How would you increase separation on PAGE? - (May/02/2006 )
QUOTE (Kathy @ May 4 2006, 06:12 AM)
thank you so much for your replies!
Don't decrease DTT, I think it won't do anything good.
i dont want to decrease DTT, i didnt add it, im thinking if I should add it...
i have attached the photo here....as you can see very poor resolution....sorry that marker (right) and my protein (left) have so much space in between...i wanted to load something in between but it didnt work out...
...so in the protein you might see two bands...lower one is not my interest....but the upper one (which should be two) is what i want to see clearly..in the marker two bands 97Kda and 66Kda are smashed together in one band (usually they are far away in SDS-PAGE..other bands are fine i guess..
Missele,was also thinking about waiting a little bit longer, i turn off the gel when BPB reaches the end...how do i know for how long to wait?
N.B sorry it seems im not able to load the attachment anyway all the bands are in the most upper part of the gel....they are not even in the middle....
thank you for the suggestions again....
QUOTE (K.B. @ May 3 2006, 03:06 AM)
Don't decrease DTT, I think it won't do anything good.
i dont want to decrease DTT, i didnt add it, im thinking if I should add it...
i have attached the photo here....as you can see very poor resolution....sorry that marker (right) and my protein (left) have so much space in between...i wanted to load something in between but it didnt work out...
...so in the protein you might see two bands...lower one is not my interest....but the upper one (which should be two) is what i want to see clearly..in the marker two bands 97Kda and 66Kda are smashed together in one band (usually they are far away in SDS-PAGE..other bands are fine i guess..Missele,was also thinking about waiting a little bit longer, i turn off the gel when BPB reaches the end...how do i know for how long to wait?
N.B sorry it seems im not able to load the attachment anyway all the bands are in the most upper part of the gel....they are not even in the middle....
thank you for the suggestions again....
you can follow migration by using pre-stained molecular weights. you can either use blue pre-stained, they are easy to see, or you can use kaleidoscop molecular weights marker, it's easier to recognize the band corresponding to your protein, because they all have a different color, but the bands are difficult to see on the gel while it's migrating. after transfer you see it again, but while it migrates in the gel, it can be difficult t see.
-Missele-
QUOTE (mdfenko @ May 5 2006, 05:32 AM)
btw, when i said, in the previous post, that the standards didn't run 'true', i meant that the distance between bands did not properly reflect their respective sizes, not that they appeared in the wrong order (they were too disparate in size to do that). sorry for any confusion that statement may have caused.
thank you for the clarifying....i did run the gel for 1 hour more and it is nicer now...
however im having problems in identifying what I see....what you said about not moving true acc to size so i cant actually compare my protein with molecular weight marker right? my protein's charge is -20 at ph8.8 (pH of the gel) and 66 Kda marker charge is -41 at that pH so as it is smaller and has greater negative charge it should migrate MUCH faster that my protein right? ....but from what ive got it just seems to be rather MUCH slower... ...please correct me if im wrong, am having such a hard time in explaining these results 
(hate UREA-PAGE 
) -Kathy-
QUOTE (Kathy @ May 7 2006, 12:49 AM)
thank you for the clarifying....i did run the gel for 1 hour more and it is nicer now... however im having problems in identifying what I see....what you said about not moving true acc to size so i cant actually compare my protein with molecular weight marker right? my protein's charge is -20 at ph8.8 (pH of the gel) and 66 Kda marker charge is -41 at that pH so as it is smaller and has greater negative charge it should migrate MUCH faster that my protein right? ....but from what ive got it just seems to be rather MUCH slower... ...please correct me if im wrong, am having such a hard time in explaining these results (hate UREA-PAGE )
however im having problems in identifying what I see....what you said about not moving true acc to size so i cant actually compare my protein with molecular weight marker right? my protein's charge is -20 at ph8.8 (pH of the gel) and 66 Kda marker charge is -41 at that pH so as it is smaller and has greater negative charge it should migrate MUCH faster that my protein right? ....but from what ive got it just seems to be rather MUCH slower... ...please correct me if im wrong, am having such a hard time in explaining these results (hate UREA-PAGE )i'm not sure that i can help you explain the results or why the results are as you see them. what you can do to make more sense of the results is run a second dimension with sds-page. one of our neighboring labs routinely ran urea-sds 2d page (as opposed to ief-sds 2d). you can either run a strip in the second dimension or cut out your bands of interest and load them onto sds-page. it's a little more work but it may help you make some sense of your experiment.
sorry, i wish i could be of more help than this.
-mdfenko-
QUOTE (mdfenko @ May 7 2006, 05:20 PM)
i'm not sure that i can help you explain the results or why the results are as you see them. what you can do to make more sense of the results is run a second dimension with sds-page. one of our neighboring labs routinely ran urea-sds 2d page (as opposed to ief-sds 2d). you can either run a strip in the second dimension or cut out your bands of interest and load them onto sds-page. it's a little more work but it may help you make some sense of your experiment.
sorry, i wish i could be of more help than this.
sorry, i wish i could be of more help than this.
thank you so much for the suggestion! yes i think this is the best way to make sense of what im seeing...
...sorry for being so naive but can i just cut out the band (stained with coomasie) boil it in SDS sample buffer and load onto the SDS gel? won't coomasie effect its migration? will gel dissolve in SDS sample buffer? thank you so much again -Kathy-
QUOTE (Kathy @ May 8 2006, 12:49 AM)
...can i just cut out the band (stained with coomasie) boil it in SDS sample buffer and load onto the SDS gel? won't coomasie effect its migration? will gel dissolve in SDS sample buffer? thank you so much again
yes, you can (we have). the coomassie will be removed from the protein by the current and will run ahead of the protein. the gel will not dissolve in the sample buffer (in fact, it may swell) so you may have to cut it into smaller pieces to fit into the well.
you're welcome.
-mdfenko-
QUOTE (mdfenko @ May 8 2006, 06:15 AM)
QUOTE (Kathy @ May 8 2006, 12:49 AM)
...can i just cut out the band (stained with coomasie) boil it in SDS sample buffer and load onto the SDS gel? won't coomasie effect its migration? will gel dissolve in SDS sample buffer? thank you so much again
yes, you can (we have). the coomassie will be removed from the protein by the current and will run ahead of the protein. the gel will not dissolve in the sample buffer (in fact, it may swell) so you may have to cut it into smaller pieces to fit into the well.
you're welcome.
thank you so much again! you can't imagine how much you are helping me...
im so new to all this.. ill run the gel and let you know about the results! have a great day! 
-Kathy-
Dear mdfenko, just wanted to let you know what is heppening...after running my protein on SDS_PAGE and the bands from UREA-PAGE i understaood that the bands were actually the degraded form of my protein which had thousands of bands (instead of just two) so it degraded fully after i left in at 4C for almost one month... 
...yes i know stupid me... ....anyway now i have prepared new protein from inclusion body by dissolving it in 6M urea....it is ok...but when i ran it on 6M urea-PAGE no bands.... ...so im trying to do 7M urea-PAGE (as I did before but for protein solubilized in 8M urea),....so thats all very strange....anyway....now the gel is running let's see what happens....
-Kathy-