pH-sensitive PEG lipids containing orthoester linkers: new potential tools for nonviral gene delivery

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pH-sensitive PEG lipids containing orthoester linkers: new potential tools for nonviral gene delivery Christophe Masson, Marie Garinot, Nathalie Mignet, Barbara Wetzer, Philippe Mailhe, Daniel Scherman, Michel Bessodes* Unite´ de Pharmacologie Chimique et Ge´ne´tique, Faculte de Pharmacie, Universite Rene Descartes, FRE 2463-U640, CNRS/INSERM/Univ. Paris V/ENSCP, 4, avenue de l’Observatoire, Paris Cedex 06 75270, France Received 15 March 2004; accepted 13 July 2004 Available online 25 August 2004

Abstract The synthesis and properties of pH-sensitive polyethylene glycol (PEG) lipids are described. The sensitivity of these conjugates to slightly acidic pH was clearly related to the structure of the orthoester linkage involved. It was found that pHsensitive PEG lipids stabilized cationic lipid/DNA isoelectric complexes as efficiently as their non-pH-sensitive PEG analogs at neutral pH. Lowering the pH resulted in the precipitation of the complexes bearing pH-sensitive PEG lipids as a consequence of their degradation. In contrast, insertion of non-pH-sensitive PEG lipids maintained the complex colloidal stability even at lower pH. In vitro results showed a significant increase in transfection with formulations containing pH-sensitive PEG lipids versus non-pH-sensitive analogs. These conjugates show promising properties as lipoplex-stabilizing agents at neutral pH, which could be triggered by a mild acidic environment such as that occurring in solid tumors, inflammatory tissues, and intracellular endosomal compartments. D 2004 Elsevier B.V. All rights reserved. Keywords: Gene delivery; Lipoplex; pH-sensitive PEG lipids; Tumor; Colloidal stabilization

1. Introduction Liposomes have been extensively used for the encapsulation of therapeutic drugs [1] and, more

* Corresponding author. Tel.: +33-1-53-73-95-66; fax: +33-143-26-69-18. E-mail address: [email protected] (M. Bessodes). 0168-3659/$ - see front matter D 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.jconrel.2004.07.016

recently, as vectors for gene delivery [2]. Among them, cationic liposomes associated with plasmid DNA, namely lipoplexes [3], have shown very promising results for in vitro transfection of various cell lines [4]. However, their application in vivo has so far led to rather disappointing results. It is currently acknowledged that these cationic particles interact in a nonspecific manner in vivo with anionic species, such as plasmic membrane components and circulating proteins. This leads to undesirable effects such as

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platelet aggregation or complement activation, and to rapid clearance from the bloodstream [5,6]. Coating these cationic particles with polyethylene glycol–lipid (PEG lipid) conjugates has reduced these effects by sterically shielding the lipoplexes peripheral cationic charges, therefore increasing the circulation time of the particles as a consequence of the reduction of nonspecific interactions [7]. Ideally, these particles should circulate until they reach the target organs or tissues, either by specific targeting [8] or by passive accumulation, and be triggered to release their therapeutic content. The lower pH in some therapeutic targets such as tumors, and inflammation or ischemia sites, has been exploited to help the release of encapsulated pharmaceutical drugs [9–11]. The pH variations in these tissues are slight and the chemical trigger should respond efficiently to these variations, while being stable at neutral pH. To this end, different pHsensitive linkers have been studied [12]. Among the different pH-sensitive linkers, the orthoester group is quite easy to handle and should provide enough sensitivity to respond to such pH variations as those encountered in the therapeutic targets already mentioned. Furthermore, the orthoester hydrolysis rate is dependent upon their chemical structure and substituents [13,14], thus offering a convenient means to modulate sensitivity. Thus, the introduction of structural variations, such as a six-membered ring structure or methyl substitution, enhances the rate of hydrolysis by several tens or hundreds of times, respectively. [15] We

therefore selected this group as a pH-sensitive chemical linker between the PEG and the hydrophobic chain of PEG lipids used for lipoplex shielding. In this way, we expected to release the PEG from the stabilized lipoplex, thus revealing the cationic entities at their surface. This would in turn induce lipoplex internalization in the targeted cells, thus favoring gene delivery and, ultimately, protein expression (Scheme 1). In the course of our present work [16], an article describing the use of a PEG–diorthoester lipid conjugate with conclusive results appeared [13]. We describe here the synthesis of PEG–orthoester conjugates with different hydrophobic moieties and orthoester structures, their physicochemical properties, and the transfection efficiency of coformulated lipoplexes.

