Physical Chemistry, Biophysics, and Analytical Chemistry; Magnetic Resonance Spectroscopy; Investigation of dynamic biological and materials systems, with particular interest in hydration water.

Elings Prize Postdoctoral Fellow, 2009-2013

Our goal: Our lab develops unorthodox spectroscopic techniques to investigate how water interacts with the surfaces of proteins and other macromolecules. The resulting insight will improve our ability to design drugs and synthetic materials. In particular, we specialize in magnetic resonance techniques that communicate with both electron and nuclear spins to probe nanometer-scale structure and dynamics.

Why hydration water? While tools have been developed for characterizing static structures, we still need techniques that can study dynamic arrangements of molecules and, most urgently, the arrangement of water molecules. Even though water appears uniform on a macroscopic scale, it behaves quite differently on the nanoscale. If we zoom in to the first few layers of water molecules near a surface, we see that hydrogen bonds break and individual water molecules move at rates anywhere from a couple times slower to hundreds of times slower than those in the bulk liquid. In both biomolecules and synthetic materials, we know that these differently behaved “hydration waters” are key to determining interactions.

Initial examples: As a fundamental example, we looked at lipid bilayers – structures that make up cell membranes. We developed (at other institutions) a specialized technique to characterize water near the bilayer surfaces. Our technique – called Overhauser Effect Dynamic Nuclear Polarization (ODNP) [1] – tracks the motion of the water nuclei, and allows us to zoom in on the “hydration water” near the surface of the lipid bilayer (figure below). It allowed us to see that the hydration water moves about 5 times slower than bulk water [2]. Furthermore, by dramatically changing the characteristics of the bulk solution, we learned that an order of magnitude change in the viscosity barely affects the diffusion in the hydration layer [2]. We have also implemented this technique in systems with membrane proteins [3], DNA [4], and large protein folding chaperones [5]. The significant variations in the properties of the hydration water play in important role in how these surfaces interact in nature.

Caption: This image illustrates ODNP of water in the hydration layer of a lipid bilayer. This is one example of many studies that are possible with ODNP. A lipid bilayer (the basis for cellular membranes) consists of a self-organized set of molecules with hydrophilic headgroups and hydrophobic tails (see labels above). One can place a spin label (typically a nitroxide group) at a particular location near the surface of the lipid bilayer (via chemical synthesis). A combination of electron- and nuclear-spin resonance (i.e. ODNP) can selectively interrogate the motion of water molecules at a specific location – here inside the hydration layer of the lipid bilayer. For example, employing the setup shown above, we were able to verify that the motion of water molecules within the hydration layer was not sensitive to the presence of a high concentration of large molecules that are not chemically disposed to disrupt the hydration layer. This was even true when the large molecules significantly altered the viscosity of the bulk solution [2].

The future: In the Franck lab, we are extending the power of ODNP-based techniques and integrating them with a toolbox of previously developed techniques, for example: NMR techniques that can measure microscopic diffusion and flow, as well as electron spin resonance (ESR) techniques that interrogate local motions (cw ESR), that measure nanometer-scale distances (double electron-electron resonance, i.e. “DEER”) and that map out local arrangements of nuclei (electron-nuclear double resonance, i.e. “ENDOR” and electron spin echo envelope modulation, i.e. “ESEEM” [5]). Towards these ends, we will draw on expertise in spectrometer design [6–8] and magnetic resonance methods [9,10]. We will also apply techniques that we have already developed to systems that remain poorly understood, despite their central importance to understanding materials function or to understanding disease. For example, the Ras protein system is integral to cell signal pathways and the development of novel cancer drugs, polymer electrolyte membranes are essential to ion transport in H2/O2- and methanol-based fuel cells – as well as sensor devices, while biological light-harvesting systems, such as Photosystem II, are believed to integrate nanoscale water channels to aid in their function. Nonetheless, the interplay between all these systems and water has yet to be fully explored.

The Franck lab will be accepting several new graduate students and undergraduate researchers in Fall 2017, and welcomes any inquiries. (feel free to contact Dr. Franck by email). Students in the Franck lab will have the opportunity to work with materials or biological systems. In addition, they will gain expertise developing new spectroscopic methods using physics, electronics, and computer programming. Students who capitalize on these opportunities will have a unique and powerful skill-set.

Franck JM, Pavlova A, Scott JA, Han S. Quantitative cw Overhauser effect dynamic nuclear polarization for the analysis of local water dynamics. Prog Nucl Magn Reson Spectrosc. Elsevier B.V. 2013 Oct;74:33–56. PMID: 24083461

Franck JM, Scott JA, Han S. Nonlinear scaling of surface water diffusion with bulk water viscosity of crowded solutions. J Am Chem Soc. 2013 Mar;135(11):4175–8. PMID: 23347324

Hussain S, Franck JM, Han S. Transmembrane protein activation refined by site-specific hydration dynamics. Angew Chem, Int Ed Engl. 2013 Feb;52(7):1953–8. PMID: 23307344

Franck JM, Ding Y, Stone K, Qin PZ, Han S. Anomalously Rapid Hydration Water Diffusion Dynamics Near DNA Surfaces. J Am Chem Soc. 2015 Sep;137(37):12013–12023.

Franck JM, Sokolovski M, Kessler N, Matalon E, Gordon-Grossman M, Han S-I, Goldfarb D, Horovitz A. Probing Water Density and Dynamics in the Chaperonin GroEL Cavity. J Am Chem Soc. 2014 Jul;136(26):9396–403. PMID: 24888581

Franck JM, Barnes RP, Keller TJ, Kaufmann T, Han S. Active cancellation – A means to zero dead-time pulse EPR. J Magn Reson. 2015 Dec;261:199–204.

Kaufmann T, Keller TJ, Franck JM, Barnes RP, Glaser SJ, Martinis JM, Han S. DAC-board based X-band EPR spectrometer with arbitrary waveform control. J Magn Reson. 2013 Oct;235:95–108. PMID: 23999530

Demas V, Franck JM, Bouchard LS, Sakellariou D, Meriles CA, Martin R, Prado PJ, Bussandri A, Reimer JA, Pines A. “Ex situ” magnetic resonance volume imaging. Chem Phys Lett. 2009 Jan;467(4-6):398–401.

Franck JM, Chandrasekaran S, Dzikovski B, Dunnam CR, Freed JH. Focus: Two-dimensional electron-electron double resonance and molecular motions: The challenge of higher frequencies. J Chem Phys. 2015;142(21):212302.

Franck JM, Demas V, Martin RW, Bouchard L-S, Pines A. Shimmed matching pulses: simultaneous control of rf and static gradients for inhomogeneity correction. J Chem Phys. 2009 Dec;131(23):234506. PMID: 20025334