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ABSTRACT: What are the thermodynamic driving forces that influence the free energy of membrane protein folding and association in lipid bilayers? For soluble proteins, the burial of hydrophobic groups away from aqueous interfaces is a major driving force, but membrane-embedded proteins cannot experience hydrophobic forces, as the lipid bilayer lacks water. A fundamental conundrum thus arises: how does a greasy protein surface find its greasy protein partner in the greasy lipid bilayer to fold faithfully into its native structure? Recently, a structurally stable and functional monomeric form of the normally homodimeric Cl^-/H⁺ antiporter CLC-ec1 was designed by introducing tryptophan mutations at the dimer interface. We have used this to develop a new model system for studying reversible dimerization in membranes for free energy measurements, which encompasses the thermodynamic properties of protein interactions in the membrane environment. To quantify monomer vs. dimer populations across a wide range of protein densities, we developed a method that quantifies the capture of subunits into liposomes from large equilibrium membranes single-molecule photobleaching by total internal reflection microscopy.  With this, we are able to determine that CLC-ec1 has a free energy of dimerization of -11 kcal/mole in 2:1 POPE/POPG membranes.  We are now investigating why this complex is so stable, dissecting the changes in enthalpy and entropy while varying protein interactions or the composition of the lipid solvent.  The results from this study will provide a physical foundation for the development of informed strategies aimed at correcting protein mis-folding or regulating protein interactions in membranes in physiologically and pathological situations.

Abstract:

Size-selective cryogenic photoelectron spectroscopy (cryoPES) coupled with electrospray ionization source (ESI) has been demonstrated to be a powerful experimental technique to investigate electronic structures and energetics of a wide variety of solution phase species and chemistry in the gas phase. In this talk, I will present the latest results probing various novel molecular clusters ranged from closo-dodecaborate dianions [B₁₂X₁₂]^2- to atmospherically relevant species by employing this technique. Transition state dynamics of unimolecular isomerization and chemical reactions via photodetachment of corresponding precursor anions will be reported as well. Future directions of ESI-cryoPES, leading to high resolution photoelectron imaging spectroscopy and time-resolved pump-probe experiments will also be briefly discussed.

Abstract:

Magnetic particle imaging (MPI) is an emerging imaging modality that enables the direct mapping of iron oxide nanoparticle tracers.  In MPI, the development of tailored magnetic nanoparticle tracers is paramount to achieving high sensitivity and good spatial resolution. This talk will provide a general overview of the progress in MPI tracer development over the past decade, and will also focus on emerging directions and new opportunities for iron oxide-based tracer design and applications. The presentation will cover magnetic nanoparticle relaxation in MPI and discuss key aspects to consider in tailoring tracers for MPI applications. Emphasis will be given on how structural changes (size, composition, shape, surface chemistry) and inter-particle interactions affect the MPI signal generation process. Moreover, the presentation will discuss emerging research directions in color-MPI (cMPI) and MPI-guided hyperthermia (hMPI).

Abstract

Lignin remains one of the most significant barriers to the efficient utilization of lignocellulosic substrates, in processes ranging from ruminant digestibility to indus­trial pulping, and in the current focus on biofuels production. Inspired largely by the recalcitrance of lignin to biomass processing, plant en­gineering has routinely sought to alter lignin quantity, composition, and structure by exploiting the inherent plasticity of lignin biosynthesis.



More recently, researchers are attempting to strategically design plants for increased degradability by incorporating monomers that lead to a lower degree of polymerization, reduced hydrophobicity, fewer bonds to other cell wall constituents, or novel chemically labile linkages in the polymer backbone.1,2 The incorporation of value-added structures could also help valorize lignin. Designer lignins may satisfy the biological requirement for lignification in plants while improving the overall efficiency of biomass utilization. Although possibilities abound, maintaining plant health is paramount and, ultimately, the plants themselves will dictate which of these approaches can be tolerated.



