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Workshop: The future of integrative structural biology

April 29th, 2017

Watt Family Innovation Center

Clemson University

The grand challenge for structural biologists is to build a complete molecular atlas of the cell. Rather than the mere acquisition and accumulation of high-resolution structural data from any one of multiple biochemical, biophysical and computational approaches, the complete molecular atlas of the cell will require “integrative structural biology” approaches relying on data from multiple methodologies. Structural biologists must integrate high resolution and dynamic data through comprehensive analyses of large, dynamic, macromolecular machines. During this workshop, we will discuss current developments in integrative structural biology though a combination of Electron Microscopy (EM), Electron Paramagnetic Resonance (EPR), Förster Resonance Energy Transfer (FRET), X-ray scattering, and computational methods. Participants will be invited to make recommendations on the field to tackle upcoming challenges such as bridging multiscale models and their corresponding representation, and to consider the fact that biomolecules are in constant motion.

Organizers

Hugo Sanabria, Ph.D. Joshua Alper, Ph.D.

Department of Physics Clemson University

Session Chairs

Laxmikant Saraf, Ph.D. Director Electron Microscope Laboratory

Kerry S. Smith, Ph.D.

Director of the Eukaryotic Pathogens Innovation Center

Feng Ding, Ph.D. Department of Physics and Astronomy

Terri Bruce, Ph.D.

Director Light Imaging Facility

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Sponsors and Acknowledgements

Oak Ridge Associated Universities

Mark D. Leising, Ph.D. Interim Dean of the College of Science

Terry M. Tritt, Ph.D.

Interim Chair of the Department of Physics and Astronomy

William R. Marcotte, Ph.D. Chair of the Department of Genetics and Biochemistry

Kerry S. Smith, Ph.D.

Director of the Eukaryotic Pathogens Innovation Center

American Friend of the Alexander von Humboldt Foundation

Administrative Support

Amanda Crumpton Graduate Student Service Coordinator

Lori Rholetter

Administrative Assistant

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The purpose of the Workshop: The future of Integrative biology is to address the ever-changing landscape within integrative structural biology. The logo highlights five methodologies that have either stood the test of time, or are just now bursting onto the scene of structural biology. Improvement in old techniques and the possibilities to integrate various sources of information to resolve a complete molecular atlas of the cell at various length scales. Credit of Background picture: Xiaowei Zhuang, HHMI, Harvard University, Nature Methods, and Nature Publishing Group. Designed by Zachary Disharoon (B.S. Physics Spring 2017)

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Agenda 8:00 am Welcome Remarks Lesly Temesvari Ph.D., Interim Associate Dean for Research, College of Science. 8:15 am -10:00 am Morning Session I Session Chair: Lax Saraf Ph.D., EM Lab Director Catherine L. Lawson, Ph.D. Rutgers University Daniela Nicastro Ph.D. University of Texas Southwestern Medical Center Elizabeth R. Wright Ph.D. Emory University School of Medicine Bin San Chan, Ph.D. Clemson University * 10:00 am – 10:30 am Coffee Break 10:30 am – 12:15 pm Morning Session II Session Chair: Kerry Smith, Ph.D., EPIC Director Daniel Keedy, Ph.D. University of California San Francisco Satyanarayana Lagishetty, Ph.D. Clemson University * Tatyana Smirnova, Ph.D. North Carolina State University Mark Bowen, Ph.D. Stony Brook University 12:15 pm -2:00 pm Lunch brown bag / Poster session 2:00 pm -3:00 pm Afternoon Session I Session Chair: Feng Ding Ph.D., Physics and Astronomy Mark Bathe, Ph.D. Massachusetts Institute of Technology Sichuan Yan, Ph.D. Case Western Reserve University 3:00 pm -3:30 pm Coffee Break / Poster Session 3:30 pm -4:15 pm Round Table I Session Chair: Terri Bruce, Ph.D., Clemson Light Imaging Facility Director Small groups discussions and panel summary. 4:15 pm – 4:30 pm Coffee Break Poster session 4:30 pm - 5:15 Round Table II Session Chair: Joshua Alper Ph.D., Physics and Astronomy Small group discussion and panel summary. 6:00 pm Dinner Twelve Mile, Hartwell Lake

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Watt Family Innovation Center No food or beverages are allowed in the auditorium. Accommodations Visitors will be staying at the James F. Martin Inn located on the campus of Clemson University. The conference rate is $120 (double) and is available for Friday, April 28 and Saturday, April 29. Transportation from the hotel to the venue will be provided. Transportation The closest airport to the Clemson area is the Greenville-Spartanburg International Airport (GSP), which is 45 minutes by car. You may find it more convenient to fly to the larger Hartsfield-Jackson Atlanta International Airport (ATL) or Charlotte Douglas International Airport (CLT), which are 2-3 hours away by car. The Amtrak Crescent train, which connects New Orleans to New York, stops twice a day (once northbound and once southbound) in Clemson. In the town of Clemson and surrounding areas, there is free public transportation via CAT Bus. The Greenville Greenlink bus system has a Clemson Commuter route which connects Clemson to Greenville. Twelve Mile Recreation Area 12 Mile Park Rd, Clemson, SC 29631 Limited shuttle service to the dinner location at Twelve Mile Recreation Area will be provided after the end of the last session. Getting There: GPS Info. (Latitude, Longitude): 34.70444, -82.83444 34°42'16"N, 82°50'4"W

Directions: From the corner of 93 and 133, right in front of Bowman Field head north on 133 through downtown Clemson. Follow 133 for about 1.6 miles. The entrance to the Twelve Mile Recreation Area is on your left.

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Once in the park, continue straight about 200 yards. The pavilion is located on the right.

