We are primarily a skeletal muscle lab but have collected other tissues (liver, intestine, adipose)...not exactly on a 'whim' but without much research into whether they should be handled different from muscle. I have successfully run liver and skeletal muscle western blots with even loading.
We collect all tissues in cell lysis buffer with HALT protease inhibitor, homogenize with inert beads and spin down @ 10K g's for 20 minutes at 4C before collecting supernatant. We use BCA to measure protein concentration in the supernatant, then dilute with water to reach the same concentration for each sample before mixing 1:1 with laemmli. We then boil for 10 minutes. Before loading into our pre-cast Tris HCL gels, we vortex every sample (and we do this for the BCA and dilutions...lots of vortexting). I load the same volume into each well, which *should* result in the same ng of protein in each well. Gels are run at 150V for 90-120 minutes and transferred cold at 100V for 1 hr.
I have small and large intestine which was handled the same way, but for the life of me I canNOT get even loading. I have redone everything from the BCA onward 3 times. (It may even look worse now). I have sample that has not been diluted, but I no longer have actual intestinal tissue.
We use ponceau staining to determine even loading. Skeletal muscle and liver are clearly even across the gel, and intestine is just all over the place. I would say the middle samples tend to end up with the biggest blots after I ponceau stain.
I can certainly invest in a loading control and compare densiometries to come up with a value, but I can't publish these ugly blots!