âZero-Lengthâ Dimers of Ribonuclease A - American Chemical Society

Bioconjugate Chem. 2010, 21, 635–645

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“Zero-Length” Dimers of Ribonuclease A: Further Characterization and No Evidence of Cytotoxicity Francesca Vottariello, Chiara Costanzo, Giovanni Gotte, and Massimo Libonati* Dipartimento di Scienze Morfologico-Biomediche, Sezione di Chimica Biologica, Facolta` di Medicina e Chirurgia, Universita` di Verona, Strada Le Grazie 8, I-37134 Verona, Italy. Received September 15, 2009; Revised Manuscript Received February 10, 2010

“Zero-length” dimers of ribonuclease A, a novel type of dimers formed by two RNase A molecules bound to each other through a zero-length amide bond [Simons, B. L., et al. (2007) Proteins 66, 183-195], were further characterized and tested for their possible in vitro cytotoxic activity. Results obtained are the following. Besides dimers, also trimers and higher oligomers could be identified among the products of the covalently linking reaction, and the “zero-length” dimers prepared by us appear not to be a unique species. The product was indeed heterogeneous, and results obtained with two RNase A mutants, E9A and K66A, indicated that amino and carboxyl groups others than those belonging to Lys66 and Glu9 are involved in the amide bond. As for their functional properties, the “zero-length” dimers degrade poly(A) · poly(U) (dsRNA) with an activity that increases with the increase of the oligomer’s basicity and yeast RNA (ssRNA) with an activity that instead decreases with the increase of oligomer’s basicity, which is in agreement with previous data. No cytotoxicity of the RNase A “zerolength” dimers could be evidenced in assays performed with various tumor cells lines; the dimers, instead, become cytotoxic if cationized by conjugation with polyethylenimine (PEI) [Futami et al. (2005) J. Biosci. Bioengin. 99, 95-103]. However, PEI derivatives of RNase A “zero-length” dimers and PEI derivatives of native RNase A resulted to be equally cytotoxic. In other words, protein “dimericity” does not play any role in this case. Moreover, the acquired cytotoxicity does not seem to be specific for tumor cells: PEI-cationized native RNase A was also cytotoxic toward human monocytes.

INTRODUCTION Many ribonucleases were found to be endowed with remarkable biological activities, among which an interesting in vitro and in vivo cytotoxic action against tumor cells (1-5). This finding aroused new interest in this class of enzymes and opened a new chapter in the field of potential anticancer therapy. Among cytotoxic monomeric ribonucleases, a well-known example is onconase, an enzyme found in the oocytes and early embryos of Rana pipiens (2). Among mammalian pancreatic-type ribonucleases, well-known is the ribonuclease from bovine semen, BS-RNase1. Bovine seminal RNase is a unique natural dimer, with its two subunits sharing 83% identity with bovine pancreatic ribonuclease (6, 7). This dimeric RNase is endowed with various biological activities, among which is a strong antitumor action (7, 8). Artificial dimers of RNase A were casually obtained for the first time in 1962 by Crestfield, Stein, and Moore (9). These authors envisaged the way in which an RNase A dimer could * Corresponding author. Tel: +39-045-8027166. Fax: +39-0458027170. E-mail: [email protected]. 1 Abbreviations: BS-RNase, bovine seminal ribonuclease; RNase A, bovine pancreatic ribonuclease A; N-dimer (ND) or C-dimer (CD), N-terminal or C-terminal swapped dimer of RNase A; C-trimer (CT), cyclic RNase A trimer; zl, “zero-length”; zl-D, zl-T or zl-HO, “zerolength” dimers, trimers or higher oligomers of RNase A; H-wt M, H-E9A M, H-K66A M, heated native (monomeric) RNase A and RNase A variants; PEI, polyethylenimine; PEI-M, PEI-conjugated wt (monomeric) RNase A; PEI-RNase A-zl-D, PEI-conjugated RNase A zerolength dimers; NaP, sodium phosphate buffer; TNBS, trinitrobenzensulfonic acid; ssRNA, single-stranded RNA; dsRNA, double-stranded RNA; dsRNase, an RNase active on dsRNA; IPTG, isopropyl β-D-1thiogalactopyranoside; PMSF, phenylmethylsulfonyl fluoride; EDC, 1-ethyl-3-(dimethylaminopropyl)-carbodiimide; cRI, cytosolic ribonuclease inhibitor; LB, lysogenic broth; FBS, fetal bovine serum.

form and proposed a model consisting in the exchange between two ribonuclease molecules of their N-terminal ends (9, 10). This mechanism was later identified and confirmed by Eisenberg and colleagues in studying the structure of the diphtheria toxin (11, 12), and named “three-dimensional (3D) domain swapping”. In 1998, Eisenberg et al. (13) found it to determine the structure of one (the N-dimer, as it was called later) of the two dimeric conformers of RNase A that had been identified by Crestfield and colleagues (10). Dimerization of RNase A had also been obtained by cross-linking two enzyme molecules with the bifunctional reagent dimethyl suberimidate, which reacts with the ε-amino group of one lysine residue of each protein molecule (14). These cross-linked RNase A dimers exhibited antitumor action similar to that of BS-RNase (15, 16). All ribonuclease dimers, natural or artificial, degrade doublestranded RNA (14, 17), DNA/RNA hybrids (18), and poly(A) (19) under conditions where native, monomeric RNase A does not. On this basis, the idea grew that “dimericity” could be a determinant factor in conferring novel and functionally important side properties to a ribonuclease (14-19). Evidence further supporting this idea also came from dimeric mutants of human pancreatic RNase (20) or bovine pancreatic RNase A (21), which were shown to be selectively cytotoxic toward tumor cells, while the corresponding native, monomeric proteins are devoid of any biological activity. With the procedure described by Crestfield et al. (9), i.e., by lyophilizing RNase A from 40-50% acetic acid solutions, trimers, tetramers, and higher-order oligomers also form besides dimers, each species comprising, like dimeric RNase A (10), at least two conformers, one more basic, one less basic, one more abundant, and one less (22-27). The structure of the second, more abundant and more basic RNase A dimer (C-dimer), formed through the exchange of the C-termini between two protein molecules (28), and the structure of the

