Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M405655200 on June 9, 2004

J. Biol. Chem., Vol. 279, Issue 35, 36753-36760, August 27, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/35/36753    most recent
M405655200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sica, F.
Right arrow Articles by Mazzarella, L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sica, F.
Right arrow Articles by Mazzarella, L.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Structure and Stability of the Non-covalent Swapped Dimer of Bovine Seminal Ribonuclease

AN ENZYME TAILORED TO EVADE RIBONUCLEASE PROTEIN INHIBITOR*

Filomena Sica{ddagger}§, Anna Di Fiore{ddagger}, Antonello Merlino{ddagger}, and Lelio Mazzarella{ddagger}§||

From the {ddagger}Dipartimento di Chimica, Università degli Studi di Napoli "Federico II," Via Cynthia, 80126 Napoli, Italy, the §Istituto di Biostrutture e Bioimmagini, CNR, Via Mezzocannone 6, 80134 Napoli, Italy, and the Dipartimento di Scienze Farmaceutiche, Università degli Studi di Salerno, Via ponte don Melillo, 84084, Fisciano (Sa), Italy

Received for publication, May 20, 2004 , and in revised form, June 8, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
A growing number of pancreatic-type ribonucleases (RNases) present cytotoxic activity against malignant cells. The cytoxicity of these enzymes is related to their resistance to the ribonuclease protein inhibitor (RI). In particular, bovine seminal ribonuclease (BS-RNase) is toxic to tumor cells both in vitro and in vivo. BS-RNase is a covalent dimer with two intersubunit disulfide bridges between Cys31 of one chain and Cys32 of the second and vice versa. The native enzyme is an equilibrium mixture of two isomers, MxM and M=M. In the former the two subunits swap their N-terminal helices. The cytotoxic action is a peculiar property of MxM. In the reducing environment of cytosol, M=M dissociates into monomers, which are strongly inhibited by RI, whereas MxM remains as a non-covalent dimer (NCD), which evades RI. We have solved the crystal structure of NCD, carboxyamidomethylated at residues Cys31 and Cys32 (NCD-CAM), in a complex with 2'-deoxycitidylyl(3'-5')-2'-deoxyadenosine. The molecule reveals a quaternary structural organization much closer to MxM than to other N-terminal-swapped non-covalent dimeric forms of RNases. Model building of the complexes between these non-covalent dimers and RI reveals that NCD-CAM is the only dimer equipped with a quaternary organization capable of interfering seriously with the binding of the inhibitor. Moreover, a detailed comparative structural analysis of the dimers has highlighted the residues, which are mostly important in driving the quaternary structure toward that found in NCD-CAM.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
The RNA population in cells is controlled post-transcriptionally by ribonucleases (RNases) of varying specificity. These secretory enzymes operate at the cross-roads of transcription and translation by catalyzing RNA degradation (1, 2). RNases exhibit diverse biological activities, including specific toxicity to cancer cells (3). RNase activity in serum and cell extracts is elevated in a variety of cancers and infectious diseases. The antitumor effect of ribonucleases has been studied using pancreatic RNase (RNase A) and seminal RNase (BS-RNase)1 from Bos taurus, and onconase and RC-RNase from amphibians (2, 3–6). RNase A does not display appreciable cytotoxic effects toward tumor cells, whereas BS-RNase (6–9), onconase (6, 10–12), and RC-RNase (13) are effective cytotoxic agents both in vitro and in vivo. It is generally accepted that the cytoxicity of these RNases is mainly related to their ability to evade the cytoplasmic RNase inhibitor (RI) and to degrade cellular RNA (5, 14). The antitumor activity exhibited by cytotoxic RNases varies depending on the type of tumor. Factors that could determine susceptibility are the ability of RNases to bind to the cell surface and the efficiency of their internalization (15).

In the vertebrate RNase family, for which RNase A is the prototype, BS-RNase holds a unique position, as it is the only native dimeric member, and the two subunits are cross-linked by disulfide bonds between Cys31 of one chain and Cys32 of the other and vice versa (16). Another unique feature is the coexistence of two dimeric forms: MxM, in which the two subunits interchange the N-terminal helical segment (residues 1–15) through a linker peptide spanning residues 16–22 (hinge peptide) and M=M with no interchange (17). Despite the swapping of the N termini, the two isomers have been shown by x-ray analysis to have similar quaternary structure (18–20), so that they are practically indistinguishable by their external shape. The MxM form, accounting for about two-thirds of protein molecules, has allosteric properties, whereas M=M shows a typical Michaelis kinetics (17, 21, 22). Moreover, the swapped form is endowed with an unusual multiplicity of biological action: it exhibits antispermatogenic, antiviral, embryotoxic, immunosuppressive, and antitumor properties (7, 16). Only a minor antitumor action has been detected for M=M (23). Since both MxM and M=M forms cannot be accommodated into the cavity of RI, the different behavior of the two isomers has been explained with their different fate inside the cell (24). In the reducing environment of the cytosolic compartment, M=M dissociates into monomers, which readily bind to RI. By contrast, in MxM the swapping of the N termini is sufficient to stabilize the dimeric form even when the interchain disulfide bridges are broken; in this case a non-covalent dimer (NCD) is formed, which is supposed to be the species responsible for the unique biological properties of MxM (17). This explanation of the remarkable difference in the biological properties of the two isomers implies that NCD would adopt a quaternary structure capable of evading RI.

Beside BS-RNase, other ribonucleases, such as RNase A (25, 26) and human pancreatic ribonuclease (HP-RNase) (27), are able to form in vitro non-covalent dimers through the swapping of a structural element among subunits. In particular, under well defined experimental conditions, monomeric RNase A produces two different non-covalent dimeric forms (26), both studied crystallographically: ND-RNase A, where the N-terminal {alpha}-helices are interchanged (28) and CD-RNase A, where the swapping involves the C-terminal {beta}-strands (29). Furthermore, the x-ray analysis of a variant of monomeric HP-RNase, in which the region 1–22 and residue 101 are substituted for those of BS-RNase, revealed the presence in the crystal of an N-terminal domain-swapped dimer (PM8), whose quaternary organization is different from ND-RNase A (27). The HP-RNase variant does not exhibit cytotoxic activity (27), and the two dimers of RNase A show low antitumor activity compared with BS-RNase (30). These results raise the problem of clarifying how the two subunits are assembled in the non-covalent dimer obtained from MxM.