One such method, via the so-called ‘zip-lignin’ approach, is showing particular promise.3-5 Poplar has been engineered to incorporate monolignol ferulate conjugates into the lignification process, by using an exotic transferase gene, FMT, and a xylem-specific promoter.5 This results in the introduction of readily cleavable ester linkages into the backbone of the polymer, and delivers significantly improved processing. Various applications for which these altered trees appear superior are only just beginning to be explored. Now that we have sensitive methods for determining if/when/whether plants are making monolignol ferulate conjugates and using them for lignification, it appears that Nature herself may have already been exploring this avenue.6 In attempting to modify monocot lignins, another transferase appears to be useful for improving cell wall digestibility. This involves p-coumaroylation of the lignin. Although this occurs naturally in monocots, it is not evident in dicots, but providing dicots with that pathway has interesting implications.7,8

Nature continues to inspire us with new avenues toward lignin modification that have potential value for various processes. For example, the ramifications of finding that grasses are using tricin, a flavone from beyond the monolignol biosynthetic pathway, to start lignin chains are interesting; tricin itself is valuable, and plants with tricin knocked out have higher lignin and lower CW digestibility.9-13 

1.    Mottiar Y, Vanholme R, Boerjan W, Ralph J, & Mansfield SD (2016) Designer lignins: harnessing the plasticity of lignification. Current Opinion in Biotechnology 37(1):190-200.



2.    Rinaldi R, Jastrzebshi R, Clough MT, Ralph J, Kennema M, Bruijnincx PCA, & Weckhuysen BM (2016) Paving the way for lignin valorisation: Recent advances in bioengineering, biorefining and catalysis. Angewandte Chemie (International Edition) 55(29):8164-8215.



3.    Grabber JH, Hatfield RD, Lu F, & Ralph J (2008) Coniferyl ferulate incorporation into lignin enhances the alkaline delignification and enzymatic degradation of maize cell walls. Biomacromolecules 9(9):2510-2516.



4.    Ralph J (2010) Hydroxycinnamates in lignification. Phytochemistry Reviews 9(1):65-83.



5.    Wilkerson CG, Mansfield SD, Lu F, Withers S, Park J-Y, Karlen SD, Gonzales-Vigil E, Padmakshan D, Unda F, Rencoret J, & Ralph J (2014) Monolignol ferulate transferase introduces chemically labile linkages into the lignin backbone. Science 344(6179):90-93.



6.    Karlen SD, Zhang C, Peck ML, Smith RA, Padmakshan D, Helmich KE, Free HCA, Lee S, Smith BG, Lu F, Sedbrook JC, Sibout R, Grabber JH, Runge TM, Mysore KS, Harris PJ, Bartley LE, & Ralph J (2016) Monolignol ferulate conjugates are naturally incorporated into plant lignins. Science Advances 2(10):e1600393.



7.    Smith RA, Gonzales-Vigil E, Karlen SD, Park J-Y, Lu F, Wilkerson CG, Samuels L, Mansfield SD, & Ralph J (2015) Engineering monolignol p-coumarate conjugates into Poplar and Arabidopsis lignins. Plant Physiology 169(4):2992-3001.



8.    Sibout R, Le Bris P, Legee F, Cezard L, Renault H, & Lapierre C (2016) Structural redesigning Arabidopsis lignins into alkali-soluble lignins through the expression of p-coumaroyl-CoA:monolignol transferase PMT. Plant Physiology 170(3):1358-1366.



9.    del Río JC, Rencoret J, Prinsen P, Martínez ÁT, Ralph J, & Gutiérrez A (2012) Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods. Journal of Agricultural and Food ����vlog�ٷ���� 60(23):5922-5935.



10. Lan W, Lu F, Regner M, Zhu Y, Rencoret J, Ralph SA, Zakai UI, Morreel K, Boerjan W, & Ralph J (2015) Tricin, a flavonoid monomer in monocot lignification. Plant Physiology 167(4):1284-1295.



