I use my own gel, and my own blotting stack and i have had no problems so far. The transfer seems comparable. I usually use the 7 min mixed MW transfer protocol. The transfer buffer i use is, Bjerrum and Schaefer (0.1% SDS). The tricky part is to replicate the blotting pads/paper. I had tried different kind of filter papers and non of them seem to work. So we switched to a material, which is more fabric than paper, with similar absorbing capabilities.

Hi David - the Bio-rad trans-blot manual (attached) has instructions for making your own transfer stacks using standard PVDF membranes and chromatography grade filter paper. According to their instructions you have to use a 30 min transfer program when you make your own stacks, however I was able to get reasonable transfer using the 7 min "Mixed-MW" mode. I am working with small peptides and proteins though (

My current lab (just joined and have not tried this myself yet) uses its own stacks and have not had any issues (gather they have probed upto 70kda).

I use my own gel, and my own blotting stack and i have had no problems so far. The transfer seems comparable. I usually use the 7 min mixed MW transfer protocol. The transfer buffer i use is, Bjerrum and Schaefer (0.1% SDS). The tricky part is to replicate the blotting pads/paper. I had tried different kind of filter papers and non of them seem to work. So we switched to a material, which is more fabric than paper, with similar absorbing capabilities.

Aman, may I ask what fabric-like material you have used? We tried blotting paper stacks in conjunction with the BioRad buffer, works ok, but still not as good as the BioRad stacks.

Hi Chinten, in our lab we re-use the fabric that comes with the BioRad stacks. The "fabric" is very durable and can handle vigorous rinsing. The western transfers look just as good as the BioRad stacks. Its all in the "fabric" :o) Good luck!

@ Aman: I also would like to know which filter papers you use, because I tried some and I only get really good blots when I reuse the BioRad stacks rinsed and soaked with our buffer system (Kyhse-Anderson). Reusing them all the time is not an option, because we do not plan to buy them.

I use Wypall X60. it is not a filter paper. it is a kind of wipers with hydroknit technology. WE observed that it has similar retention compared to the stacking papers from bio rad.

Thanks! Since these wipes are very thin, how many layers do you use on each side?

i have had success using 6 on each side. just make sure that the Nitrocellulose/PVDF forms a complete barrier in between two stacks.

When using your own materials and buffer when transferring with the transblot, does anyone realize the amperage/current is 0.1A instead of 1.3A for a HIGH MW protocol? Or is it just me who have this problem?

First of all, thanks to Aman. My transfers are brilliant now with these tissues :)

@ Vivian: I had the problem with 3M Whatman and oder filter paper. This seems due to a high resistance. With more fabric like filter paper as mentioned by Aman, I get now 0.5 A and 25V in the beginning of my 30 min transfer. After 10 min the ampere increases to 1 A and the Voltage goes down to 17-18 V.

According to our ordering system:

Distributor: VWR

Cat. no.: 115-2166

Description: WYPALL* X60, white, 420×310 mm

Measure: 200

Cost: £43.80

1 ply and white

P.S. They still work fine, no problems with the blots since I use them.

Thanks to Aman as we have now tried Wypall X60 and transfers appear comparable to the BioRad Transblot Turbo stacks. Nicholas, we used this (Amazon):

Kimberly-Clark Wypall X60 Teri Reinforced Wipe -126/Box -9-Inch x16.50-Inch -White

at 6 sheets(or ply) per stack, cut to size. Maybe the important thing is the X60 white.

Excellent! Now I wonder if anyone has had luck recreating the Transblot Turbo buffer? We still use the provided buffer (with the Wypall X60) since each kit comes with enough buffer to last for 2-3x longer than after the provided stacks are long gone. Please post solutions here or redirect me to relevant post if applicable. Thnx!

I use Kyhse Andersen (link attached), but with 0.05 SDS in the Cathode buffer and only 5% Methanol instead of 20%. Howver, at the moment I have a problem that sometimes my blots are just perfect and the next time sporadically some bands are missing. Last week I did blots which went perfect, this week I repeated them with the same sample (since I needed more membranes to probe) and transfer seems to be awful despit I did not change anything. Does this happen to somebody as well?

http://books.google.co.uk/books?id=xVSsRgCZIJIC&pg=PA249&lpg=PA249&dq=kyhse+andersen+1984+6+aminohexanoic+acid&source=bl&ots=eG2HACB6Pw&sig=E-K-0ngAmtUAWutARKnezZNfERs&hl=de&sa=X&ei=wZt4VMjcCYH2O4izgdgF&ved=0CCQQ6AEwAA#v=onepage&q=kyhse%20andersen%201984%206%20aminohexanoic%20acid&f=false

I am starting using the Turbo transfer system, and I realized that  the consumables are quite expensive. I'll try your suggestions about using the Wypall X60 instead, and get back to you with the results. However I have a question.  The manual indicates that you have to use the midi gel transfer stacks if you are transferring 2 mini gels at once in one cassette. Has anyone transferred 2 mini gels per cassette? Can you please tell me how do you get optimal results?

