Efficient Calf Thymus DNA Condensation upon Binding with Novel

Bioconjugate Chem. 2002, 13, 481−490

481

Efficient Calf Thymus DNA Condensation upon Binding with Novel Bile Acid Polyamine Amides Andrew J. Geall, Dima Al-Hadithi, and Ian S. Blagbrough* Department of Pharmacy and Pharmacology, University of Bath, Bath BA2 7AY, U.K. Received August 10, 2000

Polyamine amides have been prepared from lithocholic and cholic acids (5β-colanes) by acylation of tri-Boc-protected tetraamines spermine and thermine. These designed ligands for DNA are polyammonium ions at physiological pH. In NMR spectra, they display 14N-1H 1J ) 51 Hz, 1:1:1 triplets, due to the symmetry of the R14NH3+ cations. The binding affinities of these conjugates for calf thymus DNA were determined using an ethidium bromide fluorescence quenching assay and compared with spermine and polylysine. DNA-binding affinities were dependent upon both salt concentration and the hydrophobicity or intermolecular bonding (facial effects) of the lipid moieties in these conjugates. Light scattering at 320 nm was used to determine DNA condensation and particle formation. The observed self-assembly phenomena are discussed with respect to DNA charge neutralization and DNA bending with loss of ethidium cation intercalation sites, ultimately leading to DNA condensation. These polyamine amides are models for lipoplex formation with respect to gene delivery (lipofection), a key first step in gene therapy.

INTRODUCTION

Among polyamine-containing natural products (1-7), polyamino-steroids form a novel, small group whose members and their analogues display a variety of interesting biological activities. Walker and co-workers have recently reported the DNA-binding affinity and in vitro gene delivery potential of various polyamines conjugated to the steroidal bile acids lithocholic 1 and cholic 2 acids

(8, 9). Although most of their transfection agents contained a cationic headgroup (tetraamine spermine 3, a polyammonium ion at physiological pH) attached to a hydrophobic tail (e.g., cholic acid spermine amide 13), there were significant differences in the transfection activities across the series (8, 9). Diamino steroids cyclobuxine (10), irehdiamine A, and malouetine have long been known to interact with DNA (10-13). The natural product bufotoxin, isolated from the venom of the * To whom correspondence should be addressed. Phone: 441225-826795. Fax: 44-1225-826114. E-mail: [email protected].

European toad Bufo vulgaris (B. bufo bufo Linne´), contains an arginyl guanidine functional group pendant from a steroid (bufotalin) nucleus and displays digitalislike cardiac effects (14-16). Recently, Regen and coworkers reported their (so-called) molecular umbrella from cholic acid 2 and triamine spermidine, creating structures that can mask an attached agent (dansyl as a drug mimetic) from the surrounding environment (1719). Polyamino-steroid squalamine, isolated from liver and gallbladder tissues of the dogfish shark, Squalus acanthias, is a spermidine-containing sterol sulfate which displays antimicrobial and fungicidal properties, and induces osmotic lysis of protozoa (20-23). As part of our continuing studies on polyaminemediated DNA condensation (24-32), we have synthesized polyamine amide conjugates of lithocholic 1 and cholic acids 2 in order to investigate the effects on changes in hydrophobicity on their binding affinity to DNA. Structure-activity relationships (SAR) for the binding of polyamines to DNA (13, 33-38), and the subsequent condensation of DNA (37-41), indicate that polyammonium ions are suitable for use as gene delivery systems (42-54). Following DNA-binding studies with a short series of synthetic polyamino-steroids, up to four structural features have been shown to contribute to the strength and type of DNA interactions: cation type, total number of positive charges, regiochemical distribution of the ammonium groups, and steroid hydrophobicity (5557). The mechanism by which these compounds cause lipofection is still poorly understood (50-54). Therefore, it is important to determine their physicochemical properties for the design of lipoplexes capable of efficient lipofection (58). We are investigating how changes in the hydrophobicity of a lipopolyamine affect the condensation of DNA and hence lipoplex formation (24-32). Herein we report the design and synthesis, using our orthogonal protection strategy, of unsymmetrical polyamine amides from bile acids, two amides of lithocholic acid 1, 11 and 12 (at position 24), and two of cholic acid 2, 13, and 14 (also at

10.1021/bc000093+ CCC: $22.00 © 2002 American Chemical Society Published on Web 04/26/2002

482 Bioconjugate Chem., Vol. 13, No. 3, 2002

position 24) (24). The 5β-cholane ring structure was chosen as the lipid moiety, it has found previous use in lipoplex formation (8, 9), and a variety of derivatives of these bile acids are readily available substituted with differing numbers of hydroxyl groups at varying positions on the ring system. Two readily available polyamines were used as the cationic headgroups: 1,12-diamino-4,9diazadodecane 3 (spermine, 3.4.3) and 1,11-diamino-4,8diazaundecane (thermine, norspermine, 3.3.3) 4. Differences in the distribution of positive charges along these two polyamine headgroups and their effects on binding affinity, including salt-dependent binding of these polyamine amides of lithocholic and cholic acids have been investigated. To investigate the SAR for their binding affinities for calf thymus (CT) DNA, we used a modified ethidium bromide displacement assay (59, 60). Changes in binding affinity for CT DNA with respect to variations in the hydroxyl substituents on a lipopolyamine are investigated with respect to the use of bile acids in lipoplex formation, a key first step in gene therapy (5058). EXPERIMENTAL PROCEDURES

General Details and Reagents. General details, column, RP-HPLC, solvents and other chromatographic details, NMR, MS, and other spectroscopic details are as previously described (25, 26). Poly-L-lysine hydrobromide (RMM 4-15 kDa) was purchased from SAF and had an average RMM of 9.6 kDa (PL 9600) (DP 46) when measured by viscosity and 8.0 kDa (DP 38) measured by LALLS. CT DNA sodium salt and all other reagents and chemicals were obtained from SAF and were of the highest grades available. General Procedure A: Poly-Boc Protection of Polyamines. To a cold (-78 °C) methanolic solution of the polyamine was added ethyl trifluoroacetate (1 equiv) dropwise over 30 min, and then without isolation, the remaining amino functional groups were protected quantitatively by the dropwise addition of an excess of di-tertbutyl dicarbonate in methanol (10 mL). Trifluoroacetyl functional group removal at pH 11 with concentrated aqueous ammonia, workup, and purification as previously reported (25-27) afforded the title compound as a colorless homogeneous oil. General Procedure B: Amide Formation. To a solution of the poly-Boc-protected polyamine in CH2Cl2 (10 mL) the bile acid, 1-hydroxybenzotriazole (HOBt) (0.2 equiv) and dicyclohexylcarbodiimide (DCC) (1.5 equiv) were added. Then the reaction mixture was stirred at 25 °C, under nitrogen, for 24 h. The precipitate of dicyclohexylurea was then removed by filtration. The filtrate was concentrated in vacuo and the residue purified over silica gel (CH2Cl2-MeOH) to afford the title compound as a white foam. General Procedure C: Boc Removal. To a stirring solution of lipopolyamine dissolved in CH2Cl2 (180 mL), under nitrogen at 25 °C, was added TFA (20 mL). After 2 h, the solution was concentrated in vacuo, lyophilized, and purified by semipreparative RP-HPLC over Supelcosil ABZ+Plus (5 µm, 25 cm × 10 mm, MeOH-0.1% aqueous TFA) to afford the title compound as a white solid (poly-TFA salt). (N1,N4,N9-Tri-tert-butoxycarbonyl)-1,12-diamino4,9-diazadodecane, 5. 1,12-Diamino-4,9-diazadodecane 3 (spermine, 3.4.3) (1.0 g, 5.0 mmol) was reacted according to general procedure A to afford the title compound 5 as a homogeneous oil (1.24 g, 50%). Purified over silica gel (CH2Cl2-MeOH-concentrated NH3(aq) 70:10:1 to 50:

Geall et al.

