Quantifying and controlling the spatial molecular heterogeneity on biofunctionalized particles is essential for understanding and improving their functionality in bioscience applications. Here, we describe an analysis framework based on single-molecule localization microscopy that can quantitatively assess the spatial molecular properties of affinity molecules conjugated to particles. We performed 3D DNA-PAINT imaging on biofunctionalized particles and established analysis methods to correlate single-molecule data to the particle outer surfaces, count the number of conjugated molecules, and quantify the spatial distributions of the conjugated molecules. We show that imaging data combined with simulation-based molecular counting gives access to high densities of conjugated molecules and enables quantification of their spatial distributions. The analysis is exemplified for particles with a diameter of 1 μm functionalized with single-stranded DNA molecules via two bioconjugation methods, namely, streptavidin-biotin coupling and PLL--PEG-based click chemistry. The data reveal interparticle and intraparticle spatial heterogeneities that are dependent on the bioconjugation methods and conditions. With the analysis framework, 3D DNA-PAINT imaging becomes a versatile characterization technique to study biofunctionalized particles and guide future biofunctionalization strategies for a wide range of applications.