The Hong group develops and applies magic-angle-spinning solid-state NMR spectroscopy to elucidate the structure and dynamics of biological macromolecules. We seek to understand how molecular conformation, motion, and intermolecular interactions enable proteins and carbohydrates to carry out their functions such as ion conduction across lipid membranes, membrane fusion between viruses and cells, membrane scission during virus budding, and maintenance and expansion of plant cell walls. We also investigate the structure and dynamics of amyloid proteins in neurodegenerative diseases. To answer these mechanistic biophysical questions, we innovate multinuclear (1​​H, 1​3​C, 1​5N, 1​9​F, 2​​H, & 3​1​P) and multidimensional solid-state NMR techniques that measure molecular structure, inter-atomic distances, and amplitudes and rates of molecular motions with high sensitivity.


Figure 1. Influenza M2 structure and dynamics investigated using solid-state NMR. (A) Influenza BM2 has a distinct TMD sequence from AM2, and contains two histidine residues in the TMD. 15N NMR spectra show that the proton dissociation constants (pKa’s) of the proton-selective His19 differ significantly from the pKa’s of the corresponding histidine in AM2 due to the influence of the peripheral histidine residue, His27 1,2. (B) 2D 13C-13C correlation spectra reveal millisecond-timescale motions of the TMD at high temperature, demonstrating that the rate-limiting step of proton conduction is a conformational change between a closed and open form of the four-helix bundle 3.  (C) The antiviral drug amantadine binds M2 constructs containing the intact cytoplasmic domain, but does not bind a shorter construct comprising only the TMD and an amphipathic helix. This anomalous loss of drug binding can be attributed to the curvature-inducing function of the amphipathic helix4. (D) Cholesterol-complexed structure of M2 to mediate membrane scission during virus budding. The structure is determined using 13C-19F distance measurements between the protein and cholesterol 5.

Ion channels and transporters provide conduits for the passage of ions and polar compounds across the hydrophobic barrier of lipid membranes. The influenza M2 protein is a low-pH activated proton channel, which acidifies the endosomally trapped virion to cause virus uncoating. Since 2010 it was discovered that M2 also mediates membrane scission in the last step of virus budding and release. The proton-channel function is mostly carried out by the transmembrane domain (TMD) of the protein, whereas the membrane-scission function is conducted by an amphipathic helix C-terminal to the TMD. We investigate the structure and dynamics of M2 bound to phospholipid bilayers to elucidate the mechanisms of proton conduction and membrane scission. The M2 protein of influenza A viruses, which are responsible for all flu pandemics in history, is also the target of one of two classes of antiviral drugs, but currently circulating seasonal flu viruses have evolved drug-resistant mutations. Influenza B virus M2 is so far not druggable, thus structure elucidation of AM2 and BM2 is important for drug development.


Using solid-state NMR, we determine the three-dimensional structures and oligomeric assembly of AM2 and BM2 in lipid bilayers, elucidate proton-transfer equilibria (pKa’s) and kinetics (Fig. 1A), investigate how protein sidechain motions and global motions mediate proton transfer (Fig. 1B), elucidate the drug-binding equilibria (Fig. 1C), and investigate the structures of protein-cholesterol complex (Fig. 1D) and protein-protein complexes. This wealth of information is obtained from chemical shifts, long-range intermolecular distances, and motionally averaged NMR spectra.


Membrane curvature is essential to many biological processes such as endocytosis, vesicle trafficking, and cell division. Proteins can sense, stabilize, and induce membrane curvature. An important class of membrane proteins that induce membrane curvature is viral fusion proteins, which merge the virus envelope and the target membrane to enable virus entry into cells. They accomplish this task by undergoing complex conformational rearrangements, as seen in X-ray crystal structures of water-soluble ectodomains of these proteins (Fig. 2A). These protein conformational changes presumably lower the free energy barriers for membrane dehydration and membrane structural changes from the lamellar state to hemifused intermediates to the final fused state. However, this conceptual framework excludes two key hydrophobic domains: the N-terminal fusion peptide (FP) domain and the C-terminal transmembrane domain (TMD), which play important roles in destabilizing the lamellar structures of the two lipid membranes.


