The role of electrostatic interactions in the antitumor activity of dimeric RNases ´ ˜ Eugenio Notomista1, Jose Miguel Mancheno2, Orlando Crescenzi3, Alberto Di Donato1, ´ Jose Gavilanes4 and Giuseppe D’Alessio1 ` Dipartimento di Biologia Strutturale e Funzionale, Universita di Napoli Federico II, Napoli, Italy ´ ´ Grupo de Cristalografıa Macromolecular y Biologıa Estructural, Instituto Rocasolano, Madrid, Spain ` Dipartimento di Chimica, Universita di Napoli Federico II, Napoli, Italy ´ ´ Departamento de Bioquımica y Biologıa Molecular I, Universidad Complutense, Madrid, Spain Keywords antitumor RNases; electrostatic interactions; electrostatic interaction energy; RNases; transport through membranes Correspondence G D’Alessio, Dipartimento di Biologia ` Strutturale e Funzionale, Universita di Napoli Federico II, Via Cinthia, I-80126 Napoli, Italy Fax: +39 081 679159 Tel: +39 081 679157 E-mail: dalessio@unina.it (Received 12 April 2006, revised 31 May 2006, accepted 12 June 2006) The cytotoxic action of some ribonucleases homologous to bovine pancreatic RNase A, the superfamily prototype, has interested and intrigued investigators Their ribonucleolytic activity is essential for their cytotoxic action, and their target RNA is in the cytosol It has been proposed that the cytosolic RNase inhibitor (cRI) plays a major role in determining the ability of an RNase to be cytotoxic However, to interact with cRI RNases must reach the cytosol, and cross intracellular membranes To investigate the interactions of cytotoxic RNases with membranes, cytotoxic dimeric RNases resistant, or considered to be resistant to cRI, were assayed for their effects on negatively charged membranes Furthermore, we analyzed the electrostatic interaction energy of the RNases complexed in silico with a model membrane The results of this study suggest that close correlations can be recognized between the cytotoxic action of a dimeric RNase and its ability to complex and destabilize negatively charged membranes doi:10.1111/j.1742-4658.2006.05373.x The superfamily of pancreatic-type RNases [1] includes several members capable of carrying out ‘special’ actions, i.e., actions other than catalytic, although strictly dependent on their catalytic, RNA degrading action [2] These actions could be linked to physiological functions, as in the case of the angiogenic action of angiogenins [3], or due to the mere reflection in the laboratory assays, mirrors proposed in the experiment, of unknown functions, as may be the case of the antifertility action of seminal RNase [4] Particular attention has been given to the cytotoxic action of some RNases, especially because they often appear to be selective for tumor cells [5] Many studies have been devoted to the mechanism of action of these antitumor RNases However, a con- clusive understanding of why these RNases kill cells, especially why some of them selectively kill tumor cells, has not been obtained The correlation has been stressed [6] between the ability of certain RNases to display a cytotoxic action and their ability to evade the strong, neutralizing action of the cytosolic RNase inhibitor (cRI), a 50-kDa protein containing 16 leucine-rich repeat motifs [7,8] In fact, onconase from Rana pipiens eggs [9], and seminal RNase from Bos taurus seminal vesicles [10], among the most studied natural cytotoxic RNases, not bind cRI, and are both totally resistant to its inhibitory action [8] On the other hand, RNases with no cytotoxic action, such as bovine or human pancreatic RNase, and a very high affinity for cRI, acquire the ability to kill Abbreviations ANTS, 1,3,6-trisulphonate-8-aminonaphtalene; cRI, cytosolic RNase inhibitor; DMPG, dimyristoylphosphatidylglycerol; DMPS, dimyristoylphosphatidylserine; DPH, 1,6-diphenyl-1,3,5-hexatriene; DPX, p-xylenebispyridinium bromide; ECP, eosinophil cationic protein; EDN, eosinophil-derived neurotoxin; EIE, electrostatic interaction energy; HP-RNase, human pancreatic RNase; kT, product of the Boltzmann constant by absolute temperature in K; PG, phosphatidylglycerol; RET, resonance energy transfer; SVT2, malignant murine fibroblasts FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS 3687 Electrostatic interactions and antitumor RNases E Notomista et al cells when they are engineered into cRI-resistant RNases [6,11–14] These results led to the description of the cytosolic RNase inhibitor as a sentry to guard cells from, and inactivate, potentially cytotoxic foreign RNases [6] Recently however, it has been reported [15] that HeLa cells deprived through RNA silencing of any cRI in their cytosol, are still not affected by noncytotoxic RNases This indicates that noncytotoxic, cRI-sensitive RNases not become cytotoxic when cRI, the postulated ‘sentry’, is absent