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.


Ion channels and transporters are integral membrane proteins that transport ions and polar compounds across the hydrophobic interior of lipid bilayers. Viral ion channels, also called viroporins, are important for the lifecycles and pathogenicity of viruses. Bacteria use transporters to pump out toxic compounds to cause antibiotic resistance. Elucidating the conformation and dynamics of these membrane proteins is critical for advancing our fundamental knowledge of membrane transport and for combating infectious diseases.

Figure 1. Schematics of the tetrameric influenza M2 proton channel, the pentameric SARS-CoV-2 E cation channel, and the dimeric E. Coli transporter EmrE.  We employ solid-state NMR spectroscopy to investigate the structure and dynamics of these membrane proteins.

We are currently studying three ion channels and transporters: the influenza M2 proton channel, the Severe Acute Respiratory Syndrome coronavirus 2 (SARS-CoV-2) E cation channel, and the bacterial transporter EmrE (Fig. 1). All three proteins are all oligomeric a-helical bundles that require proton transfer for function. The acid-activated and amantadine-targeted M2 proton channel is essential for the influenza lifecycle and is a prototype of viroporins 1. The SARS-CoV-2 E protein forms a cation channel 2 whose calcium-conducting activity stimulates the host inflammasome. EmrE is a proton-coupled transporter that exports polyaromatic cations to cause drug resistance 3.

Elucidating the structures of these integral membrane proteins holds the key for designing better antiviral drugs and antibiotics. Describing the conformational dynamics of these membrane proteins is essential for understanding why some of these proteins conduct ions exclusively down the concentration gradient while others use alternating access to transport ions either down or up the concentration gradient.


Solid-state NMR spectroscopy is well suited to the structure determination of small integral membrane proteins in lipid bilayers. Moreover, solid-state NMR provides information about the dynamics of these conformationally plastic proteins, through the measurement of motionally averaged spectral lineshapes and nuclear spin relaxation times. This dynamical information is not easily accessible to cryoEM and X-ray crystallographic analysis. We have developed NMR techniques to measure inter-atomic distances to 2 nm 4 to constrain the three-dimensional structures of these membrane proteins and their drug-binding sites. We measure rotationally averaged N-H dipolar couplings to determine the helix orientations of these proteins. We design polarization transfer experiments to determine which residues line the channel pore or face lipids. We also investigate the channel-water dynamics 5 and ligand dynamics by measuring the relaxation times of channel water and motionally averaged lineshapes of small-molecule ligands.


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 fundamental biological questions, we continue to develop and expand the capabilities 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, we are now extending the measurable distance upper limit of NMR to ~2 nm (Fig. 5a). Combining 2D correlation experiments with 1H-19F and 13C-19F distance measurements, we can now extract tens to hundreds of nanometer-range distances in 2-3 pairs of 2D spectra 4, 15-16 (Fig. 5b, c). These 19F MAS NMR techniques open the avenues for elucidating the structures of fluorinated compounds bound to their target proteins 17 (Fig. 5d, e), structures of homo-oligomeric membrane proteins and other biological complexes. 1H-detected fast MAS 3D correlation experiments enable resonance assignment in a short amount of time, thus allowing this distance extraction.

Figure 5. Solid-state NMR methods for rapid measurement of nanometer-range distances. (a) Distance rulers in NMR depend on the nuclear spin gyromagnetic ratio g. Commonly used spin pairs in biomolecules with 30-Hz dipolar couplings are shown. 1H and 19F spins have the highest g, and hence longest distance reach. (b) Dipolar coupling strength as a function of internuclear distance for common spin pairs. The longest distances measurable for the intermediate difficulty regime of 30-150 Hz are ~7 Å for 1H-15N and 19F-15N spin pairs, ~10 Å for 1H-13C and 19F-13C spin pairs, and ~15 Å for 1H-1H and 1H-19F spin pairs. (c) Pulse sequence for the 2D hNH resolved 1H-19F REDOR technique. (d) 1H-19F REDOR dephasing measured for the bacterial membrane protein EmrE. These REDOR dephasing curve are obtained from 2D REDOR control (S0) and difference (DS) spectra and are fit to give precise distances. (e) High-resolution structure of EmrE bound to a fluorinated substrate, TPP+. EmrE transports such cationic aromatic antibiotic compounds to cause multidrug resistance.


(1) Mandala, V. S.;  Loftis, A. R.;  Shcherbakov, A. A.;  Pentelute, B. L.; Hong, M, Nat. Struc. Mol. Biol 2020, 27, 160–167.
(2) Mandala, V. S.;  McKay, M. J.;  Shcherbakov, A. A.;  Dregni, A. J.;  Kolocouris, A.; Hong, M., Nat. Struc. Mol. Biol 2020, 27, 1202-1208.
(3) Shcherbakov, A. A.;  Hisao, G.;  Mandala, V. S.;  Thomas, N. E.;  Soltani, M.;  Salter, E. A.;  Davis Jr., J. H.;  Henzler-Wildman, K. A.; Hong, M., Nat. Commun. 2021, 12, 172.
(4) Shcherbakov, A. A.;  Mandala, V. S.; Hong, M., J. Phys. Chem. B. 2019, 123, 4387-4391.
(5) Gelenter, M. D.;  Mandala, V. S.;  Niesen, M. J. M.;  Sharon, D. A.;  Dregni, A. J.;  Willard, A. P.; Hong, M., Commun. Biol. 2021, 4, 338.
(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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