The zip tip protocol is used to clean up the biological samples after protein digestion. Its main purpose is to get rid of the chemicals used in protein digestion, such as DTT, IAA, salts, SDS, etc.
PREPARATION OF SOLUTIONS
Three solutions need to be prepared in order to proceed with the procedure: solution A, solution B, and solution C.
Solution A: Rinse Solution (0.1% formic acid 100%)
Final volume:1000 ul
(100%)V=(0.1%)(1000 ul)
V=1 ul FA 100%
*Obtain 1 ul FA 100%, 999 ul of water*
Solution B: Wet Solution (0.1% FA 100%, 50% acetonitrite)
Final Volume: 1000 ul
*Obtain 1 ul FA 100%, 500 ul ACN, 499 ul water*
Solution C: Elution Solution (0.1% FA 100%, 60% ACN)
Final volume: 1000 ul
*Obtain 1 ul FA 100%, 600 ul ACN, 399 ul water*
PROCEDURE
(Added 70 ul of water to the dried biological samples)
1. Set pipette to 10 ul.
2. Equilibrate tip with wet solution B. Aspirate wet solution into the tip. Dispense to waste. Repeat twice.
3. Aspirate rinse solution A. Dispense to waste. Repeat once.
4. Bind peptides. Aspirate and dispense the sample 10 to 20 times (depends on sample complexity).
5. Aspirate rinse solution A and dispense to waste. Repeat 7 to 10 times.
6. Elute peptides. Set pipette to 5 ul. Aspirate 5 ul of elution solution C into the tip and place in a clean vial. Repeat once or twice.
Dilute to desired final concentration with spraying solution: 0.1% FA, 50% ACN (never actually made this because for my lab it is unnecessary, my mentor and I just dried up the samples again after these steps)
Tuesday, February 15, 2011
Monday, January 24, 2011
Trypsin Digestion Protocol~
Proteins are digested and chopped up into little pieces for further experiments. In order to digest proteins, three solutions are used: DTT, IAA, and NH4HCO3. A type of enzyme will be used to cut the protein at a specific amino acid.
PREPARATION FOR DIGESTION
1. DDT (Dithiothreitol) breaks up disulfide bonds formed by the bonds of cysteine. So, DTT is used to "unfold" the proteins and eliminate the bonds that make the protein folds.
Prepare 1 ml of 100 ml of DTT
molar weight: 154.25 g/m
100 mM in 1000 ml= x mM in 1 ml
x= 0.1 mM= 1*10^-4 M
x=0.01542 g =15.42 mg
2. IAA (Iodoacetamids) stabilizes the disulfide bonds broken up by DTT. IAA prohibits the disulfide bonds to bond again.
1. DDT (Dithiothreitol) breaks up disulfide bonds formed by the bonds of cysteine. So, DTT is used to "unfold" the proteins and eliminate the bonds that make the protein folds.
Prepare 1 ml of 100 ml of DTT
molar weight: 154.25 g/m
100 mM in 1000 ml= x mM in 1 ml
x= 0.1 mM= 1*10^-4 M
x=0.01542 g =15.42 mg
2. IAA (Iodoacetamids) stabilizes the disulfide bonds broken up by DTT. IAA prohibits the disulfide bonds to bond again.
Prepare 1 ml of 100 ml of IAA
molar weight: 184.96 g/m
100 mM in 1000 ml= x mM in 1 ml
x= 0.1 mM= 1*10^-4 M
x=0.018496 g =18.496 mg
3. NH4HCO3 is a buffer that stabilizes and helps the enzyme to rise to its maximum capability to clip and cut the protein. A certain enzyme functions better with certain pH. In this case, NH4HCO3 has a pH that works best with trypsin.
Prepare 1 ml of 100 ml of NH4HCO3
molar weight: 79.06 g/m
100 mM in 1000 ml= x mM in 1 ml
x= 0.1 mM= 1*10^-4 M
x=0.007906 g =7.906 mg
PROCESS OF PROTEIN DIGESTION
Two proteins used: BSA and apomyoglobin
1. BSA contains disulfide bonds; thus DTT and IAA are added to break the bonds.
10 ul of BSA (3 ug/ul)given, add:
5 ul DTT,
25 ul NH4HCO3, and
20 ul water.
>incubate at 50 C for 45 minutes
25 ul IAA, and
25 ul NH4HCO3.
