Heya guys, long time no see. In a blink of an eye, 13 weeks of SIP/MP have already past. Next week is our 3rd campus discussion and hope to see you all soon. By the way, please refer to Miss Chew's blog for updates regarding whether there is a blog quiz.
Today, I am going to share on one of two methods of pre-electrophoresis preparation steps that are required for my project. The gels that i will be running are 1D-Zymogram, 2D-gel and 2D-Zymogram and the pre-electrophoresis preparation is essential for running the gels mentioned above. To refresh your memory, Zymography is an electrophoresis technique that is used in the detection of protease activity under non-denaturing conditions. It is performed on a zymogram gel, which incorporates the use of a substrate that is copolymerized with polyacrylamide gel . Proteases that catalyze Gelatin , Caesin or Fibrin as a substrate will show up as clearings against a dark blue background after staining with Commassie brilliant blue. (Please read on previous post entry for more information regarding zymogram )
Wthout further ado, the two Pre-electrophoresis preparation steps are Bradford assay and TCA (Trichloroacetic acid) precipitation. In this post, the focus will be on Bradford assay.
Principle of Bradford assay: Bradford assay is a protein colormetric assay that will produces a colour change if proteins are present. The coomassie dye is originally red in colour. However in the presence of protein binding, it changes colour and stabalises into coomassie blue, resulting in an absorbance shift. This happens because of 2 bond to bond interactions taking place. The red form of commassie dye donates free proton to ionized groups on protein disrupting its conformation. This leads to hydrophobic heads of the proteins being exposed. The expose hydrophobic pockets on protein chain bind to non-polar region of the dye by van der waals force. Hence, this positions the positive amine groups closely to negative charge of the dye. Ionic interaction further strengthens the bond and ther is blue coomassie dye. Binding of the protein stabilises blue form of coomassie dye and the complex is measured for protein concentration by absorbance reading at 595nm. If no protein is bound to the dye, the cationic (unbound form) are green or red while binding stabalises the anionic (bound form) are blue in colour.
By using Bradford assay, the periplasmic protein concentration in the supernatant can be determined. The mass of periplasmic proteins remained at a constant at 10ug. By knowing the mass and protein concentration of the protein, the volume of protein sample to be loaded into the wells of the gel can be determined. This is because of the formula: Concentration (ug/ul) = Mass (ug) / Volume (ul). The volume can be found by manipulating the formula: Volume = Mass / Protein concentration (determined by Bradford assay).
Methods and Explanation
1. Warm up Bradford dye reagent to room temperature
It will not affect the sample at cold temperature and works optimally at room temperature
2. Pour Bradford dye reagent to plastic tray and cover with aluminium foil
Bradford reagent is light sensitive and cannot be exposed to light
3. Prepare the centrifuge tubes and mixed in the appopriate standards ( Milli Q + BSA)
Allows a calibration curve to be plotted
4. Prepare the sample in 5X dilutions
This ensures that there will be enough sample left after pipetting, hence need to prepare excess
5. Centrifuge standards and sample (short spin for 7 seconds)
To thoroughly mixed the milli Q and BSA/sample
6. Pipette 5ul of sample or standards in triplicates into each well using reverse pipetting (microtitre plate)
Ensures average readings can be taken after spectrophotometry for accurate results
7. Add 250uL bradford reagent into each well using multichannel pipette and reverse pipetting
Reverse pipetting to ensure exactly 250uL bradford is actually added and not more or less. It also prevents air bubbles forming
Allows binding of bradford to proteins for spectrophotometry
8. Remove air bubbles present using a pipette tip dipped with ethanol
Prevent air bubbles in samples, lead to inaccurate results
9. Cover microtitre plate with aluminium foil and incubate 30 minutes
Allow the reaction to occur at room temperature
10. Set up the spectrophotometer
To measure absorbance reading at 595 nm and to quantitate amount of proteins
The need to use BSA Standards: BSA standards are prepared at the concentration of 0, 0.1, 0.25, 0.5, 0.75mg/ml to obtain a linear range (standard curve). When the commassie blue dye binds to the protein, absorbance reading is read using spectrophotometer and absorbance reading is interpolated to the linear range of Bradford Assay. The protein concentration can thus be obtained.
That's all for now. Thanks you for reading my post and have an enjoyable next 7 weeks!