11. Lan W, Rencoret J, Lu F, Karlen SD, Smith BG, Harris PJ, del Rio JC, & Ralph J (2016) Tricin-lignins: Occurrence and quantitation of tricin in relation to phylogeny. The Plant Journal 88(6):1046-1057.



12. Lan W, Morreel K, Lu F, Rencoret J, del Rio JC, Voorend W, Vermerris W, Boerjan W, & Ralph J (2016) Maize tricin-oligolignol metabolites and their implications for monocot lignification. Plant Physiology 171(2):810-820.



13. Eloy NB, Voorend W, Lan W, Cesarino I, Vanholme R, de Lyra Soriano Saleme M, Goeminne G, Pallidis A, Morreel K, Nicomedes J, Ralph J, & Boerjan W (2017) Silencing chalcone synthase in maize impedes the incorporation of tricin into lignins and increases lignin content. Plant Physiology 173:998-1016.

Iron enzymes activate dioxygen for a large number of biomedically, agriculturally, and environmentally important oxidation reactions. Among those that use non-heme mono-iron cofactors, protein ligand sets as minimal as a pair of cis histidines enable the cofactor to coordinate and approximate as many as three different substrates, leading to an astounding array of different reaction types. Many of these reactions install key functional groups in lucrative natural-product drugs. Over the last 15 years, we have established the intermediacy of iron(IV)-oxo (ferryl) complexes in the reactions of several such enzymes with a range of distinct outcomes.¹ In general, the ferryl intermediates generate substrate radicals by abstracting hydrogen (H•) from unactivated aliphatic carbons,^2-6 initiating formation of new C–O,^2-4 ��–C��/����,^5,6 ��–S,⁷ ��–N,⁸ or C–C bonds.⁹ Hydroxylation is the default outcome, as it results from coupling between the carbon radical and the necessarily adjacent Fe(III)-coordinated oxygen that just generated it (termed “oxygen rebound” by Groves). Thus, the first imperative for enzymes that mediate outcomes other than hydroxylation is to avoid the default rebound, which generally has a low activation barrier. In our current work, we are defining the manifold of other pathways by which the substrate radical can decay and the strategies by which a given protein scaffold (or, in rare cases, the substrate itself) specifies a reaction channel. A generally important parameter is the disposition of the substrate relative to the cofactor,¹⁰ which appears to be controlled not only by the substrate pocket but also by the geometry of the ferryl complex (oxo trans to either one of the two conserved histidine ligands).¹¹ It has therefore been important to have tools to visualize intermediates (especially the ferryl complexes) through the reaction sequences.^12,13 I will summarize several of the Penn State team’s recent successes in defining reaction pathways and explaining control of outcome in this versatile enzyme family.  

1.  Krebs, C., Galonic, D.; Walsh, C. T.; Bollinger, J. M., Jr. "Non-Heme Fe(IV)-Oxo Intermediates," Acc. Chem. Res., 2007, 40, 484-492.

2.  Price, J. C.; Barr, E. W.; Tirupati, B.; Bollinger, J. M., Jr.; Krebs, C.; “The First Direct Characterization of a High-Valent Iron Intermediate in the Reaction of an aKetoglutarate-Dependent Dioxygenase: A High-Spin Fe(IV) Complex in Taurine:a-Ketoglutarate Dioxygenase (TauD) from Escherichia coli,” Biochemistry, 2003, 42, 7497-7508.

3.  Price, J. C.; Barr, E. W.; Glass, T. E.; Krebs, C.; Bollinger, J. M., Jr.; “Evidence for Hydrogen Abstraction from C1 of Taurine by the High-Spin Fe(IV) Intermediate Detected during Oxygen Activation by Taurine:a-Ketoglutarate Dioxygenase (TauD),” J. Am. Chem. Soc., 2003, 125, 13008-13009.

