Tibor PÁLI
scientific adviser

Balázs SZALONTAI retired scientific adviser
Zoltán KÓTA research associate
Krisztina SEBÕKNÉ NAGY research associate

Group webpage >>


The main projects of this research group are: the proton-transporting membrane protein complex, the vacuolar proton-ATPase (V-ATPase); soluble and trans-membrane alpha-helix and beta-strand polypeptide bundles; and molecular interactions in native and model biomembranes. These topics are strongly related to each other both thematically and methodically. Studies aim at structure-function relationships, folding and assembly of membrane proteins and plolypeptides and the unique functional and structural role of the protein-solvent interface. Our spectroscopy- and calorimetry-based, function-driven structure biology approach relies on the techniques of Fourier transform infrared (FTIR), site-specific (spin label) electron paramagnetic resonance (EPR), UV, visible and fluorescence spectroscopies, and high-sensitivity differential scanning calorimetry (DSC). The structural, dynamic and thermodynamic data on functioning bio- and model membrane-protein systems yield functionally relevant molecular and physical models – this is functional structure biology.

Proton pumping by a membranous molecular motor, the vacuolar proton-ATPase

Figure 1: The rotary mechanism of V-ATPase. Each subunit c binds a proton when in contact with lipids. Protons enter and leave the rotor in hydrophilic input and output channels, between subunits c and a (based on Ferencz et al., 2013).

The internal compartments of eukaryotic cells are more acidic than the cytoplasm. The transport protein complex that is responsible for the acidification is nature's most universal proton pump, the vacuolar proton-ATPase. It is a membrane-bound molecular rotary engine, which converts the chemical energy from ATP hydrolysis to the rotation of the rotor domain via a torque between specific subunits. This leads to trans-membrane proton pumping in the interface between the stator and rotor domains. Our studies on V-ATPase aim at subunit-subunit and subunit-lipid interactions, the effect of synthetic inhibitors on function and subunit assembly, the uncoupling of passive proton translocation and ATP hydrolysis, and the details of the rotary mechanism (Fig. 1). We have recently measured the rate of rotation of the rotor in the yeast V-ATPase in native vacuolar membrane vesicles. We were the first to report the rotation rate in a V-ATPase that is not subjected to genetic or chemical modification and it is not fixed to a solid support. We have also discovered that the activity of V-ATPase can be affected with an oscillating trans-membrane electric field. The effects of external electric field are being currently used to explore further details of the rotary mechanism.

Protein insertion, folding and assembly in biomembranes and on membrane surfaces

Figure 2: The membranous dimer conformations of gramicidin A, shown together with a phospholipid (based on Kota et al., 2004).

Membrane protein folding is a most challenging problem in biophysics today because membrane lipids and proteins are coupled structurally, dynamically and functionally. The protein-lipid interface takes several different forms, all of which are crucial to biology. Studies on the structure, dynamics and function of both membrane proteins and lipids are essential for understanding membrane protein folding. Activity measurements on purified membrane proteins require that they are inserted and assembled in the bilayer properly. Our objective is to obtain various spectroscopic and calorimetric data on factors controlling insertion, folding and assembly of selected proteins and polypeptides in membranes and on membrane surfaces. These data are then used as constraints in molecular and physical models. Currently we focus on three groups of proteins: trans-membrane helix assemblies and beta-barrels (V-ATPase subunits and E.coli pore proteins), and soluble proteins or polypeptides (gramicidin A, lysozyme) (Fig. 2) interacting with bio- and model membranes.

Molecular modelling of membrane proteins

Figure 3: The sequence and the 3-dimensional fold of the major coat protein of the M13 bacteriophage, surrounded with the solvation or first shell lipids (based on Bashtovyy et al. , 2001).