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Speakers Biographies Mark Bathe, Ph.D. (mbathe@mit.edu) Massachusetts Institute of Technology Professor Bathe obtained his Bachelor’s, Master’s and Doctoral Degrees at MIT working in the Departments of Mechanical Engineering, Chemical Engineering, Chemistry, and Biological Engineering before moving to Munich to carry out his postdoctoral research in Biological Physics at the University of Munich. He returned to MIT in 2009 to join the faculty in the Department of Biological Engineering, where he runs an interdisciplinary research group focused on developing integrated computational-experimental approaches for engineering biology. Major research thrusts of his group include programming structured nucleic acid assemblies for mRNA and CRISPR/Cas9 delivery, nanoscale energy circuits and fluorescent barcodes, and multiplexed fluorescence imaging of neuronal circuits for phenotypic profiling. Mark Bowen, Ph.D. (mark.bowen@stonybrook.edu) Stony Brook University Mark Bowen was born in Duluth, MN, on the frigid North Coast of the United States. He obtained a Bachelor of Arts in Chemistry from the University of Minnesota at Morris, a public, residential liberal arts college. After graduation, Mark worked in the chemical industry and as a home health aide. He obtained a Ph.D. from the University of Illinois at Chicago in Biochemistry under the mentorship of Dr. Peter G.W. Gettins. This work investigated alpha-2-Macroglobulin, a molecular trap found in human plasma capable of enveloping proteases within a polypeptide cage. After receiving his doctorate in 1998, Mark took a Howard Hughes Medical Institute postdoctoral position at Yale University in the Department of Molecular Biophysics and Biochemistry with Dr. Axel Brunger working on membrane protein structure and function. Mark next moved with the Brunger lab to Stanford University and joined the Department of Molecular and Cellular Physiology. At Stanford, he worked with Dr. Steven Chu on single molecule approaches to understanding the molecular mechanism of synaptic transmission. Mark Bowen is currently an Associate Professor at Stony Brook University in the Department of Physiology and Biophysics; he is also a member of the graduate programs in Biochemistry and Structural Biology, Chemical Biology, Molecular Biology, Molecular and Cellular Pharmacology & Neuroscience. Daniel Keedy, Ph.D. (Daniel.Keedy@ucsf.edu) University of California San Francisco Dr. Keedy obtained his Ph.D. at Duke University with Jane and David Richardson, where he studied protein flexibility in structure validation, prediction, and design. As a postdoctoral fellow at UCSF with James Fraser, he has exploited multitemperature X-ray crystallography and multiconformer structural modeling to identify allosteric networks and binding sites in dynamic proteins.

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Catherine Lawson, Ph.D. (cathy.lawson@rutgers.edu) Rutgers University Dr. Lawson is an Associate Research Professor at the Rutgers University Center for Integrative Proteomics Research. She is a Structural Biologist with expertise in X-ray crystallography and 3D electron microscopy (3DEM) reconstruction structure determination methods. She has contributed more than 40 structures of macromolecular complexes to public data archives, including protein-DNA complexes, DNA duplexes, vaccine targets in complex with antibody Fabs, enzymes, and viral assemblies. As a member of the Research Collaboratory for Structural Bioinformatics (RCSB) team, her recent work has focused on development and improvement of structural biology data archive resources. Key efforts have included remediation of the icosahedral and helical virus structures in the Protein Data Bank and development of resources for archiving of 3D density maps and coordinate models determined using 3DEM methods (http://emdatabank.org). Daniela Nicastro, Ph.D. (Daniela.Nicastro@UTSouthwestern.edu) University of Texas Southwestern Medical Center Daniela Nicastro received her Ph.D. in Biology from the Ludwig-Maximilians University in Munich, Germany in 2000. Following 3 years in the lab of Prof. Baumeister at the Max-Planck Institute for Biochemistry in Munich (1998-2001), she took a postdoctoral fellow position in the National Center for Research Resources for 3D Electron Microscopy of Cells at the University of Colorado in Boulder. From 2006-2015, she was an Assistant and then tenured Associate Professor of Biology and Director of the “Correlative Light and Electron Microcopy” (CLEM) facility at Brandeis University near Boston. Since July 2015, she is an Associate Professor at the University of Texas Southwestern (UTSW) Medical Center in Dallas with appointments in the Departments for Cell Biology and Biophysics. She has almost 25 years of experience in electron microscopy of cellular structures and is a leading expert in cellular cryo-electron tomography. The research interest of the Nicastro lab is focused on studying the three-dimensional structure and function of cytoskeletal assemblies, molecular motors, organelles and cells using a combination of cutting-edge methods to elucidate the structure-function relationships of macromolecular complexes in situ, i.e. in their native environment. Tatyana Smirnova, Ph.D. (tismirno@ncsu.edu) North Carolina State University Tatyana Smirnova is an Associate Professor of Chemistry at North Carolina State University. She received Ph.D. in Chemistry from the University of Illinois at Urbana-Champaign (UIUC) under the supervision of Prof. R. Linn Belford in 1997. She was a NIH postdoctoral Fellow at the UIUC before accepting Assistant Professor position at NCSU. Her research interests include applications of Electron Paramagnetic Resonance in combination with spin labeling to probe structure and dynamics of biomolecules.

