In numerous areas of biology and medicine, interference microscopy is proving to be a valuable tool for studying living cells and other biological materials.1 With this technology, it is possible to examine a living cell's biophysical properties—such as stiffness, time-dependent deformability, and mass—to infer a cell's state and its changes in response to various stimuli. In this way, the cells themselves act as biological sensors that integrate local biochemical and biophysical signals into a physiologically relevant, functional readout. A key advantage of using interference microscopy for this task is that it does not require labeling with fluorescent proteins or optically active dyes, and thus both cultured cell lines and material derived from whole organisms can be used. Interference microscopy using a broadband illumination source, also known as white-light optical profilometry, is a commonly used tool in semiconductor and microelectronics manufacturing, where it affords nanometer dimensional precision in height measurement, over wide fields of view. This feature of white-light optical profilometry makes it attractive for precise dimensional measurement of biomaterials as well, which range in size from microns to millimeters.
Recently, we have used white-light optical profilometry to conduct rapid, sub-picogram sensitivity measurements of cell mass to characterize the growth of single cells over the course of minutes and to record the instantaneous response of cells to external chemical stimuli. This technology has also allowed us to rapidly detect the chemotherapeutic response of several breast cancer cell types and of human multiple myeloma cells.2, 3 These measurements relied on the fact that the variation in phase imparted to coherent or semi-coherent light propagating through a transparent cell body is linearly proportional to the material density excluding the aqueous component of the cell.4 Figure 1(a) shows a bright field, and Figures 1(b) and (c) show interferometric phase images of an HCT116 human colon cancer cell line taken with a modified Bruker NT-9300 optical profiler.
Figure 1. Interference microscopy of living colon cancer cells.
We have demonstrated that, within just a few hours, optical profilometry quantifies the sensitivity of single-cell and colony-forming human breast cancer cell lines to the human epidermal growth factor receptor 2 (HER2)-directed monoclonal antibody trastuzumab (brand name, Herceptin).2 We compared the optically measured response to trastuzumab to that determined using traditional multi-day, cell-counting, growth-inhibition assays. In all cases, the trastuzumab sensitivity measured by optical profilometry over 6h was concordant with that measured by cell counting over 3–7 days, thus providing us with critical information about an individual cell type's relative sensitivity to trastuzumab.
Other recently developed live-cell mass profiling methods, such as microelectromechanical systems (MEMS) micro-resonators, can, depending on their configuration, measure the mass of single cells with high accuracy.5However, a drawback to this approach is that to achieve sufficient sensitivity, the active area of the resonator must be on the order of microns or smaller, which makes the continuous measurement of mixtures of single cells and larger multi-cellular colonies, as occurs in most tumor types, very difficult. We have overcome this limitation using optical profilometry. In our pilot studies, we determined the relative sensitivities of small samples (<500 cells) tens to hundreds of times faster than is possible with raditional proliferation assays. These advances in clustered sample assessment and speed open up the possibility for therapeutic response testing of patient-derived solid tumor samples, which are viable only for short periods ex vivo and are likely to be in the form of cell aggregates and clusters.
Figure 2. Drug response data for human multiple myeloma cells determined by tracking cell mass versus time.
Figure 2 shows an example of drug response data for H929 multiple myeloma cells profiled by single-cell mass accumulation.3 Heat map clustergrams of mass accumulation show a differential response in growth profiles for cells treated with 10mg/ml tunicamycin, an agent that disrupts protein processing, versus untreated control cells. In the heat map, red indicates faster growth and the mass of individual cells is normalized starting at 1h post-treatment. Using optical profilometry, we were able to detect both the population average-growth inhibitory effect of tunicamycin and the variation in response among members of the population.
Biomechanical indentation testing of tissues and soft biofilms is another area in which white-light interference microscopy has proven to be an extremely flexible and powerful technique. Mechanical indentation measurements, though widely used, are highly dependent on sample composition and geometry. Optical profilometry allows us great flexibility in sample size and configuration while conducting nanometer-precision vertical displacement measurements over large ranges (up to millimeters).6–8
For instance, conventional mechanical approaches such as uniaxial-tensile testing divide relatively large specimens into relatively long strips that are clamped at each end and pulled apart until they rupture. This geometry is not well suited to interrogate small, radially symmetric tissues such as corneal membranes. In contrast, we have used white-light interference microscopy combined with magnetically actuated force probes to measure and then quantitatively model the viscoelastic properties of all three layers of the bovine cornea, epithelium, endothelium, and stroma.9 These models, which have general validity in the mechanics of materials regardless of microscopic and molecular composition, accurately captured the viscoelastic behavior exhibited by each corneal layer. Furthermore, this type of indentation data can be interpreted in the Hertzian viscoelastic framework suitable for general mathematical characterization appropriate to approaches such as finite element analysis.
In the near future, we will continue to refine the technique of interference microscopy as applied to live-cell mass profiling and biomaterials characterization, and incorporate complementary techniques such as fluorescence microscopy.
Department of Physics
Virginia Commonwealth University
Jason Reed is an assistant professor in the Department of Physics where he develops nanotechnology-based measurement techniques for use in studying biological systems. His current research includes atomic force microscopy-based studies of single molecules and novel applications of interference microscopy in biology and medicine.
Pathology and Laboratory Medicine
University of California, Los Angeles
Los Angeles, CA
Bruker Nano Surfaces Division
1. J. Reed, J. Chun, T. Zangle, J. Hong, M. Teitell, J. Gimzewski, Cells in motion: live cell interferometry (LCI), Abs. Pap. Am. Chem. Soc. 241, 2011.
2. J. Chun, T. A. Zangle, T. Kolarova, R. S. Finn, M. A. Teitell, J. Reed, Rapidly quantifying drug sensitivity of dispersed and clumped breast cancer cells by mass profiling, Analyst 137(23), p. 5495-5498, 2012.
3. J. Reed, J. Chun, T. A. Zangle, S. Kalim, J. S. Hong, S. E. Pefley, X. Zheng, J. K. Gimzewski, M. A. Teitell, Rapid, massively parallel single-cell drug response measurements via live cell interferometry, Biophys. J.
101(5), p. 1025-1031, 2011. doi:10.1016/j.bpj.2011.07.022
4. R. Barer, Interference microscopy and mass determination, Nature 169(4296), p. 366-367, 1952.
5. M. Godin, A. K. Bryan, T. Burg, K. Babcock, S. R. Manalis, Measuring the mass, density, and size of particles and cells using a suspended microchannel resonator, Appl. Phys. Lett. 91(12), p. 123121, 2007.
6. J. Reed, M. Frank, J. Troke, J. Schmit, S. Han, M. Teitell, J. K. Gimzewski, High-throughput cell nanomechanics with mechanical imaging interferometry, Nanotechnology 19, p. 235101, 2008.
7. J. Reed, J. Troke, J. Schmit, S. Han, M. A. Teitell, J. K. Gimzewski, Live cell interferometry reveals cellular dynamism during force propagation, ACS Nano.
2(5), p. 841-846, 2008. doi:10.1021/nn700303f
8. J. Reed, W. J. Walczak, O. N. Petzold, J. K. Gimzewski, In situ mechanical interferometry of matrigel films, Langmuir 25(1), p. 36-39, 2009.
9. L. Yoo, J. Reed, J. K. Gimzewski, J. L. Demer, Mechanical interferometry imaging for creep modeling of the cornea, Invest. Ophthalmol. Vis. Sci. 52(11), p. 8420-8424, 2011.