10:1 v/v/v), Rf 0.5 (CH2Cl2-MeOH-concentrated NH3(aq) 50:10:1 v/v/v) (25). (N1,N4,N8-Tri-tert-butoxycarbonyl)-1,11-diamino4,8-diazaundecane, 6. 1,11-Diamino-4,8-diazaundecane 4 (thermine, norspermine, 3.3.3) (3.0 g, 16.0 mmol) was reacted according to general procedure A to afford the title compound 6 as a homogeneous oil (3.16 g, 41%). Purified over silica gel [CH2Cl2-MeOH-concentrated NH3(aq) 100:10:1 v/v/v], Rf 0.18 [CH2Cl2-MeOH-concentrated NH3(aq) 100:10:1 v/v/v] (25). N1-(3r-Hydroxy-5β-cholan-24-carbonyl-(N4,N9,N12tri-tert-butoxycarbonyl))-1,12-diamino-4,9-diazadodecane, 7. Polyamine 5 (500 mg, 1.0 mmol) and lithocholic acid (375 mg, 1.0 mmol) were reacted according to general procedure B to afford the title compound 7 as a white foam (610 mg, 71%). Purified by column chromatography over silica gel (CH2Cl2-MeOH; 25:1 v/v), Rf 0.22 (CH2Cl2MeOH; 20:1 v/v). 1H NMR, 400 MHz, CDCl3: 0.64 (s, 3 H, 18′-CH3); 0.83-2.26 (m, 69 H, 2-CH2, 6-CH2, 7-CH2, 11-CH2, 3 × OC(CH3)3, 1′-CH2, 2′-CH2, 4′-CH2, 5′-CH, 6′CH2, 7′-CH2, 8′-CH, 9′-CH, 11′-CH2, 12′-CH2, 14′-CH, 15′CH2, 16′-CH2, 17′-CH, 19′-CH3, 20′-CH, 21′-CH3, 22′-CH2, 23′-CH2); 2.84-3.05 (m, 12 H, 1-CH2, 3-CH2, 5-CH2, 8-CH2, 10-CH2, 12-CH2); 3.60-3.70 (m, 1 H, 3′-CH); 6.756.90 (br s, 1 H, CH2-NHCO). 13C NMR, 100 MHz, CDCl3: 12.0 (18′-CH3); 18.3 (21′-CH3); 20.7 (11′-CH2); 23.2 (19′CH3); 24.1 (15′-CH2); 25.4, 25.6, 25.9 (6-CH2, 7-CH2, overlapping); 26.3 (7′-CH2); 27.1 (6′-CH2); 27.6, 28.1, 28.4, 28.7 [2-CH2, 11-CH2, 16′-CH2, 3 × O-C-(CH3)3, overlapping]; 30.4 (2′-CH2); 31.7 (22′-CH2); 33.7 (23′-CH2); 34.5 (10′-C); 35.3, 35.4 (12-CH2, 20′-CH, 1′-CH2); 35.8 (8′-CH); 36.3 (4′-CH2); 37.3 (1-CH2); 40.1 (12′-CH2); 40.3 (9′-CH); 42.0 (5′-CH); 42.6 (13′-CH); 43.2, 43.7 (3-CH2, 10-CH2, overlapping); 46.6 (5-CH2, 8-CH2, overlapping); 56.0 (17′CH); 56.4 (14′-CH); 71.6 (3′-CH); 79.7 (3 × quaternary OC, overlapping); 156.0, 156.4 (3 × NCOO); 173.7 (CH2CON). MS, FAB+ found 861, 20% (M++1), C49H88N4O8 requires M+ ) 860. N1-(3r-Hydroxy-5β-cholan-24-carbonyl-(N4,N8,N11tri-tert-butoxycarbonyl))-1,11-diamino-4,8-diazaundecane, 8. Polyamine 6 (500 mg, 1.0 mmol) and lithocholic acid (385 mg, 1.0 mmol) were reacted according to general procedure B to afford the title compound 8 as a white foam (751 mg, 87%). Purified by column chromatography over silica gel (CH2Cl2-MeOH; 25:1 v/v), Rf 0.30 (CH2Cl2-MeOH; 20:1 v/v). 1H NMR, 400 MHz, CDCl3: 0.64 (s, 3 H, 18′-CH3); 0.84-2.24 [m, 67 H, 2-CH2, 6-CH2, 10-CH2, 3 × O-C-(CH3)3, 1′-CH2, 2′-CH2, 4′-CH2, 5′-CH, 6′-CH2, 7′-CH2, 8′-CH, 9′-CH, 11′-CH2, 12′-CH2, 14′-CH, 15′-CH2, 16′-CH2, 17′-CH, 19′-CH3, 20′-CH, 21′CH3, 22′-CH2, 23′-CH2]; 2.84-3.05 (m, 12 H, 1-CH2, 3-CH2, 5-CH2, 7-CH2, 9-CH2, 11-CH2); 3.60-3.70 (m, 1 H, 3′-CH); 6.75-6.90 (br s, 1 H, CH2-NHCO). 13C NMR, 100 MHz, CDCl3: 12.0 (18′-CH3); 18.3 (21′-CH3); 20.7 (11′CH2); 23.2 (19′-CH3); 24.1 (15′-CH2); 26.3 (7′-CH2); 27.1 (6′-CH2); 27.6, 28.1, 28.4, 28.7 [2-CH2, 6-CH2, 10-CH2, 16′CH2, 3 × OC(CH3)3, overlapping]; 30.4 (2′-CH2); 31.7 (22′CH2); 33.7 (23′-CH2); 34.5 (10′-C); 35.3, 35.4 (11-CH2, 20′CH, 1′-CH2, overlapping); 35.8 (8′-CH); 36.4 (4′-CH2); 37.3 (1-CH2); 40.1 (12′-CH2); 40.3 (9′-CH); 42.0 (5′-CH); 42.6 (13′-CH); 43.6, 43.7, 44.2, 44.7 (3-CH2, 5-CH2, 7-CH2, 9-CH2, overlapping); 56.0 (17′-CH); 56.4 (14′-CH); 71.7 (3′-CH); 79.7 (3 × quaternary OC, overlapping); 155.9, 156.3 (3 × NCOO); 173.7 (CH2-CONH). MS, FAB+ found 847, 20% (M++1), C48H86N4O8 requires M+ ) 846. N1-(3r,7r,12r-Trihydroxy-5β-cholan-24-carbonyl(N4,N9,N12-tri-tert-butoxycarbonyl))-1,12-diamino4,9-diazadodecane, 9. Polyamine 5 (500 mg, 1.0 mmol) and cholic acid (406 mg, 1.0 mmol) were reacted according

Bile Acid Polyamine Amides Condense DNA

to general procedure B to afford the title compound 9 as a white foam (532 mg, 60%). Purified by column chromatography over silica gel (CH2Cl2-MeOH; 15:1 to 10:1 v/v), Rf 0.17 (CH2Cl2-MeOH; 10:1 v/v). 1H NMR, 400 MHz, CDCl3: 0.67 (s, 3 H, 18′-CH3); 0.88 (s, 3 H, 19′-CH3); 0.91-2.40 (m, 62 H, 2-CH2, 6-CH2, 7-CH2, 11-CH2, 3 × O-C-(CH3)3, 1′-CH2, 2′-CH2, 4′-CH2, 5′-CH, 6′-CH2, 8′-CH, 9′-CH, 11′-CH2, 14′-CH, 15′-CH2, 16′-CH2, 17′-CH, 20′CH, 21′-CH3, 22′-CH2, 23′-CH2); 2.80-3.37 (m, 12 H, 1-CH2, 3-CH2, 5-CH2, 8-CH2, 10-CH2, 12-CH2); 3.37-3.50 (m, 1 H, 3′-CH); 3.83 (s, 1 H, 7′-CH); 3.96 (s, 1 H, 12′CH); 6.75-6.90 (br s, 1 H, CH2-NHCO). 13C NMR, 100 MHz, CDCl3: 12.5 (18′-CH3); 17.5 (21′-CH3); 22.5 (19CH3); 25.5, 25.7, 26.0 (6-CH2, 7-CH2, overlapping); 26.4 (9′-CH2); 27.6 (16′-CH2); 27.7, 28.1, 28.2, 28.4, 28.5, 28.7, 28.9 (2-CH2, 11-CH2, 11′-CH2, 3 × OC(CH3)3, overlapping); 30.5 (2′-CH2); 31.7 (22′-CH2); 33.6 (23′-CH2); 34.7 (6′-CH2); 34.8 (10′-C); 35.3 (1′-CH2); 35.5, 35.7 (12-CH2, 20′-CH, overlapping); 37.4, 37.7 (1-CH2); 39.5 (8′-CH); 39.6 (4′-CH2); 41.5, 41.7 (5′-CH, 14′-CH); 43.5, 43.7, 44.2 (3-CH2, 5-CH2, 8-CH2, 10-CH2, overlapping); 46.4, 46.8 (13′-C, 17′-CH); 68.4 (7′-CH); 71.9 (3′-CH); 73.0 (12′-CH); 79.6, 79.8 (3 × quaternary OC, overlapping); 156.1, 156.4 (3 × NCOO); 174.1 (CH2-CONH). MS, FAB+ found 893, 40% (M+ + 1), C49H88N4O10 requires M+ ) 892. N1-(3r,7r,12r-Trihydroxy-5β-cholan-24-carbonyl(N4,N8,N11-tri-tert-butoxycarbonyl))-1,11-diamino4,8-diazaundecane, 10. Polyamine 6 (500 mg, 1.0 mmol) and cholic acid (418 mg, 1.0 mmol) were reacted according to general procedure B to afford the title compound 10 as a white foam (824 mg, 91%). Purified by column chromatography over silica gel (CH2Cl2-MeOH; 20:1 to 15:1 to 10:1 v/v), Rf 0.20 (CH2Cl2-MeOH; 10:1 v/v). 1H NMR, 400 MHz, CDCl3: 0.67 (s, 3 H, 18′-CH3); 0.88 (s, 3 H, 19′-CH3); 0.91-2.30 [m, 60 H, 2-CH2, 6-CH2, 10-CH2, 3 × O-C-(CH3)3, 1′-CH2, 2′-CH2, 4′-CH2, 5′-CH, 6′-CH2, 8′-CH, 9′-CH, 11′-CH2, 14′-CH, 15′-CH2, 16′-CH2, 17′-CH, 20′-CH, 21′-CH3, 22′-CH2, 23′-CH2]; 2.88-3.38 (m, 12 H, 1-CH2, 3-CH2, 5-CH2, 7-CH2, 9-CH2, 11-CH2); 3.38-3.50 (m, 1 H, 3′-CH); 3.83 (s, 1 H, 7′-CH); 3.96 (s, 1 H, 12′CH); 6.75-6.90 (br s, 1 H, CH2-NH-CO). 13C NMR, 100 MHz, CDCl3: 12.5 (18′-CH3); 17.5 (21′-CH3); 22.5 (19CH3); 26.4 (9′-CH2); 27.6 (16′-CH2); 28.2, 28.5, 28.8, 28.9 [2-CH2, 6-CH2, 10-CH2, 11′-CH2, 3 × O-C-(CH3)3, overlapping]; 30.5 (2′-CH2); 31.7 (22′-CH2); 33.6 (23′-CH2); 34.7 (6′-CH2); 34.8 (10′-C); 35.3 (1′-CH2); 35.5, 35.7 (11-CH2, 20′-CH, overlapping); 37.4 (1-CH2, overlapping); 39.5 (8′CH); 39.6 (4′-CH2); 41.5, 41.7 (5′-CH, 14′-CH); 43.5, 43.8, 44.8 (3-CH2, 5-CH2, 7-CH2, 9-CH2, overlapping); 46.4, 46.7 (13′-C, 17′-CH); 68.4 (7′-CH); 71.9 (3′-CH); 73.0 (12′-CH); 79.8 (3 × quaternary OC, overlapping); 156.0, 156.1 (3 × NCOO); 174.1 (CH2-CONH). MS, FAB+ found 879, 5% (M+ + 1), C48H86N4O10 requires M+ ) 878. N1-(3r-Hydroxy-5β-cholan-24-carbonyl)-1,12-diamino-4,9-diazadodecane, 11. Amide 7 (300 mg, 0.34 mmol) was deprotected according to general procedure C. The residue was lyophilized to produce 352 mg of a white powder, 225 mg was purified by RP-HPLC (MeCN0.1% TFA(aq), 38:62 v/v) to afford the title compound 11 as a white solid (polytrifluoroacetate salt, 61 mg, 27%), tR 11.5 min by RP-HPLC (Supelcosil ABZ+Plus, 5 µm, 15 cm × 4.6 mm, MeCN-0.1% TFA(aq), 40:60 v/v). 1H NMR, 400 MHz, d6-DMSO: 0.61 (s, 3 H, 18′-CH3); 0.820.98 (m, 7 H, 1′β-CH, 19′-CH3, 21′-CH3); 0.98-1.28 (m, 9H, 2′R-CH, 6′R-CH, 7′R-CH, 11′β-CH, 14′-CH, 15′R-CH, 16′β-CH, 17′-CH, 22′β-CH); 1.28-1.44 (m, 7 H, 4′β-CH, 5′-CH, 7′β-CH, 8′-CH, 9′-CH, 11′R-CH, 20′-CH); 1.44-1.86 (m, 13 H, 6-CH2, 7-CH2, 11-CH2, 1′R-CH, 2′β-CH, 4′RCH, 6′β-CH, 15′β-CH, 16′R-CH, 22′R-CH); 1.86-2.15 (m,