Thank you in advance for your answer.

Yes, you can treat 2 mini gels as one midi. We try to butt the two gels together so as to have minimal gap between. We dont do single gels too often, but the consensus is the transfers with mini 1-gel or midi 2 mini gels are comparable and optimal.

Hi! We are starting with the tests of our turbo blot. Thanks for the tip with the paper. I found a publication focused on the change of the buffers: http://www.ncbi.nlm.nih.gov/pubmed/23872007 (Garic D et al, 2013). I tested two buffers made based on the recommendations of this paper. I am still optimizing but I can say that using Tris / Hepes/ Bicine and CAPS improved quit a lot the current from our transfer (From 0,4 A, 25V to 1,1-0,8A, 25V ) decreasing the resistance on half. I will keep you posted on the results as soon as I get the new paper that you recommended here!

I got the paper! Could you recommend and amount or thickness of the paper towards the cathode and towards the anode? It will save me a couple of try outs! Thanks!

Thanks. For some reason I do not see your previous input... I will keep you informed. Have a great day!

Ok. So I finally had time to do the test. I used the new paper, 6 sheet under and 6 sheets over. I tried the following buffer based on the publication and the material that I found (see above). It contained: 48 mM Tris, 20 mM Hepes, 20 mM Bicine, 10 mM CAPS, 1,3 mM DMF, 1 mM EDTA, 20% Methanol. The final pH measured was 8,08. With the new paper and the buffer, I reached the 1,3A 15V, (as the commercial kit reaches) but the transfer was not really complete (See my comparisons in the file attached) Any help? We have been using the AnodeI, AnodeII and Cathode buffer system because the normal "Western Transfer Buffer" left mayn of the proteins in the gel. Probably many of our proteins have a basic pI. That is the reason why I introduced CAPS. But in many protocols using CAPS, they have a stock solution 10X. Does anybody know the real reason for it? Could it be the methanol at 20%? Usually we use it at 10%... well... all comments and ideas (with reason for it, please) are welcome!

Hi everyone, i have summarized the general practice of Western blot in our lab. I hope it will help with some of the questions here:

1)      Size of the gel dictates the transfer protocol: We only use mini gels (~8.6 x 6.8 cm) for most of our western blots. We have done transfers with single mini gel or 2 mini gels. For one mini gel we use 1.3 Amp (constant), 25 V (variable) and 15 min, for 2 mini gels we use 2.1 Amp (constant), 25 V (variable) and 15 min. This transfer protocol has worked perfectly when we use Bjerrum Schafer-Nielsen buffer (48 mM Tris, 39 mM glycine containing 20% methanol) containing 0.1% SDS.

2)      Biorad’s 5X transfer buffer will make life easier: Recently we were able get hold of few bottles of the 5X transfer buffer from our collaborator. Bio rad supplies more buffer than needed in their Kit. The transfer buffer lasts for a while and we have been able to use the default 7 min protocol for our transfers using Wypall and our own membrane.

3)      Protein size : In our experience for proteins with higher molecular weight lower %acrylamide gels, 0.3 % SDS and 0.45 µm pore size supported nitrocellulose membrane works the best.

4)      Blotting Medium: We use 0.2 µm/0.45 µm pore size supported nitrocellulose membrane from bio rad. The 0.2 µm pore size membrane works very well for proteins that are below 20 kD (histones) and the 0.45 µm works great for proteins with higher molecular weight. We have only used nitrocellulose for our experiments.

5)      SDS helps: We usually transfer the gel directly to the sandwich without any equilibration step in between. The BioRad’s proprietary buffer contains SDS (I don’t know the exact concentration) and Bjerrum Schafer-Nielsen buffer that we have used also contains SDS, SDS helps with the transfer of the proteins.