Using solid-state NMR spectroscopy and complementary techniques such as small-angle X-ray scattering (SAXS), we investigate the conformations and oligomeric structures of the hydrophobic FP and TMD of viral fusion proteins in biologically relevant lipid membranes. Our studies of the parainfluenza virus fusion protein indicate that both the FP and the TMD have membrane-dependent structures (Fig. 2B), and the β-sheet conformation is responsible for generating saddle-splay curvature to the membrane. The latter is manifested in 31P NMR spectra (Fig. 2C) and SAXS data. We also measure water-lipid and water-protein interactions to obtain information about membrane dehydration during fusion. By coupling these protein structure measurements with membrane morphology experiments, we obtain comprehensive information about the protein and membrane structural changes along the fusion pathway (Fig. 2D). We are also investigating the structure of the HIV fusion protein, gp41.

Figure 2. Solid-state NMR studies of virus-cell fusion. (A) Virus-cell fusion model. The fusion protein undergoes a series of conformational changes to transition from a compact prefusion structure to a membrane-bound post-fusion structure. The N-terminal fusion peptide is encapsulated in the globular head in the prefusion state but inserts into the target cell membrane in the extended intermediate state, while the C-terminal TMD is anchored in the viral envelope. Hairpin formation by the protein ectodomain pulls the cell membrane and the virus envelope together, causing putative membrane intermediates such as the hemifusion diaphragm. Action of the FP and TMD eventually causes full merger of the two membranes.  (B) Conformations of the parainfluenza virus 5 TMD in different lipid membranes determined by 2D 13C-13C correlation NMR spectra 6,7. The TMD is α-helical in POPC membranes, adopts a mixed helix/sheet structure in DOPC/DOPG membranes, and becomes predominantly β-sheet in negative-curvature POPE membranes. These membrane-dependent conformations suggest that the local lipid composition of the membrane has significant influence on the site of virus-cell fusion. (C31P NMR spectra show that the PIV5 F TMD converts the DOPE membrane from the hexagonal phase to a cubic phase, as manifested by a strong isotropic peak 6.  (D) Model of the membrane-dependent conformations of the FP and TMD of the PIV5 fusion protein, likely in a hemifusion intermediate. Local enrichment of phosphatidylethanolamine lipids causes β-strand conformations in both the FP and TMD, which in turn cause negative Gaussian curvature and dehydration to the membrane.


Amyloids are highly aggregated β-sheet structures formed by many peptides and proteins. Protein misfolding into amyloids underlies many neurodegenerative disorders, and poses a significant problem in the formulation of pharmaceutical peptide drugs. But proteins can also assemble into amyloid fibrils to carry out biological functions.


We are interested in understanding the molecular structures and misfolding pathways of non-functional amyloid proteins. These studies aim to ultimately provide insights into ways to prevent and treat diseases. We recently determined the novel structure of the amyloid fibrils formed by the peptide hormone glucagon (Fig. 3A)8, which is used to treat diabetic hypoglycemia. This antiparallel β-sheet fibril, containing two coexisting β-sheet conformations, gives exquisite insight into the possible pathway of glucagon misfolding from its functional α-helical structure to the non-functional β-sheet structure under pharmaceutical conditions.