Bioconjugate Chem., Vol. 13, No. 3, 2002 483

5 H, 2-CH2, 12′β-CH, 23′R-CH, 23′β-CH); 2.84-3.05 (m, 10 H, 3-CH2, 5-CH2, 8-CH2, 10-CH2, 12-CH2); 3.05-3.15 (m, 2 H, 1-CH2); 3.30-3.42 (m, 1 H, 3′-CH); 4.15 [br s, 1 × OH, (+H2O)]; 7.24 (1:1:1, t, 1J ) 51, 14N-1H); 8.03, 8.71, 8.90 (3 × br s, ammonium signals). 13C NMR, 100 MHz, d6-DMSO: 11.8 (18′-CH3); 18.2 (21′-CH3); 20.3 (11′-CH2); 22.5, 22.7 (6-CH2, 7-CH2); 23.2 (19′-CH3); 23.7, 23.8 (2CH2, 15′-CH2); 26.0, 26.1 (11-CH2, 7′-CH2); 26.8 (6′-CH2); 27.7 (16′-CH2); 30.3 (2′-CH2); 31.5 (22′-CH2); 32.2 (23′CH2); 34.1 (10′-C); 34.9 (20′-CH); 35.1 (1′-CH2); 35.3 (8′CH); 35.5 (1-CH2); 36.1 (4′-CH2); 36.2 (12-CH2); 39.6 (12′CH2); 39.9 (9′-CH); 41.4 (5′-CH); 42.2 (13′-CH); 43.8 (3CH2); 44.6 (10-CH2); 46.0, 46.1 (5-CH2, 8-CH2); 55.5 (17′CH); 56.0 (14′-CH); 69.8 (3′-CH); 173.1 (CH2-CONH). MS, FAB+ found 561, 40% (M+ + 1), C34H64N4O2 requires M+ ) 560. High-resolution MS m/z, FAB+ found 561.5135, (M++1), C34H65N4O2 requires M+ + 1 ) 561.5108. N1-(3r-Hydroxy-5β-cholan-24-carbonyl)-1,11-diamino-4,8-diazaundecane, 12. Amide 8 (300 mg, 0.36 mmol) was deprotected according to general procedure C. The residue was lyophilized to produce 347 mg of a white powder, 225 mg were purified by RP-HPLC [MeCN0.1% TFA(aq), 38:62 v/v] to afford the title compound 12 as a white solid (polytrifluoroacetate salt, 71 mg, 32%). tR 7.0 min by RP-HPLC [Supelcosil ABZ+Plus, 5 µm, 15 cm × 4.6 mm, MeCN-0.1% TFA(aq), 38:62 v/v]. 1H NMR, 400 MHz, d6-DMSO: 0.61 (s, 3 H, 18′-CH3); 0.84-0.98 (m, 7 H, 1′β-CH, 19′-CH3, 21′-CH3); 0.98-1.28 (m, 9H, 2′R-CH, 6′R-CH, 7′R-CH, 11′β-CH, 14′-CH, 15′R-CH, 16′βCH, 17′-CH, 22′β-CH); 1.28-1.44 (m, 7 H, 4′β-CH, 5′-CH, 7′β-CH, 8′-CH, 9′-CH, 11′R-CH, 20′-CH); 1.44-1.86 (m, 9 H, 10-CH2, 1′R-CH, 2′β-CH, 4′R-CH, 6′β-CH, 15′β-CH, 16′R-CH, 22′R-CH); 1.86-2.15 (m, 7 H, 2-CH2, 6-CH2, 12′β-CH, 23′R-CH, 23′β-CH); 2.85-3.05 (m, 10 H, 3-CH2, 5-CH2, 7-CH2, 9-CH2, 11-CH2); 3.05-3.14 (m, 2 H, 1-CH2); 3.30-3.42 (m, 1 H, 3′-CH); 3.63 [br s, 1 × OH, (+H2O)]; 7.22 (1:1:1, t, 1J ) 51, 14N-1H); 8.01, 8.79, 8.97 (3 × br s, ammonium signals). 13C NMR, 100 MHz, d6-DMSO: 11.8 (18′-CH3); 18.2 (21′-CH3); 20.3 (11′-CH2); 22.3 (6-CH2); 23.2 (19′-CH3); 23.7 (2-CH2, 15′-CH2, overlapping); 26.0, 26.1 (10-CH2, 7′-CH2); 26.8 (6′-CH2); 27.6 (16′-CH2); 30.3 (2′-CH2); 31.4 (22′-CH2); 32.2 (23′-CH2); 34.1 (10′-C); 34.9 (20′-CH); 35.0 (1′-CH2); 35.2 (8′-CH); 35.4 (1-CH2); 36.1 (11-CH2); 36.2 (4′-CH2); 39.6 (12′-CH2); 39.8 (9′-CH); 41.4 (5′-CH); 42.2 (13′-CH); 43.8, 43.9 (3-CH2, 9-CH2); 44.7 (5CH2, 7-CH2, overlapping); 55.4 (17′-CH); 56.0 (14′-CH); 69.8 (3′-CH); 173.0 (CH2-CONH). MS, FAB+ found 547, 100% (M+ + 1), C33H62N4O2 requires M+ ) 546. Highresolution MS m/z, FAB+ found 547.4955, (M+ + 1), C33H63N4O2 requires M+ + 1 ) 547.4951. N1-(3r,7r,12r-Trihydroxy-5β-cholan-24-carbonyl)1,12-diamino-4,9-diazadodecane, 13. Amide 9 (300 mg, 0.34 mmol) was deprotected according to general procedure C. The residue was lyophilized to produce 345 mg of a white powder, 220 mg was purified by RP-HPLC [MeCN-0.1% TFA(aq), 25:75 v/v] to afford the title compound (8, 9) 13 as a white solid (polytrifluoroacetate salt, 74 mg, 34%). tR 5.0 min by RP-HPLC [Supelcosil ABZ+Plus, 5 µm, 15 cm × 4.6 mm, MeCN-0.1% TFA(aq), 27:73 v/v]. 1H NMR, 400 MHz, d6-DMSO: 0.58 (s, 3 H, 18′-CH3); 0.50-1.00 (m, 8 H, 1′β-CH, 15′R-CH, 19′-CH3, 21′-CH3); 1.05-1.50 (m, 10 H, 2′R-CH, 2′β-CH, 4′β-CH, 5′-CH, 6′R-CH, 11′-CH2, 16′β-CH, 20-CH, 22′β-CH); 1.502.00 (m, 19 H, 2-CH2, 6-CH2, 7-CH2, 11-CH2, 1′R-CH, 4′RCH, 6′β-CH, 9′-CH, 14′-CH, 15′β-CH, 16′R-CH, 17′-CH, 22′R-CH, 23′R-CH, 23′β-CH); 2.00-2.25 (m, 3 H, 4′R-CH, 9′-CH, 23R-CH); 2.78-3.01 (m, 10 H, 3-CH2, 5-CH2, 8-CH2, 10-CH2, 12-CH2); 3.01-3.12 (m, 2 H, 1-CH2); 3.12-3.22 (m, 1 H, 3′-CH); 3.43 [br s, 3 × OH, (+H2O)];


