Biomacromolecules 2005, 6, 1097-1103
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Ionic, Structural, and Temperature Effects on DNA Nanoparticles Formed by Natural and Synthetic Polyamines Veena Vijayanathan,† Jasleen Lyall,† Thresia Thomas,‡,§ Akira Shirahata,| and T. J. Thomas*,†,§ Departments of Medicine and Environmental and Occupational Medicine, University of Medicine and Dentistry of New Jersey-Robert Wood Johnson Medical School, New Brunswick, New Jersey 08903, The Cancer Institute of New Jersey, New Brunswick, New Jersey 08903, and Environmental and Occupational Health Sciences Institute, Piscataway, New Jersey 08854, and Department of Biochemistry and Cellular Physiology, Josai University, Saitama, Japan Received October 26, 2004; Revised Manuscript Received November 30, 2004
We synthesized analogues of spermine and studied the effects of chemical structure, ionic strength, and temperature on λ-DNA nanoparticle formation. Effective concentration of polyamines for DNA condensation (EC50) was lowest for hexamines (0.2 µM) and highest for spermine (tetramine, 4.2 µM). The EC50 value increased with [Na+]. Dynamic light scattering showed nanoparticles with hydrodynamic radii (Rh) of 4050 nm. Effect of temperature on Rh was measured between 20 and 70 °C. For spermine, Rh remained relatively stable until 50 °C and increased significantly at >60 °C. In contrast, the hexa- and penta-valent analogues exhibited a gradual increase in Rh between 20 and 70 °C. The nanoparticles were mainly toroidal, as revealed by electron microscopy (EM). EM studies showed changes in morphology and size of condensed structures with an increase in temperature. A possible mechanism for the differential effects of temperature on DNA nanoparticles might involve different modes of DNA-polyamine interactions. Introduction DNA compaction by biological cations has been extensively studied due to its importance in gene therapy, biotechnology, and bionanotechnology.1-8 A prerequisite for DNA transport through the cell membrane is nanoparticle formation by the condensation of DNA. Several lines of evidence suggest that the biophysical characteristics of nanoparticles, such as size, surface properties, and affinity of the condensing agent toward DNA, play important roles in gene therapy.9-14 A wide variety of multivalent cations (with charges greater than 3+), as well as monovalent cations in the presence of dehydrating or crowding agents, provoke DNA nanoparticle formation.15-20 Addition of monovalent cations or solvent dilution reverses DNA nanoparticles to wormlike chains. Interplay of several factors, such as bending rigidity, valence of the counterions, as well as hydration forces, dictate the mechanism of DNA nanoparticle formation.4 Higher valent cations are more effective in DNA condensation, indicating the overriding role of electrostatic interaction between DNA phosphate and cationic condensing agents.21 DNA condensation is a kinetically controlled process, and the size of nanoparticles formed is dependent upon salt * Corresponding author. Phone: (732) 235-8460. Fax: (732) 235-8473. E-mail:
[email protected]. † Departments of Medicine and Environmental and Occupational Medicine, University of Medicine and Dentistry of New Jersey-Robert Wood Johnson Medical School. ‡ Environmental and Occupational Health Sciences Institute. § The Cancer Institute of New Jersey. | Department of Biochemistry and Cellular Physiology.
conditions, the structure and cationicity of the condensing agent, and DNA concentration.22-24 The condensed particles appear as toroids or spheroids when observed under an electron microscope (EM). Although toroids and spheroids are considered to be the end points of DNA condensation, precursors such as flowerlike and spaghetti-like structures have been observed by atomic force microscopic (AFM) and EM techniques.25-28 Occasionally, rodlike particles are observed by EM, but their proportion to toroids is very low. Physicochemical properties and biological function of DNA complexes prepared from commercial gene transfection agents, such as polyethyleneimine, amidoamine dendrimers, and polyamine derivatized cationic lipids, depend on the chemical structure of the condensing agent, ionic conditions, and preparation techniques.9-14 Many gene delivery strategies exploit the change in the properties of the vector in response to pH gradients and redox potential between extracellular and intracellular compartments of the cell.29 Several gene delivery vectors rely on the ability of the condensing agent to undergo phase transitions in response to physical stimuli, including temperature and UV irradiation.30-33 However, systematic studies on the influence of temperature on nanoparticle stability are scant. In addition, DNA nanoparticle storage is an important aspect of developing these materials as pharmaceutical agents for gene delivery. In an effort to develop simple carrier systems for gene delivery, we synthesized several higher valent structural analogues of the natural polyamine, spermine, and studied their efficacy in condensing λ-DNA. Previous studies showed that hexamine analogues are excellent promoters of oligonucleotide transport in breast cancer cells.34 In the present
10.1021/bm0493234 CCC: $30.25 © 2005 American Chemical Society Published on Web 01/21/2005
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Figure 1. Chemical structures of polyamines used in this study. Protonated structures are shown here because these molecules are positively charged under the conditions of the experiment.
