Why do you think that the EC adhesions would be disrupted? If you are properly fixing/crosslinking the sample first then any type of permeabilization should not effect the EC adhesions.  Do not use collagenase since this will disrupt the gel and destroy the sample. 4% paraformaldehyde fixation in cytoskeletal buffer (10mM MES (pH~6.1),138mM KCl, 2mM EGTA(pH ~7),640mM sucrose: from Mitchison lab found online) for 15 minutes, wash once quickly with PHEM (60mM PIPES ,25mM HEPES ,10mM EGTA,2mM MgCl2, pH to 6.9 with NaOH pellets), then  permeabilize with 0.5%triton X-100 in CB for 5 minutes, then rinse 3X, block, etc. Pre-warm all your solutions and perform all steps at 37C since the collagen will often release from the dish.  Your biggest issue is getting the solutions into the u-slides without disrupting the gels.  The above protocol works well in open dishes, like MatTek or the Ibidi u-dish.  With open dishes be sure not to add any solutions directly on top of the gel, only on the sides.  For Ab concentrations double your primaries, but leave your 2ndaries at the same concentration as usual. DO NOT label with the 2ndary too long: 40 minutes or less is all that is needed.  Any more and you will just be fighting background fluorescence when imaging. For your final rinses (after the 2ndary only) add 0.1% Tween-20 to your rinse buffer and rinse at least 5X (10 minutes per rinse) to get rid of the 2ndary. Also do not use any mounting medium that solidifies, since this will compact your gels (I usually just use PBS--). Now being able to imaging them well that is another story.  For High resolution (40X and above) use water lenses if your samples are  greater than 10 microns from the bottom.

Why do you think that the EC adhesions would be disrupted? If you are properly fixing/crosslinking the sample first then any type of permeabilization should not effect the EC adhesions.  Do not use collagenase since this will disrupt the gel and destroy the sample. 4% paraformaldehyde fixation in cytoskeletal buffer (10mM MES (pH~6.1),138mM KCl, 2mM EGTA(pH ~7),640mM sucrose: from Mitchison lab found online) for 15 minutes, wash once quickly with PHEM (60mM PIPES ,25mM HEPES ,10mM EGTA,2mM MgCl2, pH to 6.9 with NaOH pellets), then  permeabilize with 0.5%triton X-100 in CB for 5 minutes, then rinse 3X, block, etc. Pre-warm all your solutions and perform all steps at 37C since the collagen will often release from the dish.  Your biggest issue is getting the solutions into the u-slides without disrupting the gels.  The above protocol works well in open dishes, like MatTek or the Ibidi u-dish.  With open dishes be sure not to add any solutions directly on top of the gel, only on the sides.  For Ab concentrations double your primaries, but leave your 2ndaries at the same concentration as usual. DO NOT label with the 2ndary too long: 40 minutes or less is all that is needed.  Any more and you will just be fighting background fluorescence when imaging. For your final rinses (after the 2ndary only) add 0.1% Tween-20 to your rinse buffer and rinse at least 5X (10 minutes per rinse) to get rid of the 2ndary. Also do not use any mounting medium that solidifies, since this will compact your gels (I usually just use PBS--). Now being able to imaging them well that is another story.  For High resolution (40X and above) use water lenses if your samples are  greater than 10 microns from the bottom.

Thank you very much for taking your time to post the detailed protocol Mr. Doyle. I will try the protocol .  I am using µ-Slide 4 well Ph+ . So i can directly image my cells using confocal microscope. //pre-warm all your solutions and perform all steps at 37C since the collagen will often release from the dish.//..Its a very important tip..and also the primary antibody concentration. Thank you once again  

Andrew can I have a copy of the publication -thank you.

You have detailed protocol from Mr.Doyle, so that some points in addition to his method. We use culture medium without serum with 20uM Hepes at 37oC to wash before fixation and PFA in the same (see ref). Fixation in 2%PFA (better concentration for actin isoforms and adhesion proteins) for 30 min and permeabilization in 0,5% Triton X-100. To detect a-SMA, incubate overnight at 4oC or 2h at room temperature in monoclonal anti-aSMA antibody (clone 1A4, you can use conjugated with fluorescent probes) and other antibodies, that you need. Minimal wash by PBS between all steps 30 min with 3-5 change. In case of secondary antibodies, don’t incubate them more then 30-40 min. Wash 1h after all steps before mounting. I can propose also to use cold Methanol after PFA instead of Triton (Dugina et. Al., J.Cell Sci, 2009), but it is more difficult to use with collagen. 

Good luck.

Thank you very much Ms. Dugina. I will update how it turned out soon..

So as an update, i followed the protocol with some minor modifications and got really nice results..Stained the ECs for Ve-cadherin and SMCs for alpha actin..