3.61 (s, 1 H, 7′-CH); 3.79 (s, 1 H, 12′-CH); 7.22 (1:1:1, t, 1 J ) 51, 14N-1H); 8.01, 8.67, 8.87 (3 × br s, ammonium signals). 13C NMR, 100 MHz, d6-DMSO: 12.2 (18′-CH3); 17.0 (21′-CH3); 22.5, 22.7 (6-CH2, 7-CH2, 15′-CH2, 19′CH3, overlapping); 23.7 (2-CH2); 26.0 (11-CH2); 26.1 (9′CH2); 27.2 (16′-CH2); 28.5 (11′-CH2); 30.3 (2′-CH2); 31.6 (22′-CH2); 32.3 (23′-CH2); 34.3 (10′-C); 34.8 (6′-CH2); 35.1, 35.2 (1′-CH2, 20′-CH); 35.4 (1-CH2); 36.1 (12-CH2); 38.8 (4′-CH2, 8′-CH); 41.3, 41.4 (5′-CH, 14′-CH); 43.8 (3-CH2); 44.5 (10-CH2); 45.6 (13′-C); 46.0, 46.1 (5-CH2, 8-CH2, 17′CH); 66.1 (7′-CH); 70.3 (3-CH); 70.9 (12′-CH); 173.2 (CH2CONH). MS, FAB+ found 593, 100% (M+ + 1), C34H64N4O4 requires M+ ) 592. High-resolution MS m/z, FAB+ found 593.5010, (M+ + 1), C34H65N4O4 requires M+ + 1 ) 593.5006. N1-(3r,7r,12r-Trihydroxy-5β-cholan-24-carbonyl)1,11-diamino-4,8-diazaundecane, 14. Amide 10 (300 mg, 0.34 mmol) was deprotected according to general procedure C. The residue was lyophilized to produce 352 mg of a white powder, 225 mg was purified by RP-HPLC [MeCN-0.1% TFA(aq), 27:73 v/v] to afford the title compound 14 as a white solid (polytrifluoroacetate salt, 124 mg, 55%). tR 6.8 min by RP-HPLC [Supelcosil ABZ+Plus, 5 µm, 15 cm × 4.6 mm, MeCN-0.1% TFA(aq), 30:70 v/v]. 1H NMR, 400 MHz, d6-DMSO: 0.58 (s, 3 H, 18′-CH3); 0.50-1.00 (m, 8 H, 1′β-CH, 15′R-CH, 19′-CH3, 21′-CH3); 1.05-1.50 (m, 10 H, 2′R-CH, 2′β-CH, 4′β-CH, 5′-CH, 6′R-CH, 11′-CH2, 16′β-CH, 20′-CH, 22′β-CH); 1.50-2.05 (m, 17 H, 2-CH2, 6-CH2, 10-CH2, 1′R-CH, 4′RCH, 6′β-CH, 9′-CH, 14′-CH, 15′β-CH, 16′R-CH, 17′-CH, 22′R-CH, 23′R-CH, 23′β-CH); 2.05-2.28 (m, 3 H, 4′R-CH, 9′-CH, 23R-CH); 2.85-3.05 (m, 10 H, 3-CH2, 5-CH2, 7-CH2, 9-CH2, 11-CH2); 3.05-3.15 (m, 2 H, 1-CH2); 3.153.25 (m, 1 H, 3′-CH); 3.60 (s, 1 H, 7′-CH); 3.79 (s, 1 H, 12′-CH); 5.00 [br s, 3 × OH, (+H2O)]; 7.21 (1:1:1, t, 1J ) 51, 14N-1H); 8.01, 8.78, 8.97 (3 × br s, ammonium signals). 13 C NMR, 100 MHz, d6-DMSO: 12.2 (18′-CH3); 17.0 (21′CH3); 22.4, 22.5, 22.7 (6-CH2, 15′-CH2, 19′-CH3, overlapping); 23.8 (2-CH2); 26.0 (10-CH2); 26.2 (9′-CH2); 27.2 (16′CH2); 28.5 (11′-CH2); 30.3 (2′-CH2); 31.6 (22′-CH2); 32.3 (23′-CH2); 34.3 (10′-C); 34.8 (6′-CH2); 35.1, 35.2 (1′-CH2, 20′-CH); 35.5 (1-CH2); 36.1 (11-CH2); 39.2 (4′-CH2, 8′-CH); 41.3, 41.4 (5′-CH, 14′-CH); 43.9, 44.0 (3-CH2, 9-CH2); 44.7 (5-CH2, 7-CH2); 45.6 (13′-C); 46.0 (17′-CH); 66.1 (7′-CH); 70.3 (3-CH); 70.9 (12′-CH); 173.2 (CH2-CONH). MS, FAB+ found 579, 100% (M+ + 1), C33H62N4O4 requires M+ ) 578. High-resolution MS m/z, FAB+ found 579.4849, (M+ + 1), C33H63N4O4 requires M+ + 1 ) 579.4854. In our hands, all members of this series of polyamine amides 11-14 were water soluble at 1 mg ml-1 (8, 9). RESULTS AND DISCUSSION

Polyamines spermine 3 and thermine 4 were sequentially unsymmetrically protected with di-tert-butyl dicarbonate using our orthogonal protection strategy (2431). Selective protection of one primary amine group, in each symmetrical polyamine 3 and 4, by reaction with ethyl trifluoroacetate, afforded the respective monotrifluoroacetamides. Immediately, in these solutions, the remaining amino functional groups were Boc protected, with di-tert-butyl dicarbonate, to afford the fully protected polyamines. The trifluoroacetyl protecting group was then cleaved (in situ) by increasing the pH to 11 with concentrated aqueous ammonia, to afford N1,N2,N3-triBoc protected polyamines 5 and 6. Acylation of the free primary amine of these unsymmetrically protected polyamines 5 and 6 with lithocholic 1 and cholic acids 2 (DCC, cat. HOBt), afforded the fully protected polyamine

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Figure 1. Synthesis of target polyamine amides 11-14.

amides 7-10. Deprotection (TFA-CH2Cl2, 1:9 v/v) and purification to homogeneity by RP-HPLC afforded target polyamine amides 11-14, as their polytrifluoroacetate salts (Figure 1). The proposed structures were unambiguously assigned using 1H, 13C and HETCOR NMR and accurate FAB-MS. The pKas of the polyamine headgroups on polyamino steroids 11-14 were assumed to be the same as those measured potentiometrically for the cholesterol carbamates we have previously reported (29). The net positive charge was then calculated using the HendersonHasselbach equation at pH 7.4. Therefore, polyamine amides 11 and 13 (conjugates of spermine) have been assigned a net positive charge of 2.4 and polyamine amides 12 and 14 (conjugates of thermine) have been assigned a net positive charge of 2.3. We have named the target compounds 11-14 as their corresponding polyamine derivatives and have outlined the numbering system we used in the NMR assignments with N1-(3Rhydroxy-5β-cholan-24-carbonyl)-1,12-diamino-4,9-diazadodecane 11 selected as an example. The complete NMR spectroscopic assignment of the polyamine headgroups in this series of polyamine amides 11-14 is based upon calculations using additivity rules (61), 1H, 13C chemical shift correlation spectroscopy, and detailed comparisons with cholesterol carbamates previously characterized (25, 31). The assignment of the 5βcholane ring structure is based upon literature assignments (62), and the expected changes in the carbon chemical shifts due to substituent effects are entirely consistent with these assignments. Conformational isomers (populations interconverted by σ-bond rotation) are observed for the poly-Boc-protected polyamines, and therefore, two signals for each carbon on the methylene


backbone of the polyamine and for each carbon on the Boc groups were typically observed. Generally 14N-1H couplings are not observed, but in the case of ammonium compounds, the combination of quadrupole relaxation and exchange of NH-protons is not sufficiently large to eliminate completely the coupling across one bond (61). Therefore, amides 11-14 display broad ammonium signals above δ 7.0 ppm (typically 8.0, 8.8, and 9.0 ppm, exchanged with D2O). In addition, signals at δ 7.2 (1:1:1 t, 1J ) 51 Hz, 14N-1H) were observed for these ammonium ions which we interpret as due to the symmetry of the R14NH3+ cations (61). Ethidium Bromide Displacement Assay. Relative binding affinities for CT DNA of the target compounds 11-14 were measured using a modified ethidium bromide fluorescence quenching assay, a refined displacement assay (59, 60), and plotted against charge ratio (CR) as defined in the literature (58). The decrease in fluorescence was critically compared against polylysine (PL, average RMM 9.6 kDa, PL 9600) and spermine 3 (Figure 2) for compounds 11-14 at 20 mM NaCl as a function of CR (ammonium:DNA phosphate ions) (58). At physiological pH, spermine carries a net positive charge of 3.8 (25, 29), PL 9600 a net positive charge in excess of 30. In Figure 2, polyamine amides 11 and 12, and polycationic PL 9600 exhibit similar binding affinities for DNA. With PL 9600, complete displacement of ethidium bromide occurs as the conjugate-DNA complexes approach a charge ratio of 1. A slight excess of positive charge is required to displace the ethidium cation with the lithocholic acid conjugates 11 and 12. This fluorescence quenching assay measures the relative binding affinities for DNA between compounds of comparable structure (59, 60). Therefore, compared to spermine 3, conjugates 11 and 12 have a greater binding affinity for DNA (Figure 2). There was no visible precipitation of the DNA. These data are consistent with complex formation between the DNA and the polyamine amide conjugates. This binding of polyamines and their conjugates to DNA is not a trivial process (27-41, 55-57, 63-68); they may bind preferentially to GC-rich major groove and to AT-rich minor groove regions (63-66). Covalent attachment of a lipid moiety, such as an aliphatic chain or a steroid, further enhances polyamine-mediated DNA condensation (3949). Cholic acid 2 is a sterol nucleus with a hydroxylated hydrophilic surface and an all-hydrocarbon hydrophobic surface, possessing the 5β-cholane ring structure (a cisfused A,B-bicycle). Polyamine N1-acylation with cholic acid 2 affords more hydrophilic polyamine amides 13 and