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Elizabeth R. Wright, Ph.D. (erwrigh@emory.edu) Emory University School of Medicine Elizabeth Wright received her B.S. in Biology and Chemistry from Columbus State University, Columbus, GA in 1995 and 1997, respectively. She received her Ph.D. from the Department of Chemistry at Emory University, Atlanta, GA in 2002. She was a postdoctoral associate in the laboratory of Professor Anupam Madhukar at the University of Southern California to develop approaches for bridging nanoscale biological and quantum-based materials. From there, she was a postdoctoral scholar in the cryo-EM laboratory of Professor Grant Jensen and Caltech. During that time, she received an NIH NRSA postdoctoral fellowship to study the 3D structure of HIV. While there, she determined the 3D structure of the immature virus and proposed that six-helix bundles formed by the SP1 domain of the Gag polyprotein stabilize the immature capsid protein lattice. In 2008, she joined the faculty at Emory University in the Department of Pediatrics. She is also the director of the University-wide EM facility. She has pioneered the development and application of numerous technologies including: affinity grid methods for capturing viruses and other complexes on EM grids; self-pressurized rapid freezing and CEMOVIS of bacteria; conventional and cryo-preservation methods for structural studies of the lipopolysaccharide (LPS) and capsular polysaccharide (capsule) of bacteria; enhanced phase contrast (both Zernike and hole-free) cryo-EM/cryo-ET; and cryo-correlative light and electron microscopy (cryo-CLEM) methods for studying bacteria, virus, and eukaryotic cell structure and function. Her present work is focused on structural studies of pathogenic enveloped viruses and bacteria, as well as the development of methods for improving correlative structural biology. Sichun Yang, Ph.D. (sichun.yang@case.edu) Case Western Reserve University Dr. Sichun Yang started his biophysical training at the University of California San Diego after receiving BS and MS in Physics from China. He obtained a Ph.D. in Biophysics from UCSD (2006), where his research with Prof. Herbert Levine and Prof. Jose Onuchic has been focused on protein folding. Dr. Yang pursued his postdoctoral training with Prof. Benoit Roux at the University of Chicago, where he showed that combining computer simulation and synchrotron X-ray scattering is well positioned to study the structural dynamics of biomolecular complexes. This work has been featured in a News & Views article. Around this time, Dr Yang has started studying estrogen receptor in collaboration with Dr. Geof Greene at UChicago, which had received the idea award from the DoD Breast Cancer Research Program. Dr. Yang joined the School of Medicine at Case Western Reserve University in 2010. He focuses on the biophysics and translational studies of estrogen receptor (ER), a key driver for breast cancer growth. He established a multi-technique iSPOT platform for integrative structure modeling of protein-protein complexes that are unattainable by traditional approaches. Using iSPOT, his lab has provided a first molecular view of the multidomain ER complex, thereby addressing the long-standing mystery about its allosteric domain cross-talk. Dr. Yang has contributed to technology development of multiple data analysis algorithms for small-angle X-ray scattering (SAXS) and protein footprinting techniques (supported by NIH, DoD and ACS), further leading to collaboration with colleagues at Cleveland and other parts of the globe.

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Abstracts

Archiving Structures Derived from Diverse Experimental Methods Catherine L. (Cathy) Lawson

EMDataBank and RCSB-PDB, Center for Proteomics Research at Rutgers University,

Piscataway, NJ USA Abstract The Protein Data Bank (PDB; www.pdb.org) now holds nearly 130,000 structures of biological macromolecules based on X-ray crystallography, NMR, and electron microscopy (3DEM) experimental data. Considerable effort has gone into understanding how to curate data coming from these methods. Over the past decade, the Worldwide Protein Data Bank [1] (wwPDB) established expert Task Forces [2-5] to advise as to which primary data should be archived from each method and how the data should be used to validate the resulting structures. Today, structures of important large macromolecular machines are being determined by hybrid methods that may include restraints derived from mass spectrometry, small angle scattering, chemical crosslinking, antibody localization, fluorescence resonance energy transfer (FRET), electron paramagnetic resonance, and/or H/D exchange. A Hybrid Methods Task Force convened by the wwPDB in October 2014 recommended that such structures should be archived in the PDB, and that archiving of supporting experimental data from additional methods, which is not currently done, should be explored [6]. Here, I will give an overview of data deposition and curation policies and practices for X-ray and 3DEM structures, with the goal of generalizing to a framework for archiving of supporting experimental data from diverse methods such as FRET. References [1] Berman, H.M., Henrick, K., and Nakamura, H., “Announcing the worldwide Protein Data Bank”, Nature Structural Biology, 10, 980 (2003). [2] Read, R.J., Adams P.D., Arendall III W.B., Brunger A.T., Emsley P., et al., “A New Generation of Crystallographic Validation Tools for the Protein Data Bank”, Structure, 19, 1395 (2011). [3] Montelione, G.T., Nilges M., Bax A., Güntert P., Herrmann T., et al., “Recommendations of the wwPDB NMR Validation Task Force”, Structure, 21, 1563 (2013). [4] Henderson, R., Sali A., Baker M.L., Carragher B., Devkota B., et al., “Outcome of the First Electron Microscopy Validation Task Force Meeting”, Structure, 20, 205 (2012). [5] Trewhella, J., Hendrickson W.A., Kleywegt G.J., Sali A., Sato M., et al., “Report of the wwPDB Small-Angle Scattering Task Force: Data Requirements for Biomolecular Modeling and the PDB”, Structure, 21,875 (2013) [6] Sali A., Berman H.M., Schwede T., Trewhella J., Kleywegt G.J., et al., “Outcome of the First wwPDB Hybrid/Integrative Methods Task Force Workshop”, Structure, 23, 1156 (2015).

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Probing the Molecular Organization of Cells and Organelles using Cryo-Electron Microscopy

Daniela Nicastro

Departments of Cell Biology and Biophysics, UT Southwestern Medical Center, Dallas, TX, USA Abstract Rapid freezing of cells can provide outstanding structure preservation and time resolution of dynamic cellular processes. Electron tomography of vitreous specimens (cryo-ET) is a powerful technique for imaging biological structures in their native state and in an unperturbed cellular environment. We integrate high resolution imaging by either cryo-ET and sub-tomogram averaging or TYGRESS (Tomography-Guided 3D Reconstruction of Subcellular Structures), with comparative genetics, biochemical methods and EM-visible labeling to deconstruct the 3D structure, functional organization and molecular mechanisms of protein complexes inside cells. Among different model systems, we use e.g. cilia and flagella to advance techniques and approaches for high-resolution imaging of complex cellular structures. Cilia and flagella are conserved and ubiquitous eukaryotic organelles that are composed of more than 600 different proteins and have important biological roles in motility and sensation. Our cryo-ET studies visualize the three-dimensional structures of intact wild-type and mutant flagella, and dissect the organization of key macromolecular complexes in different functional states. Such information can provide detailed insights into the structural basis and ultimately the function of many cellular processes.