In this study, we report the crystal structure of NCD, carboxyamidomethylated at the residues Cys31 and Cys32 (NCD-CAM) and complexed with 2'-deoxycitidylyl(3'-5')-2'-deoxyadenosine (dCpdA), at 2.0 Å resolution. The thermal and kinetic stability of this BS-RNase form in solution has also been investigated. Despite the lack of disulfide bridges, the quaternary assembly of NCD-CAM turned out to be much closer to MxM than to ND-RNase A and to PM8. Furthermore, model building of the complex between RI and the N-terminal swapped dimeric RNases show that NCD is the only known non-covalent dimer whose quaternary structure hampers the formation of the complex with RI. The comparative analysis of RNase A, PM8, and NCD-CAM structures has also allowed the identification of key residues that are important to stabilize for the latter enzyme a quaternary structure, which is capable of evading the protein inhibitor.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Materials—RNase A (type XII-A) from bovine pancreas, the oligonucleotide dCpdA, as well as yeast RNA were purchased from Sigma and used without further purification. Other chemicals were of the highest purity available. Superdex G-75, Superdex 75 10/30, and Source 15S HR 10/10 columns were from Amersham Biosciences. A column of Superdex G-75 was calibrated with molecular mass standards from Bio-Rad and Sigma.

NCD-CAM Preparation—The BS-RNase non-covalent dimer was prepared according to the protocol previously described by Piccoli et al. (17). Briefly, a sample of native mixture of the BS-RNase was dissolved in 0.1 M Tris-HCl buffer, pH 8.4, to the final concentration of 7.1 mg/ml. The interchain disulfide bridges were selectively reduced by adding, under a nitrogen flux, a 10-fold molar excess of dithiothreitol with respect to the protein molar concentration. The sulfurs of cysteines 31 and 32 were then alkylated in the dark with iodoacetamide, which was added to the reaction mixture in 2-fold molar excess with respect to dithiothreitol. This treatment furnishes two BS-RNase derivatives: the monomer and the non-covalent dimer, which were obtained from the M=M and MxM, respectively. The products were separated at 4 °C by HPLC (model 410 BIO distributed by PerkinElmer Life Sciences) gel-filtration chromatography, on a column of Superdex G-75, equilibrated, and eluted with 0.2 M ammonium acetate buffer, pH 5.1.

Protein homogeneity was judged by native and denaturing gel electrophoresis performed on a Phast System (Amersham Biosciences). The native electrophoresis of the purified NCD-CAM revealed a single band with a molecular mass comparable to that of the BS-RNase control. The denaturing gel showed, instead, a single band located at the height of the monomeric RNase A standard. The ribonucleolytic activity was measured by the Kunitz assay (31).

ND-RNase A Preparation—The oligomerization of RNase A was obtained, as previously described (26), by lyophilizing a solution of the enzyme (5 mg/ml) in 40% acetic acid. Briefly, the aqueous acetic acid solution of RNase A was first kept at room temperature for 60 min under stirring, then frozen in liquid nitrogen, and lyophilized overnight. The lyophilized sample was dissolved in 0.2 M sodium phosphate buffer, pH 6.7, at a protein concentration of 30 mg/ml and was subjected to ion exchange chromatography, carried out with an Amersham Biosciences Source 15S HR 10/10 column. The monomeric enzyme was eluted using 0.085 M sodium phosphate buffer (pH 6.7) at the flow rate of 0.4 ml/min, whereas the two dimers, ND-RNase A and CD-RNase A, were separated using a 0.085 ± 0.180 M sodium phosphate (pH 6.7) linear gradient.

Crystallization and Diffraction Data Collection—The non-covalent dimer of BS-RNase was crystallized in the presence of dCpdA at 4 °Cby using the sitting drop vapor diffusion method. The best crystals of NCD-CAM were grown from a crystallization solution containing 22% (w/v) PEG 8000, 5% (v/v) isopropyl alcohol, 0.2 M sodium acetate, and 0.1 M cacodilate buffer, pH 6.5. In particular, a protein sample (5 mg/ml) containing a 10-fold molar excess of the ligand (1.8 mM) was mixed with an equal volume of the precipitant solution and equilibrated against the reservoir in 2-µl drops.

The x-ray data collection has been performed at the ELETTRA Synchrotron (Trieste, Italy) using a 30-cm diameter MAR-Research image plate system. Prior to data collection, the crystal was transferred into a cryo-protectant solution containing a higher concentration of PEG 8000 (24% w/v) in comparison with the mother liquor and 15% (v/v) glycerol as cryogenic agent, and frozen at 110 K under a N2 stream. Data were indexed, processed, and scaled with the DENZO/SCALEPACK package (32). Intensities were truncated to amplitudes using the TRUNCATE program (33).

The crystal diffracted up to 2.0 Å and belongs to space group P212121 with unit cell dimensions of a = 41.47 Å, b = 69.69 Å, and c = 110.80 Å. Assuming one molecule per asymmetric unit, the Matthews coefficient (Vm) (34) is estimated to be 2.82. This value corresponds to a solvent content of 56.4%. Details of the data statistics are given in Table I.


View this table:
[in this window]
[in a new window]
  		TABLE I
Data collection and refinement statistics

 
Structure Solution and Refinement—The crystal structure was solved by the molecular replacement method with the X-PLOR program.2 As a search model, a structural unit (residues 1–15 of one subunit and 23–124 of the second one) of MxM (PDB code 1BSR [PDB] ) (18) was used, as this unit was supposed to be the conservative region of the molecule. The cross-rotation function, calculated using data in the 10–3.4 Å resolution range, clearly shows two prominent peaks, which were ascribed to the two protein chains. The two maxima are related by a non-crystallographic 2-fold rotation, in agreement with the results of the self-rotation function. After a run of the X-PLOR PC-Refinement routine,2 the dimer was finally assembled in the asymmetric unit through a stepwise translation search.