14 (moreso than 11 and 12) as they contain three hydroxyl groups (compared to the one alcohol functional group of lithocholic acid) that are all located regiospecifically on the R-face of the 5β-cholane ring system. Amides 13 and 14 have a decreased binding affinity for DNA, compared to their lithocholic acid counterparts 11 and 12 (Figure 2), and they require a large excess of positive charge to displace the ethidium cation from its CT DNA intercalation sites. The differences in binding affinities between the spermine 11 and 13 and thermine 12 and 14 polyamine moieties were small (Figure 2). At 150 mM NaCl salt concentrations the binding affinity for CT DNA of PL 9600 is unaffected, but that of spermine 3 shows marked salt-dependent binding (Figure 3). Spermine amide 11, which contains the lithocholane ring structure (with one hydroxyl moiety), requires an excess of positive charge (CR approximately 8) to displace the ethidium cation (Figure 3). The binding behavior of spermine amide 13 mimics that of spermine 3 at high salt concentrations, and the displacement of ethidium is almost completely inhibited even at a CR of >10. DNA Condensation (Light Scattering) Assay. To investigate whether CT DNA is condensed into particles by spermine cholic acid amide 13, the UV absorbance at 320 nm has been measured (33). In Figure 4, we show the apparent increase in UV absorption at 320 nm of DNA as aliquots are added at low and high salt concentrations (20 and 150 mM NaCl, respectively). At low salt concentrations, these data are consistent with particle formation and the absorption reaches a plateau at the same charge ratio (approximately 4) as essentially maximal ethidium bromide exclusion (Figure 2). However, due to the lack of sensitivity of this light-scattering assay, the DNA concentration was in a 10-fold excess compared to that used in the ethidium bromide assay. Significantly, at high salt concentrations, there are no particles formed by DNA condensation and hence no light scattering, with no visible precipitation of the DNA. These data (Figures 3 and 4) are consistent with inhibition of complex formation between the DNA and polyamine amide conjugate 13 at elevated salt concentrations. From detailed comparisons of light-scattering assay data (Figure 4 and data not shown) together with ethidium bromide fluorescence quenching data (Figure 2 and Figure 3), we conclude that gross morphological changes occur to CT DNA as a result of polyamine amide binding. These effects are sensitive to the number of hydroxyl groups on the lipid moiety (reflecting its hydrophobicity) and are also salt concentration dependent. They are consistent with charge neutralization of CT


Figure 2. Ethidium bromide displacement assay of spermine 3, amides 11-14, and polylysine at 20 mM NaCl. CT DNA (6 µg in buffer, 3 mL, 20 mM NaCl, 2 mM HEPES, 10 µM EDTA, pH 7.4) was mixed with ethidium bromide (3 µL of 0.5 mg/mL) and aliquots of compound (5 µL of 0.25 mg/mL, 1 min equilibration time) were added, the fluorescence (%) determined, and plotted against the charge ratio of ammonium:DNA phosphate ions (58).

Figure 3. Ethidium bromide displacement assay of spermine 3, amides 11 and 13, and polylysine at 150 mM NaCl. CT DNA (6 µg in buffer, 3 mL, 150 mM NaCl, 2 mM HEPES, 10 µM EDTA, pH 7.4) was mixed with ethidium bromide (3 µL of 0.5 mg/mL) and aliquots of compound (5 µL of 0.25 mg/mL, 1 min equilibration time) were added, the fluorescence (%) determined, and plotted against the charge ratio of ammonium:DNA phosphate ions (58).

DNA, the loss of ethidium cation binding sites within the DNA double helix, and ultimately with DNA condensation and particle formation. These latter effects are the first key steps in gene therapy for the formation of lipoplexes. Measurement of the ability of a drug to displace ethidium cation from DNA has been shown to be a valid measurement of DNA binding ability of both intercalative and nonintercalative drugs, including free polyamines and their conjugates (55-57, 59, 69-74). Displacement of ethidium cation from DNA provides an indirect method of measuring the binding affinity of drugs that lack a chromophore. It does not provide a direct measure of the binding constant, but offers a qualitative comparison of binding affinities within a series of compounds with similar structures. Molecular modeling studies of ethidium cation intercalation into DNA have shown that binding is accompanied by a helical screw axis displace-

Geall et al.

Figure 4. Light-scattering assay of polyamine amide 13 at low (20 mM NaCl) and high (150 mM NaCl) salt concentrations. CT DNA (60 µg in buffer, 3 mL, 20 mM NaCl, 2 mM HEPES, 10 µM EDTA, pH 7.4) was stirred and aliquots of compound (5 µL of 1.0 mg/mL, 1 min equilibration time) were added, the absorption (320 nm) measured, and plotted against the charge ratio of ammonium:DNA phosphate ions (58).

ment (or dislocation) in its structure (75). The helical axes are displaced approximately +1.0 Å (for B DNA), base pairs in the immediate region are twisted by 10°, giving rise to an angular unwinding of -26° and the intercalated base pairs are tilted relative to one another by 8°. These changes in DNA conformation mean that intercalation is limited to every other base pair at maximal drugnucleic acid binding ratios, i.e., a neighbor exclusion model (75). These modeling studies also indicate that the conformational flexibility of DNA allows intercalation of the ethidium cation at kinked regions of the double helix. Thus, intercalation of ethidium cation occurs at regions where the base-pairs are unwound (kinked), which induces a conformational change in the double helix, restricting the total number of intercalation sites. DNA collapse, by charge neutralization of cationic lipids, is thought to be a key step in lipoplex formation (58). This collapse is termed condensation, and affords compact, highly ordered toroids that have a possible relationship to the packaging of DNA in viruses, in particular they represent models for DNA organization within bacteriophage heads (76-78). The fluorescent intensity of the intercalated ethidium cation is not affected by increasing concentrations of cationic lipid until a specific lipid:DNA ratio is reached, upon which a large and sharp decrease of the intensity is observed. Ha¨rd et al. have demonstrated that the binding constant of ethidium cation is dependent on the molecular flexibility of DNA in linker regions of chromatin and that this flexibility is altered through cationic compaction (79). Thus, DNA condensation might be expected to lower the affinity of ethidium cation for DNA, and therefore, its exclusion cannot be considered to be a direct measure of a drug’s binding affinity. Basu et al. also concluded that simple polyamine-DNA association was not entirely responsible for the release of ethidium cation, but that there is an associated conformational (DNA bending) change (33, 80). DNA bending, induced by polyamine binding above a critical concentration, caused conformational changes within the double helix that facilitated the release of bound ethidium cation. The model for ethidium cation intercalation proposed by Sobell et al. shows the need for flexibility within the double helix of DNA to allow intercalation (75). Ethidium bromide exists in equilibrium between the intercalated sites and free in