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Bridging Resolutions: New Insights in Structural Virology from Cryo-Correlative Light and Electron Microscopy

Elizabeth R. Wright

Division of Infectious Diseases, Department of Pediatrics, Emory University School of Medicine, Children's Healthcare of Atlanta, Atlanta, GA, USA

Abstract Developments in technologies surrounding cryo-EM and light microscopy have transformed the fields of correlative microscopy and structural virology. It is now possible to achieve atomic and near atomic resolution structures of macromolecular complexes and viruses via cryo-EM and cryo-ET. In addition, technologies have provided a means for probing the spatiotemporal landscapes of cells and viruses at high resolution and in four-dimensions. My laboratory uses cryo-EM and molecular biology approaches to explore the three-dimensional (3D) structures of viruses and cells. In this talk, I will discuss some of our contributions to the fields of electron microscopy and structural virology. In particular, I will present data on our cryo-EM structural investigations of pleomorphic enveloped viruses, namely respiratory syncytial virus (RSV) and measles virus. I will also present results of our cryo-EM methods development as applied to studies of viruses and cells. I will conclude by briefly outlining some of our future directions and challenges facing the field.

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Chitin Facilitated Mineralization in the Eastern Oyster Vera Bin San Chan, Marybeth Johnston, Alfred P. Wheeler, Andrew S. Mount

Department of Biological Sciences, Clemson University, Clemson, SC, USA

Abstract Chitin is known to be an important component of molluscan shell for mechanical strength. We investigated the origin of chitin and extracellular matrix with an implantation method developed to study shell repair. To locate the site of chitin deposition during the experimentally induced oyster shell repair, we have cloned a chitin specific fluorescent probe useful for immunofluorescence observation on the laser confocal microscopy. Chitin fibrils were observed by electron microscopy. We tested the chemical reactivity of the fibril’s dimension to acid and bleach, and noted a size reduction after both treatments. Our results confirmed the model that chitin is supplied by chitin synthase on the lipid membrane, and chitin interacts with inorganic minerals and shell proteins. The source of lipid membrane may contain microvesicles, such as exosomes and chitosomes. Cellular mediated nucleation of nanocrystals plays an important first step in overcoming the first energy barrier of mineralization essential to the assembly and growth of the biomaterial. It was observed that calcification follows the deposition of chitin and lipid membrane during the remineralization process of the Eastern oysters. We suggest that correlative microscopy methods are reliable to understand the dynamic cellular process during shell growth and regeneration.

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From the Biophysics of Allostery to Conformational Control of a Dynamic Phosphatase Daniel Keedy1, Zachary Hill2, Emily Kang2, Justin Biel1, Tobias Krojer3, Nicholas Pearce3,

Patrick Collins4, Jose Brandao-Neto4, Frank von Delft3,4, Jim Wells2, James Fraser1

1 Department of Bioengineering and Therapeutic Sciences, 2 Department of Cellular & Molecular Pharmacology, University of California, San Francisco, USA 3 Structural Genomics Consortium, University of Oxford, UK

4 Diamond Light Source Ltd, Harwell Science and Innovation Campus, Didcot, UK Abstract Proteins have the inherent capability to send conformational signals through their structures. This process, known as allostery, is important for regulating function in many proteins. However, we lack an understanding of how allostery works at the basic atomic level. This gap limits our understanding of natural allosteric regulatory strategies, and prevents us from designing allosteric modulators to control the functions of specific proteins, such as human phosphatases. To overcome this hurdle, we have combined novel automated X-ray crystallographic, multiconformer modeling, and chemical biology methods to elucidate allosteric mechanism in the human phosphatase PTP1B, a well-validated but challenging therapeutic target for diabetes. The results reveal small molecules that bind at new allosteric sites, alter PTP1B’s conformational ensemble, and in some cases inhibit enzyme activity. This work sets the stage to deeply interrogate how perturbations bias the conformational ensemble of a protein to control its function, with broad implications for allosteric drug design and protein engineering.

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Crystal structures Phosphofructokinases Family Members: Structural Variation Based on Enzyme Functions

Satyanarayana Lagishetty

Department of Genetics and Biochemistry, Clemson University, Clemson, SC, USA Abstract Phosphopfructokinases (Pfk-B) is large kinase family with different enzymatic functions, phosphorylates variety of sugar/non sugar substrate molecules, crucial five carbon ribose to six carbon fructose or 2-Keto-3-deoxygluconate and also nucleosides adenosine. The entire PfkB family members are classified based on main “GAGD” motif at ATP binding region. The sub-families are classified based on “GG” and “NxxE” motifs at the substrate-ATP binding interface. Even though PfkB family members share common two-three domains structural organization, they activate diverse metabolic products by phosphorylation with interesting structural and biochemical properties. Several Pfk-B family genes from pathogenic and non-pathogenic organisms were cloned, expressed, purified, crystallized and solved the structure using Single/Multiple wavelength Anamolous Dispersion method (SAD/MAD) and Molecular Replacement (MR) method. Crystal structures of several Pfk-B members shows variations at the substrate binding pocket to accommodate different molecular structural scaffolds of substrates keeping overall three 3D fold similar (PDB id: 3Q1Y, 3KTN, 3LKI, 3LJS, 3KTN, 3K5W, 3K9E, 3KD6, 3JUL, 3IN1 etc). Also, there were significant secondary structural variations results in various oligomerization states and domain flexibility/moments during the substrate and cofactor binding. The structural functional relationships and evolution of PfkB family members in to various sub-family levels were discussed.

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Spin-labeling EPR as a Tool for Structural Biology: from Disordered Proteins to Membrane Protein Complexes in Native Bilayer Environment

Keith Weninger1, Alex Smirnov,2 Sergey Milikisiyants,2 Leonid S. Brown,3 Vladimir Ladizhansky3, Shenlin Wang4, Tatyana Smirnova2

1 Department of Physics, 2 Department of Chemistry, North Carolina State University

Raleigh, NC, USA 3 Department of Physics and Biophysics Interdepartmental Group, University of Guelph, Guelph,

Ontario, Canada 4 Beijing Nuclear Magnetic Resonance Center and College of Chemistry and Molecular

Engineering, Peking University, Haidian, Beijing, People's Republic of China Abstract Spin-labeling EPR is now widely used for biophysical studies of protein structure and dynamics. Molecular volume of EPR labels is comparable with that of protein sidechains and, therefore, such probes are generally low perturbing to protein structure. While continuous wave (CW) EPR experiments report on dynamics of the protein backbone, local polarity, and site accessibility, time-domain dipolar electron-electron resonance (DEER) spectroscopy is capable of obtaining long-range distance constraints (typically in 2 to 70 nm range) and the distance distributions between strategically placed paramagnetic tags. Here we report on the distance measurements using both smFRET and DEER on the same protein system. Although the distance ranges of these two methods overlap, the direct comparison of FRET and DEER data is rarely found in the literature. We attached probes to a unique pair of cysteines in the neuronal SNARE protein SNAP-25. Results of smFRET and DEER distances and distance distribution are compared for disordered SNAP-25 and folded SNAP-25 within the SNARE complex. Another challenging direction is structure-function studies of membrane proteins. There is growing evidence that many membrane proteins adopt functionally active conformational states only when embedded in or interacting with lipid bilayers of specific composition representative of the native cellular membrane environment. We report on recent results of utilizing spin-labeling DEER measurements to refine the solid-state NMR structure of an oligomeric integral membrane protein, Anabaena Sensory Rhodopsin (ASR), reconstituted in a lipid environment.