study, we evaluated the ionic, structural, and temperature effects on the compaction of λ-DNA to nanoparticles in the presence of spermine and synthetic polyamines with five and six positive charges, as well as bisethyl polyamine analogues. Our results show that pentamines and hexamines are more efficient in the induction and stabilization of DNA nanoparticles. Temperature studies indicate that the stability of DNA nanoparticles depends on the nature of the polyamine analogue used for DNA compaction. Experimental Procedures Materials. Highly purified λ-phage DNA was purchased from Sigma Chemical Co. (St. Louis, MO). Stock solutions were prepared by DNA being dissolved in 10 mM sodium cacodylate buffer containing 0.5 mM EDTA (pH 7.2), and the concentration was adjusted to 1 mg/mL. The A260/A280 absorbance ratio was 1.9, indicating that the DNA was free from protein contamination. Spermine‚4HCl and cobalt hexamine‚ 3HCl (Co(NH3)63+) were purchased from Sigma Chemical Co. 1,17-Diamino-4,9,14-triazaheptadecane (3-4-4-3), 1,17-diamino-5,9,13-triazaheptadecane (4-3-3-4), 1,15diamino-4,8,12-triazapentadecane (3-3-3-3), 1,19-diamino4,8,12,16-tetraazanonadecane (3-3-3-3-3), 1,21-diamino4,9,13,18-tetraazahenicosane (3-4-3-4-3), 3,7,12,16tetrazaoctadecane (BE-3-4-3), and 3,7,11,15,19-pentazahenicosane (BE-3-3-3-3) were synthesized by us, as described previously.35,36 Protonated chemical structures of the polyamines are shown in Figure 1. The polyamines are abbreviated by a number system, indicating the number of methylene (-CH2-) groups between the primary and the secondary amino groups). The purity and structural integrity of polyamines was confirmed by NMR, HPLC, and mass spectrometry. Polyamine stock solutions were prepared in double distilled water, and appropriate dilutions were made in 10 mM Na cacodylate buffer. Total Intensity Light Scattering. Total intensity light scattering was performed as described previously.21 Briefly, different volumes of polyamine stock solutions (90% DNA charge neutralization, and only cations of charges 3 or higher could condense DNA to nanoparticles in aqueous media. However, DNA condensation was achieved by divalent cations by changing the dielectric constant of the medium (e.g., water-alcohol mixtures).37,40 Critical concentration of polyamines required for DNA nanoparticle formation increased with monovalent salt concentration.37 Manning’s counterion condensation theory predicts a power law relationship between the concentration of free polyamines in solution and monovalent ion concentration.38 While the salt dependence of EC50 values followed a straight-line relationship for spermine, hexamines and pentamines exhibited two linear regions when log[EC50] values were plotted against log[Na+]. At monovalent concentrations >50 mM, the plot obeys Manning’s power law with a slope value of 3, which falls between the predicted value of 1 and 6. At lower monovalent ion concentrations (10-25 mM), it deviates from the power law. This deviation from the power law equation at low salt concentrations may be due to the higher binding affinity of pentamines and hexamines for DNA.21 However, caution is needed in applying the counterion condensation theory on the dependence of EC50 values on the monovalent salt concentration. Counterion condensation theory predicts a linear dependence of the binding constant of the ligand with the logarithm of salt concentration. The theory treats and distinguishes ions based on the charge and neglects any specific ion effect based on size, hydration, and symmetry. These specific ion effects may be important in the condensation of DNA by higher valent polyamine analogues. A more detailed theory based on these parameters is not currently available. Specificity of polyamine-DNA interactions continues to be a matter of considerable debate. Early models based on electrostatic theory for polyamine-DNA interactions focused on nonspecific interactions between positively charged amino groups and negatively charged phosphate groups of DNA and assumed a lack of site binding interactions.37-39,41