Furthermore, artificial dimers of RNase A, obtained through lyophilization from acetic acid, have been found to bind with high affinity the cytosolic RNase inhibitor [16], but also to be cytotoxic on malignant cell lines [17] These results have cast shadows on the prospect that the antitumor action of RNases be based solely, or primarily, on its resistance to the cytosolic RNase inhibitor RNase catalytic activity is an absolute requirement for the cytotoxic action of all cytotoxic RNases tested [5] Moreover, when the effects of cytotoxic RNases on cellular RNA were studied, cytosolic rRNA [18] or tRNA [19] were found to be the targets of cytotoxic RNases Thus, we can conclude that to exert its cytotoxic action, an RNase must reach the cytosol Furthermore, given that cRI is a cytosolic protein [6,7], the characterization of an RNase as a cRI-resistant or -sensitive RNase can only occur if the RNase reaches the cytosol These considerations lead to the conclusion that to approach the cytosol, where their cytotoxic action is exerted, cytotoxic RNases must cross not only the plasma membrane, but intracellular membranes as well A study of the intracellular journey of cytotoxic, dimeric seminal RNase has revealed that it is internalized by various types of malignant cells, and has access to endosomes, whereas noncytotoxic bovine pancreatic RNase A is not internalized [18,20] Interestingly, when RNase A was made dimeric and cytotoxic by site-directed mutagenesis, it gained access to endosomes [20] It should be added that endocytosed sem- inal RNase is found in endosomes both in malignant and normal cells, but only in malignant cells does the RNase reaches the trans-Golgi network, and then the cytosol, where it degrades rRNA [18,20] These data suggest that cell membranes can discriminate RNases, and may play an important role in determining the cytotoxicity of an RNase by simply allowing, or not, an RNase to pass from one cell compartment to another On the other hand, the RNase itself is expected to present to the membrane which it should permeate structural elements that determine recognition and favor its access through that membrane For a systematic study of the interactions between membranes and cytotoxic RNases, we report here the effects on the stability of artificial membranes of natural and engineered dimeric RNases with different degrees of cytotoxic activity Their electrostatic interactions with a model membrane are then compared through in silico analyses The results of these investigations indicate that close correlations can be recognized between the ability of an RNase to destabilize a membrane, its attraction to a negatively charged membrane, and its cytotoxic action Results The RNases The RNases investigated in this study are described in Table They are cationic proteins with cytotoxic activity: BS-RNase [10] is a natural dimeric RNase; RNaseAA [21], RNase-AA-G [21], and RNase-AA-GG [22] are dimeric variants engineered from bovine pancreatic RNase A; HHP-RNase [11] is a dimeric variant engineered from human pancreatic RNase (HP-RNase) Effects on vesicle aggregation All RNases investigated are cationic proteins, with high isoelectric point values Thus we investigated the effects Table Natural and engineered RNases investigated in this study IC50 is the RNase concentration producing half-maximal cytotoxicity on SVT2 malignant cells [5,11,14] Abbreviations Natural or modified RNase Structure pI IC50 (lgỈmL)1) RNase A RNase-AA RNase-AA-G RNase-AA-GG HP-RNase HHP-RNase BS-RNase Bovine pancreatic RNase A A19P,Q28L,K31C,S32C-RNase A D38G-RNase-AA D38G,E111G-RNase-AA Human pancreatic RNase Q28L,R31C,R32C,N34K-HP-RNase Bovine seminal RNase Monomeric Dimeric Dimeric Dimeric Monomeric Dimeric Dimeric 9.8 9.6 9.8 9.9 10.1 9.9 10.3 > 200 82 64 27 > 200 26 22 3688 FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS E Notomista et al of these RNases on the aggregation of negatively charged dimyristoylphosphatidylglycerol (DMPG) and dimyristoylphosphatidylserine (DMPS) vesicles Aggregation was measured from the increase of absorbance at 360 nm due to the enhanced turbidity as engendered by vesicle aggregation The results with DMPG vesicles are presented here, for the higher availability to outside interactions of the negatively charged glycerol-linked phosphate [23–25] In DMPS vesicles, the bonding of the serine carboxylate ion to the adjacent ammonium group renders the phosphate negative charge less available [23–25] In fact, when DMPS vesicles were tested (data not shown), similar results were obtained in terms of effects exerted by the RNases under investigation, only to some extent less powerful As previously reported [26], seminal RNase (BSRNase), a naturally dimeric RNase, exerts a very strong effect