Few cycles of rigid body refinement of the two structural units were followed by several runs of positional and temperature refinement alternated with manual building using the O program (36). The building of the two hinge peptides was not difficult as the associated Fo–Fc electron density was well defined. A randomly selected 10% of data was set aside as a free data set. Reflections with Fo<2{sigma}(Fo) were also excluded from the structure refinement. Water molecules were placed at stereochemically reasonable positions if they displayed electron density in the Fo–Fc map greater than 3{sigma}.

The final model contains 1898 protein atoms, 74 nucleotide atoms, and 120 water molecules, and has an Rfactor and an Rfree of 20.9 and 27.3%, respectively. The correctness of stereochemistry was finally checked using PROCHECK (37); calculations of r.m.s.d. values from ideal values for bonds, angles, dihedral, and improper angles were performed using X-PLOR. The relevant parameters of the refinement are presented in Table I.

Circular Dichroism Analysis—The thermal denaturation of NCD-CAM, monomeric carboxyamidomethylated derivative of BS-RNase (MCAM), RNase A, and ND-RNase A was monitored by using CD spectroscopy. The spectra were recorded with a Jasco J-710 spectropolarimeter equipped with thermostatic cuvette holder (JASCO PTC-348).

Specifically, the minimum centered around 211 nm of a 0.2 mg/ml solution of ribonuclease was monitored as the temperature of the solution was increased from 4 to 73 °C with a scan rate of 0.5 °C/min. Data were fitted to a two-state transition model, and the resulting curve was used to calculate the value of Tm, which corresponds to the temperature at the midpoint of the thermal denaturation.

Kinetics of Thermal Dissociation of NCD-CAM—The dissociation kinetics of NCD-CAM and ND-RNase A were studied by incubating solutions of protein (0.42 mg/ml) in 50 mM Tris-HCl buffer, pH 7.5, containing 130 mM NaCl, at 37 °C. At various time intervals, aliquots of the samples were withdrawn, cooled, and analyzed by gel filtration on an Amersham Biosciences FPLC system, equipped with a Superdex 75 10/30 column, equilibrated with 0.1 M Tris-HCl buffer, pH 7.5, containing 0.3 M NaCl (flow rate 0.6 ml/min). Elution was performed at room temperature. The amount of the dimeric enzyme, measured as a percentage of the initial value, was monitored as function of time.

Subunit Superposition and Construction of the Complex with the Protein Inhibitor—For superposition and comparison of the various chains, only the C{alpha} carbon atoms of residues 25–35, 40–60, 72–86, 95–110, and 116–123 were used. This selection includes the {beta}-sheet core of the protein and two {alpha}-helices. Exposed loops and the N-terminal helix involved in the swapping were not included in the calculations. For the detection of the breathing skeleton motion of each subunit, the V-shaped {beta}-sheet motif was separated in two arms, V1 and V2, which includes the C{alpha} carbon atoms of residues 61–63, 71–75, 105–111, 116–124, and 42–46, 82–87, 96–101, respectively.

Models of PM8, ND-RNase A, and NCD-CAM complexed with the protein inhibitor RI were obtained by superimposition of the atomic coordinates of one subunit of each dimer onto the crystallographic coordinates of RNase A bound to RI (Ref. 38; PDB access code 1DFJ [PDB] ). In the case of NCD-CAM and PM8, which possess a stringent 2-fold symmetry, the choice of the chain to superimpose on the RNase A coordinates is not relevant, and only one model was analyzed. On the contrary, for ND-RNase A (28), which deviates considerably from the 2-fold symmetry, two different models can be built, using either chain A or chain B (as reported in the PDB file, access code 1A2W [PDB] ).


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
CONCLUSIONS
 REFERENCES
 
Enzyme Characterization—Among the swapped proteins, the MxM isomer of BS-RNase represents a singular case because the subunit assembly is governed by two different types of constraints: the swapping of the N termini and the two consecutive antiparallel intersubunit disulfide bridges (18, 20). The selective reduction of these disulfides by titration with dithiothreitol, followed by alkylation with iodoacetamide, produces a non-covalent dimer (NCD-CAM), whose quaternary structure is held together only by the swapping (17). The nature of NCD-CAM has been assessed by gel electrophoresis under denaturing and non-denaturing conditions, in which this derivative migrates as a single band with a molecular mass of 14 and 28 kDa, respectively. The enzymatic activity of NCD-CAM, determined on yeast RNA as a substrate, in agreement with previous data (17), is almost twice that of the native enzyme and comparable with that of MCAM (39).

The conformational stability of NCD-CAM has been measured by CD spectroscopy in the range from 4 °C to 85 °C and compared with that of MCAM, RNase A, and ND-RNase A. The reversible thermal transitions of the various enzymes have been fitted to equations describing a two-state model. The values of Tm, obtained at the midpoint of the thermal denaturation curve, are 56.2 and 61.7 °C for the monomers MCAM and RNase A, respectively, 56.0 and 61.9 °C for the corresponding dimers, NCD-CAM and ND-RNase A. These data indicate that the dimers have the same conformational stability of the monomeric counterpart thus suggesting that the dissociation of the oligomers precedes the onset of the thermal unfolding.

The stability of NCD-CAM as a dimer has also been determined at 37 °C and compared with that of ND-RNase A. The amount of the dimeric protein, recovered by gel filtration at regular time intervals, was fitted with first order plots and gave t1/2 of 7 and 24 h for NCD-CAM and ND-RNase A, respectively. For the latter enzyme the t1/2 obtained in our experiments is practically consistent with literature data (26). Despite the higher dissociation rate at 37 °C, NCD-CAM is stable up to 2 months at 4 °C as judged by non-denaturing gel electrophoresis. Thus, crystals of this dimer could be grown at 4 °C from a freshly prepared sample without serious problems.

General Structure—The structure of NCD-CAM in complex with dCpdA has been refined using diffraction data extending up to 2.0 Å resolution. The final model comprises two subunits, which interchange their N termini, as in native MxM isomer (Fig. 1). The electron density map is well defined for all residues (see Fig. 2A), including the four carboxyamidomethylated cysteine residues (Ycm) 31 and 32 (Fig. 2B), which form the interchain disulfide bridges in the native protein.



