The Hong laboratory focuses on the development and application of solid-state NMR spectroscopy to elucidate the structure and dynamics of biological macromolecules, especially membrane proteins. Our research spans fundamental physical chemistry and applications to biology, pharmacology, and biomaterials. We have a long-term interest in ion channels and curvature-inducing membrane proteins. We are also intrigued by complex biomaterials such as the polysaccharides and glycoproteins of plant cell walls. Magic-angle-spinning (MAS) solid-state NMR (SSNMR) spectroscopy is the principal tool for our biophysical studies, because it provides atomic-resolution structural and dynamical information about noncrystalline and insoluble biomolecules in their native environments.

Influenza M2 protein
            The M2 protein of influenza A and B viruses is an essential and multifunctional protein of the flu viruses. It forms an acid-activated proton-selective channel in the virus envelope that is important for the virus lifecycle. After viral endocytosis into the acidic endosome, the M2 channel opens, acidifies the virion, and causes the release of the viral RNA into the cell. A single transmembrane helix in the protein oligomerizes into a four-helix bundle to form the heart of the proton channel. For influenza A viruses, the M2 (AM2) proton-channel activity is inhibited by the adamantane class of antiviral drugs, which is one of only two classes of antiviral drugs against flu. A second function of the M2 protein is to mediate release of newly assembled viruses from the host cell. A conserved amphipathic helix in the cytoplasmic tail of the protein causes high membrane curvature in a cholesterol-dependent fashion to mediate membrane scission at the neck of the budding virus. Elucidating the structural basis for these functions of the M2 protein is important for developing anti-influenza antiviral drugs as well as for elucidating the fundamental phenomena of proton conduction, protein-lipid interactions, and protein-drug interactions.
            Using solid-state NMR spectroscopy, we have been investigating the detailed structural bases and mechanisms of proton conduction, drug inhibition, and membrane-curvature induction by the M2 protein. To measure intermolecular distances, orientation, mobility and proton-transfer dynamics of the M2 protein in the lipid bilayer, we have developed a number of SSNMR techniques such as 19F spin diffusion, 13C-2H multi-spin REDOR, and rotational-diffusion based extraction of protein orientation without membrane alignment. Our results have revealed where amantadine as well as newer drugs bind in wild-type and mutant AM2 proteins to inhibit the proton-channel activity (Figure 1), how the proton-selective residue, a histidine, moves to shuttle protons into the virus, how the gating residue, a tryptophan, orients in the channel and moves in synchrony with histidine to regulate proton flux. We conduct protein-lipid correlation experiments to understand the nature of the lipid interactions of the amphipathic helix that causes membrane curvature. We are determining the structure of the N-terminal ectodomain and the C-terminal cytoplasmic domain in the full-length M2 to understand how these domains modulate the proton-channel function and membrane-scission function. With our collaborators we continue to examine new drugs that target M2 mutants that are resistant to amantadine drugs.

Figure 1. Inhibition of the influenza A M2 proton channel. (a) Amantadine binds wild-type M2 in the pore near residue S31, whereas the amantadine-resistant S31N M2 protein is inhibited by an isoxazole drug, which binds at the same pocket. (b) The drugs that have been studied by us using solid-state NMR spectroscopy, in collaboration with the Bill DeGrado group. The small adamantane class of drugs inhibits wild-type M2 with Ser31, while the bulkier isoxazole drugs inhibit the S31N mutant of M2. (c) Histidine-37 in the M2 transmembrane domain undergoes microsecond ring reorientation and tautomerization to exchange protons with water, hence shuttling protons into the virus.

