel cassette from the gel holder. 

9. Before going any further, lay the gel cassette down on the counter with the short side down. Using a UV light, look for the glowing bands of GFP on the gel. You may need to turn the lights off to see it. If you see a green band light up than this is the properly folded GFP. (think about what properly folded means in terms of the ability to glow.) Take a picture or take note of which band(s) is(are) glowing. Also make note as to the approximate size based on the protein molecular weight marker. 

10. Use a flathead tool to crack open the gel casing and carefully (gel is thin and delicate-don’t poke it with the tool) separate the two halves of the cassette. Be careful not to jab the gel, do not put the tool in between the casing more than you need. The gel will attempt to cling to one side or the other of the plastic casing. Touch it gently to get it to lay down on the bottom plastic half. BE CAREFUL WITH GEL, IT IS VERY EASY TO TEAR!

11. Carefully cut the gel into two halves separating lanes 1-6 from 7-12. Do this by cutting right between the markers in lanes 6 and 7. After the halves are separated, move the gel halves to separate staining trays. Make sure you don’t flip the gels over when you move them. The groups can now work separately.

12. Pour enough Coomassie blue stain to cover your gel in the staining tray. This is a powerful dye, be careful not to splash it on self or clothes, as it will stain any and all proteins it touches (including yours!).

13. Stain the gel, shaking gently for 30 minutes. After 30 minutes, recycle the stain by pouring it back into the bottle.

14. Now pour destain into the tray completely covering the gel.  Add two pieces of balled up kimiwipes to your destain tray to absorb the dye.  Destain for 60 min to overnight.  Pour the used destain into the destain waste container.

15. Place the gel on a light box for examination. Take note of the difference in the bands that are different between the “Green” and the “White” and between the “+heat” and “no heat”

Remember – The distance the protein band has traveled is related to the size of the protein. Also, how dark the band is tells the concentration of the protein– the darker the band, the more of that protein there is.

How many green fluorescent proteins bands did you see? Why do you think that is?

What is the approximate size of GFP?

Did you find that GFP was expressed in your sample? Why or why not?