10.1021/bc900407v  2010 American Chemical Society Published on Web 03/04/2010

636 Bioconjugate Chem., Vol. 21, No. 4, 2010

C-terminal domain-swapped cyclic trimer (C-trimer) (25, 26, 28) have been solved, and plausible models were proposed for another trimer and several other RNase A oligomers (23-29). The 3D domain-swapped RNase A oligomers larger than dimers are also able to degrade dsRNA (22, 24, 27). Dimers and higher oligomers of RNase A are cytotoxic (30) like BS-RNase (7, 8) and the RNase A cross-linked dimers (15, 16) and trimers (31), the extent of these activities increasing as a function of the oligomers’ complexity (24, 30, 32). Therefore, not only the “dimericity”, but in general the “oligomericity” of a ribonuclease, characterized by the increase of the basicity of the complex, was thought to be an important structural determinant for the acquisition by the enzyme of novel functional properties (24, 32). In 2006, Leich et al. designed a fusion protein consisting of two RNase A molecules bound to each other by a peptide linker of varying length and composition between the C-terminus of one and the N-terminus of a second ribonuclease molecule by gene duplication (33). This tandem RNase A dimer is a potent cytotoxic agent against human erytroleukemia cells (further evidence of the importance of ribonuclease dimericity), although its ribonucleolytic activity is severely reduced. More recently, Simons et al. prepared a novel type of covalently linked RNase A dimer, which consists of two enzyme molecules bound to each other through a zerolength amide bond (34). This dimer also is able to degrade double-stranded poly(A) · poly(U) besides ssRNA (34). We do not know whether the cytotoxic tandem dimer of Leich et al. is also able to attack double-helical polyribonucleotides. On the other hand, Simons et al. did not show whether their “zero-length” RNase A dimers might acquire cytotoxic activity in parallel with their ability to degrade double-stranded RNA. Theoretically, both an additional enzymatic activity for the tandem dimer of Leich et al. and a biological action for the “zero-length” dimer of Simons et al. could reasonably be expected. Here, we have examined the latter possibility, i.e., whether RNase A dimerized through the zero-length amide bond, according to the procedure of Simons et al. (34), could show a cytotoxic action. The results we obtained were negative, i.e., the “zero-length” dimers do not show, at least under the experimental conditions and with the cell lines used by us, any significant cytotoxicity. They acquire a high cytotoxic activity against tumor cells cultures if cationized with polyethylenimine (PEI) according to the procedure outlined by Futami et al. (35). The extent of this cytotoxicity is, however, not higher than that, already evidenced by Futami et al. (35), exerted by PEIcationized monomeric RNase A. Moreover, the latter appears to be cytotoxic also against human monocytes, i.e., normal cells.

EXPERIMENTAL PROCEDURES Materials. All chemicals were of the highest purity available. Their origin is specified in text and legends to figures and tables. Bovine seminal ribonuclease was a kind gift of Dr. J. Matousek (Institute of Animal Physiology and Genetics, The Academy of Sciences of the Czech Republic, Libechov 27721, Czech Republic). Isolation and purification of the enzyme were performed as described elsewhere (30). In Vacuo Cross Linking of RNase A. According to the procedure outlined by Simons et al. (34, 36), bovine pancreatic RNase A (Type XII-A, purchased from Sigma Chem. Co., lot no. 124K7680, R5500) was dissolved in ddH2O at a concentration of 5-40 mg/mL (when necessary, the pH of the solution was adjusted to 7.0 with 1 N NaOH) and lyophilized. The glass tube was sealed under vacuum (∼0.5 mBar) and incubated in an oven at 85 °C for 96-144 h. After releasing the vacuum, the protein sample was reconstituted with 40 mM sodium phosphate buffer, pH 6.7, to obtain a final protein concentration of 10 mg/mL. Then, the sample was incubated at 50 °C for 24 h to dissociate RNase A noncovalent oligomers formed.

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Conjugation of Bovine RNase A and Its “Zero-Length” Dimers to Polyethylenimine. We followed the procedure described by Futami et al. (35). Native RNase A was dissolved in a 60 mg/mL solution of polyethylenimine (PEI, Mn 1200, M.W. 1300 Da, Sigma #482595, lot no. 05329KH), pH 5.0; protein final concentration, 1 mg/mL. EDC (Sigma #E7750, lot no. 107K0664) was added, final concentration, 0.1 mg/mL. The mixture was incubated for 16-24 h at room temperature. The same procedure was followed for the conjugation of the zerolength dimers. Then, the mixture was dyalized against water, and concentrated with Amicon Ultra filters (Millipore), cutoff 3000 Da. At the end, the material was lyophilized. Otherwise, it was partially purified performing two-step cation-exchange chromatography (see next paragraph). Purification and Analysis of the Covalent RNase A Oligomers. Size-Exclusion Chromatography. Purification of the covalent RNase A oligomers was accomplished by two sizeexclusion chromatography steps performed with a Superdex 75 26/60 column (GE-Healthcare) attached to an a¨KTA FPLC system, at room temperature. The column was equilibrated with 0.2 or 0.4 M NaP (pH 6.7)/0.15 M NaCl. The flow rate was 1 mL/min. To determine the yield of the reaction and the purity of the products the mixture of oligomers was analyzed with two size-exclusion chromatography steps using a Superdex 75 HR 10/300 column (GE-Healthcare) attached to an a¨KTA FPLC system, at room temperature. The column was equilibrated with 0.2 and 0.4 NaP, pH 6.7, in the first and second purification step, respectively. Elution was performed at a flow rate of 0.15 mL/min in the first and 0.07 mL/min in the second purification step. Ion-Exchange Chromatography. Analyses of the covalent RNase A “zero-length” oligomers were also performed with cation-exchange chromatography using either Source 15S HR 10/10 or Mono S strong cationic columns (GE-Healthcare) and the same a¨KTA FPLC system mentioned above, at room temperature. Three different gradients were applied to the columns. The first was a preliminary stepwise gradient of NaP (70-90 mM, pH 6.7), followed by a linear (90-200 mM, pH 6.7) gradient at a flow rate of 1 mL/min (total, 44 min). The second was a linear 40-200 mM NaP (pH 6.7) gradient at a flow rate of 1 mL/min (total, 60 min). The third was a stepwise 40-200 mM NaP (pH 6.7) gradient. To separate the RNase A-PEI conjugate(s) from nonconjugated RNase A and purify the products, two different elution schemes of cation-exchange chromatography were used: (i) a stepwise procedure, increasing by 30 mM, from 70 to 400 mM, the NaP (pH 6.7) concentration, and (ii) a two-step cationexchange chromatography, each step under isocratic conditions (first step: 70 mM NaP, pH 6.7, to elute the nonconjugated protein; second step: 400 mM NaP, pH 6.7). Quantification of the “Zero-Length” Oligomers. Each oligomeric species produced was quantified by measuring the area of its peak appearing in the gel filtration pattern of the first sizeexclusion purification step and calculating its percentage relative to the sum of the areas of all peaks eluted. The purity of the “zero-length” species was also determined by measuring the intensity of the corresponding gel band with the use of the ImageJ program and calculating its percentage relative to the sum of the intensities of all bands. Gel Electrophoresis. Cathodic gel electrophoresis under nondenaturing conditions was performed according to Goldenberg (37), with slight modifications (22), using β-alanine/acetic acid buffer, pH 4.0. 12.5-15% polyacrylamide gels were run at 20 mA for 100-140 min at 4 °C. Fixing and staining were performed with 12% trichloroacetic acid and 0.1% Coomassie Blue R-250.