6)      Developing the blot: We usually use two different kind of developer: SuperSignal West Pico Chemiluminescent Substrate for general use and SuperSignal West femto Chemiluminescent Substrate for very low abundant proteins. We use Biorad’s Chemi Doc MP system to image our blots, which provides more control to avoid saturation of the signal

7)      Transfer Efficiency: The wet transfer method works very well for most of the transfer. The semi dry method might not provide similar or better transfer as compared to wet tank method all the time but it is surely fast, easy and efficient. I think what matters is uniformity during transfer with acceptable transfer efficiency since signal is amplified later on due to antibodies (I usually make sure that I use larger blot that the size of the pads so that there is no direct contact between the pads, I don’t know how much of a difference this approach makes but I have never had any non-uniform transfers so far).

Dear Aman,

Thanks for the nice overview! I use only PVDF membranes, I hope it will be no problem. As for transfer efficiency, I need to have all proteins transfered: from 250 kDa to 25 kDa, since we do semi-quantitative analysis of western blot signals, and our gels run as gradient gels (They are other kind of gel: Single gel system Ahn et al, 2001). Next week I will prepare you buffer and give it a try. About the SDS, is the SDS in the gel not enough to help for the transfer?

I repeated now the transfer with 15 min. It is better for the bacteria proteins (The 4 with very little protein signal over in the upper part) but I am still having problems with the big ones.

In my experience, I have tried to transfer my 250 kDa protein to membranes using the wet transfer chambers, and it has never worked as good as it works with our buffer system using Anode and Cathode buffers (see first slide of last document).

@Nicolas

This info is very helpful. I was already suspecting the alcohol. I will be making some buffers and I will keep you all posted. Have a nice week!

Thanks for Wypall X60 white. I was looking for a while for the filter paper same as from RTA transfer kit. I use bolt empty cassettes with stain free fast cast acrylamide solutions. I didn't had any problem detecting the protein from 10 to 150kd with 7 min transfer. Even with bis-tris gels transfer came out pretty good. I think 0.2um PVDF works better than 0.45um. 

Dear All, I finally got a 97% transfer of all my proteins to the membrane usind the Wypall X60 paper and our old buffers (independently of alcohol type). The proteins transfered were from 20 kDa to 270 kDa in 20 min (our old protocol was 1h20min). If someone wants to have it, just send me a message through ResearchGate.

Have a great weekend!

Hi! There have been so many people interested that I decided to place our protocol in a repository for Protocols. You can find it here: https://www.protocols.io/view/trans-blot-turbo-transfer-with-home-made-buffers-ghhbt36

Please reference this DOI and the original papers. That way we support the repository and as well those who worked so hard to find the things we were able to adapt!

Have fun!

@Luisa.

Thanks for the protocol.  I am intrigued you were able to transfer such a large range of proteins.  I will be giving your method a go this coming week.

@Greg

Great! please let me know if it worked for you. It will be great if it works outside of our lab! 

Have a great weekend!

@Greg

We use a single gel system. I published it today in the same repository as Single gel system. those gels run as gradient gels. in a 5% I am able to see my CagY (270-220 kDa) and the GFP (26 kDa). just do not let the blue front to get out. if it does you xan only see 35 kDa proteins 

The link for the single-gel system that we used for the Turbo-Blot system

https://www.protocols.io/view/single-gel-system-based-in-ahn-t-et-al-2001-gipbudn

@Luisa.

I ran a quick test run today.  Interestingly, I found the single-gel-system paper last week and I was already planning on using it.  I used a 4-14% single cast gel. My protein (150kDa) transferred across, the ladders (Precision Plus) however did not come across very nicely.  I loaded 5ul of ladder on a 1.0mm gel, and transferred onto a 0.4um PVDF membrane.  The transfer started at 0.9A and ended at the 1.3A.

I will try it again tomorrow.  It was also my first time using wypall x60.  Couple quick questions: How wet do you keep the wypall, it seemed like my membrane was on the dry side after the transfer? Do you equilibrate the membrane in your Anode solution I or II?  

Hi Greg,

Sorry... I had a mistake in the protocol. I equilibrate the membrane in Anode II. I tried to change it and I think I placed it under annotations...;-)

The wypall I completely submerge in the buffers. I never remove the excess. I know some protocols recommend to remove the excess buffer in the paper but I have never done it. Not even with the "official" filter paper.

I have been using the AllBlue high-range protein marker from Bio-Rad (3,5 µl in a 20-well gel, 1 mm thickness). And to test if the proteins have been transfered, I do not look at the marker transfer (it can be misleading!) but I look at the previously activated stain-free gels after the transfer. You can do a traditional coomassie staining from your gel after transfer to verify that it is working fine for you and all proteins have been transfered.