Structural biology of membrane proteins faces the challenges of isolating, solublising and crystallising proteins (that are natively hosted by the lipid matrix) in a foreign environment. In addition, there is always the intriguing question: to what extent the obtained structures and features of isolated membrane proteins correlate with those under native, in-membrane conditions. Theoretical approaches have therefore enormous importance. Fig. 3 shows the essence of the fundamental and most challenging problem of structure biology: if we know the sequence of a protein, how can we determine its structure? Since the native fold of a membrane protein assumes a native environment, namely the lipid bilayer, the sequence-to-fold coding, illustrated in Fig. 3, is valid only in the native membranous environment. Therefore, our approach is to measure structural data on the target membrane proteins in their functional, native membrane environment (see above), and combine the so-obtained data with bioinformatics, homology-based predictions, loop databases using molecular mechanics in order to predict the 3-dimensional fold. A good example is our prediction of the basic fold of the conserved four-helix trans-membrane bundle of the cytochrome b561 membrane protein family: even after more than two decades of failed efforts for crystallizing any protein from the family, our published structures are still the only ones available. In the future we will develop and add codes to existing coarse grained molecular mechanics algorithms in order to implement experimentally derived knowledge-based potentials, aiming at improved structure prediction of membrane proteins.

Lipid-protein interactions in biological and model systems

Figure 4: Ordered (gel) and disordered (fluid) phospholipids. The hydrophobic and hydrophilic membrane regions are indicated too.

The lipids assure the insulation capacity of the membranes, and their hydrophobic double layer provides the proper conditions for the membrane proteins. Therefore, the role of the lipid-protein interface is crucial in all biomembrane functions, and isolated membrane proteins are very difficult to study. We use both non-invasive and labelling techniques for studying the lipid-protein, or, more generally, the solvent-protein interface: we apply and further develop spin-label EPR and the non-invasive FTIR spectroscopies to study model and biological membranes. These methods have the advantage of displaying separated lipid- and protein-related spectral regions. The separation is extremely good in the FTIR spectra. Thus, both lipids and proteins can be studied individually. Furthermore, the lipid-protein interaction in the membranes can also be addressed via the correlations between changes in the lipid- and protein-related regions. Based on our novel evaluation techniques, we could determine the key elements in the coupling between lipid and protein dynamics in biological membranes among cold- and heat-stress conditions. We are moving forward to explore the membrane structural consequences of heat stress by studying living cells cultivated on the surface of attenuated total reflection crystals. Our other present interest is to collect specific data on “almost single“ molecules by using surface enhanced infrared absorption spectroscopy, which enables the detection of infrared absorption from only 8-10 nm thick layers. With this technique, and keeping in mind the Hofmeister phenomenon, we want to study (in a cooperation within the Institute) the structure of water around biological molecules, which, according to our hypothesis, might also exert ordering or disordering effect on the interfacial water. Such an effect could play important role in molecular interactions, e.g., in the action mechanism of heat-stress proteins. For the observation of a very small number of molecules, we use nanotechnological methods as well, like functionalizing surfaces by polyelectrolyte films.

Free radicals in membranes, tissues and food products

Since EPR spectroscopy is the most reliable and most established technique for detecting free radicals, we are involved in several collaborative projects aiming at detecting free radicals in biological samples, including biomembranes. The diverse biological functions of nitric oxide (NO) keep it in the focus of biochemical research. Many functions of NO are membrane-associated. We have therefore determined the membrane penetration profile of NO from the perturbation that it exerts on spin-labeled lipids. In collaboration with the Department of Biochemistry of the Szeged University, we are quantitating NO in tissues from different mammalian organs using spin trapping techniques. In collaboration with the Institute of Plant Biology, we investigate the role of monodehydroascorbate radical in photosynthetic membranes. In addition, we are routinely measuring non-specific and cellulose based radicals in food products, such as paprika and milk powder, dried onion and, currently, milled coffee.

Selected publications

Holzenburg, A., Jones, P.C., Franklin, T., Pali, T., Heimburg, T., Marsh, D., Findlay, J.B.C., and Finbow, M.E. (1993). Evidence for a Common Structure for a Class of Membrane Channels. European Journal of Biochemistry 213(1): 21-30.

Csonka, C., Pali, T., Bencsik, P, Gorbe, A., Ferdinandy, P. and Csont, T. (2015) Measurement of NO in biological samples. British Journal of Pharmacology 172(6), 1620-1632.