“Zero-Length” Dimers of Ribonuclease A

SDS PAGE was performed according to Laemmli (38) using Tris/glycine buffer, pH 8.3. 12.5-15% polyacrylamide gels and 18% acrylamide gels were used to analyze the “zero-length” species or the products conjugated to PEI. Gels were run at 20-22 mA for 100-120 min at room temperature. Staining was made with a solution containing 0.025% Coomassie Blue R-250, 40% methanol, and 7% acetic acid. DNA gel electrophoresis was performed with a 1% agarose gel, using 0.04 M Tris-acetate buffer, pH 8, containing 1 mM EDTA (TAE buffer). Gels were run at 100 V for 60 min, and a 500 bp Molecular Ruler (Bio-Rad #170-8203) was used as a molecular weight standard. Production of RNase A mutants. Construction of the Mutants. The gene coding for wild-type RNase A, cloned into the expression vector pT7-7 (containing also the gene that codes for ampicillin resistance) between the NdeI and HindIII restriction enzyme sites, was kindly provided by Professor A. Di Donato (Faculty of Sciences, the University of Naples). Two mutants of the RNase A gene (E9A and K66A) were obtained using the QuikChange II Site-Directed Mutagenesis Kit (Stratagene, #200523, lot no. 1260135). pT7-7 template (with the RNase A wt gene) was added to the reaction buffer containing dNTPs and mutagenic primers. The sequences of the primers were as follows: Mutant E9A

FW 5′-GCAGCAGCCAAGTTTGCCGCGGCAGCACATGGAC-3′ RV 5′- GTCCATGTGCTGCCGCGCAAACTTGGCTGCTGC-3′ Mutant K66A FW 5′-CCCAGAAAAATGTTGCCTGCGCGAATGGGCAGACC-3′ RV 5′-GGTCTGCCCATTCGCGCAGGCAACATTTTTCTGGG-3′

DNA polymerase was added and PCR performed. After denaturation of the double-stranded DNA template at 95 °C for 30 s, the first cycle was started: 95 °C for 30 s, annealing at 57 °C for 1 min, and polymerization at 68 °C for 12 min. These steps were repeated 16 times. Finally, the sample was cooled to 4 °C. The DNA template was digested with DpnI, and the mutated and amplified plasmid was used to transform the XL1-Blue supercompetent cells. A colony of these cells was isolated and inoculated in 5 mL of LB containing Ampicillin (100 µg/mL). This overnight culture was used for plasmid DNA miniprep purification, using the Wizard Plus SV Minipreps DNA Purification System (Promega, #A1330, lot no. 182970). A DNA gel electrophoresis was performed to verify the presence of the amplified pT7-7 plasmid after PCR and DNA miniprep purification. Furthermore, the plasmidic DNA was sequenced and the presence of the desired mutation confirmed. Production and Purification of RNase A from E. coli Inclusion Bodies. The PT7-7 plasmid containing the RNase A mutated gene was added to E. coli strain BL21-Gold (DE3) cells and incubated on an ice bath for 30 min. Cells were then transformed by thermal shock at 42 °C for 20 s. After 1 min on ice, the transformed E. coli was regenerated in 1 mL of LB (without ampicillin) and incubated at 37 °C for 1 h with shaking. After centrifugation of the culture, the pellet was resuspended in 100 µL of LB broth. Cells were seeded on LB Agar plates containing 100 µg/mL ampicillin, and incubated at 37 °C overnight. A colony was then isolated and inoculated in 5 mL of LB with 50 µg/mL ampicillin. This culture was incubated at 37 °C with shaking for at least 6 h, then inoculated in 150 mL of LB with 50 µg/mL ampicillin and incubated overnight at 37 °C with shaking. Fifteen milliters of this culture was used to inoculate 500 mL of LB containing 50 µg/mL ampicillin. The inoculated culture was shaken at 37 °C until the early log phase (A600 ) 0.6 O.D.) was reached. It was then induced to express the cDNA coding for RNase A by adding IPTG (0.5 mM, final concentration). Induction of protein expression was performed overnight under the same conditions of the incubation.

