you use 0.1% sds in the transfer buffer?

the literature recommends to not exceed 0.05% sds in the transfer buffer, the methanol (or ethanol, in your case) will not be able to remove enough sds to ensure good capture by the membrane.

but, if it works for you then why change?

Well, that recommendations comes from the fact that everything will be soaked in SDS-Methanol containing buffer but in this case the methanol containing layer doesn't have SDS(of course it will flow in during transfer).

I think that the semidry transfer system also uses 0,1% on one side (I may be wrong) I even read of some very efficient transfer with up to 0,5% SDS.

Hey all.. i have been using PVDF all my life... but now we are out of stock and have a NC membrane..
i was wondering if i can follow the same protocol for blotting as i do with PVDF.. except for the activation with methanol part of course...
but my transfer buffer also has 20% methanol.. can i continue using tat or remove it from my buffer for a NC??!!!
i usually run at 100V and 400mA max current for an hour!!!

I may be wrong but I think I remember that at least 10% methanol was required in transfer buffer for nitrocellulose, not the case of PVDF.

methanol is only required in the transfer to strip sds from the protein being transferred and to help prevent (or reduce) swelling of the gel during the transfer.

it has no direct effect on the nc (it does not require "activation" like pvdf). you can transfer without using methanol at all (with non-denaturing gels).

Yep, probably it has to be with the fact that nc has less retention capabilities for some proteins and that it is not activated, so methanol is plain zero. I read somewhere that 1% or less may be more than enough for transfers, but I haven't tried that.

I was actually just reading about the Tris-Tricine gels. I have never used them. I am using Nupage Bis-Tris gel from Invitrogen.

it may be relatively insignificant if there is only one biotin bound per protein molecule but if more biotin binds then the effect will become more significant.

also, which page formulation do you use? if you are using tris-tricine then one biotin may make a significant difference on the gel.

You can use your Nupage Bis-Tris gels with MES instead of MOPS or HEPES which will give a similar result as tricine.

Search for MES monohydrate it's much cheaper than just MES.

Ok, now it makes sense, but anyways, the gel almost melted. I used 20V though.

No, it's actually quite low - approx. 10~15V.

Yep, when heated it looks almost melted .

Can you re-use secondary Abs-HRP @4C in TBS-T?

Long term, nope, HRP breaks down very fast in water. During 1-3 days probably, but activity will start to decline.

Reusing your secondary can help you to remove unspecific binding if you have some.

If you pretend to prepare a replenish stock or something like this, it should be in 1% PVA to stabilize de HRP.
The good thing about this is that such a high amount of PVA will give very clean background.

I just started to prepare PBS-A. My PBS-A recipe came from Freshney's cell culture text book. However, I discovered that after i dissolved the mixture in water and autoclave, the PBS-A became cloudy with precipitation.

I'm sure the recipe is ok, as most of other protocol available online also suggested the same. The only modification I made is I used 1.14g Na2HPO4 anhydrous instead of the one suggested (not in anhydrous form).

Could the anhydrous form cause the problem? Is there anyway to solve it?

I compared my recipe with other labmates, and found out that they used NaH2PO4 instead of Na2HPO4. I am thinking if I can use their protocol, then adjust Na+ ion concentration using NaOH? Assuming the pH will be quite alkaline, then I will just adjust the pH back to 7.4 with HCl. Is that ok?

There is a big difference between NaH2PO4 and Na2HPO4. One is acid and the other basic, you mix them to arrive to the appropiate pH.

I remember one of them fall out of solution when frozen, thus changing the pH, the same probably occurs autoclaving. If you pretend to freeze or heat strong switch to 50mM Tris-Hcl.

To prepare PBS
1)Prepare NaH2PO4 solution 0,5M
2)Prepare Na2HPO4 solution 0,5M
3)Mix (1)&(2) to obtain a stock of 0,5M PB with pH 7,2 (6,8-7,6).
4)Make a 5M stock of NaCl (a litle heat will make it dissolve fast)

To prepare a 20X PBS stock:

5)Mix 400 ml of (3) with 600ml of (4)

To prepare 1x PBS:

Mix 50ml of 5 with 950 ml of distilled water. Ready to use.

This is the standard PBS: 10mM phosphate with 150mM salt.

To prepare 1X PBS you can use distilled water. For the stocks use miliQ water and store them in fridge to avoid fungal grow.

Hello, I have been doing western blotting a lot recently. When I transfer my poly acrylimide gel to the transfer setup, it always rips and folds on itself. Is their a trick to getting it off the glass panel and on to the filter paper without falling to pieces or tearing?


Thank you.

Mike

What lab rat told you is the best way to go, also you can add 10% glicerine to the gel to make more stable.

