Abstract
The burgeoning fields of in vivo three-dimensional (3D) microscopy and endomicroscopy, as well as ex vivo tissue cytometry have introduced new challenges for tissue preparation and staining with exogenous molecular contrast agents. These challenges include effective delivery of the agents, and once delivered, distinguishing between bound verses unbound molecular probes. If applied topically, there are additional issues with rinsing off unbound probe, which can be nonuniform and inefficient in thick tissues, thus leading to ambiguous contrast and a large nonspecific background that may obscure molecule-specific staining. Therefore, we have developed a ratiometric 3D microscopy scheme that not only reduces the effects of nonspecific sources of contrast, but also enables quantification of the relative binding affinity of imaging probes to their biomarker targets. Here we demonstrate this ratiometric approach by simultaneously imaging a HER2/neu (erbB2)-targeted monoclonal antibody labeled with one fluorophore and an isotype-matched negative control antibody labeled with another fluorophore. By taking a pixel-by-pixel calibrated ratio between the signals from each fluorescent image channel, accurate quantification of specific versus nonspecific binding affinity is achieved with cultured cells, yielding data that are in agreement with analyses via flow cytometry. We also demonstrate quantitative 3D microscopic imaging of biomarker expression in tissue models and in thick human biopsy samples of normal, HER2-negative, and HER2-positive breast tumors. This strategy enables rapid, quantitative, and unambiguous volumetric microscopy of biomarker expression in thick tissues, including whole biopsies, and will enable real-time optical assessment of disease markers in the living body.