>sit in darkness for 1 hour
The enzyme is then added according to a protein to enzyme ratio. In this case, I used 50:1 (protein to enzyme in ug)
BSA given: 10 ul of 3ug/ul
50:1 = (3 ug/ul)*10 ul: x
x= 0.6 ug
trypsin stock concentration: 0.1 ug/ul
0.1 ug/ul*x= 0.6 ug
x=6 ul
>add 6 ul of trypsin in the BSA solution and incubate at 37 C for 10 hours
2. Apomyoglobin only undergoes the protein digestion process
Apomyoglobin given: 20 ul of 1 ug/ul, add:
2 ul trypsin, and
22 ul NH4HCO3.
> incubate at 37 C for 10 hours
Sunday, January 16, 2011
Bradford Protein Assay~
Happy LATE New Years~ I am sorry, I been busy studying for school (finals is coming up) and organizing data from the lab into my notebook. However.....
Recently, in my lab at CalTech, I did a lab on Bradford protein assay. It is a procedure based on an absorbance shift of the Coomassie Brilliant Blue G250 dye. When the dye binds with proteins, it convert from its original red form into a bluer form. The bound form of the dye has an absorption spectrum maximum held at 595 nm. Certain concentration of detergents, which is used to lyse cells, interferes with the Bradford assay.
I used Bovine Serum Albumin (BSA) as a known protein to create a calibration graph of concentration versus absorbance.
Prepare BSA solutions:
I was given BSA solution of 1 ug/ml, and I need to prepare BSA concentrates at 2 ug/ml, 5 ug/ml, 7 ug/ml, 10 ug/ml, 20 ug/ml, and 30 ug/ml.
All of the solutions have the final volume of 1000 ul or 1 ml.
I, then, input 2 ul of the solution into the nanometer, which has the Bradford protein assay procedure pre installed into the computer.
A calibration graph is formed on the concentration versus absorbance. The graph uses the Beer-Lambert Law:
Possible sources of error:
I performed the calibration graph and the actual experiment on two different days, so the results are not as accurate as they possibly are.
There are some samples with 0 concentration. One thing is that I did not sanitize the nanometer correctly or air went into the machine while the experiment is performed.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
This conclude the experiment. XD
Recently, in my lab at CalTech, I did a lab on Bradford protein assay. It is a procedure based on an absorbance shift of the Coomassie Brilliant Blue G250 dye. When the dye binds with proteins, it convert from its original red form into a bluer form. The bound form of the dye has an absorption spectrum maximum held at 595 nm. Certain concentration of detergents, which is used to lyse cells, interferes with the Bradford assay.
I used Bovine Serum Albumin (BSA) as a known protein to create a calibration graph of concentration versus absorbance.
Prepare BSA solutions:
I was given BSA solution of 1 ug/ml, and I need to prepare BSA concentrates at 2 ug/ml, 5 ug/ml, 7 ug/ml, 10 ug/ml, 20 ug/ml, and 30 ug/ml.
Concentration (ug/ml) | BSA solution (based on 1 ug/ml) | Bradford reagent dye |
2 | 2 ul | 998 ul |
5 | 5 ul | 995 ul |
7 | 7 ul | 993 ul |
10 | 10 ul | 990 ul |
20 | 20 ul | 980 ul |
30 | 30 ul | 970 ul |
All of the solutions have the final volume of 1000 ul or 1 ml.
I, then, input 2 ul of the solution into the nanometer, which has the Bradford protein assay procedure pre installed into the computer.
^ Instrument I used.
^The display.
A calibration graph is formed on the concentration versus absorbance. The graph uses the Beer-Lambert Law:
Absorbance=Ebc
(where Eb are constant, sometimes written as k and c as the concentration)
The calibration graph look something like this:
Notice the r squared variable. In statistics, it is known as the coefficient of determination. It determines the linearity of the graph. In other words, the closer the variable is to 1, the more linear it is.
Using the calibration graph, I was able to determine the concentration of several biological samples based on the absorbance of the samples and the corresponding absorbance over concentration.
The following are a list of biological samples, which I performed the Bradford assay on in order to find the concentration:
Possible sources of error:
I performed the calibration graph and the actual experiment on two different days, so the results are not as accurate as they possibly are.
There are some samples with 0 concentration. One thing is that I did not sanitize the nanometer correctly or air went into the machine while the experiment is performed.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
This conclude the experiment. XD
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