Key references
  1. S.D. Cady, C. Goodman, C. Tatko, W.F. DeGrado, and M. Hong, “Determining the Orientation of Uniaxially Rotating Membrane Proteins Using Unoriented Samples: a 2H, 13C, and 15N Solid-State NMR Investigation of the Dynamics and Orientation of a Transmembrane Helical Bundle”, J. Am. Chem. Soc. 129, 5719-5729 (2007).
  2. S. D. Cady, K. Schmidt-Rohr, J. Wang, C.S. Soto, W. F. DeGrado, and M. Hong, “Structure of the Amantadine Binding Site of Influenza M2 Proton Channels In Lipid Bilayers”, Nature, 463, 689 (2010). PMCID: PMC2818718
  3. F. Hu, W. Luo, and M. Hong, “Mechanisms of proton conduction and gating by influenza M2 proton channels from solid-state NMR”, Science, 330, 505-508 (2010). PMCID: PMC4102303
  4. F. Hu, K. Schmidt-Rohr and M. Hong, “NMR Detection of pH-Dependent Histidine-Water Proton Exchange Reveals the Conduction Mechanism of a Transmembrane Proton Channel”, J. Am. Chem. Soc. 134, 3703-3713 (2012). Cover of JACS volume 134, issue 8. PMCID: PMC3288706.
  5. S. Liao, Y. Yang, D. Tietze and M. Hong, “The Influenza M2 Cytoplasmic Tail Changes the Proton-Exchange Equilibria and the Backbone Conformation of the Transmembrane Histidine Residue to Facilitate Proton Conduction”, J. Am. Chem. Soc., 137, 6067–6077 (2015). PMCID: PMC4554341.
  6. J. K. Williams, D. Tietze, M. Lee, J. Wang, and M. Hong, “Solid-State NMR Investigation of the Conformation, Proton Conduction, and Hydration of the Influenza B Virus M2 Transmembrane Proton Channel”, J. Am. Chem. Soc. 138, 8143-8155 (2016).
Viral fusion proteins
            A wide variety of membrane peptides and proteins cause membrane curvature for function. An example is the family of viral fusion proteins, which merge the virus envelope and the target-cell membrane during virus entry into cells. Existing models of virus-cell fusion (Figure 2) postulate specific oligomeric structures and lipid interactions of two hydrophobic domains in these proteins: the fusion peptide (FP) domain and the transmembrane (TM) domain. Using solid-state NMR spectroscopy, we are measuring the conformation, oligomeric assembly and lipid- and water-interactions of these two domains in the parainfluenza virus and related viruses, in order to elucidate how the fusion peptide and the transmembrane domain work in concert to destabilize the lipid membranes and reduce the hydration pressure to cause full membrane fusion.
Figure 2. (a) Conventional 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 (green) 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 TM domain (grey) is anchored in the viral envelope. Subsequent hairpin formation by the protein pulls the cell membrane and the virus envelope together, causing putative membrane intermediates such as the hemifusion diaphragm, whose structure is not well known. Action of the FP and TM domains eventually causes the two membranes to fully fuse, resulting in a fusion pore. (b) Conformations of the parainfluenza virus 5 fusion peptide in different lipid membranes as determined using two-dimensional solid-state NMR spectroscopy. The fusion peptide is a-helical in POPC/POPG membranes, adopts a mixed sheet/helix structure in the more fluid DOPC/DOPG membranes, and switches to a predominantly b-sheet conformation in the negative-curvature DOPE membrane. This membrane-dependent conformational plasticity may serve to regulate the time and location of viral fusion. (c) Representative 2D 13C-13C correlation spectra that show the membrane-induced conformational changes of the FP of the parainfluenza virus. The 13C chemical shifts are diagnostic of the secondary structures of the various residues in the three different lipid membranes.
Key references
  1. H. Yao, M.W. Lee, A.J. Waring, G.C.L. Wong, and M. Hong, “A Viral Fusion Protein Transmembrane Domain Adopts β-Strand Structure to Facilitate Membrane Topological Changes For Virus-Cell Fusion”, Proc. Natl. Acad. Sci. USA, 112, 10926-31 (2015). PMCID: PMC 4568205. 