on the aggregation of DMPG vesicles, whereas monomeric RNases, namely RNase A and a monomeric derivative of BS-RNase, exert no significant effects However, the results illustrated in Fig indicate that the quaternary structure of an RNase is not a prerequisite for its ability to aggregate vesicles, as RNase A does not acquire this ability upon dimerization into RNase-AA The replacement in dimeric RNase-AA of one negatively charged residue (Asp38) with an uncharged Gly residue generates an RNase variant (RNase-AA-G) capable of aggregating vesicles The result is even more striking when two negatively charged residues (Asp38 and Glu111) are replaced with Gly residues, as in RNase-AA-GG As shown in Fig 1, the effects on Fig Aggregation of DMPG vesicles induced by RNases Symbols: (e) BS-RNase; (+) RNase AA-GG; (n) HHP-RNase; (n) RNase AA-G; (h) RNase AA; (m) HP-RNase; (·) RNase A Electrostatic interactions and antitumor RNases DMPG vesicles of RNase-AA-GG are similar to those obtained with BS-RNase, the RNase with the most powerful aggregating effect When DMPS vesicles were used, similar effects were observed, but were less evident (data not shown) It should be added that CD spectra of the three RNase A dimers (RNase AA, -AA-G, and -AA-GG) in the far-UV and near-UV regions were virtually identical, with a slightly different spectrum for RNaseAA-G in the near-UV region Thermal denaturation, measured through the variation of ellipticity at 218 nm, gave an identical midpoint of denaturation at 62 °C for all three RNase dimers, and identical far-UV CD spectra of the thermally denatured states These data (not shown) indicate that the three proteins apparently possess an identical folded structure under the conditions used in the experiments with membranes, hence their different effects on the model membranes may not be ascribed to different conformations of the three proteins Effects on bilayer fluidity Figure illustrates the effects of RNases on the phase transition profile of DMPG vesicles labeled with 1,6diphenyl-1,3,5-hexatriene (DPH), a fluorescent probe used for measuring fluorescence polarization Bilayer fluidity was measured by comparing the difference in fluorescence anisotropy (Dr) as a function of protein : DMPG ratio Fig Effect of RNases on the thermotropic behavior of DMPG vesicles Symbols: (e) BS-RNase; (+) RNase AA-GG; (n) HHP-RNase; (n) HP-RNase; (h) RNase AA-G; (m) RNase AA; (·) RNase A FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS 3689 Electrostatic interactions and antitumor RNases E Notomista et al The data show that most investigated RNases affect the bilayer fluidity of DMPG membranes, as evidenced by a decrease of the amplitude of their thermotropic transition Only RNase A and RNase AA show no effects on the thermotropic behavior of the tested membranes Surprisingly, also monomeric HP-RNase, with no effect on membrane aggregation (see above), was found to affect to a certain extent the bilayer fluidity (Fig 2) Similar, albeit minor effects, were observed when the experiments were carried out on DMPS vesicles (data not shown) Effects on membrane fusion This parameter was evaluated by determining the intermixing of DMPG phospholipids between unlabeled vesicles and vesicles labeled with a donor ⁄ acceptor pair of fluorescent probes When the RNase under test promoted intermixing, a decrease of the energy transfer between the two probes was measured as resonance energy transfer (RET) Under the test conditions (see below), a RET value of 80% indicated that the RNase under test did not promote any perturbations of the vesicles bilayer Figure shows that all the RNases that promoted membrane aggregation also engendered membrane fusion Even native, monomeric HP-RNase, with no effects on membrane aggregation, produced a modest intermixing RNase-AA-IG instead, which induced a low, but significant degree of membrane aggregation, did not show any effects on vesicle intermixing No effects of membrane fusion were observed with RNase A and RNase-AA, or when DMPS vesicles were tested Correlations between the effects of RNases on membranes and their electrostatic charges In Table the results described above on the effects on membrane stability induced by the investigated RNases are normalized to those obtained with BSRNase, taken as 100% For the normalization the highest values of Dr, DA360 and percentage RET were used for effects on bilayer fluidity, membrane aggregation and fusion, respectively (see above and Figs 1–3) When the effects of RNases on negatively charged membranes were compared with the net charge values of the RNases, the comparison suggested (Table 2) that the ability of RNases to destabilize lipid vesicles is due to an electrostatic component, as the higher positive net charge of