Recent studies suggest a role for autocrine insulin signaling in beta cells, but the mechanism and function of insulin-stimulated Ca²⁺ signals is uncharacterized. We examined Ca²⁺-dependent insulin signaling in human beta cells. Two hundred nanomolar insulin elevated [Ca²⁺]_c to 284 ± 27 nM above baseline in ≈30% of Fura-4F-loaded cells. Insulin evoked multiple Ca²⁺ signal waveforms, 60% of which included oscillations. Although the amplitude of Ca²⁺ signals was dose-dependent between 0.002 and 2,000 nM, the percentage of cells responding was highest at 0.2 nM insulin, suggesting the interaction of stimulatory and inhibitory pathways. Ca²⁺-free solutions did not affect the initiation of insulin-stimulated Ca²⁺ signals, but abolished the second phase of plateaus/oscillations. Likewise, inositol 1,4,5-trisphosphate (IP₃) receptor antagonists xestospongin C and caffeine selectively blocked the second phase, but not the initiation of insulin signaling. Thapsigargin and 2,5-di-tert-butylhydroquinone (BHQ) blocked insulin signaling, implicating sarcoplasmic/endoplasmic Ca²⁺-ATPase (SERCA)-containing Ca²⁺ stores. Insulin-stimulated Ca²⁺ signals were insensitive to ryanodine. Injection of the CD38-derived Ca²⁺ mobilizing metabolite, nicotinic acid-adenine dinucleotide phosphate (NAADP), at nanomolar concentrations, evoked oscillatory Ca²⁺ signals that could be initiated in the presence of ryanodine, xestospongin C, and Ca²⁺-free solutions. Desensitizing concentrations of NAADP abolished insulin-stimulated Ca²⁺ signals. Insulin-stimulated Ca²⁺ signals led to a Ca²⁺-dependent increase in cellular insulin contents, but not secretion. These data reveal the complexity of insulin signal transduction and function in human beta cells and demonstrate functional NAADP-sensitive Ca²⁺ stores in a human primary cultured cell type.