Plant cell walls
            Plant cell walls shape plant cell morphology, maintain ionic balance, protect plants from environmental stress, and allow plant cell growth.  These functions are carried out by a variety of polysaccharides and glycoproteins in the cell wall, whose three-dimensional structures and packing have long evaded high-resolution structural characterization due to the insoluble and amorphous nature of the wall macromolecules. We are pioneering the first 2D and 3D MAS correlation solid-state NMR studies of the cell walls of key model plants such as Arabidopsis thaliana and Brachypodium distachyon, which represent the dicotyledonous (dicots) and monocotyledonous (monocots) families of flowering plants. These multidimensional 13C SSNMR studies are made possible by isotopic enrichment of whole plants and by the use of sensitivity-enhancing techniques such as high-field dynamic nuclear polarization (DNP), developed by the group of Bob Griffin at MIT. Our results reveal, for example, that cellulose, hemicellulose and pectins form a single spatial network in the Arabidopsis cell wall, which revises the long-held “tethered network” model of the primary cell wall. With our collaborators, we are investigating how cellulose interacts with matrix polysaccharides, the detailed conformation of crystalline and amorphous cellulose and how the conformation is modified by mutation, how proteins bind to polysaccharides to achieve functions such as wall loosening during plant growth, and how wall polysaccharides are hydrated. To address these questions, we employ a wide range of SSNMR techniques involving intermolecular correlation, spin diffusion, and dynamics measurements.
Figure 3. Solid-state NMR investigations of the structure and dynamics of plant cell wall polysaccharides and proteins. (a) A cartoon representation of the primary cell walls of dicots such as Arabidopsis thaliana, in which cellulose microfibrils act as the scaffold of the cell wall, surrounded by matrix polysaccharides, pectins and hemicelluloses.  The exact three-dimensional structures formed by these polysaccharides are still poorly understood. (b) Representative 2D 13C-13C correlation spectra of intact primary cell walls of Arabidopsis. Assignment of the 13C peaks allows the detection of intermolecular contacts between different polysaccharides. (c) Representative relaxation NMR data that provide information on the heterogeneous dynamics of the wall polysaccharides. Cellulose is the most rigid polymer, showing the longest 13C T1 relaxation times, while pectins are the most dynamic with the shortest T1. (d) A schematic of the single-network model of the dicot primary wall based on our solid-state NMR data. (e) Cell-wall loosening during plant growth is catalyzed by the protein, expansin. By measuring expansin-to-polysaccharide 1H spin diffusion spectra, we showed for the first time that expansin’s binding target in the cell wall is cellulose rather than matrix polysaccharides. An active expansin mutant, RKK, shows strong protein-to-cellulose spin diffusion, while an inactive mutant, WWY, shows little spin diffusion to cellulose. (f) A model of expansin docking onto the cellulose microfibril based on our solid-state NMR spin diffusion data.
Key references
  1. M. Dick-Perez, Y. Zhang, J. Hayes, A. Salazar, O.A. Zabotina and M. Hong, “Structure and Interactions of Plant Cell-Wall Polysaccharides by 2D and 3D Magic-Angle-Spinning Solid-State NMR”, Biochemistry, 50, 989-1000 (2011). PMID: 21204530.
  2. T. Wang, O. Zabotina, and M. Hong, “Structure and Dynamics of Brachypodium Primary Cell Wall Polysaccharides from Two-Dimensional 13C Solid-State Nuclear Magnetic Resonance Spectroscopy”, Biochemistry, 53, 2840-2854 (2014). PMID: 2472037.
  3. T. Wang, Y.B. Park, M.A. Caporini, M. Rosay, D.J. Cosgrove and M. Hong, “Sensitivity-enhanced solid-state NMR detection of expansin's target in plant cell walls”, Proc. Natl. Acad. Sci. USA, 110, 16444-9 (2013). PMCID: PMC3799313
  4. T. Wang and M. Hong, “Solid-State NMR Investigations of Cellulose Structure and Interactions with Matrix Polysaccharides in Plant Primary Cell Walls”, J. Expt. Botany, 67, 503-514 (2016). 