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The culture was centrifuged at 5500 rpm for 10 min and bacteria were resuspended in 0.1 M Tris-acetate buffer, pH 8.4 (10 mL/1000 OD600 units), containing PMSF as inhibitor of serine proteases and cysteine protease (4 µL of 0.1 M PMSF/g E. coli). To verify the presence of RNase A in the cell culture before starting protein extraction, an analysis with SDS PAGE was performed (18% polyacrylamide gel, Tris/glycine buffer, pH 8.3). Amounts corresponding to 0.126 OD600 of E. coli collected before and after induction with IPTG were run at 20-22 mA at room temperature. Cell lysis and inclusion bodies extraction were performed by incubation with lysozyme (0.8 mg/g E. coli) at room temperature for 20 min, and by freezing (in liquid nitrogen) and thawing (at 37 °C). After these last steps, the extract was treated as described elsewhere (39) to perform RNase A renaturation. To verify the presence of RNase A in the fractions eluted during these two purification steps, a Kunitz assay (40) was performed (purified yeast RNA as substrate), using native RNase A (Sigma Aldrich, type XII-A, R5500) as a control. Determination of Free Amino Groups. 150 µg of protein were dissolved in 1 mL of 0.15 M sodium tetraborate (Na2B4O7), pH 9.5. 0.2 mL of 0.015 M TNBS was added to the solution (“blank” consisted in 1 mL of 0.15 M sodium tetraborate, pH 9.5, and 0.2 mL of TNBS (14)). The reaction mixtures were maintained in the dark at 37 °C for 45 min. Reaction was stopped by adding 2 mL of 85-90% formic acid. Spectrophotometric measurements were performed at 340 nm, and the native, monomeric RNase A value was taken as a standard corresponding to 11 free amino groups (14). Enzymatic Assays. A. Ribonucleolytic activity on ssRNA (purified yeast RNA) as substrate was determined spectrophotometrically at 300 nm, according to Kunitz (40). One unit is the amount of enzyme that is capable of causing a decrease of the RNA absorbance at 300 nm (measured in the initial linear tract) per minute at 25 °C. The substrate is a 0.05% solution of purified yeast RNA in 0.05 M sodium acetate, pH 5.0. B. The activity of the various RNase A species on the doublehelical poly(A) · poly(U) complex (Sigma-Aldrich, P1537) as substrate (40 µg/mL) was determined spectrophotometrically at room temperature by measuring, as described elsewhere (17), the increase of the absorbance at 260 nm after addition of the various RNase species to the assay mixtures. One enzyme unit is the amount of enzyme that catalyzes the cleavage of one µmol of phosphodiester linkages of the double helical substrate per minute at 25 °C (22, 45). Specific activities are expressed as units per milligram protein. Cell Lines and Proliferation Assays. PSN1 (human pancreas adenocarcinoma), K562 (human chronic myelogenous leukemia), and U937 (human leukemic monocyte lymphoma) cell lines were grown in RPMI 1640 medium supplemented with 2 mM L-glutamine, 10% FBS, and 50 mg/mL gentamicin sulfate (BioWhittaker, Cambrex). Human cervix carcinoma HeLa cells were cultured in DMEM supplemented with 2 mM glutamine, 10% FBS, and 50 mg/mL gentamicin sulfate. All mentioned cell lines were incubated at 37 °C with 5% CO2. Human monocytes were separated by centrifugation on Ficoll-Paque Plus under endotoxin-free conditions from buffy coats of healthy donors. Monocytes (99.8% pure), obtained as described in ref 41 and with the MACS Human Monocyte Isolation Kit II (Miltenyi Biotec), were resuspended in RPMI 1640 supplemented with 10% low endotoxin FBS and 2 mM L-glutamine (BioWhittaker). At the beginning of each experiment, PSN1 cells were plated at 4 × 103 cells/well, K562 cells at 8 × 103 cells/well, and monocytes at 2 × 104 cells/well. All cell lines were plated in


Figure 1. Purification and analysis of RNase A “zero-length” oligomers by size-exclusion chromatography. Twenty milligrams of the oligomers’ mixture prepared by the procedure outlined by Simons et al. (34, 36) were applied onto a Superdex 75 26/60 (GE-Healthcare) column. Buffer: 0.2 M NaP, pH 6.7/0.15 M NaCl (34). Flow rate: 1 mL/min, room temperature. M: monomeric RNase A. zl-D, zl-T, zl-HO: “zero-length” dimer, trimer, and higher oligomers of RNase A.

96-well plates and after 24 h treated as described in the figure legends. At the end of the treatments, cell viability was determined by a metabolic assay using the CellTiter 96 AQueous One Solution reagent (Promega) following the manufacturer instructions. Cultures were incubated for 2 h at 37 °C with 5% CO2, and conversion of MTS into formazan was read at 490 nm.

RESULTS AND DISCUSSION Preparation of “Zero-Length” Dimers of RNase A. RNase A “zero-length” dimers were obtained with the procedure described by Simons et al. (34). The products of the reaction were heated at 50 °C for 24 h to dissociate the noncovalent aggregates possibly formed, purified, and analyzed by gel filtration and SDS PAGE. The gel filtration profile (Figure 1) shows the elution of a great amount of monomeric protein (M), preceded by a clear peak of dimer (zl-D), a smaller peak of trimer (zl-T), and a region of unidentified higher oligomers (zlHO). By measuring the areas of the peaks and/or the intensity of the bands, the following amounts could be calculated: 15.2 ( 1.9% for the dimer, 5.3 ( 2.0% for the trimer, and 3.7 ( 2.6% for the higher oligomers. The SDS PAGE analysis (Figure 2A) shows several clearly visible bands, of which the densest one is the residual monomeric RNase A. After gel filtration, the purity of the various oligomeric species was determined by electrophoretic analyses. Their purity values decreased as a function of the oligomers’ sizes (Figure 2B): the dimer was 76.8 ( 6.9% pure, the trimer 46.9 ( 6.7%, and the higher oligomers 44.8 ( 14.8%. The dimeric species, isolated from the gel filtration pattern, was subjected to an ion-exchange chromatographic purification (22, 24, 34), the results of which indicated that the product was actually very heterogeneous (data not shown). All species collected from the gel filtration were also analyzed by cathodic gel electrophoresis (Figure 2C). It is clearly visible that they display an electrophoretic mobility similar to that of the corresponding domain-swapped oligomers, but their spreading bands suggest that the material is heterogeneous. Focusing on zl-D, its heterogeneity could be due to the presence of not only a single dimer (34), but to more than one product of the reaction. In other words, this could mean the possible involvement of amino and carboxyl groups other than only those belonging to Lys66 and Glu9. Are Lys66 and Glu9 the Only Reactive Residues? An Answer from Two RNase A Mutants. To test this hypothesis, we prepared two mutants of RNase A, E9A and K66A, through

