Recognition of signaling phospholipids by proteins is a critical requirement for the targeting and initiation of many signaling cascades. Most biophysical methods for measuring protein interactions with signaling phospholipids use purified proteins, which do not take into account the effect of post-translational modifications and other cellular components on these interactions. To potentially circumvent these problems, we have developed a single-molecule fluorescence approach to analyzing lipid-protein interactions in crude cell extracts. As a proof of principle for this assay, we show that a variety of lipid-binding domains (LBDs) can be recruited from cell lysates specifically onto their target phospholipids. With single-molecule analysis in real-time, our assay allows direct determination of binding kinetics for transient lipid-protein interactions and has revealed unique assembly properties and multiple binding modes of different LBDs. Whereas single-copy LBDs display transient interaction with lipid vesicles, tandem-repeat LBDs, often used as lipid biosensors, tend to form stable interactions that accumulate over time. We have extended the assay to study a cellular protein, Akt, and discovered marked differences in the lipid binding properties of the full-length protein compared to its PH domain. Importantly, we have found that phosphorylation of Akt at T308 and S473 does not affect the lipid binding behaviors of Akt, contrary to the long-standing model of Akt regulation. Overall, this work establishes the single-molecule lipid pulldown assay as a simple and highly sensitive approach to interrogating lipid-protein interactions in a setting that at least partly mimics the cellular environment.