Hi
i am using a molecular weight marker from Bangalore Genie (India)(Old catalog no. PMWM, new catalog no. 105979). In the image below A is the pattern which the company has given in the catalog, whereas B is the pattern which I got on same- 12% SDS-PAGE. As you can see clearly, there is a difference in the spacing between bands of the 2 patterns? Can anyone comment on why is it so? Is such case can I just count number of the bands and then compare the pattern order wise?

It may be also that you and the company use different acrylamide stocks, lets say you use 19:1 and the company 37.5:1 that would also explain it.

Hi there,

I just began my bachelor thesis and have to extract a nuclear protein from mouse fibroblasts (to perform a Western Blot afterwards). Is it a good idea to use mouse ear tissue or do you have any better ideas? Maybe you have experience with that kind of experiment?!

Thanks in advance!
Isi

Yep, any connective tissue will due. As it is a nuclear protein use loading buffer for lysis, and adding deoxycholic acid may help. Use a harsh method such as rotor-stator or bead beating.

Hello

I am not very good with calculations. I used the nanodrop instrument to determine my sample protein concentration as 2 mg/ml. I would like to add 20 ul into each well such that the final protein amount is 20 ng (per one well). Would any one please show in a step by step manner the calculation to derive this (I am confused with mg, ug, ng and the amount per well).

Since I need to load the protein in duplicate or triplicate, what would the calculations be for three wells: i.e 20 ul per well and there are three wells and the protein amount is still 20 ng per well. Remember, my original stock is 2 mg/ml. If this is too low, just use a suitable original concentration as an example. Please, can any one help me with this, as I am rather confused.

Thanks
lab2009

Just keep in mind 1mg/ml =1ug/ul=1ng/nl
You have 2ng/nl. If you want 20ng that is 10 times that amount of ng and 10 times that amount of nanoliters, ie 10nl.

As you can't probably pippete less than 1ul I recommend you to dilute 1 ul of your sample in 100ul of buffer and load 1ul of that.

I recommend doing the less serial dilutions as possible as low concentrated proteins adhere to surfaces leading to incorrect dilutions.

Hi everyone,

I'm researching a secreted protein in culture medium from cell lines of prostate cancer. I run a WB without problems, however I need a secreted protein to normalize the results and loading control (like actin in protein extract).

Are there any protein that always be released to culture medium?

Thank you!
Valentina

Loading controls are fake-prone.

I would recommend you simply performing ponceau stain, scan and lane densitometry.

sorry, I am a fresh man of biology, could anybody tell me what is the sensitive amount of protein for western blot?
I purified GST-fusion protein and checked by SDS-PAGE, bands were clear ,
but for western blot I think I have to dilute the GSTs , now I need to know the sensitive amount of it so that I can determine how many times the protein to be diluted.
thank you everybody, best regards to you!

You can choose to dilute your sample or the ab. There are some tables that compare sensitivities look at pierce supersignal.

Hi all,
I've been using Amersham Full-Range Rainbow Molecular Weight marker as my ladder in my gels. However, this doesn't show up too well when stained with Coomassie Blue. Do you have any recommendations as to other types of ladders that will show up brightly? Thanks.

There are ladder's ready to be coomasie stained and they are the best choice for accurate weight stimation.
I remember Sigma has some, probably every company has some.

Just wondering if anyone knows of any good antibody which detects laminin-5 secreted protein during western blot analysis? I had a really good abcam antibody which detected the three laminin 5 chains of 165kDa, 140kDa and 105kDa, however it has just been discontinued:'(
I'd really appreciate any help.

Take a look at hybridoma bank, you may be lucky.

Hey guys,
So it was my first time doing a WB. Since I had to rush to an important thing, I did not have time to develop chromogen after 2ndry antibody. I was told that its OK to dry the membrane and later develop color, which I did. We then found out that nobody has ever done that and people dry either before they start the antibody washes or once its completely done, not in the middle. Is there anyway to salvage the situation? People told to try "activate" the membrane with methanol, wash it couple of times with PBS/TBS and then try developing color. But I wanted to ask the community if anybody has done that. By the way I'm using PVDF membrane.
Thanks in advance for your tips and assistance!

Yep, in fact HRP lives better dry than in water.

Hi tehre.. anyone aware of prestined markers (other than bio-rad-precision plus) broad range to be used for Western blot analysis
better if available in india.. loaly.. not imported.. cause i don have much time to wait for the shipment!!!

Sigma has quite a variety for better price than BIO-RAD, but the shippment will probably arrive in 1 week, your best bet, ask a local supplier or a friend.
Also I think making your own blue stained mix is rather easy.

Hi
i am using a molecular weight marker from Bangalore Genie (India)(Old catalog no. PMWM, new catalog no. 105979). In the image below A is the pattern which the company has given in the catalog, whereas B is the pattern which I got on same- 12% SDS-PAGE. As you can see clearly, there is a difference in the spacing between bands of the 2 patterns? Can anyone comment on why is it so? Is such case can I just count number of the bands and then compare the pattern order wise?