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site-directed mutagenesis (see Experimental Procedures). On the basis of the work of Simons et al. (34), these two protein species should not react in the cross-linking reaction developed by these authors, that is, no “zero-length” dimers of RNase A should form. Before testing this possibility, we verified the nature of the two mutants by sequencing their plasmidic DNA. In addition, the elution profiles of the two mutants obtained with a cationexchange chromatography performed under isocratic conditions (Supporting Information Figure 1) confirmed that the sitedirected mutagenesis had been successful. Indeed, the K66A variant is more acid than the wild type lacking one basic amino acid residue and elutes before native RNase A, whereas the E9A mutant, lacking an acid residue, becomes more basic and elutes definitely later. The two mutants were now subjected to the identical experimental conditions used with native RNase A to produce “zero-length” dimers, and the results obtained are shown in Figure 3 and Table 1. It clearly appears that the two variants (Figure 3A,B) are capable of forming dimers, trimers, and higher oligomers in amounts and relative proportions (Table 1) similar to those obtained with native RNase A (Figure 3C). Therefore, differently from what observed by Simons et al. (34), Lys66 and Glu9 are not the only residues of RNase A reacting in the procedure. Determination of Free Amino Groups. To improve the understanding of the results presented above, we determined (by the reaction with TNBS; see Experimental Procedures, and ref 14) the number of free amino groups present in the protein species deriving from the two variants. They were analyzed before (monomers) and after (monomers M, and dimers, zl-D) subjecting them to the in vacuo heating reaction outlined by Simons et al. (34). Table 2 shows the results obtained. On the basis of the eleven NH2 groups present in native, monomeric RNase A (14), it clearly appears that between two and four lysine residues are involved in the amide bond(s) forming in the procedure (34) in all types of dimers examined. The analysis was also performed on the monomers (M) of the three RNase A species. In the case of the monomeric K66A variant, both the unheated and heated (H, i.e., subjected to the in vacuo heating reaction) mutant have one NH2 group less than native RNase A. The missing amino group is obviously that of the Lys66 residue which was mutated in to alanine. Notably, no intramolecular bonds form in the K66A variant. On the contrary, in the E9A variant one amino group is missing only after the in vacuo heating reaction (H E9A M), as it occurs in native RNase A (H-wt M). This might suggest that an intramolecular amide bond could form in these species, involving the Lys66 ε-NH2 (34). The acid partner to react should be the carboxyl group of Asp121 (42), whose distance from Lys66 (42, 43) is suitable for the reaction to occur. Enzymatic Assays. All RNase A oligomers obtained so far, either by cross-linking reactions or by one of the other ways by which the enzyme protein can aggregate (24, 44), acquire the ability to degrade double-stranded polyribonucleotides. The extent of the degrading activity increases with the size of the oligomer, i.e., with the number or density of positive charges present on the molecule (24, 45). The ability of a ribonuclease to digest dsRNA was in fact ascribed to the occurrence of a high protein basicity (17, 45, 46). Along with this novel catalytic action, basic ribonucleases and the RNase A oligomers maintain their activity against ss-RNA, the extent of which, however, decreases as the size and/or the basicity of the protein molecule increase (24, 45, 47), this being just the opposite of what happens with double-helical RNA substrates. On this basis, we tested the “zero-length” dimers and trimers, and the higher, indistinct oligomers for their action on dsRNA. Table 3 shows the activities of the “zero-length” oligomers of native RNase A (wt zl-D, wt zl-T, wt zl-HO) and of the “zero-length” dimers



Figure 2. Denaturing and nondenaturing PAGE analysis of the RNase A “zero-length” oligomers. (A) 12.5% polyacrylamide SDS PAGE. Lane 1, 12 µg of the total mixture obtained by the procedure outlined by Simons et al. (34); see also Experimental Procedures; lane 2, 2 µg of native RNase A (Sigma Type XII-A) as a standard. (B) 12.5% polyacrylamide SDS PAGE purity assay of the various oligomeric species. Oligomers were purified by two successive gel filtrations. Densities of bands were determined by the ImageJ (1.36-6 MB) program. (C) 12.5% polyacrylamide cathodic PAGE of the RNase A “zero-length” species collected from gel filtration and compared with the domain swapped RNase A oligomers (22, 23). “Zero-length” RNase A oligomers: M ) monomer; D ) dimer; T ) trimer; HO ) higher-order oligomers. In cathodic PAGE: RNase A, native monomer (2 µg); ND, CD, N- or C-terminus swapped RNase A dimers (3 µg each); NCT, N+C-termini swapped trimer (3 µg); CT, cyclic C-terminus swapped trimer (2 µg).

Figure 3. Size-exclusion chromatography of “zero-length” oligomers formed by the two RNase A mutants. The reaction mixture, after heating at 50 °C and cooling, was applied on to a Superdex 75 10/300 (GE-Healthcare) column; buffer, 0.2 M NaP, pH 6.7, containing 0.15 M NaCl; flow rate, 0.15 mL/min, at room temperature. (A) K66A, 1 mg applied; (B) E9A, 1.5 mg applied. (C) WT (wt RNase A), 2 mg applied.

obtained from its two variants (E9A zl-D, K66A zl-D) on dsRNA (poly(A) · poly(U)) or ssRNA (yeast RNA), compared to the action of the corresponding monomeric species. The activities of BS-RNase and of the 3D domain-swapped RNase A oligomers are also included for comparison. The activity values determined with both the ds- and ssRNAs as substrates are in agreement with previous results (23, 24, 42, 45, 47); however, it is worth pointing out the lower capacity of degrading

dsRNA shown by the K66A variant and its oligomers, compared with native RNase A, which can be ascribed to the one positive charge missing in the mutant. While the activity values of the “zero-length” dimers against poly(A) · poly(U) determined by Simons et al. (34) and by us here are similar, we could not confirm the activity values on ssRNA as reported in ref 34: in our hands, the zl-D, zl-T, and zl-HO are progressively less active as a function of the increase of their size (or positive charges


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Table 1. Yields (%) of the “Zero-Length” Oligomers formed by the RNase A Speciesa oligomersb

WT

K66A

E9A

zl-D zl-T + zl-HO

15.2 ( 1.9 9.0 ( 2.9

17.0 ( 0.2 10.6 ( 0.8

14.7 ( 0.9 5.9 ( 0.1

a Yields were determined by measuring the area of each peak and calculating its percentage relative to the sum of the areas of all peaks eluted. The values reported for the two mutants are means of the measured areas from three different gel filtration experiments ( s.d., while the values relative to the wt are means of 11 gel filtration experiments (see Experimental Procedures) ( s.d. b WT, native RNase A; zl-D, “zero-length” dimer; zl-T, “zero-length” trimer; zl-HO, “zero-length” higher oligomers.

Table 2. Free Amino Groups Present in Each RNase A Species RNase A species

free NH2 groupsa

RNase A H-wt Mb wt zl-D E9A M H-E9A M E9A zl-D K66A M H-K66A M K66A zl-D

11.0a 10.3 ( 1.2 7.5 ( 0.9 11.4 ( 0.9 10.0 ( 1.0 7.9 ( 0.9 9.3 ( 0.8 10.0 ( 0.8 7.9 ( 0.7

a In each experiment, the absorbance value determined for native, monomeric RNase A was made equal to 11 amino groups, based on 11 free amino groups present in the native protein (14). The values reported are means of four experimental values ( s.d. b H, “heated”, i.e., RNase A monomeric species recovered after being subjected to the in vacuo heating procedure reported (34) and used to prepare the cross-linked, “zero-length” oligomers. wt, native RNase A; M, monomeric species; zl-D, “zero-length” dimers.