an RNase appears to enhance its ability to aggregate, fuse, or affect bilayer fluidity of the negatively charged membranes However, a hydrophobic component may not be excluded in the RNase– membrane destabilizing interactions This can be deduced: (a) from the RNases aggregating effects, which could be generated by the relief of hydration repulsion among vesicles occurring upon adsorption of the RNases to the membrane bilayer; (b) from the thermotropic behavior of lipids in the presence of RNases, which can be assigned to lipid molecules removed from participating in their gel-to-liquid phase transition To further investigate this aspect, values of electrostatic interaction energy (EIE) were calculated for the interactions between the dimeric RNases and a model membrane constructed with DMPG lipids (see below) A series of EIE values were determined by varying the orientation of each RNase towards the membrane (Fig 4) The results of these analyses revealed that a single highest negative value of EIE (EIEmax) could be estimated for each dimeric RNase This value was found for all RNases at the unique orientation with Table Effects on membrane vesicles and properties of the dimeric RNases investigated in this study RNase Fig Lipid mixing of DMPG vesicles induced by RNases Symbols: (e) BS-RNase; (+) RNase AA-GG; (n) HHP-RNase; (n) HP-RNase; (h) RNase AA-G; (m) RNase AA; (·) RNase A 3690 Aggregation Fluidity Fusion Positive EIEmax (%) alteration (%) (%) net charge (kT) BS-RNase HHP-RNase RNase AA-GG RNase AA-G RNase AA 100 51 71 28 12 100 103 109 58 100 100 100 0 18 10 10 )63.8 )67.3 )44.3 )27.3 )14.0 FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS E Notomista et al Fig Scheme of the initial complex dimeric RNase–model membrane used to generate the set of orientations for DELPHI calculations through rotation of the RNase around its major axis (rotation angle ‘r’) and variation of the inclination (inclination angle ‘i’) with respect to the plane of the membrane (parallel to the xy plane) Cter and Nter are the C-terminal and N-terminal structural regions, respectively, of the two A and B monomers of the dimeric RNase The black circle between the monomers indicates the mass center of the dimer Electrostatic interactions and antitumor RNases r ¼ 180° and i ¼ 90° In this orientation, characterized by the strongest electrostatic attraction of the RNases with the membrane, all RNases pointed their N-terminal regions towards the membrane Figure illustrates the results for some of the RNases (also supplementary Fig S1) It should be added that both models of RNase A or HP-RNase dimers, either swapping their N-terminal ends between subunits, or not swapping dimers, led to virtually identical EIE values (data not shown) The analysis of the electrostatic field generated by dimeric RNases showed that all dimers possess a positively charged end located at the N-terminal region and a negative end located at the C-terminal one Hence, they can be described as dipoles with an orientation parallel to the direction from the RNases C-terminal surfaces (negatively charged) to the N-terminal surfaces (positively charged) In the supplementary material, Fig S2 shows the shape of the electrostatic field of BS-RNase seen from different directions Figure S3 compares the electrostatic field of different Fig Plot of the EIE values as function of rotation (r) and inclination (i) angles of RNases The hatched axis at the bottom and right of the plots show schemes of some conformations with i ¼ 90° and r ¼ 180°, respectively FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS 3691 Electrostatic interactions and antitumor RNases E Notomista et al RNases oriented with the negative end of the dipoles toward the bottom, and the positive one toward the top of the figure Figure S4 shows instead the N-terminal surfaces colored by charge As expected, in the orientation that provides the strongest attraction between the RNase and the model membrane the RNases dipoles are perpendicular to the negatively charged membrane In Table the EIEmax values of the RNases investigated in this study are tabulated and compared to the RNases’ net charge values, and their ability to destabilize negatively charged membranes A satisfactory correlation is evident not only between the RNases’ EIEmax values and their positive net charges, but also with their ability to destabilize a membrane We then considered the possibility of a correlation between the cytotoxic action of the investigated dimeric RNases and their ability to destabilize and permeate a membrane, which in turn appears to be linked to the attraction of the RNase to a membrane, as evidenced by the RNase negative EIEmax values As a measure of the RNase cytotoxicity we used the reciprocal of the