Just plot each protein weigh vs mobility you will probably obtain a curve intead of a straight line, that's ok.

It seems that you got higher resolution of the lower weight proteins this may be due to:
-The inclusion of urea or glicerine in the gel.
-Different buffer in the + and - pole
-Reusing buffers.
This is usually an effect of different molarity between the gel buffer and +pole buffer.
By the time the gel is inmersed in the tank, if your gel has lower molarity than the tank buffer it will start to slowly equilibrate by more molecules (Tris probably) entering the gel. This will increase the resistance and thus current on that end of the gel, thus your proteins will run faster on the border of the gel.

If you want to be sure the marker is alright, which it probably is, run BSA or other know weight protein on a parallel lane. Keep the same salt, SDS-detergents, protein qt content on each lane.

Hey all.. i have been using PVDF all my life... but now we are out of stock and have a NC membrane..
i was wondering if i can follow the same protocol for blotting as i do with PVDF.. except for the activation with methanol part of course...
but my transfer buffer also has 20% methanol.. can i continue using tat or remove it from my buffer for a NC??!!!
i usually run at 100V and 400mA max current for an hour!!!

I may be wrong but I think I remember that at least 10% metanol was requiered in transfer buffer for nitrocelulose, not the case of PVDF.

I never had this during western blotting. What type of milk are you using? Has it only just started happening?

Does it effect the outcome of the blotting? If it doesn't then I don't worry about it.

Does it happen with both TBS and TBS-T?
Milk will go clear when the micelle structure is destroyed. Typically this will be because the Ca has been removed (a handy tip if you need to filter sterilize the milk) but in your case probably the micelle proteins are being dissolved by tween or bound to the membrane.

Thank you people. I use TBS, and milk powder. I prepare a solution freshly and incubate with different antibodies o/n. The next morning the milk turns clear however only with some antibodies, whereas with other antibodies it does not turn. I guess it is due to the antibody (Sigma anti-flag, M2), because the same milk does not turn with another antibody.

May your antibody contain some chelators like EDTA or EGTA?

May it be that you use different containers for different antobodies?

Hi,

I have a suspicion for why none of our bands look strong using polyclonal antibodies, even those for so-called abundant proteins.

We have a good scanner (Fuji LAS-3000) that we use for densitometry of EtBr-labeled DNA gels, which has emission filters at both 520 and 580. So we use it to visualize bands on Western blot by using fluorescently labeled secondaries (Alexa 488). Should we expect this to inherently lead to less sharp bands, or even to less intense bands, compared to the intensity of background staining? (this could be the tradeoff in which fluorescent-labeling being an easier and less messy procedure than HRP)

Any information you can give about experience with fluor-conjugated antibodies versus HRP- or AlkPhos- conjugated antibodies, for visualizing protein blots, would be interesting. Thanks.

Do you have the apropiate PVDF, most have autofluorescence.

Alexa 488 captures at 520?

Also you may need to try to let the scanner record for a long while and superimpose the images.

Still, ECL is usually more powerful than fluorescence. You should at least make a try with ECL to see that everything is ok but the fluorecence at that point.

Hello

I am targeting HSP70 and since I was having some issues with the blotting, I decided to Coomasie Brillian stain the gel after the blot to check its efficiency. I loaded the gel like this:
1.Marker
2.HSP70 pure standard
3.sample 1(liver homogenate) total protein
4.sample 1(duplicate)
5.sample 2(liver homogenate) total protein
6. sample 2 (duplicate)
7. negative control (only sample buffer)
(etc as mirror until the 14th lane since the gel was to cut in 2 to test different antibodies)
I realized that my blot wasn't efficient since the proteins were still in the gel, but what I found strange was that the marker and positive control were in the gel, but from the liver homogenate ONLY the 70kD (which I assume is actually my target protein) were in the the gel. I haven't stained the membrane to see if something passed or not (although at least half of the marker passed). My question is, in a case of a bad blotting when the proteins stay in the SDS gel, a total protein homogenate should have many lanes and not only and exclusively on the 70kD. I use as extraction buffer 1 ml (of 1mMEDTA 1mM PMSF in PBS ice cold with protease inhibitor cocktail) for 100 mg liver (of fish by the way), homogeneized, centrifuged, and supernatant stored at -80C until sds page/western blot.

Any thoughts about this are very welcome since I think I am missing something very basic here...

My best regards to all

Anatg

True, that buffer is very mild, if you just pretend to perform sds-PAGE get used to lyse your samples in loading buffer+inhibitors.

To improve your transfer you may take a look at his: http://www.protocol-...showtopic=10851 .

Is heat shock protein heat labile? If so instead of using heat for the transfer use DHEBA for polymerizing the gel.

