Malte U Hammer

Annemarie Brauser

Claudia Olak

Gerald Brezesinski

Torsten Goldmann

Thomas Gutsmann

Jörg Andrä

Lipopolysaccharide interaction is

Keywords: AFM, atomic force microscopy, AMP, antimicrobial peptide, CD, circular dichroism, CFU, colony forming unit, TEM, transmission electron microscopy, FRET, Förster resonance energy transfer, LPS, lipopolysaccharide, MIC, minimal inhibitory concentration, MBC, minimal bactericidal concentration, PE, phosphatidylethanolamine, PG, phosphatidylglycerol, surface acoustic wave, TFE, trifluoroethanol antimicrobial peptide, atomic force microscopy, biosensor, endotoxin, membrane permeabilization, NK-lysin Biochemical

come

Lipopolysaccharide interaction is decisive for the activity of the antimicrobial peptide NK-2 against Escherichia coli and Proteus mirabilis † Malte U. Hammer * † , Annemarie Brauser * , Claudia Olak ‡ , Gerald Brezesinski ‡ , Torsten Goldmann * , Thomas Gutsmann * , and Jörg Andrä *1

INTRODUCTION

The emergence of a rising number of bacterial strains resistant against common antibiotics has urged the search for potent alternatives [START_REF] Levy | Antibacterial resistance worldwide: causes, challenges and responses[END_REF][START_REF] Norrby | Lack of development of new antimicrobial drugs: a potential serious threat to public health[END_REF]. Over the last decades natural antimicrobial peptides (AMPs) and derivatives thereof have attracted the interest of researchers and pharmacologists [START_REF] Zasloff | Antimicrobial peptides of multicellular organisms[END_REF][START_REF] Hancock | Antimicrobial and host-defense peptides as new anti-infective therapeutic strategies[END_REF]. AMPs are effective against bacteria, including multiresistant strains, prevent biofilm formation [START_REF] Overhage | Human host defense peptide LL-37 prevents bacterial biofilm formation[END_REF], and thus have gained high interest as lead structures for new anti-infectives. To adopt these compounds for clinical applications it is of utmost importance to understand their mode of action on a molecular level. Cationicity and amphipathicity enable interaction with the cell wall of bacteria, and a dysfunction of this barrier as a direct consequence of peptide membrane interaction has been widely accepted as important step in peptide-mediated killing of bacteria [START_REF] Shai | Mode of action of membrane active antimicrobial peptides[END_REF][7][8]. This eventually results either in direct killing of bacteria or allows the uptake of the antibiotic peptide into the bacteria [9,10]. The molecular basis behind this mechanism, however, is still a matter of debate and far from being set.

We selected the peptide NK-2 [11], representing the cationic core region of NK-lysin, as a model compound to understand the molecular principles of bacterial killing and membrane interaction of AMPs. NK-lysin has been isolated originally from porcine NK cells. It exhibits antibacterial as well as tumourolytic activities [12]. Homologue proteins have been described in human (granulysin [13]), pathogenic amoeba (amoebapores [14]), and other species [15][16][17]. They are all membrane-interacting proteins [18] and belong to the saposinlike protein family (SAPLIP) [19], which is characterized by a common -helical fold stabilized by three disulfide bridges [20,21]. Structure-function analysis of amoebapores showed that its central -helical region is the key factor for membrane interaction [22,23]. Inspired by the homology of amoebapore to NK-lysin [24], we synthesized the respectivehelical fragment of NK-lysin, termed NK-2, and investigated its biological activities and membrane-interaction properties.

The peptide NK-2 consists of 27 amino acid residues with an overall positive net charge due to arginine and lysine residues. It adopts an amphipathic, -helical secondary structure in TFE and upon phospholipid interaction [11,[25][26][27]. NK-2 kills bacteria [26], fungi [11], parasites [28,29], as well as certain cancer cells [30]. Additionally it is an effective neutralizing agent of bacterial endotoxin (lipopolysaccharide, LPS) [31]. In contrast to this, human erythrocytes [11], glioblastoma cells [28], and primary lymphocytes [30] have been found to be protected against cytotoxic NK-2 effects. Because of this selectivity for bacteria over normal human cells, NK-2 is a potential ´peptide antibiotic´ for therapeutic usage.

The selectivity of NK-2 could be assigned to differences in the membrane phospholipid composition of the target cells [25,26,30]. Whereas bacterial membranes are characterized by a substantial amount of negatively charged lipids, e.g. LPS in the outer membrane and phosphatidylglycerol (PG) and cardiolipin in the cytoplasmic membrane, the outer layer of the plasma membrane of human cells consists of the zwitterionic phosphatidylcholine (PC) and sphingomyelin and lacks anionic phospholipids [32]. NK-2 has been shown to bind to negatively charged phosholipid model membranes and to permeabilize them, but not to zwitterionic ones. However, neither conductivity, voltage dependency, nor life time of NK-2-induced lesions in phosholipid membranes or any other membranes have been characterized yet.

In a previous paper we screened the activities of NK-2 and of a set of peptides inspired from the NK-2 sequence against a panel of Gram-negative bacterial strains [26]. Although PG is the main target of NK-2 in membranes mimicking the cytoplasmic bacterial membrane [25,33], only a qualitative but no quantitative correlation has been found between the antibiotic activity and the degree of peptide intercalation into PG liposomes. Apparently, in the case of Gram-negative bacteria, the outer layer of the outer membrane is the first barrier against all antibiotic compounds. Described targets for AMPs are the destruction of the cytoplasmic membrane or further intracellular targets [9,10]. Either way, peptides have to attach to the LPS layer and permeate through the outer membrane to reach their final target [34,35]. Therefore, the overcome of the LPS containing membrane is crucial for the mode of action of AMPs against Gram-negative bacteria. The lipid matrix of the outer layer of the highly asymmetric outer membrane is composed almost exclusively of the glycolipid LPS, which is considered to be the first target structure for AMPs in the case of Gram-negative bacteria. In particular, the chemical compositions of the LPS carbohydrate chains determine its interaction with NK-2 and lactoferricin-derived peptides [31,36]. Thus, we extended our studies and dissected the different steps of NK-2 interactions (association, insertion, permeabilization) with artificial membranes composed of LPS and with live bacteria more detailed. We focused on two Gram-negative bacterial strains which exhibit pronounced differences in the susceptibility to various AMPs including the membrane permeabilizing antibiotic polymyxin B. This differential susceptibility could be assigned to variations in the degree of positively charged arabinose attached to phosphate groups of the LPS headgroup and to the Kdo core carbohydrates resulting in differences in net charge and charge distributions of the individual LPS structures [35,37,38]. Our working hypothesis is that these structural changes are decisive for differences in the antibiotic action of AMPs as they prevent or reduce the electrostatic attraction of the polycationic AMPs and the anionic LPS. To bridge the gap between the interaction of peptides with bacteria and artificial membranes, we isolated LPS from the bacterial strains we used for the biological assays. This chemically well-defined LPS was used for the reconstitution of biologically relevant model membranes of different complexity: we comprehensively investigated the interaction of NK-2 and its biologically inactive derivative NK11 [26] with LPS aggregates, immobilized LPS bilayers, and non-supported Montal-Mueller asymmetric LPS/phospholipid planar bilayers. Various biophysical techniques, including circular dichroism (CD) spectroscopy, surface acoustic wave (SAW) biosensor, trans-membrane conductivity measurements, and FRET spectroscopy were used to characterize the effects of the used peptides to these reconstituted model membranes. These biophysical studies were complemented by experiments using intact bacteria, which provide information about the direct binding of peptides to bacteria, permeabilization of bacterial membranes, bacterial viability and change of the bacterial morphology upon peptide treatment.

EXPERMIMENTAL

Peptides

NK-2 (KILRGVCKKIMRTFLRRISKDILTGKK-NH 2 ), NK11 (KISKRILTGKK-NH 2 ), and melittin (GIGAVLKVLTTGLPALISWIKRKRQQ-NH 2 ) were synthesized with an amidated C terminus by the Fmoc solid-phase peptide synthesis technique on an automatic peptide synthesizer (model 433 A; Applied Biosystems) as described [31]. NBD-labelled NK-2 was prepared as follows: Peptide coupled to the resin and with protected sidechains was suspended in dimethylformamide, supplemented with trace amounts of pyridine. Then, a 4-fold molar excess of 4-chloro-7-nitro-1,2,3-benzoxadiazole (NBD chloride, Fluka, Germany) was added and the suspension was stirred gently overnight at 20°C. The resin was filtered, washed and the peptide was cleaved from the resin as described above. Polymyxin B was purchased from Sigma Aldrich (Deisenhofen, Germany).

Bacterial strains

All used bacterial strains were LPS rough or deep rough mutants: Salmonella enterica (serovar Minnesota) strain R60 (LPS Ra) and strain R595 (LPS Re), Escherichia coli strain WBB01 (LPS Re), and Proteus mirabilis strain R45 (LPS Re). Bacteria were grown over night in Luria-Bertani (LB) medium composed of 1% tryptone, 0.5% yeast extract, and 1% NaCl under constant shaking at 37°C and subsequently inoculated in the same medium to reach the mid-exponential phase.