A central focus in this project is the investigation of the structures and dynamics of the intrinsically disordered microtubule-binding protein tau, whose misfolding into β-sheet fibrils is one of the two hallmarks of Alzheimer’s disease (AD). Tau neurofibrillary tangles also occur in many other disorders such as chronic traumatic encephalopathy. Tau has six isoforms in human brains, which mainly differ in whether three or four microtubule-binding repeats exist in the protein (Fig. 3B). Cryoelectron microscopy studies have shown the β-sheet core structures of patient-brain tau in several diseases, but do not give information about the structures and dynamics of the rest of the protein. Also unknown is the pathway of misfolding from the microtubule-bound state to the β-sheetaggregated state. We are employing and further developing the full arsenal of high-resolution multidimensional solid-state NMR spectroscopy to understand the β-sheet core structures of tau and to delineate the dynamics of domains outside the β-sheet core. Our first study9 of in vitro fibrillized four-repeat tau has resulted in a low-resolution structural model of the β-sheet core (Fig. 3C-E), and have shown that the remainder of the protein exhibits a pronounced mobility gradient. This gradient ranges from semi-rigid domains near the β-sheet core to semi-mobile proline-rich domains and to isotropically mobile termini. Future studies will aim to elucidate 1) whether tau fibril structures are mainly dictated by the isoform or by fibril-forming conditions, 2) how in vitro and in vivo tau fibrils differ, 3) how three-repeat and four-repeat tau proteins are structured in brains, and 4) how tau evolves from its functional intrinsically disordered state to dysfunctional β-sheet states.

Figure 3. Amyloid fibril structure and dynamics investigated by solid-state NMR. (A) Atomic structure of glucagon fibrils, showing antiparallel β-sheets with two coexisting molecular conformations that are stabilized by steric zipper interactions. The solid-state NMR spectra resolve two sets of chemical shifts for all residues in this peptide hormone. (B) Amino acid sequence map of the longest isoform of tau. (C) Negative-stain electron micrograph of heparin-fibrillized four-repeat tau. (D) Solid-state NMR structural model of the rigid β-sheet core of four-repeat tau fibrils obtained from 2D and 3D SSNMR data. (E) Representative 2D 13C-13C correlation spectrum. A single set of chemical shifts is observed for the only two cysteine residues in the protein, indicating that the β-sheet core is monomorphic.


Figure 4. Solid-state NMR studies of the structure and dynamics of plant cell walls. (A) Cartoon representation of the primary cell walls of dicots, in which cellulose microfibrils act as the scaffold of the cell wall, surrounded by pectins and hemicelluloses. (B) Representative 2D 13C-13C correlation spectra of primary cell walls of Arabidopsis. High magnetic fields enhance spectral resolution and allow resonance assignment, thus permitting the detection of intermolecular contacts between different polysaccharides 10,11. (C) Expansin binding to cellulose in plant cell walls 12. By measuring expansin-to-polysaccharide 1H spin diffusion under DNP, we showed for the first time that expansin’s binding target in the cell wall is cellulose rather than matrix polysaccharides. Molecular dynamics simulation shows how expansin may dock onto the cellulose microfibril. (D) Schematic of the single-network model of the dicot primary wall based on our solid-state NMR data 13,14.

Plant cell walls provide mechanical strength to plant cells while at the same time allowing plants to grow rapidly. Plant cell walls primarily contain three types of polysaccharides: cellulose, hemicellulose, and pectins (Fig. 4A). Although the chemical composition of plant cell walls is relatively well known, the three-dimensional architecture and the dynamics of cell wall polysaccharides have long been elusive due to the lack of high-resolution structural techniques to characterize this insoluble and disordered material.


We are pioneering the application of multidimensional solid-state NMR to elucidate the structures and dynamics of the polysaccharides of intact primary cell walls. By enriching whole plants with 13C, we are able to employ 2D and 3D correlation MAS NMR techniques to detect and resolve the signals of the complex mixture of polysaccharides in intact cell walls (Fig. 4B), and determine their spatial contacts and mobilities. By using sensitivity-enhancing dynamic nuclear polarization (DNP) and paramagnetic relaxation enhancement NMR techniques, we elucidate how polysaccharides interact with proteins to loosen the cell walls during growth (Fig. 4C). Using model plants of both dicot (e.g. Arabidopsis thaliana) and grass (e.g. Brachypodium distachyon and Zea mays) families, we have shown that cellulose, hemicellulose and pectins form a single three-dimensional network (Fig. 4D) instead of two separate networks, thus revising the long-held view of the plant cell wall structure. With our collaborators, we also investigate the structural polymorphism of cellulose microfibrils, hydration of wall polysaccharides, interactions of cellulose with matrix polysaccharides, and the effects of genetic mutations on cell-wall structure.