In theory, your membrane pore size should not be a source of discrepancy, but just to make sure: Our membranes (PVDF) have 0,22µm pores.

@nicolas

I wash really good the cassettes after transfer. but what I meant is that I have seen people removing excess of buffer from their filter paper by placing them shortly onto another paper to absorb any dripping excess before placing in it on the stack. This I do not do it. sorry about the confusion.

But you are right: thr surfaces of the Turbo blot are really sensitive and they have to be washed carefully and thtoughly with a lot of water to remove any salt.

@Luisa.

Second attempt at the transfer went well using a 8% single-gel.  Yesterdays attempt: I removed as much excess buffer as I could and the membrane seemed to dry out. This time, the membrane was equilibrated in the correct buffer, and I kept everything quite wet.  The transfer dropped once to 1.1A, but for the most part stayed at 1.2-1.3A.  The 250kDa to 75kDa bands of the ladder seemed to transfer really well. The smaller bands, however, seem to get progressively fainter. These bands were not present in the gel or Wypall.  There is also a couple yellowish marks on the edges of the membrane (see picture).  My protein of interest transferred across fine, and there wasn't much left on the post transfer coomassie either.  

I am reasonably happy with the transfer, and I will perform another Western tomorrow. Is this quality of transfer from a single-gel system what you normally see in your lab?

Hi Greg!

I am sooo happy it works for you too!

I have seen the same effect of the small marker when I use the 0,4µm PVDF. I have the experience that they get through. Some of my proteins tend to disappear with that pore size membrane ( for example my 35 kDa RecA). That is the reason we buy the 0,22 (Bio-Rad or Millipore, they are the same!). It is a bit more expensive, but I am not really afraid of loosing anything while blotting.

The yellow happens because they get hot! I got it twice now, but with the drying of the membranes after blotting and then activation proceedure for western blot, it did not seem to bother in the signals for chemiluminiscence.

Thank you soo much for letting me know! I love to get feedback!

have a great day!

Luisa

 @Aman/ Luisa,

Do you use self-cast Tris-HCl gels? Using the WypallX60 and the trans-blot turbo system, I see a pattern of the filter paper (etched) on the blot after transfer and Ponceau-S stain. Anyone come across similar problems?

Thanks in advance!

Poornima

@Poornima

Hi. No, we use self-cast Single Gel system. I have seen the pattern but until now it has cause no problems in detection. We do not do Ponceau. We do Stainfree staining of the gel and verification of full transfer through detection of proteins left in the gel. If you want to check the protocols, I placed them in the repository Protocols.io.

Did you have probkems with detection?

@Poornima

We self cast standard PAGE gels; have not observed strange patterns on NC blots. We also do not Poncaeu stain.

Thank you Luisa and Chinten.

I never saw this problem when using pre-cast TGX gels and Biorad pre-assembled stacks. Trying to figure out what might cause this imprinting on the membrane. We have now switched to casting our own gels and using WypallX60 as transfer stacks. When developing low abundance proteins, I see a pattern on the band which makes the western look ugly. Any thoughts will be appreciated!

I should add that we use both Wypall X60 and the Biorad supplied transfer stacks. The Biorad transfer stacks stand up very well to extensive washing (mostly just water rinse) and reuse, which on routine use is more convenient than cutting X60 stacks. It is possible that if you routinely load high amounts of protein in your gels and have 'overtransfer' then, the proteins may remain on the stack sufficiently to produce the pattern background. Do you get the same problem with both a 'fresh' stack of X60 or might they be due to re-use? Just a thought...

Sorry! No, until now we have no problem with the bands. How think is your gel? And your paper stack? Our is 1 mm gel and 1.4 mm thick paper stack.

Could it be that it is too thick and the paper is squizzing the gel  when you place them in the cassete and close them? Our I have tu press just a little bit and then turn the green wheel which drives the pins for closure of the cassette...

That is the only thing I can think of at the moment

I have been using Bio-Rad-s nitrocellulose, but we buy it by the roll and cut it to the size we need it. I also use Wypall X60 as blotting paper, and it works just fine. I haven't had any problems. The deal breaker for us was that you have to buy the whole kit just to get the buffer, too expensive. I will try the buffer suggested by Aman.  Thank you all.

Hello,

I was just wondering if anyone could recommend which sponges to use please?