Table 3. Enzymatic Activities of the Various RNase A Monomeric and “Zero-Length” or Domain-Swapped Oligomeric Species on Single-Stranded (Yeast) or Double-Stranded (poly(A) · poly(U)) RNA RNase species

specific activity,a poly(A) · poly(U) as substrate

specific activity,a Yeast RNA as substrate

BS-RNase RNase A H-wt M wt zl-D wt zl-T wt zl-HO E9A M H-E9A M E9A zl-D K66A M H-K66A M K66A zl-D ND CD NCT CT HO

7.8 ( 1.3 2.4 ( 0.5 1.3 ( 0.5 4.6 ( 1.1 6.3 ( 2.1 11.3 ( 2.7 1.1 ( 0.2 n.d.b 2.0 ( 0.2 0.5 ( 0.1 n.d.b 0.6 ( 0.2 2.8 ( 0.1 8.6 ( 0.7 16.1 ( 3.6 20.5 ( 2.2 24.2 ( 2.1

15.5 ( 3.4 70.3 ( 8.8 62.9 ( 8.6 44.7 ( 6.3 29.0 ( 4.4 5.1 ( 0.6 73.6 ( 7.0 66.4 ( 8.3 36.0 ( 8.4 66.8 ( 6.2 68.5 ( 4.1 40.4 ( 5.6 40.7 ( 18.1 16.0 ( 3.5 33.2 ( 7.7 25.7 ( 2.2 15.5 ( 2.7

a Specific activity values were calculated as described in refs 17, 45 (see also Experimental Procedures). Each value is the mean of results obtained in several different assays ( s.d. Amounts of enzyme used with poly(A) · poly(U) as substrate: monomer, 10 µg; dimer, 5 µg; trimer, 2.5 µg; higher oligomers, 1 µg. Amounts of enzyme used with yeast RNA as substrate: monomer or dimers, 0.5 µg; trimer, 1.0 µg; higher oligomers (HO), 1.5 µg. ND, CD, N- or C-terminus swapped RNase A dimers; NCT, N+C-termini swapped trimer; CT, cyclic C-terminus swapped trimer; HO, mixture of N+C-swapped higher-order RNase A oligomers. b Not determined. H (heated) wt M: see legend to Table 2.

density), which agrees with all previous results (24, 42, 45, 47). The enzymatic activity values of the three RNase A monomeric variants determined with yeast RNA as substrate before and after the in vacuo heating reaction show that the activities of the wt enzyme and E9A monomeric variant slightly decrease after the reaction, while the activity of the K66A variant does

Figure 4. Cytotoxic activity of the various RNase A “zero-length” species on two different tumor cell lines. The various “zero length” oligomers were purified, lyophilized, and dissolved in sterile, twicedistilled water just before their addition to the cell cultures for 48 h (see Experimental Procedures). H-M indicates native, monomeric RNase A subjected to, and recovered after, the in vacuo heating (H) procedure of Simons et al. (34); zl-D, “zero-length” dimers of RNase A. (A) PSN1 cell cultures. The various “zero length” oligomers and spermine were used at 10 µg/mL, 25 µg/mL, 50 µg/mL, and 100 µg/mL. (B) K562 cell cultures. The various RNase A ”zero-length” species were used at 10 µg/mL, 25 µg/mL, 50 µg/mL, and spermine at 20 µg/mL. Cell viability was calculated as a percentage of treated versus untreated cells. Each value shown is the mean of three measures from three independent experiments ( s.d.

not. Although modest, that lowering in activity might be ascribed to the modification of D121, whose catalytic role in RNase A is well-known (48). The activity of monomeric K66A, on the contrary, does not substantially change, remaining slightly lower than that of the wt enzyme and E9A mutant either before or after the reaction. This is in line with the known catalytic role of K66 P0 subsite (49). Although the differences of the enzymatic activities (see Table 3) are modest and lower than what could be expected for species that potentially lose D121 (48), it has to be taken into account that possibly yeast RNA is not the most appropriate ssRNA substrate to highlight them (48). Nevertheless, the trend shown by the enzymatic activity of the three RNase A monomeric variants agrees with the hypothesis that the K66-D121 intramolecular bond could form in the in vacuo heating of wt RNase A. Are the RNase A “Zero-Length” Dimers Cytotoxic? This novel type of dimers was tested on PSN1 human adenocarcinoma cell line grown in culture. The results are shown in Figure 4A. We tested various concentrations of (i) native RNase A (NH-M), (ii) the RNase A that was recovered from the in vacuo heating reaction, indicated as H-M, (iii) the “zero-length” dimers (zl-D), and in addition (iv) spermine as a positive control. After 48 h incubation, no significantly higher cytotoxic action of the “zero-length” dimers, compared to that of the two monomeric



Figure 5. Purification and analysis of monomeric RNase A conjugated with polyethylenimine. (A) Stepwise cation-exchange chromatography of the PEI-RNase A complex. The reaction mixture was dialyzed against Milli-Q water and concentrated by Amicon filters (cut off 3000 Da). 750 µg of material were applied on to a Source 15S HR 10/10 (Ge-Healthcare) column. Eluting buffer, NaP, pH 6.7. Starting NaP concentration, 70 mM; at each step, it was raised by 30 mM. Final NaP concentration, 400 mM. Before the end of chromatography, 1 M NaCl was also added to ensure a complete elution of all conjugated species. (B) Partial purification of the PEI-RNase A complex (1 mg): cation-exchange chromatography with a Source 15S HR 10/10 (GE-Healthcare) column. Buffer: NaP, pH 6.7. Two-step chromatography under isocratic conditions: first step, 70 mM NaP buffer, with elution of nonconjugated monomeric RNase A; second step, 400 mM NaP buffer, with elution of the PEI-conjugated species. Percentage values were calculated by measuring the areas of single peaks (see Experimental Procedures). M, RNase A monomer; D, dimer; T, trimer; HO, higher oligomers of RNase A; PEI-M, polyethylenimine-conjugated monomeric RNase A.

