The purpose of this study was to develop and characterize a functional air-interfaced primary culture of rabbit conjunctival epithelial cells grown on a permeable support for drug transport studies. Conjunctival epithelial cells from the pigmented rabbit were isolated, seeded at 1.2×10 6 cells cm −2 on permeable Transwell filters, and cultured at the air interface using a modified PC-1 medium. Conjunctival epithelial cell layers showed a transepithelial resistance of 1.1±0.1 kΩ cm 2, a potential difference of 17.0±0.5 mV, and an equivalent short-circuit current ( I eq) of 16.1±0.4 μA cm −2. The I eq was reduced by 35% using 0.01 mM bumetanide, 66% using 0.1 mM ouabain, 46% using 2 mM barium chloride (all three in the basolateral fluid), and 63% using 0.3 mM NPAA in the apical fluid, consistent with active Cl −-secretion across the conjunctival epithelial barrier. Amiloride-sensitive Na + channels were absent. The permeability of the cell layers to polar solutes decreased with increased solute size, and the calculated equivalent pore size was about 8.0 nm. The Papp of β-blockers varied with lipophilicity in a sigmoidal fashion. Uridine transport showed temperature sensitivity and directionality, favoring transport in the apical-to-basolateral direction. Apical l-carnosine uptake was reduced by 46% in the absence of an inwardly directed proton gradient, and lowering the temperature to 4°C abolished direction-dependent l-carnosine uptake. Furthermore, uptake was inhibited by 73% using apical 10 mM glycyl sarcosine (a dipeptide transporter substrate) and by 60% using 1 mM l-valacyclovir (a dipeptide prodrug). In conclusion, a functional air-interfaced primary culture of rabbit conjunctival epithelial cell layers was established. This air-interfaced primary culture model may be useful for studying passive and active transport processes for ion and solute translocation in the mammalian conjunctival epithelial barrier in a defined experimental setting.