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Fluorescence Resonance Energy Transfer is an Accurate Molecular Ruler in vivo and in vitro

James J. McCann1, Orchi Annanya1, Patrick Rock2, Laura Dougherty1, Zhuojun Guo1, R. Bryan Sutton2, Mark E. Bowen1.

1 Department of Physiology & Biophysics, Stony Brook University, Stony Brook, NY, USA 2 Department of Cell Physiology and Molecular Biophysics, Texas Tech University Health

Sciences Center, Lubbock, TX, USA Abstract Measuring protein structure in biological environments is critical to fully understanding their biological functions. Fluorescence Resonance Energy Transfer (FRET) is one of few methods available that can be used in vivo and in vitro. However, there is disagreement regarding the reliability of FRET as a probe of protein conformation. Many use concerns regarding fluorophore rotational dynamics to dismiss the interpretation of FRET as molecular distance. Despite this widespread dismissive belief, numerous studies continue to appear using FRET as indicating the direction if not magnitude of protein conformational changes. We used a series of fluorescent protein tandems that differ in linker length along with a DNA FRET ruler to benchmark the reliability of FRET for both organic dyes and fluorescent proteins through a combination of biophysical measurements, single molecule fluorescence and molecular modeling. For organic dyes, we found that individual measurements have high uncertainty but sub-nanometer precision can be achieved with a sufficient number of restraints. For fluorescent proteins, we validated the calculated FRET distances with two hydrodynamic measurements. We confirmed the predicted distribution of “static” FRET states with single molecule fluorescence and used torsion angle refinement to show that the orientational distribution comes close to the standard approximation used in calculating distances from FRET. Lastly, to ensure that these findings are relevant to measuring protein conformation in vivo, we confirmed the constancy of FP-FRET in live cells. This comprehensive study shows that when sufficient conformational freedom is achieved, FRET measurements can accurately measure molecular distance even in live cells.

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Programming and probing biomolecular machines Mark Bathe

Laboratory for Computational Biology & Biophysics, Department of Biological Engineering,

Massachusetts Institute of Technology, Cambridge, MA, USA Abstract Nucleic acids offer a high degree of programmability that enables the rational design and synthesis of structured three-dimensional molecular architectures that mimic aspects of highly evolved, natural protein assemblies, as well as the interrogation of messenger RNA and protein structure and dynamics in living systems using super-resolution fluorescence imaging. In the first part of my talk I will present work in our group to enable the design and synthesis [1, 2, 3] of structured nucleic acid assemblies to engineer synthetic viral capsid mimics for high-resolution imaging, metallic nanoparticle synthesis, and therapeutic delivery. In the second part of my talk I will present the application of nucleic acids to the multi-scale confocal and super-resolution fluorescence imaging of neuronal synapse proteins using an approach that overcomes the four-color spectral limit of conventional fluorescence imaging [4]. References [1] Pan, K., Kim, D.N., Zhang, F., Adendorff, M., Yan, H., et al. “Lattice-free prediction of three-dimensional structure of programmed DNA assemblies “, Nature Communications, 5: 5578 (2014). [2] Veneziano, R., Ratanalert, S., Zhang, K., Pan, K., Zhang, F., et al., “Designer Nanoscale DNA Assemblies Programmed From the Top Down“, Science, 352: 1534 (2016). [3] Sun, W., Boulais, E., Hakobyan, Y., Wang, W., Guan, A., et al., “Casting Inorganic Structures With DNA Molds”, Science, 346: 717 (2014). [4] Guo, S-M., Veneziano, R., Gordonov, S., Li, L., Park, D., et al. “Multiplexed Confocal and Super-resolution Fluorescence Imaging of Cytoskeletal and Neuronal Synapse Proteins”, bioRxiv, doi: 10.1101/111625 (2017).

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Integrative iSPOT Modeling of Estrogen Receptor Sichun Yang

Center for Proteomics and Department of Nutrition, Case Western Reserve University,

Cleveland, OH, USA Abstract Estrogen receptor alpha (ER), a dominant driver of most breast cancers, consists of two functional (DNA-binding and ligand-binding) domains responsible for hormonal signaling and gene regulation. A fundamental question, however, is still unanswered regarding how these domains interact with each other at the molecular level because structural determination of such a large protein-protein complex has been a daunting task for conventional techniques. To address this issue, we have established a multi-technique iSPOT platform by integration of scattering, footprinting, and docking simulation. iSPOT has revealed a first median-resolution structure of the multidomain ER complex, a completely new protein fold that is different from that of any other existing hormone receptors. This iSPOT application uncovers a hidden cross-talk mechanism between ER's domains that is of high promise for the design of new drugs, but also demonstrates a strong potential of iSPOT as a niche method to advance our capability to characterize the structures of many other protein-protein complexes.