Lipids and reagents

Phospholipids (cardiolipin (bovine heart), L--phosphatidylethanolamine (PE, Escherichia coli), L--phosphatidyl-DL-glycerol (PG, chicken egg)) were purchased from Avanti Polar Lipids (Alabaster, AL, USA). Fluorescently labelled phospholipids N-(7nitrobenz-2-oxa-1,3-diazol-4yl)-phosphatidylethanolamine (NBD-PE) and N-(lissamine rhodamine B sulfonyl)-phosphatidylethanolamine (Rh-PE) were from Molecular Probes (Eugene, OR, USA). Rough type LPS were extracted by the phenol/chloroform/petrol ether method from bacteria grown at 37°C, purified, and lyophilized [39]. All other chemicals were analytical grade and acquired from Merck (Darmstadt, Germany).

Preparation of LPS aggregates

LPS dispersed in buffer (see respective method) by vortexing, was sonicated in a water bath for 30 min at 60°C. The samples were heated up to 60°C and cooled down to room temperature three or four times. After this cycles, the lipid samples were stored at least 12 h at 4°C before used. The size of the LPS aggregates in water was determined by dynamic light scattering. There are mainly two populations with 72 nm and 256 nm in a ration of 2:1, respectively.

Assay for antibacterial activity

The peptides were dissolved at the desired concentration in assay buffer (20 mM HEPES, pH 7.0 with or without additional 150 mM NaCl), and 180 µl of these solutions were filled in the wells of the first row of a microtiter plate. For a two-fold serially dilution, 90 µl of each peptide solution were transferred to the well in the next row, which was filled with 90 µl buffer before. Subsequently, a suspension of exp-phase bacteria in LB was added (10 µl, containing 10 4 CFU) to the peptide solution (90 µl). The constantly shaken plates were incubated in a wet chamber over night at 37°C. Bacterial growth was monitored by measuring the optical density at 620 nm in a microtiter plate reader (Rainbow, Tecan, Crailsham, Germany). The minimal inhibitory concentration (MIC) was defined as the lowest peptide concentration at which no bacterial growth was measurable. Portions of each well (10 µl) were diluted with buffer and plated out in duplicates on LB-agar plates. After incubation over night at 37°C bacterial colonies were counted. The minimal bactericidal concentration (MBC) was defined as the lowest peptide concentration at which no colony growth was observed. Determination of the time course of bacterial killing was performed in 20 mM HEPES, 150 mM NaCl, pH 7.0 without additional nutrient as described previously in detail [41].

Electron microscopy

Transmission electron microscopy (TEM) was performed as described previously in detail [26]. Bacteria (5x10 8 CFU/ml) were incubated with indicated concentrations of NK-2 and NK11 in buffer (20 mM HEPES, 150 mM NaCl, pH 7.0) supplemented with 10% LB medium for 30 min at 37°C. Images shown are representative for the respective sample.

Atomic force microscopy

Bacteria and bacteria/peptide samples as prepared for TEM were placed on mica, excess liquid was removed with filter paper and the bacteria were air dried at room temperature for 24 h. Dried bacteria were rinsed with 3 ml H 2 O and imaged with an atomic force microscope in air using a MFP-3D (Asylum Research, Santa Barbara, CA, USA). Imaging in air was performed with NSG11-B cantilever (k= 5.5 N/m; NT-MDT, Moscow, Russia) in AC (tapping) mode using frequencies of about 150 kHz. The frequency was chosen to result in an amplitude 5% lower than the amplitude at the resonance frequency. The set point was always adjusted to guarantee minimum forces applied to the sample. Further image processing (flattening and plane fitting) was done with the MFP-3D software under IGOR Pro (Lake Oswego, OR, USA). Images shown are representative for the respective sample.

Binding of fluorescently labelled peptide to live bacteria

Exp-phase bacteria were washed in Dulbecco's phosphate buffered saline without Ca 2+ , pH 7.4 (PBS, Biochrom AG, Berlin, Germany) and adjusted to 5x10 8 CFU/ml. NBD-NK-2 was added to the bacteria (800 µl, 10 µM final peptide concentration) and the suspension was incubated for 20 min at 37°C. The bacterial suspension was then analyzed with a fluorescence assisted cell sorter (FACS) flow cytometer (FACSCalibur, BD Biosciences, Heidelberg, Germany) with computer-assisted evaluation of data (CellQuest software). Bacteria were selected by Forward Scattered (FSC) and Side Scattered (SSC) signals. Then, we measured the intensity of green fluorescence (FL-1) of 10,000 single bacteria with and without fluorescently labelled peptide NBD-NK-2. By adding the measured fluorescence intensity multiplied with the corresponding number of observed events, we measure the relative amount of bound peptide. To take care of the self-fluorescence of used bacteria, the measured values of bacteria without peptide were subtracted and afterwards normalized to fluorescence intensity of NBD-NK-2 bound to bacteria of E. coli strain WBB01. Data presented are the mean of two independent experiments each performed in duplicates.

SYTOX green uptake assay

Bacteria with compromized membranes were detected by monitoring the fluorescence of the DNA-intercalating dye SYTOX Green (Invitrogen, Molecular Probes). Bacteria (E. coli strain WBB01) were grown to mid-exponential phase in LB medium and washed twice with and resuspended in 5 mM HEPES, 100 mM NaCl, pH 7.0 at a density of 10 7 CFU/ml. Peptides were diluted in the same buffer and incubated with bacteria (5x10 5 CFU) and with 2 µM of the fluorescent dye SYTOX Green in a total volume of 200 µl at 37 °C for 30 min. Permeabilization of the bacterial membranes allows the dye to enter the cytoplasm and to intercalate into the DNA. After incubation, the mixture was diluted 5 times and a fluorescence spectrum of the samples was measured after excitation at 488 nm in acryl cuvettes with a spectrofluorometer (Fluorolog-3, Horiba, USA). Total fluorescence intensity was obtained by curve integration from 504-600 nm. Membrane-permeabilizing activity of the peptides is expressed as percentage of permeabilized bacteria. For spontaneous lysis of the bacteria (0% value), incubation was done in buffer, for maximum permeabilization (100% value), cells were incubated with 0.5% triton x-100 in buffer. Experiments were done at least twice. Data shown represent mean +/-standard deviation.

Surface acoustic wave (SAW) biosensor measurements

Binding of peptides to immobilized lipid bilayers was analyzed using the S-sens K5 biosensor device (Biosensor GmbH, Bonn, Germany). The procedure has been described in detail elsewhere [42]. Note that in contrast to the previously described procedure, here we used a different type of dextran (see below). Functionalization of gold-coated chips (S-sens K5 Biosensor Quartz Chips, Biosensor GmbH, Bonn, Germany) was achieved by epichlorhydrine (Sigma) coupling of dextran (3 mg/ml in 0.1 M NaOH, average molecular mass 400-500 kDa, Sigma) to a self-assembled monolayer of 11-mercapto-1-undecanol (Aldrich), and finally carboxylation of dextran with bromoacetic acid (Sigma). The dry chip was stored until use at 4°C. Measurements were performed at a continuous buffer ( 5 A c c e p t e d M a n u s c r i p t HEPES, 100 mM KCl, pH 7.0, or, if indicated, 20 mM HEPES, pH 7.0 without or with additional 150 mM NaCl) flow of 20 µl/min at 22°C. By injection of 100 µl poly-L-lysin (PLL, 60 µg/ml, Fluka, Basel, Switzerland) a positively charged layer was formed on top of the negatively charged carboxymethylated-dextran matrix. Formation of a LPS bilayer was done by two consecutive injections of LPS aggregates (180 µl, 100 µM) each followed by 5 min buffer rinse. Peptide injections (= adsorption, 200 µl) were followed by 10 min buffer rinse (= desorption). Peptide binding to the PLL layer was negligible. All injections, but those containing peptide, were performed with an initial burst, a brief increase in flow to 150 µl/min. All binding experiments were performed at least with three independent preparations of LPS aggregates. Representative traces are shown. Phase shift of an acoustic surface wave corresponds roughly to a change in mass loading on the surface and is presented here normalized to the phase shift induced by the formation of the lipid bilayer.

CD spectroscopy

The LPS dispersion was prepared as described above. CD data were acquired with a Jasco J-715 CD spectrophotometer using quartz cuvettes with an optical path length of 0.1 cm. The response was measured from 190 to 300 nm with 0.2 nm step resolution and 1 nm bandwidth at 20°C. The rate was 50 nm/min using a 2 s response time. Each spectrum is a sum of at least four scans to improve the signal-to-noise ratio. All spectra are reported in terms of mean residue molar ellipticity ([ ] R = deg cm 2 dmol -1 ). Spectra were collected for samples of 63 µM peptide and 0.1 mM LPS in 10 mM potassium phosphate at pH 7.0 and 20°C.

Förster resonance energy transfer (FRET) spectroscopy

Intercalation of peptides into LPS aggregates was determined in 20 mM HEPES, pH 7.0 or in 20 mM HEPES, 150 mM NaCl, pH 7.0 at 37°C by FRET spectroscopy applied as a probe dilution assay [43]. The peptide was added to LPS aggregates, which were labelled with 1% of the donor NBD-phosphatidylethanolamine (NBD-PE) and 1% of the acceptor Rhodamine-PE. NBD-excitation wavelength was 470 nm. Intercalation was monitored as the increase of the ratio of the donor fluorescence intensity I D at 531 nm to that of the acceptor intensity I A at 593 nm (FRET signal) in dependence on time. This ratio depends on the Förster efficiency, therefore a rising value means that the mean distance separation of donor and acceptor dyes is raising.