species, could be evidenced, while the unspecific cytotoxicity of spermine was remarkably high. A second experiment was carried out with K562 cells, which previously were shown to be susceptible to the action of BSRNase (50). The cells were subjected for 48 h to the action of all RNase A species at the concentrations indicated (see Figure 4B), but the zl-D were inactive, while BS-RNase and spermine were considerably cytotoxic. By extending this experiment up to 72 h, the results did not change (data not shown). Further experiments were performed with HeLa and U937 cells using the same protein concentrations and times of incubation tested with PSN1 cells. In these cases also, the various RNase A species resulted to be totally inactive (data not shown), while spermine was cytotoxic like in the previous experiment. In conclusion, therefore, the “zero-length” oligomers of RNase A appear to be devoid, at least under the experimental conditions and with the cell lines used by us, of any significant cytotoxic activity. Cationization of Native RNase A and Its “Zero-Length” Dimers with Polyethylenimine and Analysis of the Products of the Conjugation Reaction. The cytotoxicity of RNase A cationized with polyethylenimine (PEI) has been shown by Futami et al. (35). Could the PEI-cationized RNase A “zerolength” dimer(s) acquire a cytotoxic action higher than that of the PEI-cationized monomer, which could be ascribable to the dimericity of the RNase A molecule? To answer this question, we used the procedure outlined by Futami et al. (ref 35; see also Experimental Procedures) to cationize with PEI the zl-D prepared by us. The reaction was first carried out with native, monomeric RNase A, to obtain the species called PEI-M. The products of the reaction were purified and analyzed by cationexchange chromatography and SDS PAGE. Figure 5A shows the profile of a cation-exchange chromatogram in which a stepwise increase of NaP concentration was applied (see Experimental Procedures). A sharp peak appeared at a NaP concentration of 70 mM (pH 6.7). From its position, it can be deduced that it corresponds to unreacted RNase A contained in the reaction mixture. At higher salt concentration, several irregular peaks emerged successively. This in turn suggests that the product of the PEI-RNase A conjugation is a heterogeneous mixture of PEI-M species deriving from the reaction of the various carboxyl groups available on the protein. To make the

Table 4. Enzymatic Activity of PEI-Conjugated or Not Conjugated RNase A on Double-Stranded (poly(A) · poly(U)) or Single-Stranded (yeast) RNA RNase species

Specific activitya, poly(A) · poly(U) as substrate

Specific activitya, yeast RNA as substrate

BS-RNase RNase A PEI-M (NPb) zl-D PEI-zl-D (NPb)

6.9 ( 0.8 1.9 ( 0.1 2.6 ( 0.5 4.3 ( 0.3 8.4 ( 1.1

14.0 ( 1.6 75.3 ( 8.4 41.3 ( 5.3 59.7 ( 2.5 38.0 ( 8.2

a Specific activity values calculated as described (refs 17, 45; see also Experimental Procedures). Each value is the mean of three different assays ( s.d. Concentrations of the various enzyme species in the assay: 10 µg monomer, and 5 µg dimer with poly(A) · poly(U) as substrate; 0.5 µg monomer or dimer with yeast RNA as substrate. b NP: not purified, total reaction mixture (35) dialyzed against water, concentrated, and then dissolved (1 mg/mL) in Milli-Q water.

isolation of the PEI-RNase A products easier, a different chromatographic approach was followed (Figure 5B): after the elution of monomeric unreacted RNase A at 70 mM NaP, pH 6.7 (22-24), the NaP concentration was brought to 400 mM. Two sharp, but not completely resolved, peaks of conjugated species eluted immediately. They were collected, dialyzed against water, concentrated with Amicon ultrafilters, and electrophoresed or kept at 4 °C until use. The SDS-PAGE analysis carried out on these PEI-M products (Supporting Information Figure 2) confirmed that they were heterogeneous. Now, we carried out the conjugation reaction of the RNase A “zerolength” dimers with polyethylenimine. The results (data not shown) were qualitatively similar to those obtained with native RNase A. Considering that the starting dimeric protein species to be PEI-conjugated was actually a mixture of “zero-length” dimers and not the only E9-K66 zl-D as described in ref 34, the product of the conjugation reaction could have obviously been even more heterogeneous than the one obtained with monomeric RNase A. We decided therefore to use the entire, unpurified mixture of PEI-RNase A “zero-length” dimer(s) (PEIzl-D) to determine its enzymatic and biological activity. Enzymatic Assay of the “Zero-Length” RNase A Dimers Conjugated to Polyethylenimine. Table 4 shows the enzymatic activities of the PEI-zl-D on dsRNA and ssRNA. The assay comprised the action of (i) native RNase A, (ii) the PEI-native RNase A complex (PEI-M), (iii) the unconjugated


Vottariello et al.

Figure 6. Cytotoxic action of the PEI-RNase A monomeric and dimeric complexes on K562 cell line, as a function of protein concentration. Interaction of the various RNase species with cells lasted 24 h. The two RNase A species (PEI-M and PEI-zl-D) were not purified samples. M, monomeric RNase A; zl-D, RNase A “zero-length” dimers. BS-RNase, bovine seminal RNase, PEI-M, PEI-conjugated monomeric RNase A; PEI-zl-D, PEI-conjugated RNase A “zero-length” dimers; PEI, polyethylenimine. PEI alone was used at 50% concentration that of the RNase A species, as it had been estimated to represent about half of the mass of the PEI-RNase A complexes (see text for details). Cell viability was calculated as a percentage of treated versus untreated cells. Each value shown is the mean of three measures from three independent experiments ( s.d.

“zero-length” RNase A dimers (zl-D), (iv) the PEI-RNase A “zero-length” dimers (PEI-zl-D) complex(es), and (v) BS-RNase for comparison. As for the dsRNase activity, while that of PEI-M was only about 30% higher than the activity of native RNase A, the degrading actions of the “zero-length” dimers as such (220% higher), BS-RNase (360% higher), and PEI-zl-D (440% higher) were remarkable, and in line with results presented in this work or elsewhere (22-24, 27, 45, 47). The same is true for the results obtained with yeast RNA as a substrate, i.e., all RNase A derivatives and BS-RNase showed lower activity values in comparison with the activity of native RNase A. This result is also in agreement with all previous available data, and with the interpretations advanced elsewhere (24, 42, 47). PEI alone (not shown) was absolutely inactive. In conclusion, the relationship between the RNase basicity and the increased activity on dsRNA or the decreased activity on ssRNA was once again confirmed here. Indeed, the RNase A derivatives are more basic than native RNase A, as demonstrated by the chromatographic pattern of Figure 5, although their pI values are missing. Instead, the pI of BS-RNase is known to be at pH 10.3 (versus the RNase A pI at pH 9.6) as shown in 1972 by D’Alessio et al. (6), who demonstrated the higher basicity of BS-RNase compard to RNase A. Cytotoxicity on K562 Cells of RNase A Monomers and “Zero-Length” Dimers Cationized by Conjugation with Polyethylenimine. The cell line used in the following experiments was K562, which, as mentioned above, is sensitive to BS-RNase. Figure 6 shows the results obtained in a 24 h test by comparing the cytotoxic actions of various concentrations of the PEI-M and the PEI-zl-D complexes with the cytotoxic actions of native RNase A, BS-RNase (both tested in single doses), and three concentrations of PEI alone. We have chosen the amount of PEI (MW 1300 Da) to be tested alone as the one theoretically corresponding to the amount of the substance (five molecules) bound to 50% of the side-chain carboxyl groups present in RNase A. It clearly appears that PEI-M and PEIzl-D showed strong but quite similar cytotoxicity values,

