Hey everyone, I've been scratching my head over this in the lab lately. Last week I was running some Western blots trying to detect a primary that's an IgM, and the signal from the secondary just looked way weaker and spottier than when I use IgG primaries. It got me thinking—why does that pentameric setup of IgM make it trickier for secondary antibodies to bind properly compared to the straightforward monomeric IgG? Back in my postdoc days we mostly stuck to IgG stuff and never had these headaches, but now it's driving me nuts during troubleshooting. Anyone dealt with this and figured out what's going on structurally?
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Yeah, I've bumped into the exact same frustration a bunch of times. The whole pentameric thing with IgM means it's this big bulky molecule with five units stuck together, so the Fc regions aren't as nicely exposed or accessible like on the smaller, single-unit IgG. That can lead to some steric crowding where the secondary struggles to latch on effectively, especially if the primary's oriented weirdly on the membrane after transfer. I've noticed signals often come out fainter unless I tweak concentrations or blocking steps a lot more. For what it's worth if you're digging deeper into the differences, check out this page on lgm full form here—it's just a quick read that helped me wrap my head around the basics without all the hype. Personally I think sticking with mu-chain specific secondaries helps a ton instead of generic ones, but yeah it's definitely more finicky than IgG work in my experience.