Dilution of protein–surfactant complexes is an integrated step in microfluidic protein sizing, where the contribution of free micelles to the overall fluorescence is reduced by dilution. This process can be further improved by establishing an optimum surfactant concentration and quantifying the amount of protein based on the fluorescence intensity. To this end, we study the interaction of proteins with anionic sodium dodecyl sulfate (SDS) and cationic hexadecyl trimethyl ammonium bromide (CTAB) using a hydrophobic fluorescent dye (sypro orange). We analyze these interactions fluourometrically with bovine serum albumin, carbonic anhydrase, and beta-galactosidase as model proteins. The fluorescent signature of protein–surfactant complexes at various dilution points shows three distinct regions, surfactant dominant, breakdown, and protein dominant region. Based on the dilution behavior of protein–surfactant complexes, we propose a fluorescence model to explain the contribution of free and bound micelles to the overall fluorescence. Our results show that protein peak is observed at 3 mM SDS as the optimum dilution concentration. Furthermore, we study the effect of protein concentration on fluorescence intensity. In a single protein model with a constant dye quantum yield, the peak height increases with protein concentration. Finally, addition of CTAB to the protein–SDS complex at mole fractions above 0.1 shifts the protein peak from 3 mM to 4 mM SDS. The knowledge of protein–surfactant interactions obtained from these studies provides significant insights for novel detection and quantification techniques in microfluidics.