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However, our studies using a series of isovalent tetramines showed that in addition to the predominant charge effects, chemical structure of the polyamine also exerts a profound influence in precipitating and condensing DNA to nanoparticles.42-45 Theoretical studies by Zakrzewska and Pullman46 indicated that spermine-DNA interactions depend on DNA conformation as well as base sequence. On the basis of energy minimization calculations, Feuerstein et al.47 suggested that DNA stabilization by spermine is achieved by the bending of DNA over polyamines and localization of the polyamine into the major grove. X-ray crystallographic analysis of polyamine-DNA complexes indicates that polyamines occupy specific sites, especially the major and minor grooves, depending on the conformation of DNA.48,49 X-ray diffraction studies by Schellman and Parthasarathy50 observed that the Bragg spacing and interhelical distance varied systematically with the length of the methylene bridges in a series of spermidine homologues of the structure, H2N(CH2)3NH(CH2)nNH2, where n ) 2-8. However, molecular dynamic studies by Nordenskiold et al.51 indicate that while spermine and spermidine are totally absent in the major groove of DNA, putrescine and a synthetic polyamine, diaminopropane, interact with bases in the major groove. X-ray crystallographic analysis by Williams et al.52 also shows that spermine can assume different conformations and form extensive van der Waals contacts with the DNA bases in a sequence specific manner. Theoretical studies also suggest that spermine favorably binds to A-DNA as compared to B-DNA.53 Our recent studies with RNA‚DNA hybrids showed a preferential increase in the melting temperature of the hybrid as compared to the DNA‚DNA duplex at 150 mM Na+.54 Raman spectroscopic measurements indicate that spermidine and spermine interact primarily with the phosphate groups without disrupting their native structure.55 However, Ruiz-Chica et al.56 observed structural specificity for polyamine-DNA interactions, with putrescine and spermine preferentially binding to the minor groove and spermine to the major groove of DNA. Taken together, it can be assumed that polyamines appear to interact with the bases and phosphate groups, depending on DNA sequence and conformation, as well as the distance separating the positive charges within the polyamine. Despite differences in the cationicity and chemical structure, all polyamines used in this study compacted DNA to nanoparticles with hydrodynamic radii in the range of 4050 nm. The structural effect of the polyamines seems to affect the size and the temperature-dependent stability of the nanoparticles formed. Although the alkyl-substituted polyamines bear the same number of positive charges as their unsubstituted counterparts, they exhibit significant differences in the formation and stability of DNA nanoparticles. Alkyl substituted polyamines enhance aggregation of condensed DNA structures by hydrophobic interactions.57 Increase in the size of Co(NH3)63+-condensed structures with temperature can be attributed to the ability of Co3+ ions to interact with the bases and disrupt base pairing and induce aggregation at higher temperatures.58 Compactness of the Co(NH3)63+ molecule can also enable the counterions to pack more
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Figure 6. Schematic model to explain the temperature effects on DNA nanoparticles formed in the presence of polyamines. Increase in temperature destabilizes the DNA nanoparticles either by polyamine mediated aggregation or by temperature induced melting. Nanoparticles formed in the presence of spermine are relatively stable until an increase in temperature destabilized the particle by temperature induced melting, which tends to cross-link the nanoparticles. Nanoparticles formed from higher valent polyamine analogues are more sensitive to temperature. Hexamine analogues can mediate intermolecular cross-linking between nanoparticles. The effective range of these interactions depends on the chemical structure of the polyamines as well as the temperature.