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Poster Abstracts (Alphabetical order)

Engineering Robust Activity in Extremophilic Enzymes Siva Dasetty, Weigao Wang, Mark Blenner, Sapna Sarupria

Department of Chemical Engineering, Clemson University, Clemson, SC, USA

Enzymes are highly specific and environmentally friendly biological catalysts with a potential multi-billion-dollar market. Examples of their applications include a wide array of industries, such as food and beverages, paper and pulp, textiles, detergents, pharmaceuticals, and production of biofuels. The broad range of operating temperatures of these industries require the catalytic activity of enzymes to be less sensitive to temperature. Accordingly, the physical and chemical basis for the temperature adaptability of enzymes from psychrophiles and thermophiles are extensively studied. These studies indicate that the cold adaptability of psychrophilic enzymes is due to their high flexibility, specifically near their active site. In contrast, the loss of flexibility helps thermophilic enzymes to retain their stability at high temperatures. We hypothesize that the incorporation of flexibility near the active site of thermophilic enzyme can increase its activity at low temperatures without compromising its overall stability. We test this hypothesis by mutating glutamic acid by glycine near the active site of G. thermocatenulatus (GTL) – a thermophilic enzyme. We performed two such mutations, Glu316Gly and Glu361Gly and observed an increase in the specific activity at both lower and higher temperatures compared to the wild type (WT) GTL. We use all-atom molecular dynamics (MD) simulations to explore the local and global flexibilities and to understand the mechanisms through which the mutations have increased the catalytic activity of GTL. In our presentation, we will discuss these results and comment on their implications for designing enzymes with a broader range of stability and activity.

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Zinc-coordination and C-peptide Complexation: A Mechanism for the Native Inhibition of IAPP aggregation

Xinwei Ge1, Aleksandr Kakinen2, Esteban N. Gurzov3,4, Wen Yang5, Lokman Pang3,4, Emily H. Pilkington2, Praveen Govindan-Nedumpully1, Pengyu Chen5, Frances Separovic6, Thomas P.

Davis2,7, Pu Chun Ke2, and Feng Ding1

1 Department of Physics and Astronomy, Clemson University, Clemson, SC, USA

2 ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, Monash Institute of Pharmaceutical Sciences, Monash University, Parkville, VIC, Australia

3 St Vincent's Institute of Medical Research, Fitzroy, VIC, Australia 4 Department of Medicine, St. Vincent’s Hospital, University of Melbourne, Melbourne, Australia

5 Department of Material Engineering, Auburn University, Auburn, AL, USA 6 School of Chemistry, University of Melbourne, Melbourne, Victoria, Australia

7 Department of Chemistry, Warwick University, Gibbet Hill, Coventry, United Kingdom The amyloid aggregation of islet amyloid polypeptide (IAPP) is associated with β-cell death in type-2 diabetes (T2D), and in vitro studies suggest IAPP as one of the most aggregation-prone peptides, which readily forms amyloid aggregates at μM concentration. Interestingly, before its secretion to the bloodstream, IAPPs are synthesized and secreted inside beta-cell granules at mM concentration with no aggregates observed, indicating an endogenous inhibition mechanism inside beta-cell granules. Various individual granule components - e.g., low pH and high concentrations of insulin, zinc, and C-peptide - have been studied showing limited inhibition or even promotion effects on IAPP aggregation. For instance, insulin efficiently inhibits IAPP aggregation, but the two molecules do not co-localize. Here, we combined thioflavin T assay, high-resolution transmission electron microscopy imaging, circular dichroism spectroscopy with molecular dynamics simulations to show that zinc-coordination and C-peptide complexation stabilize IAPP from aggregation. The cytotoxicity of IAPP to insulin-producing NIT-1 cells is reversed in the presence of zinc and C-peptide. Our study offers a new paradigm for IAPP stabilization in vivo and suggests that mimicking or promoting zinc-coordinated complexation is a promising approach for mitigating IAPP aggregation and beta-cell death in type-2 diabetes.

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The Flagella Beat of Chlamydomonas Has Distinctly Regulated Static and Dynamic Components, Which Accord with Curvature Controlled Dynein Activity

Veikko F. Geyer1, Pablo Sartori2, Frank Jülicher3, Jonathon Howard1

1 Department of Molecular Biophysics & Biochemistry, Yale University, New Haven, CT, USA 2 Institute for Advanced Study, Princeton, NJ, USA

3 Max Planck Institute for the Physics of Complex Systems, Dresden, Germany

The axoneme, the mechanical core of eukaryotic cilia and flagella, is a huge macromolecular assembly of microtubules, cross-links and dynein motors. The coordinated activity of the dyneins produces a diversity of flagellar waveforms that drive fluid flows across tissue surfaces or propel micro-swimmers. While it is believed that the flagellar beat is a result of feedback - based on the ability of motors to produce and sense forces - the mechanism by which motors are controlled remains unknown. To elucidate how molecular motors are involved in the generation of flagellar waveforms, we studied the beat of axonemes isolated from Chlamydomonas reinhardtii, a single cell green alga. Chlamydomonas flagella can exhibit both of the most common waveforms found in nature: the asymmetrical breaststroke-like beat as well as the symmetrical sperm-like beat. Using high speed microscopy and image analysis we precisely determine axonemal shapes in space and time. A characterization of the waveform properties at different ATP concentrations shows that the static asymmetry of the breaststroke - which has an approximately circular shape - can be separated from the dynamic beat component. This analysis reveals that the breaststroke can essentially be viewed as a sperm-like beat traveling around a circular shape and allows us to examine both waveform components independently. By comparing the dynamic component of experimentally measured waveforms to a mechanical model of the axoneme, we found that the shapes were consistent with a model in which dynein motors respond to changes in axonemal curvature. We furthermore show that the static asymmetric shape underlying the beat could also result from curvature controlled dynein activity. Together our findings present novel insights into how molecular motors shape the asymmetric waveform of Chlamydomonas flagella.