View larger version (38K):
[in this window]
[in a new window]
  		FIG. 1.
Ribbon representation of the NCD-CAM peptide fold (chain A in light gray and chain B in dark gray). The carboxyamidomethylated side chains of Cys31 and Cys32 and the nucleotides are drawn in ball-and-stick representation. The figure was prepared with MOLSCRIPT (59).

 



View larger version (68K):
[in this window]
[in a new window]
  		FIG. 2.
3 {sigma}(I) level omit Fo–Fc maps, seen along the molecular local 2-fold axis. Hydrogen bonds are represented by dotted lines. Both chains A and B are shown: the hinge region (A); the four carboxyamidomethylated Cys residues (Ycm) (B), which form the interchain disulfides in the intact protein; a water molecule sits on the local axis and bridges the side chains of Ycm32 of the two chains. The figure was drawn with BOBSCRIPT (35).

 
Superposition of the C{alpha} carbon atoms of the two subunits, with exclusion of the swapped helix and exposed loops (see "Experimental Procedures" for details) gives an r.m.s.d. of 0.28 Å (0.41 for all residues). Similar values are found when each subunit is compared with those of free or liganded MxM. In all cases the largest variations are associated with the hinge peptide.

Because of the swapping, NCD-CAM has composite active sites, in which one residue (His12) of the catalytic triad belongs to one subunit, whereas the other two (His119 and Lys41) belong to the second subunit. The overall architecture of the active sites is virtually identical to that of MxM (18, 20). The two dCpdA inhibitors are well defined in the electron density, and their conformation is similar to that observed in the complex with MxM (20) (PDB code 1R5C [PDB] ) and RNase A (40) (PDB code 1RPG [PDB] ). Cytosine and adenosine moieties adopt the anti conformation. The ribose rings present the C4'-exo and C3'-endo puckering, respectively. The binding of the bases in the B1 and B2 sites is canonical (41). In particular, cytosine in B1 is H-bonded to Thr45 and to two water molecules and forms stacking interactions with Phe120. The phosphate moiety makes contacts with the side chains of the catalytic triad, with the main chain of Phe120 and with the side chain of Gln11. In B2, Asn71 and Asn67 are H-bonded to adenosine, which is also involved in stacking interactions with the side chains of His119 and Gln69.

A structural feature of RNases is a domain breathing motion corresponding to an adaptation of the protein skeleton to changed environmental conditions (42–46). In particular, upon binding of a ligand, the two arms V1 and V2 of the {beta}-sheet core (see "Experimental Procedures" for details) modify slightly their relative orientation to produce a more compact structure (42–46). This effect can be evidenced for each chain of the present structure in comparison to those of the free MxM (18, 20). Indeed, in a chain-to-chain comparison, after superimposition of V1, a further rotation of about 4° is required to best-fit V2. The rotation parameters are in good agreement with those calculated when the free and liganded MxM are compared.

Quaternary Structure—The quaternary structure displays a well preserved 2-fold symmetry. The parameters defining the chain assembly are given in Table II, together with those of two different crystal structures of MxM (18, 20) and other non covalent N-swapped dimers of RNases, such as PM8 (27) and ND-RNase A (28). The free and complexed forms of MxM were also included in Table II to give an indication of the degree of variability of the quoted parameters. NCD-CAM was first aligned with its dyad axis parallel to z, and then the other dimers were rotated to superimpose one chain. In Table II, {chi} measures the internal symmetry, r.m.s.d. is the discrepancy between the two subunits in a dimer, and {psi} is the co-latitude of the rotation axis with respect to the NCD-CAM 2-fold axis. {psi}' is the co-latitude with respect to MxM. 2 {psi} is the misalignment of the second subunit of a dimer in comparison to the reference structure, after the superposition of the first subunit. The variety of quaternary structures is best grasped by looking at the models drawn in Fig. 3A, where all the dimers are seen with one subunit in a common orientation. The dyad axis of NCD-CAM is rotated only 19° with respect to that of the covalent MxM dimer. PM8 also displays a strict 2-fold symmetry; however, the molecular axis is oriented 76° away from that of MxM. ND-RNase A has a lower symmetry, and the two subunits are related by a rotation of 162°; the orientation of the axis is very different from the other dimers as can be seen from the values of {psi} and {psi}' (in addition, the symmetry axis of ND-RNase A forms an angle of 65° with that of PM8).


View this table:
[in this window]
[in a new window]
  		TABLE II
Quaternary structures of ribonuclease swapped dimers

 



View larger version (45K):
[in this window]
[in a new window]
  		FIG. 3.
A, C{alpha} atom tracing of NCD-CAM quaternary structure (red) compared with that of MxM (cyan; PDB code 1BSR [PDB] ), ND-RNase A (blue; PDB code 1A2W [PDB] ), and PM8 (green; PDB code 1H8X [PDB] ). One subunit was used for the superimposition of the dimers. The trace of the symmetry axis for each dimer is also shown. B, van der Waals representation of Pro19 side chain (green) inserted into a cavity of the partner subunit: a contribution to the O-interface in the swapped NCD-CAM dimer. C, superimposition of hinge peptides of NCD-CAM (red) and PM8 (green; PDB code 1H8X [PDB] ). D, model of the complex between NCD-CAM and RI. The drawings were prepared with MOLSCRIPT (59).

 
The accessible area was also evaluated for the five dimers (Table II). The differences are clearly small, as expected on the basis of the elevated homology and strict similarity of their tertiary structure. However, for the same reasons, they were highly sensitive to the quaternary changes and, indeed, the calculated values are clearly correlated to the relative stability of the dimers. Thus, ND-RNase A and PM8 have the lowest and the largest exposed surface, respectively, while NCD-CAM falls somewhat in the middle and its accessible area is comparable to that of the covalent dimer MxM.

Thus, despite the lack of the two interchain disulfides, the quaternary assembly of NCD-CAM is closer to the intact MxM dimer rather than to ND-RNase A and to PM8. When NCD-CAM is compared with MxM, the amount of rotation required to superimpose the second subunit is about 40°. The rotation axis is approximately normal to the molecular dyad and passes close to the midpoint between the Leu28 residues of the two subunits. Thus the movement, which follows the disruption of the covalent linkages, increases by about 3 Å the distances between the cysteine residues involved in the interchain disulfides and decreases the distances between the hinge peptides. It leaves almost unperturbed the interactions between the Leu residues across the molecular dyad.