Preparation of planar bilayers and electrical measurements

Planar bilayers were prepared according to the Montal-Mueller technique [44] as described earlier [45]. Briefly, symmetric and asymmetric bilayers were formed by opposing two lipid monolayers, prepared in separated compartments on aqueous subphases from chloroformic solutions of the lipids, at a small aperture (ca. 150 µm) in a thin teflon septum (thickness 25 µm). The inner leaflet of the outer membrane of Gram-negative bacteria was reconstituted by a phospholipid mixture (PL) consisting of PE, PG, and cardiolipin (molar ratio 81:17:2) resembling the phospholipid composition of the inner leaflet of the outer membrane of Salmonella typhimurium being composed of the same constituents as that of the other strains used [46]. For electrical measurements, planar membranes were voltage-clamped via a pair of Ag/AgCl electrodes (type IVM E-255, Science Products, Hofheim, Germany) connected to the headstage of an L/M-PCA patch-clamp amplifier (List-Medical, Darmstadt, Germany). In all experiments, the compartment to which peptide was added is named first, and the compartment opposite to the addition was grounded. In comparison to the natural system, a positive clamp voltage represents a membrane with negative potential on the inside. All measurements were performed at a temperature of 37°C in 5 mM HEPES, 100 mM KCl, 5 mM MgCl 2 , pH 7.0. Two electrical parameters were determined: (i) current through Assuming a plate condensator model, the membrane capacitance yields information about the area, thickness, and composition of the bilayer. Using small positive and negative jumps of the clamp voltage, the maximum voltage required to determine membrane capacitance is <1 mV. Therefore, peptide membrane interaction can be investigated without the influence of a significant external potential. This method allows the determination of changes in membrane capacitance with a precision of <1 pF.

RESULTS

Antibacterial activity of peptides

We tested the activity of NK-2, of the fluorescently labelled peptide NBD-NK-2, and of the shortened variant NK11 against two bacterial strains we have established in our lab as source for the extraction of highly purified and well characterized deep rough mutant LPS. These were E. coli WBB01 and P. mirabilis R45. The latter has been reported to be resistant to many cationic antimicrobial peptides and in particular to polymyxin B. The LPS structures of these strains differ by the degree of substitution with cationic aminoarabinose residues at the head group phosphates or at the first Kdo sugar. This results in variations in charge distribution and total net charges of -4 and -3 for E. coli WBB01 and P. mirabilis R45 LPS, respectively [35]. We also tested two internal reference strains from S. enterica with variations in the LPS structure: strain R60 with LPS Ra chemotype, i.e. complete core sugar, and the deep rough mutant strain R595 with LPS Re chemotype. The minimum peptide concentrations necessary to inhibit bacterial growth (MIC) and to kill bacteria (MBC) were determined under two different salt conditions (Tab. 1). The peptide NK-2 exhibited excellent activity against all used strains in low salt buffer. In physiological salt concentration, however, the activity against P. mirabilis was considerably impaired. The fluorescently labelled NBD-NK-2 exhibited almost the same activity as NK-2, i.e. the MIC/MBC values varied within one dilution step. The shortened version of NK-2, NK11 was almost inactive. The well known AMPs polymyxin B and melittin served as references. As demonstrated before, the activity of polymyxin B against P. mirabilis was extremely salt sensitive [41]. The differences in growth inhibiting and in bactericidal concentrations of NK-2 were also reflected in the killing kinetics. E. coli WBB01 was completely eradicated after 10 min of incubation with 5 µM NK-2, whereas the growth of P. mirabilis was reduced but not inhibited even after over night incubation by the same concentration of peptide (Fig. 1).

Influence of peptides on the ultrastructure of bacteria

The morphological changes of E. coli WBB01 and P. mirabilis R45 upon treatment with NK-2 and NK11 were visualized by TEM and AFM. NK-2 induced membrane blebbing, membrane ruffling, membrane detachment, and formation of electron dense dots in-and outside the cells of both strains (Fig. 2,3). Furthermore, the surface roughness of bacteria increased in the presence of the peptide (Fig. 4 B,E). This effect was even more pronounced for P. mirabilis. Additionally, NK-2 evoked distinct processes in the cytoplasm of the bacteria. The peptide induced the release of cytoplasmic material from E. coli (Fig. 2) and the formation of electron dense fibrous structures in P. mirabilis (Fig. 3 G,I). In sharp contrast, no profound differences were evident between NK11-treated bacteria and controls (Fig. 2,3). Only minor alterations of the bacterial surface were visible (Fig. 4). Peptide-induced changes of the bacterial morphology corresponded with viability. After incubation of bacteria with peptides, aliquots of the bacterial suspensions were plated out on LB agar plates and incubated over night. Colony growth of E. coli was completely inhibited by 2 µM NK-2, while 20 µM NK-2 reduced the growth of P. mirabilis colonies by 54%. NK11 (20 µM) had

Binding of fluorescently labelled NK-2 to live bacteria

Peptide binding to live bacteria was analyzed by FACS by measuring the green fluorescence (FL1) of single bacteria after incubation with the fluorescently labelled peptide NBD-NK-2 (Fig. 5). Dotplots of the FSC signal against the green fluorescent signal (FL1-Area) for E. coli WBB01 without and with NBD-labelled NK-2 are shown in Fig. 5 A andB, respectively. The fluorescence intensity of NBD-NK-2 bound to bacteria was measured and normalized to that of E. coli. This relative fluorescence signal (FL-1 Area) was 1.0 for E. coli WBB01 and 0.36 0.14 for P. mirabilis R45, respectively (Fig. 5C). Thus the binding of NBD-NK-2 to P. mirabilis was strongly impaired compared to E. coli.

Permeabilization of bacterial membranes

As the observed ultrastructure of NK-2-treated bacteria strongly suggested a membranolytic step in the mode of action of this peptide, we investigated the uptake of a DNA-intercalating dye, SYTOX green, by a suitable bacterial strain, i.e. E. coli WBB01, during incubation with NK-2, NK11 and melittin. Dye uptake directly reflects membrane permeabilization, as SYTOX green itself cannot permeate through an intact lipid bilayer. In the case of Gram-negative bacteria, inner and outer membrane permeabilization is apparently necessary to accomplish dye uptake. A dose-dependent uptake of the fluorescent dye was observed for bacteria treated with NK-2 and melittin (Fig. 6). In contrast, no increase of fluorescence was induced by NK11. The peptide concentration necessary to induce dye uptake in E. coli WBB01 correlated with the MIC of NK-2, however, occurred at a concentration considerably below the MIC for melittin.

Secondary structure of peptides

The secondary structures of NK-2, and NK11 were analyzed by CD spectroscopy. Both peptides appeared without defined structure in buffer [26]. When added to an aggregate suspension of E. coli WBB01 LPS, however, NK-2 adopted an ordered structure with a pronounced higher -helical content than NK11 (Fig. 7). Apparently, the spectrum of NK-2 in the presence of LPS does not reflect all characteristics of a typical CD spectrum of an -helical peptide [47], which has been observed earlier for NK-2 in the presence of TFE and SDS [11,26,27]. Whereas the bands at 190 nm and 222 nm are visible in the CD spectrum of NK-2 in the presence of LPS, the negative band at 208 nm is drastically reduced due to the interference with LPS and light scattering effects due to LPS aggregates. LPS alone produces already a strong CD signal, so that the interaction between polarized light and LPS interferes with its peptide interaction. Light scattering due to LPS particles is another problem which disturbs the CD spectrum. However, a deconvolution of the CD spectrum into contributions of different secondary structure elements using CONTIN/LL shows that the -helix content is around 50 %. The CD spectrum of NK11 is very low in intensity and difficult to analyze in terms of secondary structure elements. The -helix content is clearly much lower compared with NK-2, and there are strong contributions from unordered peptides.

Binding of peptides to solid-supported LPS bilayers

Binding of NK-2 and NK11 to LPS bilayers was monitored in real time using the SAW biosensor. Injection of suspensions of E. coli WBB01 LPS and of P. mirabilis R45 LPS led to the formation of LPS bilayers on top of a PLL layer (not shown), and induced a phase shift of 82 18 ° and of 162 23 °, respectively. Phase shifts induced by subsequent peptide injections were normalized to the LPS-induced phase shift, i.e. to the mass loading of the LPS bilayer (Fig. 8). Both peptides induced a phase shift after injection (each 100 µM) to LPS bilayers constituted of E. coli LPS WBB01, or of P. mirabilis LPS R45, indicating peptide- LPS binding in all cases. The maximal normalized increase of the phase induced by NK-2 injection was 84% and 44% for bilayers of E. coli LPS WBB01, and of P. mirabilis LPS R45, respectively. By NK11 injection the corresponding values were 14% and 18%. Therefore, the amount of NK-2 bound to the bilayer of E. coli LPS was twice as much as the amount of peptide bound to the bilayer of P. mirabilis LPS under the same conditions. Off rates were low for NK-2 but high for NK11, demonstrating significant differences in LPS binding affinities of the two peptides. To address salt dependency of the peptide-LPS interaction, we compared the binding of NK-2 to LPS in 20 mM HEPES, pH 7.0 and in the same buffer with additional 150 mM NaCl. The observed binding curves were very similar (not shown), suggesting that the ionic strength has no major impact on the amount of peptide bound to the LPS bilayer.