closely, thereby changing the binding isotherm and interDNA attractions in significant ways. Increase in temperature destabilizes DNA nanoparticles either by polyamine-mediated aggregation or by temperatureinduced melting and intermolecular cross-linking. In the case of spermine, the Rh values remain fairly constant until an increase in temperature changes the actual composition of the complex due to counterion redistribution and complex dissociation. Because of differences in the spatial correlations between DNA phosphate charges and positive charges on the polyamine molecule, there are potentially several uncompensated charges on the condensed structures. The residual electrostatic repulsion tends to keep the particles apart. With an increase in temperature, the unbound portion of the hexamine molecule can mediate intermolecular crosslinking, finally forming structures involving more than one nanoparticle. Our results indicate that at higher temperatures, cross-linking might play an important role in the stability of the nanoparticles formed with hexamine. A schematic model for the mechanism of temperature-dependent aggregation is depicted in Figure 6. Toroids observed in this study (Figure 5) are comparable in size and shape to those observed by other investigators for different forms of DNA. For example, Chattoraj et al.59 observed toroids of 80 nm for spermidine-condensed structures, whereas Eickbush and Moudrianakis 60 reported larger toroids of 100 nm. Arscott et al.61 observed toroidal particles of 80 nm diameter for Co(NH)63+-condensed particles. AFM studies by Lin et al.62 show that spermidine condensed λ-DNA into toroidal structures with average outer diameter
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of 120 ( 15 nm. Hud et al.22 found that toroid dimensions could be controlled by the introduction of intrinsically bent A-tracts in the DNA sequences. Introducing two static loops in DNA provides a built in site for toroid nucleation, and the size of the toroids decreased from 80 to 68 nm, indicating size of the nucleation loop directly influences the diameter of the toroid formed.22 Several models and theories have been put forward to explain toroid formation and the origin of toroid size and thickness.6,7,41,63-66 The spool model for toroid formation assumes circumferential wrapping of DNA as strings in a spool, with DNA packed in a hexagonal lattice.63 Cryotransmission electron microscopy and polarizing microscopy provide evidence for DNA packaging in hexagonal lattice.6,64 Theoretical considerations based on a constant loop model predict that the kinetic process of spontaneous loop formation is the dominant factor in the determination of toroid dimensions.65 According to the thermodynamic model proposed by Bloomfield,17 build up of uncompensated negative charges during toroid formation produces electrostatic repulsions and limits toroid size. According to a recent report by Ray and Manning,66 the bundle size results from the balance of short-range attractive interactions and long-range repulsive interaction between polyanions. A recent study by Conwell and Hud67 suggests that the temperature effect on particle size is limited both by thermodynamic and by kinetic factors. When limited by thermodynamic factors, the toroid sizes decreases with an increase in temperature, whereas in a kinetically controlled process, the toroid size increases with an increase in temperature. Conclusions Our data demonstrate that tetra-, penta-, and hexavalent polyamine analogues condensed λ-DNA to nanoparticles of 80-100 nm in diameter. Hexavalent polyamines were more efficacious than pentamines and tetramines in provoking DNA nanoparticle formation, indicating that electrostatic interactions between polyamines and DNA are the major force governing DNA nanoparticle formation. Increase in the effective