Cationic antimicrobial and cell-penetrating peptides
            Although the interior of lipid membranes is highly hydrophobic, a surprising number of cationic membrane-active peptides exist in biology. Examples are antimicrobial peptides produced by animals and plants for immune defense against microbes, cell-penetrating peptides that act as the Trojan horse of macromolecular cargos into cells, and voltage-sensing domains of ion channels. All these membrane peptides and protein domains contain many arginine and lysine residues, whose transport from aqueous solution to lipid bilayers presumably encounters a large free-energy barrier. We are interested in the structure, dynamics and lipid interactions of these membrane peptides, to understand the mechanism with which these cationic molecules overcome the free-energy barrier to insert into the lipid membrane.  31P, 13C and 2H solid-state NMR experiments are used to measure the membrane morphology and domain structure induced by these peptides, the peptide structure and dynamics, and the interactions among the cationic residues, lipid and water.
Key references
  1. S. Yamaguchi, A. Waring, T. Hong, R. Lehrer and M. Hong, “Solid-state NMR Investigations of Peptide-Lipid Interaction and Orientation of a β-sheet antimicrobial peptide, Protegrin”, Biochemistry, 41, 9852-9862 (2002)
  2. R. Mani, S.D. Cady, M. Tang, A.J. Waring, R.I. Lehrer, and M. Hong, “Membrane-dependent oligomeric structure and pore formation of a β-hairpin antimicrobial peptide in lipid bilayers from solid-state NMR”, Proc. Natl. Acad. Sci. U.S.A., 103, 16242-16247 (2006). PMID: 17060626
  3. M. Tang, A. J. Waring and M. Hong, “Phosphate-Mediated Arginine Insertion Into Lipid Membranes and Pore Formation by a Cationic Membrane Peptide from Solid-State NMR”, J. Am. Chem. Soc. 129, 11438-11446 (2007). PMID: 17705480
  4. Y. Su, A.J. Waring, P. Ruchala, and M. Hong, “Membrane-bound dynamic structure of an Arginine-rich cell-penetrating peptide, the protein transduction domain of HIV TAT, from solid-state NMR”, Biochemistry, 49, 6009-6020 (2010). PMCID: PMC2925115
  5. M. Hong and Y. Su, “Structure and Dynamics of Cationic Membrane Peptides and Proteins: Insights from Solid-State NMR”, Protein Sci., 20, 641-655 (2011). PMCID: PMC3081543


Solid-state NMR techniques for biomolecular structure determination

            To address cutting-edge biological questions we develop a range of new solid-state NMR techniques. We have long-standing interest in multinuclear distance-measuring techniques (e.g. 19F-19F, 13C-2H, 13C-31P and 13C-1H distances); intermolecular correlation experiments that probe protein-lipid, protein-water and polysaccharide-water interactions; 2D and 3D correlation techniques and computational methods for more efficient resonance assignment of NMR spectra; anisotropic-isotropic correlation techniques to measure torsion angles and molecular motion; and novel isotopic labeling strategies for proteins and other biological molecules.

Key references
  1. M. Hong and K. Schmidt-Rohr, “Magic-Angle-Spinning NMR Techniques for Measuring Long-Range Distances in Biological Macromolecules”, Acct. Chem. Res, 46, 2154-63 (2013). PMCID: PMC3714308
  2. D. Huster, X.L. Yao and M. Hong, “Membrane Protein Topology Probed by 1H Spin Diffusion from Lipids Using Solid-State NMR Spectroscopy”, J. Am. Chem. Soc. 124, 874-883 (2002).
  3. K. J. Fritzsching, Y. Yang, K. Schmidt-Rohr, and Mei Hong, “Practical use of chemical shift databases for protein solid-state NMR: 2D chemical shift maps and amino-acid assignment with secondary-structure information”, J. Biomol. NMR, 56, 155-167 (2013). PMCID: PMC4048757
  4. K. Schmidt-Rohr, K. J. Fritzsching, S. Y. Liao, M. Hong, “Spectral editing of two-dimensional magic-angle-spinning solid-state NMR spectra for protein resonance assignment and structure determination”, J. Biomol. NMR, 54, 343-353 (2012). PMCID: PMC3656487
  5. J.K. Williams, K. Schmidt-Rohr and M. Hong, “Aromatic Spectral Editing Techniques for Magic-Angle-Spinning Solid-State NMR Spectroscopy of Uniformly 13C-Labeled Proteins”, Solid State Nuc. Magn. Reson. 72, 118-126 (2015). PMCID: PMC4674322