Hinge Peptides—In the non-swapped forms of BS-RNase (M=M dimer, Ref. 19, and monomeric species, Ref. 39) the hinge peptide was considerably disordered so that it was not possible to build a full model. In the various crystal structures of the swapped MxM (18, 20, 42, 47), the peptide is much more ordered, although it adopts different conformations for the two chains. In all cases, these conformations are characterized by the presence of a type I {beta}-turn, which encompasses residues 18–21 and is highly favored by proline in position 19 (48). In NCD-CAM (Fig. 2) this feature is maintained (Ser18-O–N-Ser21 distance is 2.93 Å in both subunits) and extended to include a hydrogen bond interaction between Ser18 N and Ser21 O (3.23 and 3.24 Å in the two subunits, respectively). In addition, the side chain atoms OD1 of Asn17 and OG of Ser23 are 2.70 Å apart (2.58 Å in the second chain). The tight {beta}-hairpin structure is followed by a type I {beta}-turn (residues 22–25), which then evolves into the helix 25–35, an invariant feature of the ribonuclease folding. In addition to the three hydrogen bonds formed within each hinge peptide, the two peptides of the dimer tightly pack across the molecular axis and form two additional main chainmain chain hydrogen bonds between the nitrogen atom of Ser21 of one subunit and the carbonyl oxygen of Pro19 of the second chain and vice versa. Among the interchain interactions, of particular interest are those involving the pirrolydine ring of Pro19, which fits well into a cavity formed by the side chains of Tyr25 and Gln101 of the other subunit (Fig. 3B). This interchain complementary at the level of Pro19 is also a characteristic of MxM (18, 20), the swapped isomer of BS-RNase. In the unswapped forms of BS-RNase (M=M, Ref. 19 and monomeric species, Ref. 39) the disorder of the hinge peptide is probably caused by the absence of low energy unswapped conformations, which could satisfy the additional constraint of properly locating the side chain of proline in the cavity.

Interestingly, in PM8 (27), which has residues 1–20 and 101 equal to those of the seminal enzyme, the two hinge peptides present structural features very similar to those of NCD-CAM (Fig. 3C) up to Pro19, whose side chain is located in the cavity formed by Tyr25 and Gln101 of the partner chain. However, a dramatic change is localized at residues 20–22, which, together with Pro19, form one turn of a helix. Thus the following {alpha}-helix, starting at residue 25, takes a completely different orientation and accounts for the differences in the quaternary structure of the two enzymes.

In the case of ND-RNase A (28) the substitution of Pro19 for Ala increases the flexibility of the hinge peptide (49), which allows a more efficient optimization of the contact surface between the two subunits. This is eventually maximized through a non-symmetric pairing of the {beta}-sheets, which produces highly stabilizing contacts. The two hinge peptides are then forced to adopt different conformations, and one of them folds in an {alpha}-helix, which would be disfavored if a proline were present in position 19.

O-Interface—In the domain-swapping terminology, the interface found only in the oligomer is termed O-interface, whereas the interactions found in the monomer, and recreated by two polypeptide chains in the oligomer, form the C-interface (50).

In NCD-CAM, the O-interface is formed by the two hinge peptides and the following {alpha}-helices. The interhinge contacts have already been discussed in a previous section. Additional contributions to the O-interface are given by the hydrogen interactions involving the carbonyl oxygen of Asn17 and OG of Ser20 of one chain and the NE2 of His48 and NE2 of Gln101 of the second chain, respectively. All these interactions are also present in PM8 (27). The most outstanding interactions between the two {alpha}-helices in NCD-CAM are provided by the Leu28 side chains of the two subunits, which face each other across the molecular axis and are completely buried at the interface, as it occurs in the structure of MxM (18, 20). This contact region is also the pivoting point for the reorientation of the chains that follows the rupture of the interchain disulfides of MxM. Despite this rotation, the side chains of Ycm31 of one chain and that of Ycm32 of the second chain, and vice versa, are in contact across the molecular axis through a water molecule. The helix-helix interaction is lost in PM8 (27), which however presents an additional contribution to the O-interface through a partial pairing of the {beta}-sheets across the 2-fold axis. This pairing produces two salt bridges between Glu103 of one chain and Arg104 of the second and vice versa. A more efficient pairing of the two {beta}-sheets is achieved in the asymmetric ND-RNase A (28), which is stabilized by several interchain H-bonds (28, 51).

Protein Inhibitor Binding—Models of PM8, ND-RNase A, and NCD-CAM complexed with the protein inhibitor RI were obtained by superimposition of the atomic coordinates of one subunit of the dimers on the coordinates of RNase A complexed with RI (38). As pointed out by Kobe et al. (37) the complex is characterized by an extensive intrusion of the exposed and highly flexible loops 35–39 and 88–94 of RNase A into the cavity of the inhibitor molecule, lined by residues of the loops L2, L3, L6, and L8. The interactions are essentially electrostatic in nature, and the interacting surfaces do not show elevated shape complementary, thus suggesting that in the complex the orientation of the ribonuclease moiety may have a certain degree of flexibility.

One subunit of PM8, the swapped variant form of HP-RNase, could be easily accommodated into RI horseshoe cavity, and the partner chain does not produce steric overlap with RI. In the case of ND-RNase, the two subunits are not related by a 2-fold symmetry, therefore two different models can be built, depending on which subunit is chosen to fit the coordinates of RNase A moiety of the complex. In one case the second subunit strongly interpenetrates the inhibitor molecule, whereas in the alternative model no severe clashes are produced. Few short distances are observed between the loop 76–78 of the mate subunit and the RI region encompassing residues 148–151. However, considering the properties of the complex and, in particular, the flexibility of the ribonuclease loop involved in the complex formation, minor modifications of the interacting loops and a slight reorientation of the dimer may easily release the few repulsive contacts. In addition, the structural characterization of RI, both in the free form and in the form complexed with RNase A (38, 52), has shown that the characteristic horseshoe shape of the molecule can be significantly modified in the presence of an external perturbation. Thus both PM8 and ND-RNase A appear to be capable to interact with RI, as well as their monomeric counterparts.