Insertion of peptides into the bilayer of LPS aggregates

FRET spectroscopy served as a sensitive tool for the insertion kinetics of NK-2 and NK11 into LPS bilayers. The peptides were added to LPS aggregates doped with the FRET pair NBD-and rhodamine-PE. A strong, dose-dependent increase of the FRET-signal was observed immediately after NK-2 addition indicating a fast insertion into the LPS aggregate. Eventually, the FRET signal reached a plateau value, similar for the two LPS (Fig. 9A,B). In contrast, addition of NK11 led to a reduced FRET signal, i.e. enhanced FRET efficacy, indicating a different mode of interaction. We suggest that NK11 did not intercalate into the lipid aggregate but bound superficially and induced the segregation of LPS and of PE. Hence, labelled PE were accumulated, which account for an increased FRET probability. Notably, differences in the ionic strength of the buffer have only a negligible influence on intercalation of the peptides into bilayers of LPS of P. mirabilis R45 (Fig. 9B,C) and of LPS of E. coli WBB01 (not shown).

Lesion formation by NK-2 in symmetric phospholipid and in asymmetric LPS/phospholipid planar lipid bilayers

The formation of membrane lesions or pores by the peptide NK-2 was done by the Montal-Mueller planar lipid bilayer technique. This system allows the reconstitution of membranes with virtually any lipid composition, including a symmetrical membrane of a defined phospholipid mixture (PL) mimicking roughly the cytoplasmic membrane of bacteria. Furthermore, it allows mimicking the asymmetrical outer membrane of Gram-negative bacteria consisting of LPS on its outer leaflet and of PL on its inner leaflet (LPS/PL). We determined the membrane capacitance as a measure of peptide/membrane interaction and measured the emerging current fluctuations while a transmembrane potential was applied. The peptide was always added to one side of the membrane, in case of the asymmetric LPS/PL membrane to the LPS side. After addition of NK-2 to a symmetric PL/PL or an asymmetric LPS/PL membrane, we observed initially a slight increase of membrane capacitance followed by a significant decrease of about 5-20% before the formation of any lesions or pores. A representative curve for a symmetric PL/PL membrane is depicted in Fig. 10. Since membrane capacitance is a result of many different factors (i.e. membrane thickness, area, and dielectricity) we cannot assign the factor or factors which are influenced by the peptide, however, the change of capacitance is a clear-cut indicator that an interaction takes place. After applying a transmembrane potential (negative inside), single current fluctuations were observed at -20 mV, which in the case of the symmetric PL/PL bilayer accumulate already at this low voltage over time. Finally, this led to the disruption of the membrane (Fig. 10). No current fluctuations were observed for NK11 under the same conditions (not shown). After the addition of NK-2 to asymmetric LPS/PL bilayers, we obtained current fluctuations of 50 pA to more than 400 pA at -100 mV (Fig. 11). These lesions were unstructured and heterogeneous without a clear gating characteristic and the induction of lesions seemed to be Eventually, the number and/or size of lesions increased dramatically and the integrity of the membrane was destroyed (not shown). In some cases, however, we observed a stable integration of the peptide NK-2 into the membrane resulting in defined conductance levels and gating (Fig. 12). They exhibited conductance levels up to 24 nS, which were relatively independent of the voltage polarity. No pronounced differences of the NK-2induced lesions between the two types of LPS membranes were observed. It should be emphasized that each planar lipid bilayer experiment is unique and we selected representative current traces for figures 10-12. However, due to the experimental setup, we cannot rule out stochastic events, which are apparently the reason for the formation of lesions with a defined conductance level (Fig. 12). Asymmetric bilayers (LPS/PL) with one monolayer of the membrane composed of LPS and the other one composed of a phospholipid (PL) mixture behave differently than symmetric PL/PL bilayers. Whereas PL/PL bilayers were immediately destabilized after the addition of NK-2 (Fig. 10), LPS/PL bilayers were of higher stability. In most experiments with asymmetric membranes, we observed heterogeneous lesions (Fig. 11), which were indistinguishable from those lesions observed in PL/PL bilayers. However, contrary to PL/PL bilayers, NK-2-induced lesions in LPS/PL bilayers sometimes closed spontaneously. This inherent membrane stability may be the reason for the stochastic formation of lesions with a defined conductance level (Fig. 12).

DISCUSSION

The NK-lysin derived peptide NK-2 exhibits activity against a wide range of Gramnegative bacteria, whereas its shortened derivative NK11 is almost inactive [26,31]. In this study, we asked three major questions: 1) Is NK-2-induced killing of Gram-negative bacteria mediated by membrane permeabilization, and what are the characteristics of NK-2-induced membrane lesions? Which types of model membranes are suitable 2) to allow a discrimination between a peptide with potent activity against Gram-negative bacteria and an inactive peptide, and 3) to explain the differential activity of a certain peptide against two Gram-negative bacterial strains? These questions have been investigated representatively with NK-2 and NK11 against E. coli WBB01 and P. mirabilis R45.

In several previous investigations, we have documented the surface activity of NK-2 [27] and its impact on artificial membranes composed of phospholipids [25,26,33] and LPS [31]. However, the direct link between artificial membranes and intact bacterial membranes is still missing. Here, we provide convincing evidences that a direct peptide-membrane interaction is involved in antibacterial action of NK-2: i) The killing of E. coli was rapid, also the delayed killing of P. mirabilis cultures does not necessarily exclude rapid killing of a single bacteria, and ii) was accompanied by dramatic changes of the bacterial morphology. The surface roughness increased upon peptide interaction and membrane effects such as membrane ruffling and detachment of the outer and inner membrane took place. Morphological membrane damage and membrane detachment have also been shown for other peptides and appear to be general features of membranolytic peptides such as melittin, lactoferricin B, and hecate 1 [48-50]. The observed changes distinguish lytic peptides from those membrane-interacting peptides acting primarily by a non-lytic mechansim such as human psoriasin [51]. iii) Even more convincing, NK-2 (and also melittin) permeabilized the membranes of viable E. coli cells as evidenced by uptake of the fluorescent DNAintercalating dye SYTOX green by the bacteria. iv) NK-2 concentrations necessary to induce bacterial killing, to permeabilize bacterial membranes, as well as to induce measurable effects on the artificial LPS model membranes were very similar. All findings together strongly support the thesis that LPS interaction and outer membrane permeabilization are crucial for bacterial killing by NK-2. Recently, we have analyzed the activity of NK-2-derived peptides against Gramnegative bacterial strains [26]. However, the applied simple artificial membrane system (PG liposomes), which was formerly sufficient to explain the target cell selectivity of NK-2 [25], failed to explain different activities of the tested peptides. In particular, PG was not a suitable model to explain differences in activity between the parent peptide NK-2 and NK11. The inactivity of NK11 was not due to its inability to interact with negatively charged lipid membranes in general. As we have shown earlier, NK11, like NK-2, intercalated into PG membranes [26]. Here, we show that peptide interactions with the bacterial outer membrane lipids, i.e. LPS, determine whether bacteria are killed or not. This is demonstrated by significant differences in amounts and affinities of NK-2 vs NK11 binding to solid supported LPS bilayers on a polymer cushion. Moreover, NK-2 desorption from the LPS bilayer was very slow indicating a high binding affinity. Upon contact with LPS containing membranes, the peptide NK-2, but not NK11, adopted a defined, mostly -helical, secondary structure. NK-2 intercalated into all tested LPS bilayer membranes, as demonstrated by FRET spectroscopy, whereas no intercalation was observed for NK11. Eventually, NK-2 induced transient, heterogeneous lesions without a clear gating characteristic in symmetric and asymmetric planar lipid bilayers mimicking the cytoplasmic and the outer membranes of E. coli and P. mirabilis. Stable integration of NK-2 was observed into asymmetric LPS/PL bilayers, but not in symmetric PL/PL bilayers, underlining the importance of the choice of lipids for peptide/membrane-interaction studies. The conductivity of these stable lesions was independent of the applied voltage polarity, which suggests a symmetrical pore structure. This is in contrast to the conductivity of membrane lesions induced by human -defensin 3 in LPS/PL bilayers, which was strongly dependent on the voltage polarity and which indicated an asymmetry in pore geometry or charge distribution [38].