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Single Molecule Study of Dynein Processivity Using Ultra-fast Long-range Optical Tweezers

Subash Godar, Joshua Alper

Department of Physics and Astronomy, Clemson University, Clemson, SC, USA Motor proteins are biological macromolecules involved in various dynamic cellular phenomena including cell division, intracellular transport, cell motility and flagellar bending. Dyneins are responsible for processive transport of a variety of cargos toward the minus end of microtubules and to drive the beat of motile cilia and flagella in a non-processive manner. As processivity is largely a function of motor-filament interaction, it is likely that procecssivity of dynein is, at least in part, regulated by its microtubule binding domain (MTBD). We are investigating the role of the MTBD in the motility of single motors and motor teams in vitro using ultra-fast long-range optical tweezers to both measure andapply forces to these reconstitutions. We aim to determine the fundamental regulation mechanisms of dynein processivity. Because the unique properties of kinetoplastid (a class of flagellated eukaryotic protists of which three are human pathogens) ciliary motility (they beat tip-to-base rather than base-to-tip) are likely due to differences in regulation of dynein processivity, we are extending our fundamental work to the investigation of the molecular mechanisms of kinetopastid ciliary waveforms. Using Trypanosome brucei as a model, our work on MTBD-microtubule interactions will help in the development of novel therapeutic measures based on exploiting unique features of the ciliary motility of Kinetoplastids

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Electrostatic Force Differences Caused by Mutations in Kinesin Motor Domains Can Distinguish Between Disease Causing and Harmless Mutations

Zhe Jia, Lin Li, Yunhui Peng, Joshua Alper, Emil Alexov

Department of Physics and Astronomy, Clemson University, Clemson, SC, USA Mutations in kinesin, the enzyme that use the energy of ATP hydrolysis to transport chemicals along microtubules, can cause various diseases. Typically, the pathogenic mutations in kinesin motor domain are located in the ATP binding and tubulin binding area. Here we investigate the effect of such mutations on electrostatic component of the forces between the kinesin motor domain and the microtubule. We demonstrate that the changes in electrostatic forces are able to discriminate between disease-causing and harmless mutations found in human kinesin motor. Not only the forces in the binding direction are correlated to the pathogenicity, but also forces on the longitude and latitude directions. This may reflect the dependence of kinesins’ functions on motility along the microtubule, which requires a precise balance of microtubule binding forces. Because diseases may originate from multiple effects not related to kinesin-microtubule binding, the prediction rate of 0.85 area under the ROC plot just the change in magnitude of the electrostatic force is remarkable.

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Dynamic Equilibrium of the TPP Riboswitch as Observed by MFD FRET Junyan Ma, Soheila Rezaei-Adariani, Feng Ding, Hugo Sanabria

Department of Physics and Astronomy, Clemson University, Clemson, SC, USA

Antibiotic-resistant infections are one of the widely-known health problems that threatens over two million people each year in United States alone. In 2013, over 23,000 people died due to antibiotic resistant bacteria. Recently, an innovative approach was suggested in which messenger RNAs (mRNA) could serve as potential targets for novel antibiotic drugs. Riboswitches are one of the most studied mRNAs that control gene function and respond to second messengers such as small molecules or proteins. Thus, serving as potential targets. However, riboswitches are highly dynamic structures and evade most common methods of structural characterization. Using a structure-guided drug design rationale, our first goal is to determine the structure-function relationship of riboswitches upon binding of effector molecules. We utilize Förster Resonance Energy Transfer (FRET) at a single molecule level in Multiparameter Fluorescence Detection mode to understand the relationship between structure and dynamic of the TPP Riboswitch. We compare our results with Discrete Molecular Dynamic (DMD) simulations and find that the TPP-riboswitch is in equilibrium between two conformational states. We alter the dynamic equilibrium by the presence TPP, MgCl2 and TPP+MgCl2. Only the open state could serve as potential target to bind new small molecules.

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Predicting Binding Free Energy Change Caused by Point Mutations in Protein-DNA Interactions

Yunhui Peng, Emil Alexov

Department of Physics and Astronomy, Clemson University, Clemson, SC, USA Protein–DNA interactions are essential for the regulations of many important cellular processes, such as transcription, replication, recombination and translation. Many disease-causing mutations occur in DNA-binding protein and have profound effect on binding interaction. In particular, the effect of mutations on binding free energy (binding affinity) is considered to be an important component of the overall disease effect. Although, the effect of missense mutations on protein-DNA binding interaction can be experimentally assessed by various techniques such as isothermal titration calorimetry, FRET and many others methods, the experimental approaches are usually time-consuming, expensive and hard to be applied on a large-scale study. Hence, accurate prediction of missense mutations' effect on protein–DNA binding is essential to our understanding of disease's molecular mechanism and capable for large scale investigation. Here, we develop a new methodology to accurately predict the effect of missense mutations on protein-DNA binding affinity. This new method will utilize MM/PBSA approach along with an additional set of knowledge based terms delivered from investigation of the physico-chemical properties of protein-DNA complexes. An experimental dataset (taken from ProNIT database and references) including experimentally determined binding free energy changes caused by 108 mutations from 13 proteins served as an ultimate benchmark for computational methods aiming at in silico predictions.

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FRET for Dynamic and Integrative Structural Biology Hugo Sanabria

Department of Physics and Astronomy, Clemson University, Clemson, SC, USA

A fundamental goal of structural studies is to unravel the relationship between structure and function. Consideration of the dynamic nature of biomolecules has taken a secondary role in the structure function paradigm. To fill this gap, we propose to use Förster Resonance Energy Transfer (FRET), which is capable of accessing a wide range of biologically relevant motions spanning over 10 decades in time. Historically, FRET has not taken an important role in structural biology, because FRET is most commonly used as a qualitative tool, where “yes” or “no” constitute the expected answer. However, for FRET to play a role on structural biology, FRET should provide detailed structural models with the information on their precision, accuracy and confidence levels. This can only be achieved if distances are measured with high accuracy and if a distance network is devised capable of differentiating representative structures that model the overall ensemble of the biomolecule. Using a Multiparameter approach, and integrating FRET measurements in various modalities, we have measured distance restraints, and dynamics of various model systems. We combine this information with pre-known structural information and multiscale dynamic simulations to show that indeed FRET could be used to differentiate representative structures in highly dynamic biomolecules.