4.  Hoffart, L. M.; Barr, E. W.; Guyer, R. B.; Bollinger, J. M., Jr.; Krebs, C. “Direct spectroscopic detection of a C-H-cleaving high-spin Fe(IV) complex in a prolyl-4-hydroxylase,” Proc. Natl. Acad. Sci. USA, 2006, 103, 14738-14743.

5.  Galonic, D. P.; Barr, E. W.; Walsh, C. T.; Bollinger, J. M., Jr.; Krebs, C. “Two Interconverting Fe(IV) Intermediates in Aliphatic Chlorination by the Halogenase CytC3,” Nat. Chem. Biol., 2007, 3, 113-116.

6.  Matthews, M. L.; Krest, C. M.; Barr, E. W.; Vaillancourt, F. H.; Walsh, C. T.; Green, M. T.; Krebs, C.; Bollinger, J. M., Jr. “Substrate-Triggered Formation and Remarkable Stability of the C-H Bond-Cleaving Chloroferryl Intermediate in the Aliphatic Halogenase, SyrB2,” Biochemistry, 2009, 48, 4331-4343.

7.  Tamanaha, E. Y.; Zhang, B.; Guo, Y.; Chang, W.-c.; Barr, E. W.; Xing, G.; St. Clair, J.; Ye, S.; Neese, F.; Bollinger, J. M., Jr.; Krebs, C. “Spectroscopic evidence for the two C-H-cleaving intermediates of Aspergillus nidulans isopenicillin N synthase," J. Am. Chem. Soc. 2016, 138, 8862-8874.

8.  Matthews, M.L.; Chang, W.-c.; Layne, A.P.; Miles, L.A.; Krebs, C.; Bollinger, J. M., Jr. “Direct Nitration and Azidation of Aliphatic Carbons by an Iron-dependent Halogenase,” Nat. Chem. Biol. 2014, 10, 209-215.

9.  Dunham, N. P.; Chang, W.-c.; Mitchell, A. J.; Martinie, R. J.; Zhang, B.; Bergman, J. A.; Rajakovich, L. J.; Wang, B.; Silakov, A.; Krebs, C.; Boal, A. K.; Bollinger, J. M., Jr. (2018) "Two Distinct Mechanisms for C–C Desaturation by Iron(II)- and 2-(Oxo)glutarate-Dependent Oxygenases: Importance of α-Heteroatom Assistance," J. Am. Chem. Soc., in review.

10. Matthews, M. L.; Neumann, C. S.; Miles, L. A.; Grove, T. L.; Booker, S. J.; Krebs, C; Walsh, C. T.; Bollinger, J. M., Jr. "Substrate positioning controls the partition between halogenation and hydroxylation in the aliphatic halogenase, SyrB2," Proc. Natl. Acad. Sci. USA, 2009, 106, 17723-17728.

11. Martinie, R. J.; Livada, J.; Chang, W-c.; Green, M. T.; Krebs, C.; Bollinger, J. M., Jr.; Silakov, A. "Experimental Correlation of Substrate Position with Reaction Outcome in the Aliphatic Halogenase, SyrB2," J. Am. Chem. Soc. 2015, 137, 6912-6919.

12. Martinie, R. J.; Pollock, C. J.; Matthews, M. L.; Bollinger, J. M., Jr.; Krebs, C; Silakov, A. “Vanadyl as a Stable Structural Mimic of Reactive Ferryl Intermediates in Mononuclear Nonheme-Iron Enzymes,” Inorg. Chem. 2017, 56, 13382-13389.

13. Mitchell, A. J.; Dunham, N. P.; Martinie, R. J.; Bergman, J. A.; Pollock, C. J.; Hu, K.; Allen, B. D.; Chang, W.-c.; Silakov, A.; Bollinger, J. M., Jr.; Krebs, C.; Boal, A. K. “Visualizing the Reaction Cycle in an Iron(II)- and 2-(Oxo)-glutarate-Dependent Hydroxylase,” J. Am. Chem. Soc. 2017, 139, 13830-13836.