solution. Therefore, loss of flexibility in the doublestranded structure of DNA through condensation will result in a shift in the binding equilibrium of ethidium cation into the solution phase, with the resultant loss in fluorescence. Wilson and Bloomfield predicted (81), using the polyelectrolyte theory of Manning (82), that when approximately 90% of the negative (phosphate) charge on the DNA is neutralized, condensation will occur (39-41, 67, 68). Thus, if binding affinities of compounds causing DNA condensation are expressed in terms of the CR at which 50% (CR50) of the ethidium cation fluorescence is quenched (58), efficient condensing agents will have CR50 values below 1. DNA condensation, at low salt concentrations (20 mM NaCl), is clearly an efficient process with lithocholic acid polyamine amides 11 and 12 (CR50 ) 0.7 and 0.6, respectively). However, an excess of positive charges is required for cholic acid polyamine amides 13 and 14 (CR50 ) 2.6 and 2.8 respectively) and for free spermine (CR50 > 4.0) to condense CT DNA, reflecting their significantly weaker binding affinities for DNA. At high salt concentrations (150 mM NaCl), DNA condensation with lithocholic acid polyamine amide 11 is salt dependent, the CR50 has increased from 0.7 to 4.7. The more hydrophilic cholic acid polyamine amide 13 exhibits even greater salt-dependent DNA condensation; the CR 50 has increased from 2.6 to >12. Basu et al. have previously shown that the association of free polyamines with DNA is more sensitive to change in ionic strength than is the association of ethidium bromide with DNA (80). The concentration of polyamine required to release all of the intercalator dye in their studies was too high for experimental use without causing DNA aggregation (37, 38, 80). Basu and co-workers have also demonstrated that this assay is suitable for the investigation of DNA binding with simple, free polyamines, e.g., spermine 3 and thermine (norspermine) 4 where there was little fluorescence quenching at a CR of 1 (33, 73, 80). We have obtained similar results with conjugates 13 and 14 and free spermine 3 at low (20 mM NaCl) (Figure 2) and especially at high (150 mM NaCl) salt concentrations (Figure 3). Thus, while hydrophobicity is important for minor groove recognition (83), DNA condensation is dependent upon hydrophobicity and distance between positive charges (84, 85) as well as total number of charges (39-41, 67, 68). These data give support to our hypotheses that both DNA binding and DNA condensation are also sensitive functions of the lipid that is covalently bound to the polyamine, as well as functions of the positively charged polyamine moiety itself. Neutral and cationic lipid-DNA interactions are a mature research area (86-90). The binding interactions between naturally occurring lipids and polynucleotides are important, and they have also been extensively studied with model membranes (86). The mechanisms by which these novel complexes are formed between cationic lipids and plasmid DNA have recently gained further significance with the potential of these particles in nonviral gene delivery (87-90). These new structures, both those involving cationic liposomes and those, like our studies, using cationic lipids have given rise to detailed freezefracture electron and atomic force microscopic studies of these particles sometimes described as “spaghetti and meatballs” (48, 53, 60, 77, 78, 88, 91-97). The formation of these compact particles is also directly related to the displacement of water molecules as a result of polyelectrolyte effects. Record and co-workers (98-101) have made detailed analytical studies of the effects of oligocationic ligand binding to DNA.


One mechanistic explanation of our observed selfassembly phenomena is that hydrophobic interactions between lithocholic acid moieties within 11 and 12, but not between the more polar cholic acid moieties in 13 and 14, cooperatively stabilize the condensed and/or condensing DNA complexes. There is some limited literature support for this possible explanation, in studies of the molecular mechanics of the complex formation of cholic acid micelles (102). The complex facial interactions observed with other arch-shaped or concave bile acids have been reviewed by Fuhrhop and Ko¨ning (103) where “micelle curvature is determined not by the hydrophobic effect, but by the molecular shape of the monomers and strong intermolecular bonding” (103). Regen and coworkers have recently reported, in a previous article in Bioconjugate Chem. (104), detailed studies on the “cloistering” of cholic acid amides due to facial effects mediated by either hydrogen bonding or van der Waals’ forces between nearest-neighbors (104). We propose that initial charge neutralization of the polyanionic polynucleic acid by our polycationic lipo-polyammonium ions (25, 26, 105) is followed by DNA bending (106, 107). After several of these bends (106), the induced curve is symmetrical, this is consistent with Hud’s constant radius of curvature model (77, 78), and adds further evidence to the microscopy pictures currently available. It has recently been shown that, depending upon the exact conditions, spermine 3 can induce condensation of DNA (97). Nevertheless, the key questions about any nucleotide sequence selectivity for the initial bending sites along the duplex CT DNA (106, 107), and the accuracy of the constant radius of curvature model will require further experiments (77, 78), preferably with designed high affinity small molecule probes and also defined DNA sequences (38). The polyamine amides designed and prepared herein should therefore contribute to those studies. ACKNOWLEDGMENT

We thank the EPSRC and Celltech Chiroscience for a CASE studentship (to A.J.G.). We are grateful to a referee for helpful directions with respect to lipid-DNA interactions. We acknowledge useful discussions with Dr. Richard J. Taylor and Dr. Michael A. W. Eaton (Celltech Chiroscience), and with Dr. Ian S. Haworth (University of Southern California). I.S.B. and I.S.H. are recipients of a NATO grant (CRG 970290). LITERATURE CITED (1) Ganem, B. (1982) New chemistry of naturally occurring polyamines. Acc. Chem. Res. 15, 290-298. (2) Tabor, C. W., and Tabor, H. (1984) Polyamines. Annu. Rev. Biochem. 53, 749-790. (3) Bergeron, R. J. (1986) Methods for selective modification of spermidine and its homologues. Acc. Chem. Res. 19, 105113. (4) Behr, J.-P. (1994) Gene transfer with synthetic cationic amphiphiles: prospects for gene therapy. Bioconjugate Chem. 5, 382-389. (5) Marton, L. J., and Pegg, A. E. (1995) Polyamines as targets for therapeutic intervention. Annu. Rev. Pharmacol. Toxicol. 35, 55-91. (6) Blagbrough, I. S., Carrington, S., and Geall, A. J. (1997) Polyamines and polyamine amides as potent selective receptor probes, novel therapeutic lead compounds, and synthetic vectors in gene therapy. Pharm. Sci. 3, 223-233. (7) Cohen, S. S. (1998) A guide to the polyamines, Oxford University Press, New York. (8) Walker, S., Sofia, M. J., Kakarla, R., Kogan, N. A., Wierichs, L., Longley, C. B., Bruker, K., Axelrod, H. R., Midha, S., Babu, S., and Kahne, D. (1996) Cationic facial amphiphiles: a

488 Bioconjugate Chem., Vol. 13, No. 3, 2002 promising class of transfection agents. Proc. Natl. Acad. Sci. U.S.A. 93, 1585-1590. (9) Walker, S., Sofia, M. J., and Axelrod, H. R. (1998) Chemistry and cellular aspects of cationic facial amphiphiles. Adv. Drug Delivery Rev. 30, 61-71. (10) Mahler, H. R., and Dutton, G. (1964) Nucleic acid interactions. V. Effects of cyclobuxine. J. Mol. Biol. 10, 157-175. (11) Mahler, H. R., Goutarel, R., Khuong-Huu, Q., and Ho, M. T. (1966) Nucleic acid interactions. VI. Effects of steroidal diamines. Biochemistry 5, 2177-2191. (12) Mahler, H. R., Green, G., Goutarel, R., and Khuong-Huu, Q. (1968) Nucleic acid-small molecule interactions. VII. Further characterization of deoxyribonucleic acid-diamino steroid complexes. Biochemistry 7, 1568-1582. (13) Waring, M. J. (1970) Variation of the supercoils in closed circular DNA by binding of antibiotics and drugs: evidence for molecular models involving intercalation. J. Mol. Biol. 54, 247-279. (14) Wieland, H., and Alles, R. (1922) Poisonous substance of toads. Chem. Ber. 55, 1789-1798. (15) Wieland, H., and Behringer, H. (1941) Toad poisons. (XI). Constitution of bufotalin. Liebigs Ann. 549, 209-237. (16) Pettit, G. R., Kamano, Y., Drasar, P., Inoue, M., and Knight, J. C. (1987) Steroids and related natural-products. 104. Synthesis of Bufalitoxin and Bufotoxin. 36. Bufadienolides. J. Org. Chem. 52, 3573-3578. (17) Janout, V., Lanier, M., and Regen, S. L. (1996) Molecular umbrellas. J. Am. Chem. Soc. 118, 1573-1574. (18) Janout, V., Lanier, M., and Regen, S. L. (1997) Design and synthesis of molecular umbrellas. J. Am. Chem. Soc. 119, 640-647. (19) DeLong, R. K., Yoo, H., Alahari S. K., Fisher, M., Short, S. M., Kang, S. H., Kole, R., Janout. V., Regan, S. L., and Juliano, R. L. (1999) Novel cationic amphiphiles as delivery agents for antisense oligonucleotides. Nucleic Acids Res. 27, 3334-3341. (20) Moore, K. S., Wehrli, S., Roder, H., Rogers, M., Forrest, J. N., McCrimmon, D., and Zasloff, M. (1993) Squalamine - an aminosterol antibiotic from the shark. Proc. Natl. Acad. Sci. U.S.A. 90, 1354-1358. (21) Moriarty, R. M., Tuladhar, S. M., Guo, L., and Wehrli, S. (1994) Synthesis of squalamine. A steroidal antibiotic from the shark. Tetrahedron Lett. 35, 8103-8106. (22) Moriarty, R. M., Enache, L. A., Kinney, W. A., Allen, C. S., Canary, J. W., Tuladhar, S. M., and Guo, L. (1995) Stereoselective synthesis of squalamine dessulfate. Tetrahedron Lett. 36, 5139-5142. (23) Sadownik, A., Deng, G., Janout, V., and Regen, S. L. (1995) Rapid construction of a squalamine mimic. J. Am. Chem. Soc. 117, 6138-6139. (24) Geall, A. J., Al-Hadithi, D., and Blagbrough, I. S. (1998) Spermine and thermine conjugates of cholic acid condense DNA, but lithocholic acid polyamine conjugates do so more efficiently. Chem. Commun. 2035-2036. (25) Geall, A. J., Taylor, R. J., Earll, M. E., Eaton, M. A. W., and Blagbrough, I. S. (2000) Synthesis of cholesteryl polyamine carbamates: pKa studies and condensation of calf thymus DNA. Bioconjugate Chem. 11, 314-326. (26) Geall, A. J., and Blagbrough, I. S. (2000) Homologation of polyamines in the rapid synthesis of lipospermine conjugates and related lipoplexes. Tetrahedron 56, 2449-2460. (27) Blagbrough, I. S., and Geall, A. J. (1998) Practical synthesis of unsymmetrical polyamine amides. Tetrahedron Lett. 39, 439-442. (28) Geall, A. J., and Blagbrough, I. S. (1998) Homologation of polyamines in the synthesis of lipo-spermine conjugates and related lipoplexes. Tetrahedron Lett. 39, 443-446. (29) Geall, A. J., Taylor, R. J., Earll, M. E., Eaton, M. A. W., and Blagbrough, I. S. (1998) Synthesis of cholesterolpolyamine carbamates: pKa studies and condensation of calf thymus DNA. Chem. Commun. 1403-1404 and 1607. (30) Blagbrough, I. S., Al-Hadithi, D., and Geall, A. J. (1999) DNA condensation by bile acid conjugates of thermine and spermine. Pharm. Pharmacol. Commun. 5, 139-144.

