Abstract
A protocol was developed for the isolation and characterization of extracellular vesicles (EVs) from Coffea arabica cell suspension cultures (CSCs). The isolation method involved differential ultracentrifugation of the CSC filtrate, yielding two fractions: the pellet after 100,000×g for 36 min (100k×g) and the pellet obtained from the previous supernatant after 125,000 g for 6 h (125k×g). Both fractions were characterized by size, morphology, and proteomic profiles (ProteomeXchange identifier PXD071909). While no significant differences in average EV size were observed between the two fractions, proteomic analysis revealed distinct quantitative and compositional variations. The 100k×g fraction was enriched in proteins associated with cell periphery, plasma membrane, and extracellular region, whereas the 125k×g fraction predominantly contained proteins from the extracellular region. Proteomic marker analysis confirmed that both fractions contained protein EV markers, such as transmembrane and transport proteins, soluble EV-associated proteins, and proteins targeted to the extracellular environment or cell wall. Conversely, negligible contamination from non-EV-related proteins was detected. Furthermore, transmission electron microscopy (TEM) showed that the average size of the fractions was consistent with that reported for plant EVs. These findings demonstrate that the protocol utilized to isolate EVs from coffee CSC applies to the release of such vesicles without mechanical harsh grinding that leads to tissue/cell rupture and consequent contamination by other cell components. EVs obtained from coffee CSC represent a valuable and scalable platform, paving the way for the development of tools for biotechnological applications.