In contrast, when one subunit of NCD-CAM dimer is fitted on the ribonuclease coordinates of the complex, a large part of the accompanying chain severely penetrates the N-terminal region of the inhibitor (Fig. 3D). This result indicates that inhibition by RI should require a drastic change in the quaternary organization of the dimer.


   CONCLUSIONS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
REFERENCES
 
The explanation of the cytotoxic properties of ribonucleases is basically linked to the hypothesis that these enzymes are left free to act on the cellular RNA by their capacity to evade inhibition by RI (5, 14). This capacity may be built on by dimerization and acquisition of a quaternary structure, which interferes with the binding of the inhibitor. The covalent quaternary structure of native MxM does indeed possess this property (17). The different antitumor activity of the two dimeric forms of BS-RNase has long been associated with the ability of MxM, the covalent swapped isomer of BS-RNase, to preserve a dimeric structure in the reducing environment of the cytosol (7, 24, 53, 54). This implies that the non-covalent swapped dimer of the enzyme is still able to adopt a quaternary association capable to interfere negatively with the inhibitor binding. We have determined the crystal structure of NCD-CAM and used these data to model its complex with RI. The model clearly indicates that the structure adopted by NCD severely hampers the binding of the inhibitor. On the contrary, PM8 can easily bind to RI, whereas ND-RNase A needs only small modifications of exposed and highly flexible loops to optimize the binding. These results are in line with the data of inhibitor binding assays performed on RNase A dimers (26), on PM8 (27), and on M=M, and indicate that these species are RI-sensitive, whereas MxM and NCD are insensitive to the inhibitor (24).

More recently, it has been shown that in a mixture of reduced and oxidized glutathione, which better mimics the cytosolic environment with respect to previous experiments, M=M dissociates into monomers, whereas MxM not only remains a dimer but also has intact interchain disulfides (53). It has been suggested that NCD maintains the proximity of the reduced cysteine residues, thus facilitating the reformation of the intersubunit disulfides (53). This explanation is well sustained by the quaternary shape of NCD revealed by the present work. In conclusion, it appears that, whichever is the effective form of the enzyme, the unique properties of seminal ribonuclease are strictly related to the uniqueness of the quaternary structure of its non-covalent swapped dimeric species.

Why should this particular organization of the NCD-CAM dimer be favored in comparison to that of PM8 and in particular to that of the highly stable structure adopted by ND-RNase A? In our opinion two contributions are equally important. The first is provided by Pro19; this residue plays a role in the destabilization of the compact structure of ND-RNase A by limiting the conformational flexibility of the hinge peptide. Furthermore, in NCD-CAM, its side chain builds on a very stringent complementary interaction on the surface of the partner chain, nicely fitting a cavity lined by Tyr25 and Gln101. This feature is common to PM8. The second contribution is provided by the hydrophobic interactions between the two Leu28 side chains, which would be exposed to the solvent in the PM8 structure (and in ND-RNaseA), but become fully buried in NCD-CAM. In PM8 this position is occupied by a Gln, whose side chain protrudes out of the surface of the protein. Pro19 and Leu28 seem then to act in a cooperative fashion to push the quaternary association of the dimer toward the one found in NCD-CAM. Pro19 prevents the hinge peptide to adopt a (ND-RNase A)-like conformation; Leu28 provides the hydrophobic interaction between the {alpha}-helices, which favors the collapse of a PM8-like quaternary association into that of NCD-CAM.

The structural features of the three non-covalent dimers are reflected in their different stability. The data on the rate of spontaneous dissociation of ND-RNase A and NCD-CAM at 37 °C show that the latter is more readily dissociated. This is paralleled by the larger O-interface and the smaller overall accessible area of ND-RNase A. On the other hand the low stability of PM8 in solution demonstrates the importance of hydrophobic interactions for the stabilization of a more compact dimer with a larger O-interface.

In conclusion, the MxM swapped isomer of bovine seminal ribonuclease is the only dimeric ribonuclease to be found in vivo and to display, both in the covalent and non covalent form, the proper shape to impair any interaction with the protein inhibitor. The evolutionary pathway that has led to such a remarkable structure has been extensively debated in the literature (55–58). The unique feature, provided by the simultaneous presence of two strong quaternary structural constraints, has raised the question of whether the swapping or the interchain disulfides were initially conceived (56–58). The swapping hypothesis has been disputed on the basis of ancestor gene sequence analysis, which indicates that the introduction of a second cysteine (Cys31 in the sequence) could, indeed, have played a central role in the process (56). We believe that the structural features, together with the relative stability of the three dimers discussed above, may provide new insights. Indeed, it seems unlikely that a swapped dimer could have progressively lost stability with respect to ND-RNase A to evolve toward a structure, which had to be stabilized at the end by the interchain disulfides. On the other hand, it is tempting to speculate that the acquisition of the covalent bonds was the primary event in the evolutionary pathway of the protein, in agreement with previous suggestions (48, 56, 58). In this respect, the intrinsic ability of ribonuclease to swap would not be relevant. However, once the covalent linkages had fixed the appropriate quaternary scaffold, this can have exerted a strong pressure and determined a "fast" adaptation of the O-interface toward a stabilization of a swapped quaternary structure that, even in the absence of the covalent restraints, was capable of neutralizing the protein inhibitor.


   FOOTNOTES
 
* This work was supported by Ministero Dell' Istruzione, Università e Ricerca (Grant PRIN2002). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The atomic coordinates and structure factors (code 1TQ9) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/). Back

|| To whom correspondence should be addressed: Dipartimento di Chimica Università di Napoli "Federico II," Complesso Universitario di Monte Sant' Angelo, Via Cynthia, 80126 Napoli, Italy. Fax: 39-081-674090; E-mail: lelio.mazzarella{at}unina.it.