The concentration of NK-2 necessary to kill P. mirabilis was higher than that to kill E. coli. This tendency was even more distinctive when NaCl was added at physiological concentration, suggesting that electrostatic interactions may be important for the antibacterial activity of NK-2. Moreover, this phenomenon was also observed with two reference AMPs used in this study, namely melittin and polymyxin B. The latter, which was completely inactive against P. mirabilis at physiological salt concentration rendered highly active in low ionic strength buffer. The used LPS from E. coli and P. mirabilis differ in the number of core carbohydrates and phosphate groups attached to aminoarabinose residues. These changes result in an altered charge distribution and slightly reduced net charge for P. mirabilis LPS [35,38]. Consistently, the enthalpy change of the binding reaction of NK-2 to LPS of P. mirabilis was lowered in the presence of 150 mM NaCl compared to buffer without salt (not shown). This is an apparent contradiction to the biosensor and FRET measurements which suggest that the binding and insertion of NK-2 into LPS is independent on the ionic strength. Currently, we have no satisfactorily explanation for this behaviour and we will investigate this issue in future studies. Besides electrostatic attraction, also other factors such as osmotic stress have to be considered to explain the salt-dependence of the antibacterial activity of NK-2 and other peptides. In Gram-negative bacteria, osmotic effects should be restricted to the cytoplasmic membrane. The presence of porins in the outer membrane enable the free flux of ions and thus prevent the emergence of osmotic pressure.

Interestingly, neither intercalation of NK-2 nor permeabilization of model membranes by this peptide appeared to be dependent on the differing LPS carbohydrate structures of the two tested LPS. Hence, these processes do not contribute to differences in biological activity. From the analysis of the interaction of fluorescently labeled NK-2 with viable bacteria and from biosensor data it is apparent that improved LPS binding of NK-2 is important for an enhanced activity against E. coli WBB01 compared to P. mirabilis. Thus, it may be concluded that electrostatic attraction of the peptide to the bacterial surface is the decisive step in the killing process, and that intercalation of NK-2 into the hydrophobic acyl chain region of the and subsequent permeabilization of the bacterial membranes are independent of the chemical structure of the LPS sugar part. A similar molecular basis for target discrimination has been described for protegrin action on Pseudomanas aeruginosa and Burkholderia cepacia [52]. Also different principles are known for other AMPs such as human -defensin 3, and polymyxin B, where the emergence and the size of LPS membrane lesions have been made responsible for activity, respectively [35,38].

Besides membranes also intracellular structures have been discussed as targets for AMPs [9,53]. For several peptides, a cytoplasmic localization has been demonstrated [54,55] or ruled out [56]. Membrane permeabilization and disturbance are important first steps in bacterial killing by NK-2, it should be mentioned, however, that this peptide also elicits bacteria-specific effects inside the cytoplasm, such as cytoplasmic retraction and formation of filamentous structures, suggesting that secondary targets of NK-2 are intracellular structures, such as DNA, or that the impact of NK-2 on the bacterial membrane results in clustering or rearrangement of these intracellular structures.

In summary, NK-2 binds to the surface of Gram-negative bacteria and also to LPS model membranes and subsequently permeabilizes bacterial membranes as well as artificial lipid bilayers composed of LPS. LPS binding affinity clearly differentiates active NK-2 from inactive NK11. We conclude that the LPS carbohydrate structures determine the binding affinity of the peptides to the bacterial surface and thus their biological activities. Our data stress the importance of membrane reconstitution systems which closely resemble the lipid compositions of their natural counterparts. A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.

FIGURE LEGENDS

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Biochemical

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  mM Biochemical Journal Immediate Publication. Published on 26 Feb 2010 as manuscript BJ20091607 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091607

  Biochemical Journal Immediate Publication. Published on 26 Feb 2010 as manuscript BJ20091607 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091607 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2010 The Authors Journal compilation © 2010 Portland Press Limited peptide-induced lesions and (ii) peptide-induced changes of the membrane capacitance.

  Biochemical Journal Immediate Publication. Published on 26 Feb 2010 as manuscript BJ20091607 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091607 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2010 The Authors Journal compilation © 2010 Portland Press Limited no effect on the viability of both strains.

Biochemical

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  membrane

Fig. 1 .

 1 Fig. 1. Time course of bacterial killing by NK-2. Bacteria ((A) E. coli WBB01, and (B) P. mirabilis R45) were incubated alone (control) and in the presence of 1 µM (open circles), 2 µM (filled circles), and 10 µM (open triangles) peptide in buffer (20 mM HEPES, 150 mM NaCl, pH 7.0) at 37°C. Viability of bacteria was assessed by plating out the bacterial suspensions at various time points and is presented as CFU (% of control) = (CFU peptide / CFU control ) * 100.

Fig. 2 .

 2 Fig. 2. Effect of peptide treatment on the morphology of bacteria (I). TEM images of E. coli WBB01. Bacteria were incubated alone (A, B), with 20 µM NK11 (D, E), and with 2 µM NK-2 (C, F-I). Length scale: each bar represents 1 µm. Arrows: (1) membrane blebs; (2) membrane ruffling; (3) membrane detachment; (4) electron dense dots; (5) release of cytoplasm; (6) electron dense fibres.

Fig. 3 .

 3 Fig. 3. Effect of peptide treatment on the morphology of bacteria (II). TEM images of P. mirabilis R45. Bacteria were incubated alone (A, B), with NK11 (D, E) and NK-2 (C, F-I) (each 20 µM). Length scale: each bar represents 1 µm. *, (C) bar represents 0.2 µm. Arrows: please refer to figure 2.

Fig. 4 .

 4 Fig. 4. Effect of peptide treatment on the morphology of bacteria (III). AFM images of (A-C) E. coli WBB01, and (D-F) P. mirabilis R45. Bacteria were incubated alone (A, D), with NK-2 (B: 2 µM, E: 20 µM) and NK11 (20 µM, C and F). Images were taken in air in AC (tapping) mode. Length (bar) and height (colour code) scales are indicated.

Fig. 5 .Fig. 6 .

 56 Fig. 5. FACS analysis of the binding of NBD-NK-2 to bacterial strains E. coli WBB01 and P. mirabilis R45. Dotplots of forward scattered light (FSC) against area of relative intensity of green light (FL-1 Area) for (A) E. coli WBB01 and (B) E. coli WBB01 incubated with 10 M NBD-NK-2. (C) The amount of NBD-NK-2 bound to both strains of bacteria is shown normalized to peptide bound to E. coli WBB01.

Fig. 7 .

 7 Fig. 7. CD spectra of NK-2 (black line) and NK11 (grey line) in the presence of a suspension of E. coli WBB01 LPS aggregates.

Fig. 8 .

 8 Fig. 8. Binding curves of NK-2 (black lines) and NK11 (grey lines) to bilayers of LPS from (A) E. coli WBB01 and (B) P. mirabilis R45. Peptides injections (start and end points are indicated by arrows) last 10 min (adsorption) followed by 10 min buffer rinse (desorption).

Fig. 9 .

 9 Fig. 9. Dose-dependent interaction of NK-2 (solid lines: black, 0.8 µM; grey 0.4 µM, light grey 0.2 µM) and NK11 (3 µM, dotted black line) with LPS membranes. Peptide intercalation into aggregates of LPS from (A) E. coli WBB01, and (B) P. mirabilis R45 in 20 mM HEPES, 150 mM NaCl, pH 7.0, and (C) P. mirabilis R45 in 20 mM HEPES, pH 7.0 was measured by by FRET spectroscopy at 37°C. Peptides were added at 50 s. An increase of the FRET-signal (I Donor /I Acceptor ) corresponds to a reduced FRET efficiency and indicates insertion of peptide into the membrane.

Fig. 10 .

 10 Fig. 10. Representation of a typical planar lipid bilayer experiment. NK-2 (0.5 µM) was added at time point 0 min to a symmetrical PL/PL membrane mimicking the bacterial cytoplasmic membrane. Time courses of applied voltages (lower panel), peptide-induced capacity changes (upper panel, dotted grey line), and current traces (upper panel, black line) are shown. An arrow indicates disruption of the membrane.

Fig. 11 .

 11 Fig. 11. Time courses of NK-2-induced (0.3 µM) current traces and applied voltages in asymmetric planar lipid bilayers mimicking the outer membrane of Gram-negative bacteria: (A) E. coli WBB01 LPS/PL, and (B) P. mirabilis R45 LPS/PL.

Fig. 12 .

 12 Fig. 12. Stable integration of the peptide NK-2 (1 µM) into an asymmetric E. coli WBB01 LPS/PL planar lipid bilayer. After setting a clamp voltage, defined conductivity levels were reached.

  Figure 2

Figure 4 Biochemical

 4 Figure 3

Figure 6 Biochemical

 6 Figure 6

2 C

 2 Figure 9

  Figure 11

Licenced copy. Copying is not permitted, except with prior permission and as allowed by law.© 2010 The Authors Journal compilation © 2010 Portland Press Limited

© 2010 The Authors Journal compilation © 2010 Portland Press Limited

Acknowledgements

We thank Rainer Bartels for peptide synthesis, Jan Demmer for AFM measurements, Christine Hamann for FRET spectroscopy, Heike Kühl for electron microscopy, and Kerstin Stephan for performing the antibacterial tests.

Funding

This study has been carried out with financial support from the Deutsche Forschungsgemeinschaft (SFB 617 ´Molecular mechanisms of epithelial defense´, project A17).