IC50 parameter The values of IC50, the RNase concentration at which 50% of the cytotoxic effect is displayed, were obtained [11,21,22] by assaying the RNase cytotoxic activity on the same cell type, namely SVT2 malignant fibroblasts When the EIEmax values of dimeric, cytotoxic RNases were plotted against the IC50 values reported for these RNases (Fig 6), a direct, Fig Correlation between antitumor activity on SVT2 cells and the highest negative EIE values for covalent dimeric RNases identified by the abbreviations listed in Table Antitumor activity was plotted as the reciprocal of IC50, the RNase concentration producing 50% cytotoxicity [11,14] The square correlation coefficient (R2) is shown 3692 linear correlation was found between the two sets of values, with a significant correlation coefficient (0.88) This strongly suggests a positive correlation between the attraction to a membrane of a dimeric RNase and its cytotoxic activity Discussion Although many years have gone by since an RNase was found to have an antitumor action [27], the structural and functional determinants which render certain RNases capable of exerting this effect are mostly obscure As mentioned above, a primary if not exclusive role of the cytosolic inhibitor in the mechanism of antitumor RNases cannot be generalized The only firmly established fact is that the cytotoxic action of RNases is based on their RNA degrading activity [5] The cytosolic localization of target RNA(s), and previous data on the intracellular journey of antitumor seminal RNase [20], indicate that cytotoxic RNases must be capable of permeating specific membrane compartments to reach the cytosol Furthermore, antitumor seminal RNase has been found to alter the stability of negatively charged membrane vesicles, with no effects on positively charged membranes [26] It has long been known that subcellular membranes from malignant tissues have a higher content of negatively charged lipids when compared to normal tissues [28] More recently, it has been found [29] that some C2 domains, folded modules from proteins involved in signal transduction and vesicle trafficking, specifically associate to negatively charged membranes As all these data are suggestive of a role of membranes, in particular of negatively charged membranes, in the mechanism of action of RNases, we investigated the ability of a series of RNases to destabilize model membranes as a representation of their potential to cross intracellular membranes through their journey to the cytosol We found that cytotoxic, dimeric RNases, natural or engineered, are able to destabilize negatively charged membranes, as they strongly affect membrane aggregation, fluidity and fusion When the RNases were tested in silico with a model membrane by determining the electrostatic energy of their interaction, we found that their ability to destabilize membranes is correlated with their electrostatic attraction to the model membrane In particular, such property was found to be strictly dependent on the ability of the RNases to approach the membranes with their N-terminal regions Any other orientations in their contacts with the membranes produced a less attractive electrostatic interaction Hence RNases that tumble about a membrane FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS E Notomista et al until their most membrane-attractive structural element, namely their N-terminal regions, interact with the membrane, are the most capable of affecting and permeating membranes Furthermore, our results also reveal a stringent correlation between (a) the ability of dimeric RNases to destabilize membranes with their specific membrane-attractive region, and (b) their cytotoxic action This finding strongly suggests that the electrostatic surface potential of a defined 3D district of these proteins has a role in determining their cytotoxic action It has been reported that when negative charges are abolished in noncytotoxic RNases, these become cytotoxic [30,31] This result, obtained with heterogeneous protein products, only indicates that molecules with a high content of positive charges are generally toxic to cells However, a high content of basic residues may not be the only basis of RNase cytotoxicity Two RNases, homologous to the RNases investigated in this study, have been isolated from human eosinophils, namely eosinophil-derived neurotoxin (EDN) and eosinophil cationic protein (ECP) They are RNases with a high sequence identity (66%), a very high content of basic residues and high isoelectric points (pI ¼ 11.9 and 10.4, for EDN and ECP, respectively) Yet only ECP, not EDN, displays cytotoxic activity on eukaryotic and bacterial cells [32] Given the recently reported cytotoxic activity of the noncovalent dimers of RNase A [17], intriguing when compared to their sensitivity to the cytosolic inhibitor (see above), we tested their interaction with the model membrane described above for RNase covalent dimers, and calculated their EIEmax values Two distinct noncovalent dimers have been described for RNase A, in which the dimeric structure is stabilized through the exchange between subunits of either their N-terminal, or their C-terminal ends [33–35] We found that the behavior with the model membrane of the dimer in which the C-terminal ends are exchanged between subunits is perfectly superimposable to the behavior described in this report for any covalently dimeric RNase Its highest negative value of EIE was calculated to be )28 kT The dimer, in which the subunits exchange their N-terminal ends, behaved differently It did not present a single, unique value of highest negative EIE, but two weak negative values of )13 and )10 kT We noted that by summing up the two EIEmax values, a value of EIE was obtained comparable to those calculated for the other dimeric RNase As shown in Fig 7, the EIEmax values obtained for the two dimers satisfactorily correlate with the dimers IC50 values as reported by Matousek Electrostatic interactions and antitumor RNases Fig Correlation between antitumor activity on HL-60 cells (n) and ML-2 cells (h) and the highest negative EIE values for BS-RNase (BS) and RNase A noncovalent dimmers Nd, the dimer with N-terminal swapping between subunits; Cd, the dimer with C-terminal swapping between subunits Antitumor activity was plotted (see legend to Fig 6) as the reciprocal of IC50 reported values [17] The square correlation coefficients (R2) are shown et al [17] Thus, a role of electrostatic membrane attraction has been identified also for noncovalent, cRI sensitive RNases In conclusion, the data presented here lead to the proposal that cytotoxic RNases must possess specific electrostatic features and structural element(s) for destabilizing and eventually permeating membranes, and that intracellular membranes play a decisive role in defining the antitumor action of an RNase Experimental procedures Assays with membranes Synthetic DMPG and DMPS were purchased from Avanti Polar Lipids (Alabaster, AL) Egg yolk phosphatidylglycerol (PG) was from Sigma (St Louis, MO) Unless indicated, lipid vesicles were prepared at mgỈmL)1 phospholipid concentration in the indicated buffer, by extrusion through two 0.1-lm polycarbonate filters (Nuclepore, Costar, Cambridge, MA) in an Extruder (Lipex Biomembranes Inc., Vancouver, Canada), as described previously [26,36–38] Protein concentration was determined from absorbance measurements on a Beckman DU-7 (Palo Alto, CA) or Uvikon 930 (Kontron Instruments, Milan, Italy) spectrophotometers using the following extinction coefficients: FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS 3693 Electrostatic interactions and antitumor RNases E Notomista et al 0.695 (E0.1%, 280 nm) for RNase A and its variants, 0.465 (E0.1%, 278 nm) for BS-RNase The absorbance variation at 360 nm produced by the addition of the proteins to a vesicle suspension was continuously measured on a Beckman DU-640 spectrophotometer equipped with a high performance temperature controller In all assays, controls with no protein were always carried out Intermixing of membrane lipids was analyzed by fluorescence RET assays [39] The RET assay monitors the relief of fluorescence energy transfer between a donor ⁄ acceptor pair as the two probes dilute from labeled into unlabeled bilayers A vesicle population containing 0.6% (v ⁄ v) N-(lissamine rhodamine sulphonyl)-diacylphosphatidylethanolamine as acceptor and 1% (v ⁄ v) N-(7-nitro-2-1,3benzoxadiazol-4-yl)-dimyristoylphosphatidylethanolamine as donor (Avanti Polar Lipids) was mixed with unlabeled vesicles at : molar ratio The increase of the fluorescence emission at 530 nm (donor emission) upon excitation at 450 nm (4 nm slit width for both excitation and emission beams), was continuously recorded on a SLM-Aminco 8000 spectrofluorimeter (Urbana, IL) Measurements were performed in a thermostated cell holder at 37 °C The percentages of RET were calculated according to the equation (%RET) ¼ (1–F ⁄ F0) · 100 where F and F0 are the fluorescence emission intensities at 530 nm upon addition of the corresponding protein, and the value measured with vesicles lacking the acceptor probe, respectively Normalization of the measurements was performed by independent experiments in the presence of 1% (v ⁄ v) Triton X-100 [26,36,39] Fluorescence depolarization measurements were performed on a SLM-Aminco 8000 spectrofluorimeter equipped with 10 mm Glan-Thompson polarizers (Urbana, IL) Cells of 0.2-cm optical path were used Slit widths were nm both for excitation and emission beams Labeling