concentration of polyamines for DNA condensation with monovalent ions suggests ion competition effects, in accordance with the counterion condensation theory. However, this theory fails to explain the dependence of log[polyamine] versus log[Na+] plots in the case of pentamines and hexamines. Our results show that subtle variation in the structure of the polyamines used in DNA nanoparticle formation can have significant effects on the morphology of nanoparticles and their stability at high temperatures. These findings might help to develop strategies for the design and synthesis of polyamine analogues for DNA nanoparticle formation and their storage as pharmaceutical agents for gene delivery. Acknowledgment. This work was supported, in part, by Public Health Service Grants CA80163, CA73058, and CA42439 from the National Cancer Institute; a grant award from the Susan G. Komen Breast Cancer Research Foundation; and a Grant-in-Aid for Scientific Research from the
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Ministry of Education, Science and Culture, Japan. J.L. was supported by a Summer Fellowship from the Robert Wood Johnson Medical School Dean’s Office. References and Notes (1) Vijayanathan, V.; Thomas, T.; Thomas, T. J. Biochemistry 2002, 41, 14085-14094. (2) Seeman, N. C. Chem. Biol. 2003, 10, 1151-1159. (3) Seeman, N. C. Biochemistry 2003, 42, 7259-7269. (4) Bloomfield, V. A. Biopolymers 1997, 44, 269-282. (5) Ibarra, B.; Caston, J. R.; Llorea, O.; Valle, M.; Valpuesta, J. M.; and Carrascosa, J. L. 2000, J. Mol. Biol. 298, 807-815. (6) Hud, N. V.; Downing, K. H. Proc. Natl. Acad. Sci. U.S.A 2001, 98, 14925-14930. (7) Luo, D.; Saltzman, W. Nat. Biotech. 2000, 18, 33-37. (8) Blessing, T.; Remy, J. S.; Behr, J. P. Proc. Natl. Acad. Sci. U.S.A 1998, 95, 1427-1431. (9) Lim, Y. B.; Choi, Y. H.; Park, J. S. J. Am. Chem. Soc. 1999, 121, 5633-5639. (10) Wagner, E.; Ogris, M.; Zauner, W. AdV. Drug DeliVery ReV. 1998, 30, 97-113. (11) Ogris, M.; Walker, G.; Blessing, T.; Kircheis, R.; Wolschek, M.; Wagner, E. J. Controlled Release 2003, 91, 173-181. (12) Tang, M.; Szoka, F. C. Gene Ther. 1997, 4, 823-832. (13) Dauty, E.; Behr, J. P.; Remy, J. S. Gene Ther. 2002, 9, 743-748. (14) Pollard, H.; Remy, J. S.; Loussouarn, G.; Demolombe, S.; Behr, J. P.; Escande, D. J. Biol. Chem. 1998, 273, 7509-7511. (15) Widom, J.; Baldwin, R. L. Biopolymers 1983, 22, 1595-1620. (16) Bloomfield, V. A. Curr. Opin. Struct. Biol. 1996, 6, 334-341. (17) Bloomfield, V. A. Biopolymer 1991, 13, 1471-1481. (18) Livolant, F.; Leforestier, A. Prog. Polym. Sci. 1996, 21, 1115-1164. (19) Koltover, I.; Wagner, K.; Safinya, C. R. Proc. Natl. Acad. Sci. U.S.A 2000, 97, 14046-14051. (20) Brewer, L. R.; Corzett, M.; Balhorn, R. Science 1999, 286, 120123. (21) Vijayanathan, V.; Thomas, T.; Antony, T.; Shirahata, A.; Thomas, T. J. Nucleic Acids Res. 2004, 32, 127-134. (22) Conwell, C. C.; Vilfan, I. D.; Hud, N. V. Proc. Natl. Acad. Sci. U.S.A 2003, 100, 9296-9301. (23) Stevens, M. J. Phys. ReV. Lett. 1999, 82, 101-104. (24) Ha, B.-Y.; Lin, A. J. Phys. ReV. Lett. 1998, 81, 1011-1014. (25) Fang, Y.; Hoh, J. H. J. Am. Chem. Soc. 1998, 120, 8903-8909. (26) Golan, R.; Pietrasanta, L. I.; Hseih, W.; Hansma, H. G. Biochemistry 1998, 18, 14069-14076. (27) Fang, Y.; Hoh, J. H. FEBS Lett. 1999, 459, 173-176. (28) Dunlap, D. D.; Maggi, A.; Soria, M. R.; Monaco, L. Nucleic Acids Res. 1997, 25, 3095-3101. (29) Quipicky, D.; Diwadkar, V. Curr. Opin. Mol. Ther. 2003, 5, 345350. (30) Nagasaki, T.; Tamiguchi, A.; Tamagaki, S. Bioconj. Chem. 2003, 14, 513-516. (31) Kurisawa, M.; Yokoyama, M.; Okano, T. J. Controlled Release 2000, 69, 127-137. (32) Nagasaki, T.; Atarshi, K.; Makiro, A.; Noguchi, T.; Matsumoto, T.; Tamagaki, S. Mol. Cryst. Liq. Cryst. 2000, 345, 227-232. (33) Qupicky, D.; Reschel, T.; Konak, C.; Qupicka, L. Macromolecules 2003, 36, 6863-6872.