Hi,
I am completely puzzled by this. Every time I try to transfer a particular protein, it wont stick to the membrane unless I run it to the point where the buffer evaporates, almost like baking the system. I am using the semi-dry blotting system from Invitrogen. Even if I use the same amount of voltage (50V, Invitrogen protocol calls for 20V) and keep it really cold that it doesnt evaporates, I won't see my protein, unless I let it go until the buffer evaporates and it smells like burnt! I don't get it! is my protein that hard to to stick? its a 17KDa protein, very small!
I have done this multiple times as to determine that the only way I can detect it is by frying the membrane!

Any comments please?? I'm about to call Invitrogen and ask about this.

Well,this is because heat impairs the gel structure and allows proteins to flow through smoothly. To achieve even transfer temperature should be over 65ºC

If I wanted to detect a protein that normally would be 9kDa as a monomer, and I got it biotinylated, would it be detected in a WB at a different molecular weight using Streptavidin-HRP for detection? And how much more heavier would it be?

You may observe differences higher than expected as biotin is a vitamin that may lead to a different ratio of weight/mobility than that of aminoacids. This is due to it's conductivity, hydrophobicity and SDS interacting characteristics.

Hi all,

Hoping you can help me troubleshoot my problems with transferring ~220kDa proteins, I scoured these forums yesterday trying to pick up relevant tips but I don't seem to have cracked it yet.

5% acrylamide SDS containing minigels (10x10) loaded with 50ug protein per lane and run @ 200V for 45 mins, 2 checked with coomassie and confirmed clean, crisp bands and even loading. Markers up to 460kDa separated nicely.

2 gels put into wet transfer overnight in Towbin containing 5% methanol and 0.1% SDS with Millipore 0.45um pore PVDF @ 90V in coldroom. Previously tried transfer at 50V and the larger proteins stayed put in the gel.

This morning I disassembled and the buffer was fairly warm (not a great surprise given the higher voltage) but there was no sign of the prestained markers on either the gel or the membrane or on the filter paper. I was always under the impression that it is quite difficult for proteins to become unbound from PVDF and pass through the other side, is this not the case? I am just in the process of staining the gel to see if there is anything still left on it.

Any ideas about where I may have gone wrong?

Thanks in advance

Following the 1h 1amp rule, 90v OvN may be too much.

As you've been told you need SDS too accelerate the protein and Ethanol to stop it. Also heat really improves transfer, and over 65º makes it even.

I developed a system that works like a charm an is as follow (tank transfer).

sponge.m-paper.m-membrane-gel-paper.g-sponge.g

Activate the membrane in ethanol

Equilibrate in tris-gly buffer without sds and without ethanol everything but sponge.m.

Equilibrate sponge.m. in the same buffer but containing 20% ethanol.

I don't use SDS in this equilibration buffer as it makes bubles.

As transfer buffer use the tris gli but in this case without ethanol and with 0,1%SDS.


The high SDS will accelerate your proteins and the ethanol will flow from the sponge to the membrane removing the SDS and making the proteins stop there.

Avoid equilibrating the membrane in ethanol containing buffer as it will contact the gel and impair protein trip.

I usually heat the transfer buffer to 75º in the microwave and transfer for 30-60min-, 400mA.

If you are afraid the your protein surpases the first membrane ad a second membrane between paper.m and membrane.

Also if your protein has high IP it will trip to the (-)pole. Usually SDS will avoid this but if you want to be sure simply place another membrane between the gel and paper.g.

If your protein is travelling the wrong way you can use higher pH transfer buffer like CAPS ph 11.

This is inspired in a semidry blotting system but works as-well.

Hi, all,

I expressed a fusion protein with mbp tag (MW: 81KDa+42KDa) and everything looks fine. Molecular weight from SDS-PAGE matches well with above. After the cleavage, it shows clearly a single band at 81 KDa and mbp single band (42KDa). After size exclusion column, I got the pure target protein (81KDa). However, from SDS-PAGE gel (16% + 5%), I got several discontinuous bands around 81KDa (they are really very close to each other). I know my protein is pure. I don't know the reason.
I made fresh APS solution and didn't make any difference. I tried to load different amount of proteins and the single bands occurs for smallest amount of samples.

The same situation happens for my another protein which has the molecular weight of 54KDa. But I haven't had the same problem before. Is there anyone who can give me some suggestions.


Thanks.

Haiying

I think that you have been very well advised, if it where doublets you would see much distant bands.

The problem seems like mild proteolysis or bad S-S reduction.

If you are sure the column is clean ad protease inhibitors as soon as you collect the sample from the column.

If you have already tried mercaptoethanol and DTT y suggest trying neutral ph TCEP for sample reduction, it is very stable and will keep your samples reduced.

If it came from a deficient gel you would probably see a smear instead of discrete bands.