Driven by our interest in answering important biological questions, we continue to expand the capability of solid-state NMR spectroscopy. We have a long-standing interest in increasing the distance reach of NMR. Using nuclear spins with high gyromagnetic ratios such as 19F and 1H, and exploiting multi-spin effects, we are pushing the distance upper limit of NMR to ~2 nm (Fig. 5A). This capability opens up many possibilities for determining the oligomeric structures and global conformational changes of proteins, and for determining the structures of protein-lipid, protein-carbohydrate and other macromolecular complexes. To determine the amplitude of molecular motion, we develop anisotropic-isotropic correlation experiments involving both dipolar interactions and 2H quadrupolar interactions (Fig. 5B) under fast MAS and in high magnetic fields.

Figure 5. Solid-state NMR methods for distance measurements and dynamics studies. (A19F-19F dipolar coupling measurements by spin exchange and dipolar recoupling now can probe distances to 1.8 nm. (B2H-13C correlation experiments to investigate small-amplitude motions under fast MAS 15.


(1) Williams, J. K.; Tietze, D.; Lee, M.; Wang, J.; Hong, M. J. Am. Chem. Soc. 2016, 138, 8143-8155.
(2) Williams, J. K.; Shcherbakov, A. A.; Wang, J.; Hong, M. J. Biol. Chem. 2017, 292, 17876-17884.
(3) Mandala, V. S.; Gelenter, M. D.; Hong, M. J. Am. Chem. Soc. 2018, 140, 1514-1524.
(4) Cady, S. D.; Wang, T.; Hong, M. J. Am. Chem. Soc. 2011, 133, 11572-11579.
(5) Elkins, M. R.; Williams, J. K.; Gelenter, M. D.; Dai, P.; Kwon, B.; Sergeyev, I. V.; Pentelute, B. L.; Hong, M. Proc. Natl. Acad. Sci. U. S. A. 2017, 114, 12946-12951.
(6) Yao, H.; Lee, M. W.; Waring, A. J.; Wong, G. C.; Hong, M. Proc. Natl. Acad. Sci. USA 2015, 112, 10926-10931.
(7) Yao, H.; Lee, M.; Liao, S. Y.; Hong, M. Biochemistry 2016, 55, 6787-6800.
(8) Gelenter, M. D.; Smith, K. J.; Liao, S. Y.; Mandala, V. S.; Dregni, A. J.; Lamm, M. S.; Tian, Y.; Xu, W.; Pochan, D. J.; Tucker, T. J.; Su, Y.; Hong, M. Nat. Struc. Mol. Biol 2019, 26, 592-598.
(9) Dregni, A. J.; Mandala, V. S.; Wu, H.; Elkins, M. R.; Wang, H. K.; Hung, I.; DeGrado, W. F.; Hong, M. Proc. Natl. Acad. Sci. U.S.A 2019, 116, 16357-16366.
(10) Wang, T.; Zabotina, O. A.; Miller, R. C.; Hong, M. Biochemistry 2012, 51, 9846-9856.
(11) Wang, T.; Hong, M. J. Exp. Botany 2016, 67, 503-514.
(12) Wang, T.; Park, Y. B.; Caporini, M. A.; Rosay, M.; Zhong, L.; Cosgrove, D. J.; Hong, M. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 16444-16449.
(13) Dick-Pérez, M.; Zhang, Y.; Hayes, J.; Salazar, A.; Zabotina, O. A.; Hong, M. Biochemistry 2011, 50, 989-1000.
(14) Wang, T.; Phyo, P.; Hong, M. Solid State Nucl. Magn. Reson. 2016, 78, 56-63.
(15) Gelenter, M. D.; Wang, T.; Liao, S. Y.; O’Neill, H.; Hong, M. J. Biomol. NMR 2017, 68, 257-270.

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