Biomacromolecules, Vol. 6, No. 2, 2005 1103 (34) Thomas, R. M.; Thomas, T.; Wada, M.; Sigal, L. H.; Shirahata, A.; Thomas, T. J. Biochemistry 1999, 38, 13328-13337. (35) He, Y.; Suzuki, T.; Kashiwaga, K.; Kusuma-Eguchi, K.; Shirahata, A.; Igarashi, K. Eur. J. Biochem. 1994, 221, 391-398. (36) Igarashi, K.; Koga, K.; He, Y.; Shimogori, T.; Ekimoto, H.; Kashiwagi, K.; Shirahata, A. Cancer Res. 1995, 55, 2615-2619. (37) Wilson, R. W.; Bloomfield, V. A. Biochemistry 1979, 18, 21922196. (38) Manning, G. S. Q. ReV. Biophys. 1978, 11, 179-246. (39) Record, M. T., Jr.; Anderson, C. F.; Lohman, T. M. Q. ReV. Biophys. 1978, 11, 103-178. (40) Arscott, P. G.; Ma, C.; Wenner, J.; Bloomfield, V. A. Biopolymers 1995, 36, 345-365. (41) Manning, G. S. Biopolymers 1972, 11, 937-949. (42) Saminathan, M.; Antony, T.; Shirahata, A.; Sigal, L. H.; Thomas, T.; Thomas, T. J. Biochemistry 1999, 38, 3821-3830. (43) Vijayanathan, V.; Thomas, T.; Shirahata, A.; Thomas, T. J. Biochemistry 2001, 40, 13644-13651. (44) Thomas, T. J.; Bloomfield, V. A. Biopolymers 1983, 22, 1097-1106. (45) Thomas, T. J.; Messner, R. P. J. Mol. Biol. 1988, 201, 463-467. (46) Zakrzewska, K.; Pullman, B. Biopolymers 1986, 25, 375-392. (47) Feuerstein, B. G.; Pattabiraman, N.; Marton, L. J. Nucleic Acids Res. 1990, 18, 1271-1282. (48) Jain, S.; Zon, G.; Sundaralingam, M. Biochemistry 1989, 28, 23602364. (49) Egli, M.; Williams, L. D.; Gao, Q.; Rich, A. Biochemistry 1991, 30, 11388-402. (50) Schellman, J. A.; Parthasarathy, N. J. Mol. Biol. 1984, 175, 313329. (51) Korolev, N.; Lyubartsev, A. P.; Laaksonen, A.; Nordenskiold, L. Eur. Biophys. J. 2004, 33, 671-682. (52) Williams, L. D.; Frederick, C. A.; Ughetto, G.; Rich, A. Nucleic Acids Res. 1990, 18, 5533-5541. (53) Zhurkin, V. B.; Lysov, Y. P.; Ivanov, V. I. Biopolymers 1980, 19, 1415-1434. (54) Venkiteswaran, S.; Vijayanathan, V.; Shirahata, A.; Thomas, T.; Thomas, T. J. Biochemistry, 2005, 44, 303-312. (55) Deng, H.; Bloomfield, V. A.; Benevides, J. M.; Thomas, G. J., Jr. Nucleic Acids Res. 2000, 28, 3379-3385. (56) Ruiz-Chica, J.; Medina, M. A.; Sanchez-Jimenez, F.; Ramirez, F. J. Biophys. J. 2001, 80, 443-54. (57) Thomas, T.; Thomas, T. J. Biochemistry 1993, 32, 14068-14074. (58) Duguid, J. G.; Bloomfield, V. A. Biophys. J. 1995, 69, 2642. (59) Chattoraj, D. K.; Gosule, L. C.; Schellman, A. J. Mol. Biol. 1978, 121, 327-37. (60) Eickbush, T. H.; Moudrianakis, E. N. Cell 1978, 13, 295-306. (61) Arscott, P. G.; Li, A. Z.; Bloomfield, V. A. Biopolymers 1990, 30, 619-630. (62) Lin, Z.; Wang, C.; Feng, X.; Liu, M.; Li, J., Bai, C. Nucleic Acids Res. 1998, 26, 3229-3234. (63) Bottcher, C.; Endisch, C.; Furhop, J. H.; Catterall, C.; Eaton, M. J. Chem. Soc. 1998, 120, 12-17. (64) Saminathan, M.; Thomas, T.; Shirahata, A.; Pillai, C. K. S.; Thomas, T. J. Nucleic Acids Res. 2002, 30, 3722-3731. (65) Hud, N. V.; Downing, K. H.; Balhorn, R. Proc. Natl. Acad. Sci. U.S.A 1995, 92, 3581-3585. (66) Ray, J.; Manning, G. S. Macromolecules 2000, 33, 2901-2908. (67) Conwell, C. C.; Hud, N. V. Biochemistry 2004, 43, 5380-5387.
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