Table 1 -Antibacterial activity

Activity of peptides against Gram-negative LPS rough mutant bacterial strains. The minimal inhibitory concentrations (MIC) and the minimal bactericidal concentrations (MBC) are given as MIC (MBC) in µg/ml. The assay was performed in (1) 20 mM Hepes pH 7.0, or in (2) 20 mM Hepes, 150 mM NaCl pH 7.0, each supplemented with 10% LB medium.

Bacterial strain


Ana R Correia

Tao Wang

Elizabeth A Craig

Cláudio M Gomes
email: gomes@itqb.unl.pt

Iron binding activity in yeast frataxin entails a trade off with stability in the 1/1 acidic ridge region

Keywords: yeast frataxin, protein folding, metallochaperone, binding affinities, protein plasticity

Frataxin is a highly conserved mitochondrial protein whose deficiency in humans results in Friedreich's ataxia (FRDA), an autosomal recessive disorder characterized by progressive ataxia and cardiomyopathy. Although its cellular function is still not fully clear, the fact that frataxin plays a crucial role in Fe-S assembly on the scaffold protein Isu is well accepted. Here we report the characterisation of eight frataxin variants having alterations on two putative functional regions -the  acidic ridge and the conserved -sheet surface. We report that frataxin iron binding capacity is quite robust: even when five of the most conserved residues from the putative iron binding region are altered, at least 2 iron atoms per monomer can be bound, although with decreased affinity. Furthermore, we conclude that the acidic ridge is designed to favour function over stability. The negative charges have a functional role, but at the same time significantly impair frataxin's stability. Removing five of those charges results in a thermal stabilization of ~24ºC and reduces the inherent conformational plasticity. Alterations on the conserved -sheet residues have only a modest impact on the protein stability, highlighting the functional importance of residues 122-124.

Introduction

Reduced expression of frataxin, or in some cases, expression of frataxin variants, leads to the development of Friedreich's ataxia (FRDA), an autosomal recessive neurodegenerative and cardiodegenerative disease characterized by progressive ataxia and cardiomyopathy [START_REF] Pandolfo | Friedreich ataxia: the clinical picture[END_REF][START_REF] Becker | Frataxin: its role in iron metabolism and the pathogenesis of Friedreich's ataxia[END_REF][START_REF] Durr | Clinical and genetic aspects of spinocerebellar degeneration[END_REF]. In spite of the availability of multi cellular models for the disease [START_REF] Puccio | Multicellular models of Friedreich ataxia[END_REF][START_REF] Huang | Elucidation of the mechanism of mitochondrial iron loading in Friedreich's ataxia by analysis of a mouse mutant[END_REF], insights into possible therapeutic strategies [START_REF] Richardson | Friedreich's ataxia: iron chelators that target the mitochondrion as a therapeutic strategy?[END_REF][START_REF] Herman | Histone deacetylase inhibitors reverse gene silencing in Friedreich's ataxia[END_REF] and broader systematic clinical studies [START_REF] Schulz | Diagnosis and treatment of Friedreich ataxia: a European perspective[END_REF], the pathogenesis of Friedreich's ataxia and its relationship with frataxin remains to be fully understood. Frataxin is a highly conserved protein localized in the mitochondria whose function is associated with iron homeostasis. Frataxin deficiency, as recently shown using conditional knockout mice, down regulates molecules involved in mitochondrial iron utilization pathways (FeS cluster and heme biosynthesis, and iron storage) while cellular iron uptake and its availability for mitochondrial uptake are increased [START_REF] Huang | Elucidation of the mechanism of mitochondrial iron loading in Friedreich's ataxia by analysis of a mouse mutant[END_REF]. This illustrates the complex interplay between frataxin deficiency and iron homeostasis in the context of FRDA [START_REF] Huang | Elucidation of the mechanism of mitochondrial iron loading in Friedreich's ataxia by analysis of a mouse mutant[END_REF]. The human and yeast frataxin (Yfh1) orthologs have a high degree of amino acid (65%) and structural identity. The phenotype observed as a consequence of frataxin's deficiency is very similar in human and yeast cells [9,[START_REF] Rotig | Aconitase and mitochondrial iron-sulphur protein deficiency in Friedreich ataxia[END_REF]. Hence, the yeast model system has been frequently used to study frataxin's function, as well as the pathological events associated with FRDA.Although a detailed understanding of frataxin function is still missing, data suggests a role as an iron metallo-chaperone [START_REF] Gakh | Physical evidence that yeast frataxin is an iron storage protein[END_REF][START_REF] Schulz | Oxidative stress in patients with Friedreich ataxia[END_REF][START_REF] Thierbach | Targeted disruption of hepatic frataxin expression causes impaired mitochondrial function, decreased life span and tumor growth in mice[END_REF][START_REF] Vazquez-Manrique | Reduction of Caenorhabditis elegans frataxin increases sensitivity to oxidative stress, reduces lifespan, and causes lethality in a mitochondrial complex II mutant[END_REF], delivering iron to its binding partners, promoting heme and Fe-S clusters biosynthesis, as well as Fe-S cluster repair [START_REF] Bulteau | Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity[END_REF][START_REF] Yoon | Iron-sulfur cluster biosynthesis. Characterization of frataxin as an iron donor for assembly of [2Fe-2S] clusters in ISUtype proteins[END_REF][START_REF] Yoon | Frataxin-mediated iron delivery to ferrochelatase in the final step of heme biosynthesis[END_REF]. While initially oligomerization was believed to be dispensable in vivo recently, it has been shown that oligomerization impairment is harmful under oxidative stress conditions and it leads to a decrease of yeast chronological lifespan [START_REF] Gakh | Physical evidence that yeast frataxin is an iron storage protein[END_REF][START_REF] Adamec | Iron-dependent self-assembly of recombinant yeast frataxin: implications for Friedreich ataxia[END_REF]19]. Yfh1 is not essential, but its absence causes severe growth defects and a reduction in the activity of Fe-S cluster-containing enzymes, such as aconitase [9,[START_REF] Rotig | Aconitase and mitochondrial iron-sulphur protein deficiency in Friedreich ataxia[END_REF]19,[START_REF] Foury | Acidic residues of yeast frataxin have an essential role in Fe-S cluster assembly[END_REF]. Iron binding by frataxins, a requirement for a role as a metallochaperone, has been well documented by several laboratories [START_REF] Yoon | Iron-sulfur cluster biosynthesis. Characterization of frataxin as an iron donor for assembly of [2Fe-2S] clusters in ISUtype proteins[END_REF][START_REF] Cook | Monomeric yeast frataxin is an iron-binding protein[END_REF][START_REF] Kondapalli | Drosophila frataxin: an iron chaperone during cellular Fe-S cluster bioassembly[END_REF]. Frataxins bind iron at micromolar binding affinities, with different stoichiometries: while monomeric human frataxin binds 6/7 iron ions (K d ~ 12-55M [START_REF] Yoon | Iron-sulfur cluster biosynthesis. Characterization of frataxin as an iron donor for assembly of [2Fe-2S] clusters in ISUtype proteins[END_REF]), monomeric frataxin from Drosphila binds 1 ferrous iron (K d ~6M, [START_REF] Kondapalli | Drosophila frataxin: an iron chaperone during cellular Fe-S cluster bioassembly[END_REF]) and yeast frataxin binds 2 ferrous irons (K d ~2.5M, [START_REF] Cook | Monomeric yeast frataxin is an iron-binding protein[END_REF]). This variability suggests a certain plasticity of the frataxin fold in respect to binding iron. In fact, an NMR based investigation of the iron binding to Yfh1 at low stoichiometry (up to 2Fe:Yfh1) denoted changes in 10 distinct residues [START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF], which could indicate long range effects upon iron binding, or lower affinity sites. More complex effects arise at higher Fe:Yfh1 stoichiometries. Like human frataxin, Yfh1 undergoes iron-dependent oligomerization, above a threshold of 2 irons per monomer, according to the progression    3  6   12   24  48 [START_REF] Gakh | Physical evidence that yeast frataxin is an iron storage protein[END_REF][START_REF] Adamec | Iron-dependent self-assembly of recombinant yeast frataxin: implications for Friedreich ataxia[END_REF]. This property has been established as important for ironinduced oxidative stress protection, but not for Fe-S cluster biogenesis [19,24]. The conserved frataxin fold consists of an  sandwich motif, with two -helices (Nand C-terminal) that construct the helical plane and five antiparallel -strands that form the -sheet plane. A sixth (seventh in the human protein) -strand intersects the helical and -sheet planes [START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF][START_REF] Cho | Crystal structure of Escherichia coli CyaY protein reveals a previously unidentified fold for the evolutionarily conserved frataxin family[END_REF][START_REF] Dhe-Paganon | Crystal structure of human frataxin[END_REF][START_REF] Musco | Assignment of the 1H, 15N, and 13C resonances of the C-terminal domain of frataxin, the protein responsible for Friedreich ataxia[END_REF][START_REF] Nair | Solution structure of the bacterial frataxin ortholog, CyaY: mapping the iron binding sites[END_REF]. The first helix (1) and the edge of the first -strand (1) form a semi-conserved acidic ridge that constitutes the putative iron binding region of the protein [START_REF] Cook | Monomeric yeast frataxin is an iron-binding protein[END_REF][START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF][START_REF] Nair | Solution structure of the bacterial frataxin ortholog, CyaY: mapping the iron binding sites[END_REF]. A set of conserved residues in the -sheet surface constitutes the A c c e p t e d M a n u s c r i p t other putative functional region of the protein, believed to be involved in the binding of frataxin to its protein partners [START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF][START_REF] Musco | Towards a structural understanding of Friedreich's ataxia: the solution structure of frataxin[END_REF]. Recently we have shown that mutations at the conserved -sheet residues 122-124 affect neither iron binding capacity nor the oligomerization properties of frataxin, although this mutation does compromise the interaction with Isu [START_REF] Wang | Binding of yeast frataxin to the scaffold for Fe-S cluster biogenesis[END_REF]. To complement previous functional characterization of YFH1 mutants, here we address the structural and conformational effects of eight functional mutants in yeast frataxin that alter either the acid ridge or conserved residues on the -sheet surface. We found that the negatively charged residues in the acid ridge that are important for iron binding reduce Yfh1's stability, indicating a trade-off between functionality and stability. In addition, alterations in the conserved residues of the portion of the acid ridge found in the -sheet, affected interaction with the scaffold protein Isu, but did not affect overall protein stability.