From: Benjamin Ma
Class: TG01
0606181F
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12 comments:
hey hey
why must the pipette tip be dipped in alcohol to remove any bubbles?
Maya
tg02
Hello maya,
THe presence of air bubbles will affect the absorbance readings by the spectrophotometer. THus, there will be inaccurate quantitation of proteins. Air bubbles in any case, must be removed and a pipette tip dipped with alcohol is usually used to burst the air bubbles. The alcohol is to make the tip sterile and prevents introduction of contaminants into the proteins.
Thanks!
From: Benjamin ma
TG01
0606181F
hello!
got a question.
What is reverse pipetting? why can't the normal pipetting be used?
Thanks.
Li Ping
TG o2
hi ben,
what's a multichannel pipette and reverse pipetting? and how does reverse pipetting ensure the exact amount is being pipetted? isn't the normal pipetting we do accurate? thanks.
Malerie
TG02
Hello Li Ping and Malarie,
Reverse Pipetting (rP): A selected volume plus excess is aspirated into the tip. After delivery the excess volume remains in the tip and is discarded.
In technical terms, it means that when you suck up the volume using the pipette, u press the plunger till the 2nd stop before fully releasing it. This will suck up the volume in excess of your intended volume. Thus when you pipette, you do not press the plunger fully to release your volume, instead you press only until the 1st stop. This ensures that only the intended volume will be pipetted out.There will be excess volume left in the pipette tip after pipetting because you would have sucked up excess volume in the 1st place.
The importance of this, is that it prevents air bubbles. Why? DO you notice that when you use normal pipetting, you depress the plunger till the 2nd stop to fully release the intended volume and the last drop will almost certainly be a drop of air bubble? Reverse pipetting prevents this from happening as the last drop that is released is not air bubble, meaning that the full intended volume will be pipetted out. This is because of the excess volume sucked up in the first place.
For Bradford assay, it is important not to have any air bubbles as it will affect the protein quantitation (incorrect quantitation). Thus Reverse pipetting is used to prevent introducing air bubbles.
Multichannel pipette is like a normal pipette with the only difference that it can fit many pipette tips all at once (can fit maximum 8 tips).
For a picture of a multichannel pipette please refer to this website:
http://www.morningtech.com.hk/biohit.htm
From: ben
hello ben, i wld like to ask if the standards u use are commercial or DIY. and what happens if the standards fail? what do you do?
Yuxuan
Hi Yu XUan,
1) The standards are commercially available from Biorad and the standards i used are 2mg/ml Bovine Serum Albumin
2)Standards can fail when pipetting skills are not done properly, BSA standards have expired, Contamination of the microtitre plates, etc
Thus, it is very important to prepare the standards correctly and minimize errors as the protein concentration is interpolated from the absorbance reading of the linear range of BSA standards. If the BSA standards are wrong, the absorbance readings for the protein will be wrong and the quantitation of protein will also be wrong.
If the standards fail, just redo the standards.
Thanks!
From: Benjamin Ma
Class: tg01
0606181F
Heys Benjamin,
regarding the reverse pipetting, does it mean that you all have to use filter tips instead of normal pipette tips? Because if i use a 200 microliters pipette and i press it to the 2nd stop to suck up more than 200 microliters of the liquid, i think it will kinda be too much inside the pipette tip and contaminate the pipettes?
Thanks for clarifying
Jean Leong
TG02
Hi Jean,
Yes for reverse pipetting, it is best to use a filter tip instead of an orignal pipette tip. For protemoics work, all work is performed usually using filter tips to reduce the chance of contamination to the pipettes.
And if we want to pipette out 200ul of sample for example, we cannot use a 200ul pipette (max vol) to suck the sample. This will spoil the pipette as it is not advisable to use a pipette at its highest volume. If we want to pipette out 200ul of sample, we should use a 100-1000ul range pipette as 200ul falls within the range. Thus, if we follow this method of selection of pipette, it is unlikely that too much volume of sample will be sucked up in the first place, if the pipettes selected are the suitable ones.
In the case whereby the pipettes are contaminated due to sample overload within the pipette tip (if a filter tip is not used), it is advisable to spray 70% ethanol to clean the pipette after use to reduce contamination
Thanks!
From: benjamin Ma
Class: Tg01
0606181F
Pasteur Pipettes: These are made of glass so that one can get a clear view of the liquid being transferred. They are reusable with the pasteurization process that is affected for cleaning.
Pipette
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