Geall et al. (31) Geall, A. J., and Blagbrough, I. S. (1999) DNA condensation by cholesterol polyamine carbamates. Pharm. Pharmacol. Commun. 5, 145-150. (32) Geall, A. J., Eaton, M. A. W., Baker, T., Catterall, C., and Blagbrough, I. S. (1999) The regiochemical distribution of positive charges along cholesterol polyamine carbamates plays significant roles in modulating DNA binding affinity and lipofection. FEBS Lett. 459, 337-342. (33) Basu, H. S., and Marton, L. J. (1987) The interaction of spermine and pentamines with DNA. Biochem. J. 244, 243246. (34) Thomas, T. J., and Messner, R. P. (1988) Structural specificity of polyamines in left-handed Z-DNA formation: immunological and spectroscopic studies. J. Mol. Biol. 201, 463-467. (35) Thomas, T. J., Gunnia, U. B., and Thomas, T. (1991) Polyamine-induced B-DNA to Z-DNA conformational transition of a plasmid DNA with (dG-dC)n insert. J. Biol. Chem. 266, 6137-6141. (36) Schneider, H.-J., and Blatter, T. (1992) Interactions between acyclic and cyclic peralkylammonium compounds and DNA. 33. Angew. Chem., Int. Ed. Engl. 31, 1207-1208. (37) Pelta, J., Livolant, F., and Sikorav, J.-L. (1996) DNA aggregation induced by polyamines and cobalthexamine. J. Biol. Chem. 271, 5656-5662. (38) Saminathan, M., Antony, T., Shirahata, A., Sigal, L. H., Thomas, T., and Thomas, T. J. (1999) Ionic and structural specificity effects of natural and synthetic polyamines on the aggregation and resolubilization of single-, double-, and triplestranded DNA. Biochemistry 38, 3821-3830. (39) Bloomfield, V. A. (1991) Condensation of DNA by multivalent cations - considerations on mechanism. Biopolymers 31, 1471-1481. (40) Bloomfield, V. A. (1996) DNA condensation. Curr. Opin. Struct. Biol. 6, 334-341. (41) Bloomfield, V. A. (1997) DNA condensation by multivalent cations. Biopolymers 44, 269-282. (42) Behr, J.-P., Demeneix, B., Loeffler, J.-P., and Perez-Mutul, J. (1989) Efficient gene transfer into mammalian primary endocrine cells with lipopolyamine-coated DNA. Proc. Natl. Acad. Sci. U.S.A. 86, 6982-6986. (43) Guy-Caffey, J. K., Bodepudi, V., Bishop, J. S., Jayaraman, K., and Chaudhary, N. (1995) Novel polyaminolipids enhance the cellular uptake of oligonucleotides. J. Biol. Chem. 270, 31391-31396. (44) Lee, E. R., Marshall, J., Siegel, C. S., Jiang, C., Yew, N. S., Nichols, M. R., Nietupski, J. B., Ziegler, R. J., Lane, M. B., Wang, K. X., Wan, N. C., Scheule, R. K., Harris, D. J., Smith, A. E., and Cheng, S. H. (1996) Detailed analysis of structures and formulations of cationic lipids for efficient gene transfer to the lung. Hum. Gene Ther. 7, 1701-1717. (45) Moradpour, D., Schauer, J. I., Zurawski, V. R., Jr., Wands, J. R., and Boutin, R. H. (1996) Efficient gene transfer into mammalian cells with cholesteryl-spermidine. Biochem. Biophys. Res. Commun. 221, 82-88. (46) Bischoff, R., Cordier, Y., Perraud, F., Thioudellet, C., Braun, S., and Pavirani, A. (1997) Transfection of myoblasts in primary culture with isomeric cationic cholesterol derivatives. Anal. Biochem. 254, 69-81. (47) Byk, G., Dubertret, C., Escriou, V., Frederic, M., Jaslin, G., Rangara, R., Pitard, B., Crouzet, J., Wils, P., Schwartz, B., and Scherman, D. (1998) Synthesis, activity, and structureactivity relationship studies of novel cationic lipids for DNA transfer. J. Med. Chem. 41, 224-235. (48) Remy, J.-S., Abdallah, B., Zanta, M. A., Boussif, O., Behr, J.-P., and Demeneix, B. (1998) Gene transfer with lipospermines and polyethylenimines. Adv. Drug Delivery Rev. 30, 85-95. (49) Hebling-Leclerc, A., Scherman, D., and Wils, P. (1999) Cellular uptake of cationic lipid/DNA complexes by cultured myoblasts and myotubes. Biochim. Biophys. Acta 1418, 165175. (50) Crystal, R. G. (1995) Transfer of genes to humans. Early lessons and obstacles to success. Science 270, 404-410. (51) Felgner, P. L. (1997) Nonviral strategies for gene therapy. Sci. Am. 276, 86-90.

Bile Acid Polyamine Amides Condense DNA (52) O’Driscoll, C. (1997) Gene geniuses. Chem. Brit. 33, 6669. (53) Mahato, R. I., Rolland, A., and Tomlinson, E. (1997) Cationic lipid-based gene delivery systems: pharmaceutical perspectives. Pharm. Res. 14, 853-859. (54) Verma, I. M., and Somia, N. (1997) Gene therapy promises, problems and prospects. Nature 389, 239-242. (55) Hsieh, H.-P., Muller, J. G., and Burrows, C. J. (1994) Structural effects in novel steroidal polyamine-DNA binding. J. Am. Chem. Soc. 116, 12077-12078. (56) Hsieh, H.-P., Muller, J. G., and Burrows, C. J. (1995) Synthesis and DNA binding properties of C3-, C12-, and C24substituted amino-steroids derived from bile acids. Bioorg. Med. Chem. 3, 823-835. (57) Muller, J. G., Ng, M. M. P., and Burrows, C. J. (1996) Hydrophobic vs Coulombic interactions in the binding of steroidal polyamines to DNA. J. Mol. Recognit. 9, 143-148. (58) Felgner, P. L., Barenholz, Y., Behr, J.-P., Cheng, S. H., Cullis, P., Huang, L., Jessee, J. A., Seymour, L., Szoka, F., Thierry, A. R., Wagner, E., and Wu, G. (1997) Nomenclature for synthetic gene delivery systems. Hum. Gene Ther. 8, 511512. (59) Cain, B. F., Baguley, B. C., and Denny, W. A. (1978) Potential antitumor agents. 28. Deoxyribonucleic acid polyintercalating agents. J. Med. Chem. 21, 658-668. (60) Gershon, H., Ghirlando, R., Guttman, S. B., and Minsky, A. (1993) Mode of formation and structural features of DNAcationic liposome complexes used for transfection. Biochemistry 32, 7143-7151. (61) Tables of Spectral Data for Structure Determination of Organic Compounds (1989) 2nd ed., pp C5-C177 and H15H80, Springer-Verlag, Berlin. (62) Waterhous, D. V., Barnes, S., and Muccio, D. D. (1985) Nuclear magnetic resonance spectroscopy of bile acids. Development of two-dimensional NMR methods for the elucidation of proton resonance assignments for five common hydroxylated bile acids, and their parent bile acid, 5β-cholanoic acid. J. Lipid Res. 26, 1068-1078. (63) Adlam, G., Blagbrough, I. S., Taylor, S., Latham, H. C., Haworth, I. S., and Rodger, A. (1994) Multiple binding modes with DNA of anthracene-9-carbonyl-N1-spermine probed by LD, CD, normal absorption, and molecular modelling compared with those of spermidine and spermine. Bioorg. Med Chem. Lett. 4, 2435-2440. (64) Rodger, A., Blagbrough, I. S., Adlam, G., and Carpenter, M. L. (1994) DNA binding of spermine derivatives: spectroscopic study of anthracene-9-carbonyl-N1-spermine with poly[d(G-C).d(G-C)] and poly[d(A-T).d(A-T)]. Biopolymers 34, 1583-1593. (65) Rodger, A., Taylor, S., Adlam, G., Blagbrough, I. S., and Haworth, I. S. (1995) Multiple DNA-binding modes of anthracene-9-carbonyl-N1 spermine. Bioorg. Med. Chem. 3, 861-872. (66) Blagbrough, I. S., Taylor, S., Carpenter, M. L., Novoselskiy, V., Shamma, T., and Haworth, I. S. (1998) Asymmetric intercalation of N1-(acridin-9-ylcarbonyl)spermine at homopurine sites of duplex DNA. Chem. Commun. 929-930. (67) Tam, S. C., and Williams, R. J. P. (1985) Electrostatics and biological systems. Struct. Bonding 63, 103-151. (68) Rowatt, E., and Williams, R. J. P. (1992) The binding of spermine and magnesium to DNA. J. Inorg. Biochem. 46, 8797. (69) Braithwaite, A. W., and Baguley, B. C. (1980) Existence of an extended series of antitumor compounds which bind to deoxyribonucleic acid by nonintercalative means. Biochemistry 19, 1101-1106. (70) Stewart, K. D. (1988) The effect of structural changes in a polyamine backbone on its DNA-binding properties. Biochem. Biophys. Res. Commun. 152, 1441-1446. (71) Edwards, M. L., Snyder, R. D., and Stemerick, D. M. (1991) Synthesis and DNA-binding properties of polyamine analogues. J. Med. Chem. 34, 2414-2420. (72) Stewart, K. D., and Gray, T. A. (1992) Survey of the DNA binding properties of natural and synthetic polyamino compounds. J. Phys. Org. Chem. 5, 461-466.