Experimental

Yeast strains, plasmids and media. Saccharomyces cerevisiae strains carrying YFH1 mutant or wild type genes on plasmids were created by sporulation and dissection of YFH1/yfh1 (W303 background) transformed with the plasmids. YFH1 mutants were generated by site-directed mutagenesis using pCM189-YFH1 as a template, having YFH1 under the control of the tetO promoter [START_REF] Gari | A set of vectors with a tetracycline-regulatable promoter system for modulated gene expression in Saccharomyces cerevisiae[END_REF]. Constructs containing the following mutations were prepared: yfh1-D86A/E90A/E93A, yfh1-D101A/E103A, yfh1-D86A/E90A/E93A/D101A/E103A, yfh1-N122A, yfh1-N122K (corresponding to the human clinical mutation N146K), yfh1-K123T, yfh1-Q124A and yfh1-N122A/K123T/Q124A. Doxycycline was used at 1 g/ml to reduce expression. Cells were grown in minimal synthetic media with -ura DO at 30ºC for 2 days.

Aconitase activity in isolated mitochondria. Mitochondria were isolated, according to the method described in [START_REF] Li | Yeast mitochondrial protein, Nfs1p, coordinately regulates iron-sulfur cluster proteins, cellular iron uptake, and iron distribution[END_REF], from cells grown in synthetic minimal media (-ura), in the absence of doxycycline. Aconitase activity was measured in isolated mitochondria by monitoring the decrease in the absorbance of the substrate isocitrate at 240 nm as described in [START_REF] Murakami | 70-kD heat shock-related protein is one of at least two distinct cytosolic factors stimulating protein import into mitochondria[END_REF].

Gene Expression and Protein purification.

The mature form of Yfh1 (amino acids 52-174) with an N-terminal His tag and a thrombin cleaving site in between was cloned into pET-3a vector (Novagen, Madison, WI, USA) [19]. Protein expression was induced over 3h at 37ºC in BL21(DE3) E. coli by adding IPTG (isopropyl-b-galactopyranoside) at a final concentration of 0.5mM. With the exception of Yfh1-K123T, which was expressed at much lower levels (approximately 30%), all other mutant variants had expression levels identical to those of the wild type Yfh1. The reduced expression levels of Yfh1-K123T had already been described when expressing this mutant in yeast [START_REF] Wang | Binding of yeast frataxin to the scaffold for Fe-S cluster biogenesis[END_REF]. Under the tested conditions, wild type Yfh1 was exclusively expressed as a soluble protein, as were most of the mutants. The only exceptions are Yfh1-86/90/93A, Yfh1-N122A and Yfh1-N122A/K123/Q124A, in which ~15% of the expressed protein was found in the insoluble fraction. After harvesting the cells they were lysed on a French press. Cell lysate was subjected to His-binding resin (Amersham) chromatography and the protein was eluted A c c e p t e d M a n u s c r i p t with 500mM Imidazole in the binding buffer (50mM Tris-HCl Ph8.0, 200mM NaCl, 10mM Imidazole). The His tag was cleaved using biotinylated thrombin and thrombin was removed by streptavidin agarose (Amersham). When necessary (purity less then 90%) the protein was further purified by applying the sample on a Superdex 75. At the end the buffer was changed to 10mM HEPES, 50mM NaCl pH 7.0 using Centricons (Millipore). Protein concentration was determined using the extinction coefficient  280nm = 15470 M -1 .cm -1 . The mature form of Isu1(amino acids 35-165) with an His tag and a TEV cleaving site in between was cloned into pKLD37 vector. Protein expression was induced over 4h at 30ºC in C41 E. coli by adding IPTG (isopropyl--galactopyranoside) at a final concentration of 1mM the purification was performed as above described for the Yfh1 variants. Protein concentration was determined using the extinction coefficient

 280nm = 14161.4 M -1 .cm -1
Spectroscopic methods. UV/Vis Spectra were recorded at room temperature in a Shimadzu UVPC-1601 spectrometer. Fluorescence spectroscopy was performed on a Cary Varian Eclipse instrument ( ex =280nm,  em =340nm, slit ex : 5 nm, slit em : 10 nm unless otherwise noted) equipped with cell stirring and Peltier temperature control. Far-UV CD spectra were recorded typically at 0.2 nm resolution on a Jasco J-715 spectropolarimeter fitted with a cell holder thermostated with a peltier. Thermal denaturation. Thermal unfolding was followed by ellipticity (  mrw at 222nm) variations. In all experiments a heating rate of 1ºC/min was used, and the temperature was changed from 10 to 90ºC. All reactions were found to be reversible under the tested conditions, as inferred from obtaining identical far-UV CD spectra of the proteins before the thermal ramping and after cooling down the heated sample; a second temperature ramp of the same sample also yielded identical melting transitions. Protein aggregation was not observed during thermal unfolding. Data were analysed according to a two-state model and the thermodynamic data parameters were determined [START_REF] Pace | Conformational stability and thermodynamics of folding of ribonucleases Sa, Sa2 and Sa3[END_REF].

Trypsin limited proteolysis and LC-MS analysis.

Frataxins were incubated with trypsin (bovine pancreas trypsin, sequencing grade, Sigma) at 37°C in 0.1 M Tris-HCl pH 8.5, in a 150 fold excess over the protease. Aliquots (300 mol of protein) were sampled at different incubation periods and the reaction stopped by adding loading buffer with 5M BSA followed by a 10 min incubation at 100ºC. The products of the proteolysis reaction were analysed by SDS-PAGE [START_REF] Correia | Dynamics, stability and iron-binding activity of frataxin clinical mutants[END_REF].

Iron-binding assays. Yeast frataxin oligomerizes in an iron-concentration dependent fashion, but at up to a stoichiometry of 2 irons per protein the protein remains in the monomer form [START_REF] Gakh | Physical evidence that yeast frataxin is an iron storage protein[END_REF][START_REF] Adamec | Iron-dependent self-assembly of recombinant yeast frataxin: implications for Friedreich ataxia[END_REF][START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF][START_REF] Bencze | The structure and function of frataxin[END_REF]. To avoid side effects from different oligomerization behaviours we have measured iron binding before oligomerization takes place, i.e. up to a iron:protein ratio of 2. Trp fluorescence emission spectroscopy was used to monitor iron binding, which quenches Trp emission as the iron-frataxin complex is being formed. As previously shown, this methods yield results identical to those obtained by isothermal titration calorimetry [START_REF] Yoon | Iron-sulfur cluster biosynthesis. Characterization of frataxin as an iron donor for assembly of [2Fe-2S] clusters in ISUtype proteins[END_REF][START_REF] Correia | Conformational stability of human frataxin and effect of Friedreich's ataxia-related mutations on protein folding[END_REF]. For the purpose, the quenching of the Trp fluorescence ( exc =290nm;  em =340 nm) of a 50M solution of apo frataxin in 100mM HEPES pH 7.0, 50mM NaCl was measured upon stepwise addition of an iron citrate solution, in 1 ml [START_REF] Correia | Dynamics, stability and iron-binding activity of frataxin clinical mutants[END_REF][START_REF] Correia | Conformational stability of human frataxin and effect of Friedreich's ataxia-related mutations on protein folding[END_REF]. This data was used to calculate the fraction of binding sites occupied. The stoichiometry, p, and apparent dissociation constant, K d , were then determined as described by Winzor and Sawyer [START_REF] Winzor | Quantitative characterisation of ligand binding[END_REF].

Isu:Yfh1 interaction assay. Holo-Yfh1 was prepared using a stoichiometry of 2 irons per frataxin to avoid interference from oligomer formation. The formation of a Isu:holofrataxin complex was followed monitoring the Trp fluorescence, taking advantage of the fact that yeast Isu does not contain any Trp residue. Thus, when Isu binds to holo-Yfh1, the Trp fluorescence emission ( exc =290nm;  em =340 nm) of the latter is quenched and this can be used to monitor binding. Holo-Yfh1 was prepared in 100 mM HEPES pH 7.0, 50mM NaCl. Aliquots of Isu were added to 2.5 mM holo frataxins under constant stirring. The stoichiometry, p, and apparent dissociation constant, K d , were then obtained as described by Winzor and Sawyer [START_REF] Winzor | Quantitative characterisation of ligand binding[END_REF].