Kostrzewa, A., Pali, T., Froncisz, W. and Marsh, D. (2000). Membrane location of spin-labeled cytochrome c determined by paramagnetic relaxation agents. Biochemistry 39(20): 6066-6074.

Bashtovyy, D., Marsh, D., Hemminga, H.M. and Pali, T. (2001). Constrained modelling of spin-labelled major coat protein mutants from M13 bacteriophage in a phospholipid bilayer. Protein Science 10(5): 979-987.

Schwinte, P., Voegel, J.C., Picart, C., Haikel, Y., Schaaf, P., and Szalontai, B. (2001). Stabilizing effects of various polyelectrolyte multilayer films on the structure of adsorbed/embedded fibrinogen molecules: An ATR-FTIR study. J. Phys. Chem. B. 47: 11906-11916.

Kota, Z., Horvath, L.I., Droppa, M., Horvath, G., Farkas, T., and Pali, T. (2002). Protein assembly and heat stability in developing thylakoid membranes during greening. Proc. Natl. Acad. Sci. USA 99(19): 12149-12154.

Bashtovyy, D., Berczi, A., Asard, H. and Pali, T. (2003). Structure prediction for the di-heme cytochrome b-561 protein family. Protoplasma 221(1-2): 31-40.

Kispeter, J., Bajusz-Kabok, K., Fekete, M., Szabo, G., Fodor, E. and Pali, T. (2003). Changes induced in spice paprika powder by treatment with ionising radiation and saturated steam. Radiation Physics and Chemistry 68(5): 893-900.

Pali, T., Garab, G., Horvath, L.I. and Kota, Z. (2003). Functional significance of the lipid-protein interface in photosynthetic membranes. Cellular Molecular Life Sciences 60(8): 1591-1606.

Kota, Z., Pali, T. and Marsh, D. (2004). Orientation and lipid-peptide interactions of gramicidin A in lipid membranes: polarized ATR infrared spectroscopy and spin-label electron spin resonance. Biophysical Journal 86(3): 1521-1531.

Marsh, D. and Pali, T. (2004). The protein-lipid interface: perspectives from magnetic resonance and crystal structures. Biochimica et Biophysica Acta - Biomembranes 1666(1-2): 118-141.

Laczko-Dobos, H. and Szalontai, B., (2009). Lipids, proteins, and their interplay in the dynamics of temperature-stressed membranes of a cyanobacterium, Synechocystis PCC 6803. Biochemistry 48: 10120–10128. Szalontai, B. (2009). Membrane protein dynamics: Limited lipid control. PMC Biophysics 2: 1.

Nagy, K., Pilbat, A., Groma, G., Szalontai, B, and Frederic J. G. Cuisinier (2010). Casein aggregates built step-by-step on charged polyelectrolyte film surfaces are calcium phosphate-cemented J. Biol. Chem. 285: 38811-38817.

Ferencz, C., Petrovszki, P., Kota, Z., Fodor-Ayaydin, E., Haracska, L., Bota, A., Varga, Z., Der, A., Marsh, D. and Pali, T. (2013). Estimating the rotation rate in the vacuolar proton-ATPase in native yeast vacuolar membranes. European Biophysics Journal 42: 147–158.

Pali, T. and Kota, Z. (2013). Studying lipid-protein interactions with electron paramagnetic resonance spectroscopy of spin-labeled lipids. In: Lipid-Protein Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 974, Jorg H. Kleinschmidt (ed.), Springer New York 2013: pp. 297-328.

Szalontai, B., Nagy, G., Krumova, S., Fodor, E., Pali, T., Taneva, S.G., Garab, G., Peters, J., and Der, A. (2013) Hofmeister ions control protein dynamics. Biochimica et Biophysica Acta - General Subjects 1830: 4564–4572.

Csonka, C., Pali, T., Bencsik, P, Gorbe, A., Ferdinandy, P. and Csont, T. (2015) Measurement of NO in biological samples. British Journal of Pharmacology 172(6), 1620-1632.

Ferencz, Cs-M., Petrovszki, P., Der, A., Sebok-Nagy, K., Kota, Z. and Pali, T. (2017) Oscillating electric field measures the rotation rate in a native rotary enzyme. Nature – Scientific Reports (accepted for publication).