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Western blot on intestine samples - (Sep/29/2011 )


Previously I've done lots of WB on the liver tissue, and it worked perfect. Now I started working on WB of intestine samples (fr. mice) and there seem to be some problems in getting a clear sharp band. I used the whole cell extracts and proteinase inhibitor was added in the Ripa buffer during the protein extraction, run 180V for 1.5h, and dry blotting for 2h before the blocking and incubation.

However, the bands of the intestine samples are always somewhat smeared, or fuzzy, feels like the membrane was covered by some ''clouds'' while developing, and I even got a double band for GAPDH (same antibody I've tried on the liver and it was perfect)!!!

So I guess the problem was in the sample prep... Has anyone here had any experience in working with intestine samples (better protein samples)? Are there anything special I should pay attention to? Thanks =)



Intestines contain quite a lot of proteases including some of the more unusual ones, which aren't covered by normal protease inhibitor cocktails. You may need to add more protease inhibitor cocktail and look for inhibitors to the unusual ones.

Intestines also contain lots of glycoproteins which may be causing some resolution issues.


I know this is an old topic, but I thought I would share what I recently discovered for Westerns using mouse intestinal tissue. Let me give you a little background first so you can see how I arrived at what worked for me.

Typically, we cryofracture our mouse tissues under liquid nitrogen with a mortar and pestle. Powdered tissue is then scraped into a microcentrifuge tube and weighed. Samples are then lysed at 3x, 4x, 5x, the dry weight in the Protein Homogenization Buffer or PHB (Recipe Below), homogenzied with a Kontes handheld pellet pestle, cleared by centrifugation, and then the Bio-Rad protein assay is used to determine protein concentrations of each supernatant. Samples are then aliquot for Western blot. Depending on the tissue 25 - 50 µg is mixed with loading dye, boiled, and loaded onto a gel. I have had great success using this method to look as phospho and total protein signal in liver, white adipose, and muscle. Kidney was a bit weird at first, but I managed to get it to work by loading less and diluting into water + loading dye. Some other tissues like salivary gland and tumor sample were okay, but problematic. At the time I blamed it on the sample, not the buffer, but in the future I will change to the "new" buffer.

Several months ago, I had to run small intestines. The first batch of proximal intestine looked horrible. We were worried that they had not been handled fast enough during the dissection leading us to believe that, perhaps, they had started to self-digest. Months past and I was asked to try again a few weeks ago with a different cohort that was handled more rapidly. Once gain, though, my gels ran strangely. Aside from occasional field effects across the whole gel, you could also see waves and/or points in the dye front for individual bands. The Western blots confirmed that I had wavy and sometimes smeared bands. I tried loading less protein. Things improved a bit, but I still had abnormal bands. It then seemed that the protein ran as a band, but the signal (on the blot) was concentrated into a point in the middle of the band. After looking at the literature, studying various protocols, and reading suggestions on this site, I decided that the culprit was either salts or lipids. Since salts were easier to address, I worked on that first.

I made new Protein Homogenization Buffer (thinking that I could have accidentally used too much in the first lysis), and used a microconcentrator to "desalt" and do a buffer exchange. I then proceeded as usual with new concentrations, gels, blots, etc. There was a little improvement, but overall, things still looked bad. After several other failed attempts, I went to the literature and protocols again. This time, I also looked up commercial companies that might sell total intestinal lysates. I found that AbCam sold two versions, one in a proprietary preparation and another in a modified RIPA (Recipe below). I made test samples of a few proximal intestine pieces from the same mice set of mice. I did everything as listed in my methods above, but this time, I used the modified RIPA instead of the PHB. My first blot looked good, but was a little overloaded (50 µg). After decreasing to 30-35 µg, I was able to get a beautiful gel. I have since used the same method for distal small intestine.

I hope my trial and error helps someone else save time in their research!

Happy blotting! : )

Final Conc. Protein Homogenization Buffer (PHB) --> This works great for many tissues, but definitely not small intestines.
50 mM HEPES, pH 7.6 (@4˚C)
150 mM NaCl
20 mM Na Pyrophosphate
20 mM ß-Glycerophosphate
10mM NaF
2 mM Na Orthovanadate (Na3VO4)
2 mM EDTA, pH 8.0
1% IGEPAL (Same as NP40)
10% Glycerol
1 mM MgCl2
1 mM CaCl2
10 µg/ml Leupeptin
10 µg/ml Aprotinin

In lieu of PMSF, Leupeptin, and Aprotinin, we use the Complete Mini (EDTA free) tablets from Roche, but we don’t usually make up the missing volume with water since it’s a negligible amount.

Modified RIPA Buffer from AbCam ab7939 - small intestine, whole lysate product literature --> This works great for small intestines.
Lyse in 5-6x dry volume 1x RIPA (small intestines)

150 mM NaCl
50 mM Tris-HCl, pH 7.4
1% Triton X-100
1% Na Deoxycholic Acid
0.1% SDS
Protease / Phosphatase Inhibitor Tablet 1 tab/10 ml

AbCam uses 5 µg/ml Aprotinin and Leupeptin and 1 mM PMSF. I used a Protease and Phosphatase Inhibitor tablet (Pierce 88668) + Pepstain A, PMSF, and Benzamidine as an extra precaution, but it was probably overkill. In the future, I may continue to add PMSF, but probably no Pepstatin A or Benzamidine.