whereas PEI alone was inactive at 25 µg/mL and slightly cytotoxic at 50 or 75 µg/mL. RNase A (150 µg/mL) was inactive and BS-RNase slightly active at 50 µg/mL. Additional experiments as a function of time were performed using two different doses of all agents tested above, i.e., 10 µg/mL (Supporting Information Figure 3A) and 25 µg/mL (Supporting Information Figure 3B). Only PEI-M and PEI-zl-D showed a significant but quite similar cytotoxic action at a concentration of 10 µg/mL, while very modest, if any, was the cytotoxic activity of BSRNase under identical conditions. At the concentration of 25 µg/mL, stronger but again quite similar to each other were the actions of both PEI-M and PEI-zl-D, whereas the cytotoxic activity of BS-RNase was now evident but still rather modest. In conclusion, the RNase A “zero-length” dimers cationized with polyethylenimine did not acquire a cytotoxic action higher than that of the PEI-cationized monomeric RNase A. Moreover, we also tested the latter complex on human monocytes, and Figure 7 shows the results obtained. It is quite clear that both the unpurified and purified PEI-RNase A complexes, at the concentration of 25 µg/mL, showed a remarkable cytotoxicity toward monocytes under conditions where BS-RNase (50 µg/mL), native RNase A (100 µg/mL) or PEI alone (12.5 µg/mL) were substantially inactive. Therefore, the cytotoxic action of monomeric RNase A (and presumably that of its “zero-length” dimers) conjugated to PEI appears not to be specific for tumor cells, which drastically decreases the significance of a potential therapeutic use of ribonucleases cationized with polyethylenimine. Why are the Non-Cationized “Zero-Length” Dimers of RNase A Not Cytotoxic? While native, monomeric RNase A is not cytotoxic (24, 30-32), other members of the pancreatictype ribonuclease superfamily display, often along with other biological actions, remarkable activity against malignant tumor cells (1, 7, 8, 51-55). Cytotoxic RNases usually present a high basic charge density and the ability to evade the cytosolic ribonuclease inhibitor (cRI). The former is a feature also linked to the ability of a ribonuclease to attack dsRNA: the higher the number of positive charges present in the proximity of the



protein, which could maintain its enzymatic activity against both ss- and dsRNA, are facts that point out the substantial complexity of the matter (65). Moreover, since from the present work it appears that the amide bond induced by the procedure outlined by Simons et al. (34, 36) may intervene not only between Lys66 and Glu9 (34), but also between amino and carboxyl groups of other amino acid residues, the reciprocal location of the protein subunits in the dimer can vary remarkably. Therefore and in conclusion, the compactness and the possibly variable features of the structure of the RNase A “zero-length” dimers may be their most characteristic structural aspect, responsible, at least partly, for the absence of cytotoxicity in them, making these dimers different in this respect from other natural or artificial ribonuclease dimers.

ACKNOWLEDGMENT

Figure 7. Action of polyethylenimine-cationized RNase A on human monocytes. Monocytes were plated in 96-well plates at 2 × 104/well and incubated overnight. Then, the following substances were added: BS-RNase, 50 µg/mL; native RNase A, 100 µg/mL; purified or unpurified native PEI-RNase A complex(es), 25 µg/mL; PEI, 12.5 µg/ mL. The cells were treated for 24 h at 37 °C. Cell viability was calculated as a percentage of treated versus untreated cells. Each value shown is the mean of three measures from three independent experiments ( s.d.

enzyme active site, the higher the ability of the RNase to degrade dsRNA (17, 22, 24, 27, 32, 45). It has to be recalled here that the preservation of enzymatic activity also seems to be mandatory for a ribonuclease to be cytotoxic or, generally, biologically active (24, 30, 33, 56). It is also worth mentioning that enzymatic activity and cytotoxicity cannot be proportionally related to each other (57, 58). This evidence suggests that ribonuclease cytotoxicity could actually be ruled by several factors, among which is the possibility for the enzyme protein to evade interaction with the cytosolic ribonuclease inhibitor. However, several exceptions to the “rule” that only RNases able to evade interaction with cRI are cytotoxic were recently found (33, 59, 60). Therefore, it could not be surprising that the “zero-length” dimers of RNase A are ineffective against tumor cells notwithstanding they are not trapped by cRI (34) and are enzymatically active toward ss- and dsRNA (ref 34, and present work). The conformational stability (61) and, above all, the location of the positive charges on an RNase molecule (62) can be determinant for it to be biologically active (61, 62) considering either the efficacy of the RNase-RNA interaction (32, 45, 61, 62) or the productive interactions that the protein can build with the cell membrane in order to be internalized (62-64). As far as the RNase A zl-D (34, 36) are concerned, it must be recalled that their molecular structure is unknown. On the basis of the study of Simons et al., it appears improbable for their structure to be similar to any of the well-known dimeric structures of RNase A (9-11, 13, 23-28) or even to that of the cross-linked dimers constructed by Wang, Wilson, and Moore in 1976 (14). Indeed, in this last case the diimidoester linker leaves remarkable freedom to the two protein molecules linked to each other. Instead, the amide bond of the RNase A zl-D makes the vicinity of the two protein subunits quite tight. In other words, these novel dimers should be characterized by a relatively strong immobility, high rigidity, and lower symmetry compared to other known dimeric ribonuclease structures (13, 28, 29). The structural characteristics of the RNase A zl-D might be a key in understanding why this novel type of RNase A dimer is not cytotoxic. On the other hand, the evident accessibility of its active site to substrates, and the positive charge density of the

This work was supported by the Italian Ministero per l’Istruzione e la Ricerca Scientifica and the University of Verona, Verona, Italy (Grant: ex60%-Gotte07). We deeply thank Dr. Monica Vincenzi, the Medical Faculty of the University of Verona, for the DNA sequencing experiments. Supporting Information Available: Supporting graphs showing (1) Superimposed Cation-Exchange chromatographies of native RNase A and its two mutants (K66A and E9A); (2) SDS PAGE (18% polyacrilamide) of PEI-RNase A complex(es); (3) Kinetics of the cytotoxic action of monomeric or PEI-“zerolength” dimeric RNase A complexes on cultures of K562 cell line. This material is available free of charge via the Internet at http://pubs.acs.org.

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