Results and Discussion

Alterations in the conserved -sheet surface result in functional effects

To gain a better understanding of the relationship between the structure and function of frataxin, we compared four Yfh1 variants. Three (Yfh1-D101/E103A, Yfh1-D86/E90/E93A and a combination of these two, Yfh1-86/90/93/101/103A) alter the putative iron binding region at the 1/1 acidic ridge and oligomerization region (Fig. 1). The fourth, Yfh1-N122A/K123T/Q124A (hereafter referred to as Yfh1-122-4), alters the conserved -sheet, the putative binding surface for interaction with Isu, the scaffold on which Fe-S clusters are built [START_REF] Wang | Binding of yeast frataxin to the scaffold for Fe-S cluster biogenesis[END_REF]. Since Yfh1 cellular levels can be significantly reduced before any growth defects are detected [39], wild type and mutant YFH1 genes were placed under the control of the tetO promoter allowing the modulation of protein expression using doxycycline. In this way, in vivo function could be tested under both normal and reduced levels of expression. At normal Yfh1 expression levels (minus doxycycline) all mutant gene were able to support growth as well as the wild type gene (Fig. 2). In addition, as we previously reported [19, 30], Yfh1-122-4 cells grew poorly when the variant was expressed at reduced levels (plus doxycycline), while cells expressing low levels of Yfh1-86/90/93A grew as well as wild type cells. However, Yfh1-101/103A was slightly compromised, and when these mutations were combined with those of Yfh1-86/90/93A, generating alterations in five of the charged residues in the acidic ridge, growth was severely compromised. Consistent with this enhanced growth phenotype, mitochondria isolated from Yfh1-101/103A had lower levels of activity of the Fe-S enzyme aconitase, even when the mutant protein was expressed at normal levels (Fig. 3). This affect was more extreme in Yfh1-86/90/93/101/103A mitochondria, which had only 40% the aconitase activity of wild type mitochondria. Consistent with the affects on growth, Yfh1-122-4 mitochondria had only 20% the aconitase activity quantified for wild type mitochondria.

None of the mutations abolish iron binding

Since iron binding is an essential functional feature of frataxin, we asked if iron-binding properties of the Yfh1 variants were affected. In order to exclude iron-induced frataxin oligomerization effects, we have investigated iron-binding at low stoichiometry (up to 2

Charge-to-neutral alterations in the  acidic ridge increase stability

In order to evaluate if the functional impairment may result from decreased protein stability, the effect of the alterations on Yfh1 folding thermodynamics was analyzed by comparing the thermal stability of mutant variants to that of wild type. According to the analysis of the far-UV CD spectra at 20ºC, before and after thermal denaturation, thermal unfolding was reversible for all protein variants studied and no aggregation was observed after thermal unfolding. The results showed that charge-to-neutral alterations in the acidic ridge result in an impressive stabilization of the protein fold: an increase of up to ~24ºC was noted for the Yfh1-86/90/93/101/103A variant, while alterations in the sheet surface had almost no effect on protein thermal stability (Fig. 4, Table 1). The protein stability decreased in the order: Yfh1-86/90/93/101/103 A > Yfh1-86/90/93 A > Yfh1-101/103 A > Yfh1-N122K > Yfh1-K123T > Wild Type > Yfh1-N122A/K123T/Q124A > Yfh1-Q124A > Yfh1-N122A. The exception for the effect on the sheet surface are the mutations in residue Asn-122: changing to an alanine (T m = -4.5 ºC) or to a lysine (T m = +4.3 ºC) had opposite effects, probably due to the effect these alterations would be expected to have on the hairpin between strands 3 and 4, which involves two hydrogen bonds (Asn122-Trp131 and Val120-A133). While the insertion of an Ala likely disrupts the hydrogen bond with Trp131, the positively charged Lys might strengthen it, stabilizing the protein. The 101/103A alterations stabilize the protein in spite of compromising 2 of the 3 hydrogen bonds involved in the hairpin connecting strands1 and 2. This suggests that minimizing repulsive interactions overcomes the stabilisation obtained by the two hydrogen bonds. Since protein conformational plasticity affects both protein function and degradation rates. We next analyzed whether the functional mutations were also affecting frataxin flexibility, by performing limited proteolysis experiments using trypsin. Our underlying rationale was that mutations resulting in an increased structural flexibility would increase trypsin access to cleavage sites and consequently increase the degradation rate. The results show two distinct patterns, depending on the region in which the alteration is located (Fig. 5). Alterations on the acidic ridge had a pronounced effect on frataxin dynamics, making the protein more rigid and substantially less susceptible to proteolysis. In fact, the Yfh1-86/89/93/101/103A variant remains essentially intact under conditions in which wild type frataxin is essentially completely digested (80% versus 10% integrity after 100 min digestion). On the other hand, alterations on the -sheet surface (Yfh1-N122K/K123T/Q124A) behave almost identically to the wild type, suggesting that modifications in this region have either a very small or no effect on the protein conformational plasticity. Overall, alterations in the acidic ridge th

Conclusions

Here we present a detailed characterization of eight yeast frataxin functional variants, that either alter the acidic ridge between -helix 1 and -sheet 1, or the conserved -sheet surface between strands 3 and 4. Changing the conserved -sheet residues Asn122-Lys-123-Gln124 had almost no effect on Yfh1 stability and plasticity, indicating that changes in this region did not disrupt overall conformation, but are relevant for the Yfh1-Isu interaction. Alteration of up to five residues in the acidic ridge region, four of which had been identified as iron binding sites [START_REF] He | Yeast frataxin solution structure, iron binding, and ferrochelatase interaction[END_REF], significantly increase frataxin stability. Thus illustrates a rather interesting trade off between activity and stability in this region. In addition, our study suggests that residues Asp101 and Glu103 are involved in the ironmediated interaction between Isu and Yfh1, but their alteration does not abrogate the interaction, as evidenced by the rescue of the yfh1 phenotype. [START_REF] Pace | Conformational stability and thermodynamics of folding of ribonucleases Sa, Sa2 and Sa3[END_REF]; for parameters see Table 2.

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Figure Legends Figure 1 :

 Legends1 Figure Legends

Figure 2 :

 2 Figure2: Yeast growth rescue of (yfh1) cells by Yfh1 variants. A 10-fold dilutions of cell suspension of wild type , yfh1 and yfh1 transformed with plasmids harbouring the indicated YFH1 mutant genes under the control of the tet-O-regulatable wild type or mutant yfh1 were plated on minimal synthetic medium -ura DO containing (+) or lacking (-) doxycycline (dox). Plates were incubated at 30ºC for 2 days.

Figure 3 :

 3 Figure 3: Aconitase activity. Extracts from mitochondria isolated from the indicated strains were prepared (150g) and aconitase activity was measured. Data is presented as percentage of wild type aconitase activity (n=3).

Figure 4 :

 4 Figure 4: Thermal denaturation curves at pH 7.0 following Trp emission. Impact of mutations on Yfh1 thermal stability. (■) Wild Type , (a) single point mutations of -sheet surface residues (122-123). (□) N122A, (∆) K123T, (○)Q124A, (b) Mutations on putative iron binding sites: (▲) D86A E90A E93A, (◊) 86/90/93/101/103 A. Lines represent fits to the two-state model[START_REF] Pace | Conformational stability and thermodynamics of folding of ribonucleases Sa, Sa2 and Sa3[END_REF]; for parameters see Table2.

Figure 5 :

 5 Figure 5: Time course of trypsin limited proteolysis. Comparison between wild type and functional mutants (A) SDS-PAGE analysis of the time course trypsin limited proteolysis experiments. (B) Evaluation/quantification of frataxin degradation during incubation with trypsin. Gels (A) densitometric analysis allowed the quantification of frataxin for the different incubation times with trypsin. (■) Wild Type, (□) N122A K123T Q124A, (▲) D86A E90A E93A, (◊) 86/90/93/101/103 A.

  Publication. Published on 14 Dec 2009 as manuscript BJ20091612 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091612 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited Publication. Published on 14 Dec 2009 as manuscript BJ20091612 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091612 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited Publication. Published on 14 Dec 2009 as manuscript BJ20091612 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091612 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited Publication. Published on 14 Dec 2009 as manuscript BJ20091612 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091612 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited Publication. Published on 14 Dec 2009 as manuscript BJ20091612 THIS IS NOT THE VERSION OF RECORD -see doi:10.1042/BJ20091612 A c c e p t e d M a n u s c r i p t Licenced copy. Copying is not permitted, except with prior permission and as allowed by law. © 2009 The Authors Journal compilation © 2009 Portland Press Limited

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Acknowledgements

This work is partly financed by grants from the Fundação para a Ciência e Tecnologia (PTDC/QUI/70101) and National Ataxia Foundation (to CMG) and grants from the National Institutes of Health Grant (GM27870) and Muscular Dystrophy Association (to EAC.). A.R.C and T.W were recipients of FCT PhD Fellowship (SFRHBD/24949/2005) and an American Heart Association Postdoctoral Fellowship 0525728Z, respectively.