1 The abbreviations used are: BS-RNase, bovine seminal ribonuclease; MxM, dimeric form of BS-RNase in which the chains swap their N-terminal tails; M=M, unswapped dimer of BS-RNase; NCD, non-covalent dimeric-swapped form of BS-RNase; NCD-CAM, S-[Cys31]carboxyamidomethyl, S-[Cys32]carboxyamidomethyl-NCD; RNase A, bovine pancreatic ribonuclease; dCpdA, 2'-deoxycitidylyl(3'-5')-2'deoxyadenosine; PDB, Protein Data Bank; r.m.s.d., root mean square deviation; RI, ribonuclease protein inhibitor; MCAM, monomeric carboxyamidomethylated derivative of BS-RNase. Back

2 A. T. Brunger (1996) Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT. Back


   ACKNOWLEDGMENTS
 
We thank Giuseppe D'Alessio (Dipartimento di Chimica Biologica, University di Napoli, Federico II, Napoli, Italy) and Giuseppe Graziano (Dipartimento di Scienze Biologiche ed Ambientali, Università del Sannio, Benevento, Italy) for critically reading the manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 CONCLUSIONS
 REFERENCES
 

  1. Schein, C. H. (1997) Nat. Biotechnol. 15, 529–536[CrossRef][Medline] [Order article via Infotrieve]
  2. Raines, R. T. (1998) Chem. Rev. 98, 1045–1065[CrossRef][Medline] [Order article via Infotrieve]
  3. Youle, R. J., and D'Alessio, G. (1997) in Ribonuclease: Structures and Functions (D'Alessio, G., and Riordan, J. F., eds) pp. 491–514, Academic Press, San Diego
  4. Leland, P. A., and Raines, R. T. (2001) Chem. Biol. 8, 405–413[CrossRef][Medline] [Order article via Infotrieve]
  5. Makarov, A. A., and Ilinskaya, O. N. (2003) FEBS Lett. 540, 15–20[CrossRef][Medline] [Order article via Infotrieve]
  6. Matousek, J., Soucek, J., Slavik, T., Tomanek, M., Lee, J. E., and Raines, R. T. (2003) Comp. Biochem. Physiol. C Toxicol. Pharmacol. 136, 343–356[CrossRef][Medline] [Order article via Infotrieve]
  7. Kim, J. S., Soucek, J., Matousek, J., and Raines, R. T. (1995) J. Biol. Chem. 270, 10525–10530[Abstract/Free Full Text]
  8. Antignani, A., Naddeo, M., Cubellis, M. V., Russo, A., and D'Alessio, G. (2001) Biochemistry 40, 3492–3496[CrossRef][Medline] [Order article via Infotrieve]
  9. Cinatl, J., Jr., Cinatl, J., Kotchetkov, R., Matousek, J., Woodcock, B. G., Koehl, U., Vogel, J. U., Kornhuber, B., and Schwabe, D. (2000) Anticancer Res. 20, 853–859[Medline] [Order article via Infotrieve]
  10. Mikulski, S. M., Costanzi, J. J., Vogelzang, N. J., McCachren, S., Taub, R. N., Chun, H., Mittelman, A., Panella, T., Puccio, C., Fine, R., and Shogen, K. (2002) J. Clin. Oncol. 20, 274–281[Abstract/Free Full Text]
  11. Mikulski, S. M. (1995) Int. J. Oncol. 7, 1415–1420
  12. Mikulski, S. M., Viera, A., and Shogen, K. (1993) Int. J. Oncol. 2, 807–812
  13. Wei, C. W., Hu, C. C., Tang, C. H., Lee, M. C., and Wang, J. J. (2002) FEBS Lett. 531, 421–426[Medline] [Order article via Infotrieve]
  14. Haigis, M. C., Kurten, E. L., and Raines, R. T. (2003) Nucleic Acids Res. 31, 1024–1032[Abstract/Free Full Text]
  15. Haigis, M. C., and Raines, R. T. (2003) J. Cell Sci. 116, 313–324[Abstract/Free Full Text]
  16. D'Alessio, G., Di Donato, A., Mazzarella, L., and Piccoli, R. (1997) in Ribonuclease: Structures and Functions (D'Alessio, G., and Riordan, J. F., eds) pp. 383–423, Academic Press, San Diego
  17. Piccoli, R., Tamburrini, M., Piccialli, G., Di Donato, A., Parente, A., and D'Alessio, G. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1870–1874[Abstract/Free Full Text]
  18. Mazzarella, L., Capasso, S., Demasi, D., Di Lorenzo, G., Mattia, C. A., and Zagari, A. (1993) Acta Crystallogr. Sect. D 49, 389–402[CrossRef][Medline] [Order article via Infotrieve]
  19. Berisio, R., Sica, F., De Lorenzo, C., Di Fiore, A., Piccoli, R., Zagari, A., and Mazzarella, L. (2003) FEBS Lett. 554, 105–110[CrossRef][Medline] [Order article via Infotrieve]
  20. Merlino, A., Vitagliano, L., Sica, F., Zagari, A., and Mazzarella, L. (2004) Biopolymers 73, 689–695[CrossRef][Medline] [Order article via Infotrieve]
  21. Di Donato, A., Piccoli, R., and D'Alessio, G. (1987) Biochem. J. 241, 435–440[Medline] [Order article via Infotrieve]
  22. Piccoli, R., Di Donato, A., and D'Alessio, G. (1988) Biochem. J. 253, 329–336[Medline] [Order article via Infotrieve]
  23. Cafaro, V., De Lorenzo, C., Piccoli, R., Bracale, A., Mastronicola, M. R., Di Donato, A., and D'Alessio, G. (1995) FEBS Lett. 359, 31–34[CrossRef][Medline] [Order article via Infotrieve]
  24. Murthy, B. S., De Lorenzo, C., Piccoli, R., D'Alessio, G., and Sirdeshmukh, R. (1996) Biochemistry 35, 3880–3885[CrossRef][Medline] [Order article via Infotrieve]
  25. Crestfield, A. M., Stein, W. H., and Moore, S. (1962) Arch. Biochem. Biophys. 1, 217–222
  26. Sorrentino, S., Barone, R., Bucci, E., Gotte, G., Russo, N., Libonati, M., and D'Alessio, G. (2000) FEBS Lett. 466, 35–39[CrossRef][Medline] [Order article via Infotrieve]
  27. Canals, A., Pous, J., Guasch, A., Benito, A., Ribo, M., Vilanova, M., and Coll, M. (2001) Structure 9, 967–976[Medline] [Order article via Infotrieve]