Bioconjugate Chem., Vol. 13, No. 3, 2002 489 (73) Delcros, J.-G., Sturkenboom, M. C. J. M., Basu, H. S., Shafer, R. H., Szo¨llo¨si, J., Feuerstein, B. G., and Marton, L. J. (1993) Differential effects of spermine and its analogues on the structures of polynucleotides complexed with ethidium bromide. Biochem. J. 291, 269-274. (74) Cai, J., Soloway, A. H., Barth, R. F., Adams, D. M., Hariharan, J. R., Wyzlic, I. M., and Radcliffe, K. (1997) Boroncontaining polyamines as DNA targeting agents for neutron capture therapy of brain tumors: synthesis and biological evaluation. J. Med. Chem. 40, 3887-3896. (75) Sobell, H. M., Tsai, C.-C., Jain, S. C., and Gilbert, S. G. (1977) Visualization of drug-nucleic acid interactions at atomic resolution. J. Mol. Biol. 114, 333-365. (76) Widom, J., and Baldwin, R. L. (1980) Cation-induced toroidal condensation of DNA. Studies with Co3+(NH3)6. J. Mol. Biol. 144, 431-453. (77) Hud, N. V., Downing, K. H., and Balhorn, R. (1995) A constant radius of curvature model for the organization of DNA in toroidal condensates. Proc. Natl. Acad. Sci. U.S.A. 92, 3581-3585. (78) Hud, N. V. (1995) Double-stranded DNA organization in bacteriophage heads: An alternative toriod-based model. Biophys. J. 69, 1355-1362. (79) Ha¨rd, T., Nielsen, P. E., and Norden, B. (1988) Molecular flexibility of extended and compacted polynucleosomes - a steady-state fluorescence polarization study. Eur. J. Biophys. 16, 231-242. (80) Basu, H. S., Schwietert, H. C. A., Feuerstein, B. G., and Marton, L. J. (1990) Effects of variation in the structure of spermine on the association with DNA and the induction of DNA conformational changes. Biochem. J. 269, 329-334. (81) Wilson, R. W., and Bloomfield, V. A. (1979) Counterioninduced condensation of deoxyribonucleic acid. A lightscattering study. Biochemistry 18, 2192-2196. (82) Manning, G. S. (1978) The molecular theory of polyelectrolyte solutions with applications to the electrostatic properties of polynucleotides. Q. Rev. Biophys. 11, 179-246. (83) Haq, I., Ladbury, J. E., Chowdhry, B. Z., Jenkins, T. C., and Chaires, J. B. (1997) Specific binding of Hoechst 33258 to the d(CGCAAATTTGCG)2 duplex: calorimetric and spectroscopic studies. J. Mol. Biol. 271, 244-257. (84) Mahler, H. R., and Mehrotra, B. D. (1963) The interaction of nucleic acids with diamines. Biochim. Biophys. Acta 68, 211-233. (85) Yoshikawa, Y., and Yoshikawa, K. (1995) Diaminoalkanes with an odd number of carbon atoms induce compaction of a single double-stranded DNA chain. FEBS Lett. 361, 277-281. (86) Budker, V. G., Godovikov, A. A., Naumova, L. P., and Slepneva, I. A. (1980) Interaction of polynucleotides with natural and model membranes. Nucleic Acids Res. 8, 24992515. (87) Smith J. G., Walzem, R. L., and German, J. B. (1993) Liposomes as agents of DNA transfer. Biochim. Biophys. Acta 1154, 327-340. (88) Sternberg, B., Sorgi, F. L., and Huang, L. (1994) New structures in complex formation between DNA and cationic liposomes visualized by freeze-fracture electron microscopy. FEBS Lett. 356, 361-366. (89) Reimer, D. L., Zhang, Y., Kong, S., Wheeler, J. J., Graham, R. W., and Bally, M. B. (1995) Formation of novel hydrophobic complexes between cationic lipids and plasmid DNA. Biochemistry 34, 12877-12883. (90) Bally, M. B., Zhang, Y. P., Wong, F. M. P., Kong, S., Wasan, E., and Reimer, D. L. (1997) Lipid/DNA complexes as an intermediate in the preparation of particles for gene transfer: an alternative to cationic liposome/DNA aggregates. Adv. Drug Delivery Rev. 24, 275-290. (91) Gosule, L. C., and Schellmann, J. A. (1976) Compact form of DNA induced by spermidine. Nature 259, 333-335. (92) Gosule, L. C., and Schellmann, J. A. (1978) DNA condensation with polyamines I. Spectroscopic studies. J. Mol. Biol. 121, 311-327. (93) Chattoraj, D. K., Gosule, L. C., and Schellmann, J. A. (1978) DNA condensation with polyamines II. Electron microscopic studies. J. Mol. Biol. 121, 327-337.

490 Bioconjugate Chem., Vol. 13, No. 3, 2002 (94) Mou, J., Czajkowsky, D. M., Zhang, Y., and Shao, Z. (1995) High-resolution atomic-force microscopy of DNA: the pitch of the double helix. FEBS Lett. 371, 279-282. (95) Hansma, H. G., Golan, R., Hsieh, W., Lollo, C. P., MullenLey, P., and Kwoh, D. (1998) DNA condensation for gene therapy as monitored by atomic force microscopy. Nucleic Acids Res. 26, 2481-2487. (96) Lin, Z., Wang, C., Feng, X., Liu, M., Li, J., and Bai, C. (1998) The observation of the local ordering characteristics of spermidine-condensed DNA: atomic force microscopy and polarizing microscopy studies. Nucleic Acids Res. 26, 32283234. (97) Bo¨ttcher, C., Endisch, C., Fuhrhop, J.-H., Catterall, C., and Eaton, M. (1998) High-yield preparation of oligomeric C-type DNA toroids and their characterization by cryoelectron microscopy. J. Am. Chem. Soc. 120, 12-17. (98) Anderson, C. F., and Record M. T. (1995) Salt nucleic acid interactions. Annu. Rev. Phys. Chem. 46, 657-700. (99) Zhang, W., Bond, J. P., Anderson, C. F., Lohman, T. M., and Record, M. T. (1996) Large electrostatic differences in the binding thermodynamics of a cationic peptide to oligomeric and polymeric DNA. Proc. Natl. Acad. Sci. U.S.A. 93, 2511-2516. (100) Record, M. T., Zhang, W. T., and Anderson, C. F. (1998) Analysis of effects of salts and uncharged solutes on protein and nucleic acid equilibria and processes: a practical guide to recognizing and interpreting polyelectrolyte effects, Hofmeister effects, and osmotic effects of salts. Adv. Protein Chem. 51, 281-353.

Geall et al. (101) Zhang, W. T., Ni, H. H., Capp, M. W., Anderson, C. F., Lohman, T. M., and Record M. T. (1999) The importance of Coulombic end effects: experimental characterization of the effects of oligonucleotide flanking charges on the strength and salt dependence of oligocation (L8+) binding to singlestranded DNA oligomers. Biophys. J. 76, 1008-1017. (102) Jou, W. S., and Chun, P. W. (1991) Molecular mechanics of the formation of cholic acid micelles. J. Mol. Graphics 9, 237-246. (103) Fuhrhop, J.-H., and Ko¨ning, J. (1994) in Membranes and Molecular Assemblies: The Synkinetic Approach, pp 34-36 and 203-204, RSC, Cambridge, U.K. (104) Zhang, L.-H., Janout, V., Renner, J. L., Uragami, M., and Regen S. L. (2000) Enhancing the “stickiness” of bile acids to cross-linked polymers: a bioconjugate approach to the design of bile acid sequestrants. Bioconjugate Chem. 11, 397-400. (105) Geall, A. J., Al-Hadithi, D., and Blagbrough, I. S. (2000) Cheno-, urso- and deoxycholic acid spermine conjugates: relative binding affinities for calf thymus DNA. Tetrahedron 56, 3439-3447. (106) Rouzina, I., and Bloomfield, V. A. (1998) DNA bending by small, mobile multivalent cations. Biophys. J. 74, 31523164. (107) Shen, M. R., Downing, K. H., Balhorn, R., and Hud, N. V. (2000) Nucleation of DNA condensation by static loops: formation of DNA toroids with reduced dimensions. J. Am. Chem. Soc. 122, 4833-4834.

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