of the vesicles with DPH (Aldrich, Milwaukee, WI) was performed as previously described [40] The degree of polarization of the fluorescence emission of DPH was measured at 425 nm upon excitation at 365 nm, after equilibration of the sample at the required temperature Independent experiments demonstrated a negligible contribution of the protein to the degree of polarization of the fluorescence probe Leakage of vesicle aqueous contents induced by the RNases was measured by the 1,3,6-trisulphonate-8-aminonaphtalene (ANTS) ⁄ p-xylenebispyridinium bromide (DPX) system [26,38,41] PG vesicles contained 12.5 mm ANTS, 45 mm DPX, 20 mm NaCl and 10 mm Tris buffer pH 7.5 Unencapsulated material was separated from the ANTS ⁄ DPX containing vesicles by gel filtration on a Sephadex G-75 column (Sigma) equilibrated in 10 mm Tris buffer pH 7.5 containing 0.1 m NaCl and mm EDTA Fluorescence emission at 510 nm was measured upon excitation at 386 nm on a SLM-Aminco 8000 spectrofluorimeter Internal calibration of the assays was as follows: 0% leakage corresponded to the fluorescence signal of the 3694 vesicles before protein addition (F0), and 100% leakage was the fluorescence intensity measured upon detergent addition [1% (v ⁄ v) Triton X-100] (Fmax) Percentages of leakage were calculated as (%L) ¼ (F–F0) ⁄ (Fmax–F0) · 100 Protein fluorescence emission was measured on a SLM-Aminco 8000 spectrofluorimeter at 25 °C with 0.2 cm optical path cells CD measurements were performed on a Jasco J-715 spectropolarimeter (Tokyo, Japan) with thermostated cylindrical cells Determination of EIE The EIE for RNase–lipidic membrane model complexes was carried out using delphi version [42,43] A model membrane in a crystal-like state was modeled containing 192 palmityl-stearyl-phosphatidylglycerol molecules per layer arranged in a parallelepiped with dimen˚ sions 117(l) · 108(w) · 54(h) A The phospholipids were arranged in a compact hexagonal lattice with the fatty acid tails and the terminal glycerol moiety completely extended Each layer was composed of seven rows of 15 molecules separated by six rows of 16 molecules Within each row the distance between the P atoms of two adjacent phosphates ˚ was 8.88 A, whereas the minimum distance between the P ˚ atoms of adjacent rows was 8.98 A The minimum distance between two P atoms of phosphates of the two different ˚ layers was 48.96 A For each RNase several RNase–lipidic membrane model complexes were generated through an automated procedure which included the following steps: (a) the model of the membrane was oriented with each layer parallel to the xy plane so that all the phosphorus atoms of each layer had the same z coordinate and the z axis was perpendicular to the center of the membrane (Fig 4); (b) the major axis of the dimeric RNase was made coincident with the z axis of the system, hence orthogonal to the membrane plane (Fig 4); (c) the dimeric RNase was rotated around the axis z (by rotation angle r); values of EIE were recorded at 30° intervals of r from to 360°; (d) the inclination of the dimer major axis with respect to the z axis, hence to the membrane plane, was changed (by inclination of angle i); EIE values were recorded at 30° intervals of i from to 180°; (e) steps (c) and (d) were repeated until i reached the maximum value (180°) It should be noted that after each rotation or inclination the RNase dimer was translated: (a) to keep the mass centre of the protein (marked with a black dot in Fig 4) onto the axis z, and (b) to conserve the minimal distance ˚ between protein and membrane at 3.5 A The EIE, i.e., the electrostatic contribution to the binding energy (DGbindingR ⁄ M) of RNase (R) to a lipidic membrane FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS E Notomista et al (M) model complexes was calculated by the method of the grid energy differences [42] through the equation: EIE ¼ DGbindingR=M ¼ GR=M À GR À GM where GR ⁄ M, GR and GM are the grid energies of the complex, of the RNase alone and of the membrane alone, respectively The following delphi parameters were used for the calculations of the grid energy values: gsize ¼ 250; scale ¼ 1.2; indi ¼ 4.0; exdi ¼ 80.0; prbrad ¼ 1.4; salt ¼ 0.1; ionrad ¼ 2.0; bndcon ¼ 2; maxc ¼ 0.0001; linit ¼ 800 Standard formal charges at pH 7.0 were attributed to protein atoms, whereas a charge of )0.5 was attributed to the nonester oxygen atoms of each phosphate group in the membrane (total charge of each phosphate group ¼ )1) RNase structures and modeling Protein–lipidic membrane complexes of BS-RNase, RNase A, RNase AA-N, RNase AA-C and HP-RNase were prepared using the crystallographic structures of these proteins (PDB codes 1BSR, 7RSA, 1A2W, 1F0V and 1Z7X, respectively) As in the 1Z7X structure of HP-RNase Lys1 is lacking, this