  28. Liu, Y., Hart, P. J., Schlunegger, M. P., and Eisenberg, D. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 3437–3442[Abstract/Free Full Text]
  29. Liu, Y., Gotte, G., Libonati, M., and Eisenberg, D. (2001) Nat. Struct. Biol. 8, 211–214[CrossRef][Medline] [Order article via Infotrieve]
  30. Matousek, J., Gotte, G., Pouckova, P., Soucek, J., Slavik, T., Vottariello, F., and Libonati, M. (2003) J. Biol. Chem. 278, 23817–23822[Abstract/Free Full Text]
  31. Kunitz, M. (1946) J. Biol. Chem. 164, 563–568[Free Full Text]
  32. Otwinowsky, Z., and Minor, W. (1997) Methods Enzymol. 276, 307–326[CrossRef]
  33. Bailey, S. (1994) Acta Crystallogr. Sect. D 50, 760–763[CrossRef][Medline] [Order article via Infotrieve]
  34. Matthews, B. W. (1968) J. Mol. Biol. 33, 491–497[Medline] [Order article via Infotrieve]
  35. Esnouf, R. M. (1999) Acta Crystallogr. Sect. D 55, 938–940[CrossRef][Medline] [Order article via Infotrieve]
  36. Jones, T. A., Bergdoll, M., and Kjeldgaard, M. (1990) in Crystallography Model Methods Molecular Design (Bugg, C., and Ealick, S., eds) pp. 189–199, Springer-Verlag Press, New York
  37. Laskowski, R. A., MacArthur, M. W., Moss, M. D., and Thorton, J. M. (1993) J. Appl. Crystallogr. 26, 283–291[CrossRef]
  38. Kobe, B., and Deisenhofer, J. (1995) Nature 374, 183–186[CrossRef][Medline] [Order article via Infotrieve]
  39. Sica, F., Di Fiore, A., Zagari, A., and Mazzarella, L. (2003) Proteins 52, 263–271[CrossRef][Medline] [Order article via Infotrieve]
  40. Zegers, I., Maes, D., Dao-Thi, M. H., Poortmans, F., Palmer, R., and Wyns, L. (1994) Protein Sci. 3, 2322–2339[Medline] [Order article via Infotrieve]
  41. Gilliland, G. (1997) in Ribonuclease: Structures and Functions (D'Alessio, G., and Riordan, J. F., eds) pp. 306–341, Academic Press, San Diego
  42. Vitagliano, L., Adinolfi, S., Riccio, A., Sica, F., Zagari, A., and Mazzarella, L. (1998) Protein Sci. 7, 1691–1699[Medline] [Order article via Infotrieve]
  43. Vitagliano, L., Merlino, A., Zagari, A., and Mazzarella, L. (2002) Proteins 46, 97–104[CrossRef][Medline] [Order article via Infotrieve]
  44. Merlino, A., Vitagliano, L., Ceruso, M. A:, Di Nola, A., and Mazzarella, L. (2002) Biopolymers 65, 274–283[CrossRef][Medline] [Order article via Infotrieve]
  45. Berisio, R., Sica, F., Lamzin, V. S., Wilson, K. S., Zagari, A., and Mazzarella, L. (2002) Acta Crystallogr. Sect. D 52, 441–450
  46. Merlino, A., Vitagliano, L., Ceruso, M. A., and Mazzarella, L. (2003) Proteins 53, 101–110[CrossRef][Medline] [Order article via Infotrieve]
  47. Vitagliano, L., Adinolfi, S., Sica, F., Merlino, A., Zagari, A., and Mazzarella, L. (1999) J. Mol. Biol. 293, 569–577[CrossRef][Medline] [Order article via Infotrieve]
  48. Mazzarella, L., Vitagliano, L., and Zagari, A. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 3799–3803[Abstract/Free Full Text]
  49. Avitabile, F., Alfano, C., Spadaccini, R., Crescenzi, O., D'Ursi, A. M., D'Alessio, G., Tancredi, T., and Picone, D. (2003) Biochemistry 42, 8704–8711[CrossRef][Medline] [Order article via Infotrieve]
  50. Liu, Y., and Eisenberg, D. (2002) Protein Sci. 11, 1285–1299[CrossRef][Medline] [Order article via Infotrieve]
  51. Merlino, A., Vitagliano, L., Ceruso, M. A., and Mazzarella, L. (2004) Biophys. J. 86, 2383–2391[Medline] [Order article via Infotrieve]
  52. Kobe, B., and Deisenhofer, J. (1993) Nature 366, 751–756[CrossRef][Medline] [Order article via Infotrieve]
  53. Kim, J. S., Soucek, J., Matousek, J., and Raines, R. T. (1995) J. Biol. Chem. 270, 31097–31102[Abstract/Free Full Text]
  54. Murthy, B. S., and Sirdeshmukh, R. (1992) Biochem. J. 281, 343–348
  55. Schlunegger, M. P, Bennett, M. J, and Eisenberg D. (1997) Adv. Protein Chem. 50, 61–122[Medline] [Order article via Infotrieve]
  56. D'Alessio, G. (1999) Eur. J. Biochem. 266, 699–708[Medline] [Order article via Infotrieve]
  57. D'Alessio, G. (1999) Prog. Biophys. Mol. Biol. 72, 271–298[CrossRef][Medline] [Order article via Infotrieve]
  58. Ciglic, M. I., Jackson, P. J., Raillard, S. A., Haugg, M., Jermann, T. M., Opitz, J. G., Trabesinger-Ruf, N., and Benner, S. A. (1998) Biochemistry 37, 4008–4022[CrossRef][Medline] [Order article via Infotrieve]
  59. Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946–950[CrossRef]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
D. Picone, A. Di Fiore, C. Ercole, M. Franzese, F. Sica, S. Tomaselli, and L. Mazzarella
The Role of the Hinge Loop in Domain Swapping: THE SPECIAL CASE OF BOVINE SEMINAL RIBONUCLEASE
J. Biol. Chem., April 8, 2005; 280(14): 13771 - 13778.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/35/36753    most recent
M405655200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Sica, F.
Right arrow Articles by Mazzarella, L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sica, F.
Right arrow Articles by Mazzarella, L.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement