I'm isolating small intestine IEL/LPL following a protocol from Jove “Isolating Lymphocytes from the Mouse Small Intestinal Immune System”, from Qui et al. (2018)

In brief, the washed intestine (cut open longitudinally then into ~2cm pieces) would be digested twice in 1mM DTE solution(with 10% FBS) while being stirred at 37 °C and ~220 rpm in an incubator for 20 min. After each digestion, I vortex the pieces for 10 s and filter the supernatant through a 70-µm cell strainer, combine and pellet the filtered supernatant, then use gradient density centrifugation (44/67 percoll) to collect IELs. The remaining pieces would be further digested by EDTA to remove IECs then collagenase to collect LPLs.

Initially, I was getting great IEL yields from mice ranging from P14 to adult, with a high proportion of CD45+ cells after density gradient centrifugation. But, after a few months my yields seem to have dropped off a cliff despite not change to the protocol or dissection technique. There is still a visible buffy coat at the layer boundary, but trypan blue staining shows a low number of viable cells.

At this point we have purchased brand new DTE (thinking the bottle we were using may have become oxidized), which did not seem to help. I have also checked and refreshed all the other reagents in the DTE solution, including adjusting the pH of the HEPES-bicarb buffer and 10X HBSS. I've also tried increasing the DTE concentration and incubation RPM to no avail. I'm at a complete loss but don't have enough technical knowledge of the protocol to further troubleshoot.

Any advice or ideas would be appreciated!