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Brushed Polyethylene Glycol and Phosphorylcholine as Promising Grafting Agents against protein binding

Bo Wang1, Thomas Blin2, Aleksandr Käkinen2, Xinwei Ge1, Emily H. Pilkington2, John F. Quinn2, Michael R. Whittaker2, Thomas P. Davis2,3, Pu Chun Ke2 and Feng Ding1

1 Department of Physics and Astronomy, Clemson University, Clemson, SC, USA

2 ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, Monash Institute of Pharmaceutical Sciences, Monash University, Parkville, VIC, Australia

3 Department of Chemistry, University of Warwick, Gibbet Hill, Coventry, United Kingdom The grafting of linear polyethylene glycol (PEG) is a common strategy in ascribing the stealth effect to nanostucture against protein adsorption for biomedical applications. However, PEG may still evoke side effects in vivo that compromises the circulation and efficacy of PEGylated drugs or nanocarrier. Recently, we demonstrated the synthesis of brushed phosphorylcholine (PC) as biomimetic alternative to linear PEG for the grafting of superparamagnetic iron oxide nanoparticles (IONP). Compared to brushed PEG (bPEG), brushed PC (bPC) rendered even better suspendability, stability, biocompatibility and cellular distribution. To further examine the structures of bPEG and bPC and directly test their antifouling properties against proteins, discrete molecular dynamics (DMD) simulations were performed. We found that brushed polymers were more rigid than the linear PEG while bPEG and bPC ligands displayed distinct globular and cylindrical morphologies. Grafting either bPEG or bPC onto IONPs led to different characteristics of the grafting layer, namely ligand coverage, height, and conformational strains both laterally and vertically. Upon mixing both bPEG and bPC onto the same IONP surface while maintaining the same grafting density, a conformational relaxation of the bPEG was observed in DMD simulations, as corroborated by fluorescence quenching of Cy5 attached to bPEGs in the experiment. Both bPEG- and bPC-grafted NPs displayed antifouling against human serum albumin (HSA), with an increased grafting density giving rise to enhanced protein avoidance. Our results suggest that, with a stronger repulsion to HSA and the capability to a higher grafting density due to its cylindrical shape, bPC is more advantage than both linear and brushed PEG for grafting NPs with minimal protein binding. These new structural and energetic insights offer a general guidance for NP synthesis and anti-fouling applications employing branched polymers.

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Effects of Homologous Proteins on IAPP Amyloid Aggregation, Fibril Remodelling, and Cytotoxicity

Yanting Xing1, Emily Pilkington2, Bo Wang1, Feng Ding1, Pu Chun Ke2

1 Department of Physics and Astronomy, Clemson University, Clemson, SC, USA 2 Monash Institute of Pharmaceutical Sciences, Monash University, Parkville, VIC, Australia

Aggregation of islet amyloid polypeptide (IAPP), a peptide hormone co-synthesized and co-stored with insulin in pancreatic cells and also co-secreted to the circulation, is associated with beta-cell death in type-2 diabetes (T2D). In T2D patients IAPP is found aggregating in the extracellular space of the islets of Langerhans. Although the physiological environments of these intra- and extra-cellular compartments and vascular systems significantly differ, the presence of proteins is ubiquitous but the effects of protein binding on IAPP aggregation are largely unknown. Here we examined the binding of freshly-dissolved IAPP as well as pre-formed fibrils with two homologous proteins, namely cationic lysozyme (Lys) and anionic alpha-lactalbumin (aLac), both of which can be found in the circulation. Biophysical characterizations and a cell viability assay revealed distinct effects of Lys and aLac on IAPP amyloid aggregation, fibril remodelling and cytotoxicity, pointing to the role of protein “corona” in conferring the biological impact of amyloidogenic peptides. Systematic molecular dynamics simulations further provided molecular and structural details for the observed differential effects of proteins on IAPP amyloidosis. This study facilitates our understanding of the fate and transformation of IAPP in vivo, which are expected to have consequential bearings on IAPP glycemic control and T2D pathology.

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List of Participants First Name Last Name Affiliation e-mail Emil Alexov Clemson University ealexov@g.clemson.edu Joshua Alper Clemson University alper@clemson.edu

Mark Bathe Massachusetts Institute of Technology mbathe@mit.edu

Mark Bowen Stony Brook University mark.bowen@stonybrook.edu Terri Bruce Clemson University terri@clemson.edu Argo Chakravorty Clemson University arghyac@g.clemson.edu Bin San Chan Clemson University vbschan@clemson.edu Robert Cohen Clemson University rscohen@clemson.edu Siva Dasetty Clemson University sdasett@g.clemson.edu Delphine Dean Clemson University finou@clemson.edu Feng Ding Clemson University fding@clemson.edu Zach Disharoon Clemson University zack_disharoon@yahoo.com Xinwei Ge Clemson University xinweig@clemosn.edu Veikko Geyer Yale University veikko.geyer@yale.edu Subash Godar Clemson University sgodar@g.clemson.edu George Hamilton Clemson University ghamil4@g.clemson.edu

Daniel Keedy University of California San Francisco daniel.keedy@gmail.com

Satyanarayana Lagishetty Clemson University slagish@clemson.edu Catherine L. Lawson Rutgers University cathy.lawson@rutgers.edu Lin Li Clemson University lli5@clemson.edu Junyan Ma Clemson University junyanm@clemson.edu William R. Marcotte Jr. Clemson University marcotw@clemson.edu Jillian Milanes Clemson University jmilane@clemson.edu Meredith Morris Clemson University mmorri3@clemson.edu James Morris Clemson University jmorri2@clemson.edu Andrew Mount Clemson University mount@clemson.edu

Daniela Nicastro University of Texas Southwestern Medical Center

Daniela.Nicastro@UTSouthwestern.edu

Yunhui Peng Clemson University yunhuip@g.clemson.edu Hugo Sanabria Clemson University hsanabr@clemson.edu Laxmikant Saraf Clemson University lsaraf@clemson.edu Sapna Sarupria Clemson University ssarupr@g.clemson.edu

Tatyana Smirnova North Carolina State University

tismirno@ncsu.edu

Kerry Smith Clemson University kssmith@clemson.edu Jimmy Suryadi Clemson University jsuryadi@clemson.edu Lesly Temesvari Clemson University ltemesv@clemson.edu

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Bo Wang Clemson University bwang4@g.clemson.edu

Elizabeth Wright Emory University School of Medicine erwrigh@emory.edu

Yanting Xing Clemson University yantingx@g.clemson.edu

Sichun Yang Case Western Reserve university sichun.yang@case.edu