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cytochrome C Western Blot


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#1 cigdem

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Posted 26 November 2009 - 02:18 PM

Hi everybody
I'm doing western blot for cytochrome c. the molecular weigth of cytochrome c is 13 kDa but i always get 2 strong bands with high molecular weight between 40-60 kDa and a very weak signal foe 13kDa. I read somewhere that cytochrome c can form dimers, tetramers even pentamers but I'm not sure if it is a complex of cytochrome c because i'm doing SDS-PAGE . I use %5 beta-mercaptoethanol in sample buffer as a reducing agent. Someone suggested me to increase the concentration of reducing agents but i don't know if i can use an upper concentration or i have no idea about the maximum concentration of this agents for sample buffer. If anyone can help me, i will be glad.
Thanks

#2 mdfenko

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Posted 26 November 2009 - 09:57 PM

you can try 10% mercaptoethanol or 10-20mM dtt.
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#3 Prep!

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Posted 26 November 2009 - 10:06 PM

you can try 10% mercaptoethanol or 10-20mM dtt.



i agree but also it will help if the aggregation is disulphide linked!!! hydrophobic interactions may be more difficult.. sometimes they dont break even by boiling!!! may be a urea gel can help.. not sure though..
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#4 cigdem

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Posted 27 November 2009 - 03:37 AM

you can try 10% mercaptoethanol or 10-20mM dtt.



i agree but also it will help if the aggregation is disulphide linked!!! hydrophobic interactions may be more difficult.. sometimes they dont break even by boiling!!! may be a urea gel can help.. not sure though..


Thank you .

#5 medchemgirl

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Posted 03 December 2009 - 10:21 AM

Did you figure out the problem, I would like to know how u solved it.

you can try 10% mercaptoethanol or 10-20mM dtt.



i agree but also it will help if the aggregation is disulphide linked!!! hydrophobic interactions may be more difficult.. sometimes they dont break even by boiling!!! may be a urea gel can help.. not sure though..


Thank you .



#6 Tai

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Posted 05 December 2009 - 02:33 AM

It is also the same problem to me.

I tried to use in various concentration of dtt or B-mercap, or try to boil in different time point, but it doesn't work <_<

I have just found this faqs, I may try soon if I have time.

Q. How do I avoid protein oxidation after reducing SDS treatment?

A. Reducing agents such as dithiothreitol (DTT) or 2-mercaptoethanol cleave disulfide bonds into free sulfhydryl (SH) groups to allow proteins to unfold completely. However, the reducing agent can be oxidized during sample heating which may allow these disulfide bonds to reform, leading to the appearance of ghost bands in the high molecular weight area or precipitation at the sample application point.
Blocking the reduced SH groups can prevent disulfide bonds from reforming. One common way to do this is to alkylate with iodoacetamide. Iodoacetamide also alkylates residual DTT to prevent point-streaking and other artifacts in horizontal flatbed gels.

The recommended amount of iodoacetamide is 2.0-2.5% (w/v) in the sample. The iodoacetamide should be added after boiling the reduced sample, but prior to loading the sample onto the SDS-PAGE gel. Alternatively, for first-dimension IPG strips, perform a second equilibration step for the IPG strips with an iodoacetamide solution in SDS equilibration buffer (without DTT). The alkylation reaction should take 15-20 minutes at room temperature.

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#7 cigdem

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Posted 10 December 2009 - 10:23 AM

Did you figure out the problem, I would like to know how u solved it.

you can try 10% mercaptoethanol or 10-20mM dtt.



i agree but also it will help if the aggregation is disulphide linked!!! hydrophobic interactions may be more difficult.. sometimes they dont break even by boiling!!! may be a urea gel can help.. not sure though..


Thank you .


I couldn't solve the the problem. I used sample buffers with different concentrations of DTT and 2-mercaptoethanol and i boiled the samples 10 minutes but it didn't work. I have 2 strong band with high molecular weigth. But also i figure out that DTT works better than 2-mercaptoethanol. at least i could get stronger signal in the right position :P

#8 mdfenko

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Posted 10 December 2009 - 12:24 PM

I couldn't solve the the problem. I used sample buffers with different concentrations of DTT and 2-mercaptoethanol and i boiled the samples 10 minutes but it didn't work. I have 2 strong band with high molecular weigth. But also i figure out that DTT works better than 2-mercaptoethanol. at least i could get stronger signal in the right position :D


rather than boiling (95-100C) you can try 65-70C for 5-15 minutes. this way proteins will denature with less possibility of aggregation.
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