residue was modeled using the 7RSA structure as template Protein–lipidic membrane complexes of RNase AA, RNase AA-G, RNase AA-GG, and HHP-RNase were prepared using models of these dimers The models of RNase A covalent dimers in the nonexchanging conformation were prepared using the 7RSA structure and the structure of nonexchanging BS-RNase (1R3M) as template Briefly, two molecules of RNase A were superimposed to the subunits of BS-RNase using the fit tools of swisspdbviewer (http://www.expasy.org/spdbv); hence residues 19, 28, 31, 32, and when necessary 38 and 111 of RNase A molecules were mutated to the corresponding residues of BS-RNase, forcing the mutated side-chains to adopt the conformation present in BS-RNase The models were optimized for energy minimization using the gromos implementation of swiss-pdbviewer (50 cycles of steepest descent followed by 50 cycles of conjugate gradients and 50 cycles of steepest descent) No clashes were detected at the dimer interface of the models or in the surroundings of the mutated residues The models of RNase A covalent dimers in the exchanging conformation were prepared using the 7RSA structure and the structure of exchanging BS-RNase (1BSR) as template The server swissmodel [44–46] was used to prepare the homology model of exchanging RNase AA dimer based on the exchanging structure of BS-RNase The hinge loop, i.e., the loop which adopts different conformations in the exchanging and nonexchanging BS-RNase dimers, was cut from the optimized homology model and ligated to the models of nonexchanging RNase A dimers as described above The gromos implementation of swiss-pdbviewer was used to optimize geometry of the hinge loop after ligation Also in this case no clashes were detected at Electrostatic interactions and antitumor RNases the dimer interface of the models or in the surroundings of the hinge loop residues Likewise, the models of nonexchanging and exchanging HHP-RNase were prepared starting from 1Z7X The programs pymol (DeLano Scientific LLC, San Francisco, CA), and swiss-pdbviewer were used to inspect RNase structures and models 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Peitsch MC (1997) Swiss-Model and the Swiss-PDB-viewer: an environment for comparative protein modelling Electrophoresis 18, 2714–2723 Electrostatic interactions and antitumor RNases Supplementary material The following supplementary material is available online: Fig S1 Structure of the BS-RNase–model membrane complex with the highest negative EIE value Lipids are shown as van der Waals spheres and colored according to the atom type Solvent accessible surface is shown for BS-RNase The two subunits are colored blue and red In (A) the solvent accessible surface is transparent to show secondary structure elements Fig S2 Electrostatic field of BS-RNase at pH 7.0 Blue meshes correspond to potential values ẳ +2 kTặe)1 and red meshes to potential values ẳ )2 kTặe)1 Secondary structure elements are shown in green Fig S3 Electrostatic field at pH 7.0 of monomeric and dimeric RNases The proteins have the same orientation with the ‘basic surface’ toward the top of the figure Cyan meshes correspond to potential values ¼ +2 kTặe)1 and red meshes to potential values ẳ )2 kTỈe)1 The solvent accessible surfaces of the RNases are shown in light gray Fig S4 Electrostatic potential at pH 7.0 mapped to the accessible surface of monomeric and dimeric RNases The proteins have the same orientation with the ‘basic surface’ toward the observer Blue color becomes saturated at +2 kTỈe)1 and the red color at )2 kTỈe)1 In RNase-AA the ‘basic surface’ is much less basic owing to the four carboxylates of residues Asp38 and Glu111 In RNase-AA-GG the ‘basic surface’ is more similar to that of BS-RNase because of the removal of the four carboxylates The ‘basic surface’ of the mutant HHP-RNase is very similar to that of RNase-AA-GG as the presence of Glu111 is counterbalanced by basic residues not present on RNaseAA-GG This material is available as part of the online article from http://www.blackwell-synergy.com FEBS Journal 273 (2006) 3687–3697 ª 2006 The Authors Journal compilation ª 2006 FEBS 3697 ... geometry of the hinge loop after ligation Also in this case no clashes were detected at Electrostatic interactions and antitumor RNases the dimer interface of the models or in the surroundings of the. .. the C-terminal and N-terminal structural regions, respectively, of the two A and B monomers of the dimeric RNase The black circle between the monomers indicates the mass center of the dimer Electrostatic. .. Electrostatic interactions and antitumor RNases r ¼ 180° and i ¼ 90° In this orientation, characterized by the strongest electrostatic attraction of the RNases with the membrane, all RNases pointed their