2. Materials and methods 2.1. Materials The PEGs and chemicals were obtained from Sigma-Aldrich. The solvents were purchased from SDS and were of synthesis grade. Analytical TLC was run on Merck silica-precoated aluminum plates, and silica for preparative chromatography was also purchased from Merck. The PEG derivatives were revealed by a 50/50 solution of 1 M iodine and 1 M H2SO4. The synthesis of the polyamine lipid RPR120535 used in the formation of liposomes has

Scheme 1. pH-triggered unshielding of PEG lipoplex.

already been described [17]. Dioleylphosphatidylethanolamine (DOPE) was purchased from Avanti Polar. The pH-stable lipid PEG analogs used as controls were, respectively, polyoxyethylene-100 stearyl ether (Brij 700; Sigma) and cholesteryl PEG (chol-PEG), whose structure is presented in Scheme 2. Chol-PEG was obtained in one step from the reaction of cholesteryl chloroformate and a-amino-N-methoxy-PEG. 2.2. Structural analysis 1

H nuclear magnetic resonance (NMR) spectra have been recorded at 300, 400, and 600 MHz on Bruker spectrometers using tetramethylsilane as internal reference. Mass spectra have been obtained using

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two different methods: electrospray (ES) with an LCTOF micromass or desorption/chemical ionization (D/CI) with a SSQ7000 Finnigan. 2.3. Synthesis 2.3.1. 2,2,2-Trifluoro-N-(2,3-dihydroxy-propyl)-acetamide (1) To a stirred solution of 3-amino-1,2-propanediol (16.8 g, 0.180 mol) in THF (125 ml), ethyl trifluoroacetate (25 ml, 0.210 mol) was added dropwise at 0 8C. After 2 h, the mixture was evaporated and the residue dissolved in ethyl acetate (150 ml). It was washed with aqueous potassium hydrogen sulfate (0.5 M, 20 ml 2) and brine (20 ml), then dried over MgSO4 and concentrated to a

Scheme 2. PEG orthoester lipids and stable chol-PEG.

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colorless viscous liquid (32 g, 95%). 1H NMR (CDCl3, 300 MHz): d (ppm) 2.12 (1H, t, J 5.5 Hz, OH), 2.61 (1H, d, J 5 Hz, OH), 3.25–3.85 (4H, m, 2 CH 2), 3.92 (1H, m, CH), 6.88 (1H, m, NH); MS (D/CI) m/z calculated for (M+H+) C5H8F3NO3 188, found 188. 2.3.2. 2,2,2-Trifluoro-N-(2-methoxy-[1,3]dioxolan-4ylmethyl)-acetamide (2) The diol 1 (29 g, 0.155 mol) was dissolved in dichloromethane (75 ml) and trimethyl orthoformate (75 ml, 0.685 mol); p-toluene sulfonic acid (PTSA; 0.3 g, 0.0017 mol) was then added. The solution was stirred at room temperature for 2 h. The solution was then diluted with dichloromethane (250 ml), washed successively with saturated aqueous sodium hydrogen carbonate (3 100 ml) and brine (100 ml), and dried over anhydrous potassium carbonate. It was filtered and concentrated to a colorless oil (30 g, 85%). 1H NMR (CDCl3, 300 MHz): mixture of two diastereomers 50/50: d (ppm) 3.33 and 3.37 (3H, s, O-CH 3), 3.35–3.80 (3H, m, N-CH 2, and O-CH 2), 4.10–4.25 (1H, m, O-CH 2), 4.50 (1H, m, CH), 5.73 and 5.78 (1H, s, (RO)3-CH), 6.66 and 7.55 (1H, s, NH); MS (D/CI) m/z calculated for (M+NH4+) C7H10F3NO4 247, found 247. 2.3.3. 2,2,2-Trifluoro-N-(2-octadecyloxy-[1,3]dioxolan-4-ylmethyl)-acetamide (3) To a solution of 2 (3 g, 13 mmol) and octadecyl alcohol (3.54 g, 13 mmol) in toluene (20 ml), pyridinium p-toluene sulfonate (0.032 g, 0.13 mmol) was added. The mixture was refluxed for 2 h, cooled, and diluted with cyclohexane (100 ml). The solution was washed successively with aqueoussaturated sodium hydrogen carbonate (3 30 ml) and brine (30 ml), dried over anhydrous potassium carbonate, filtered, and evaporated. The syrup was purified by column chromatography and eluted with cyclohexane/ethyl acetate (6:4) to give 3 as a mixture of diastereomers 50/50 (4 g, 65%). 1H NMR (CDCl3, 300 MHz): d (ppm) 0.89 (3H, t, J 7, CH 3), 1.20–1.45 and 1.5–1.7 (32H, m, CH 2), 3.35– 3.8 (5H, m, N-CH 2, O-CH 2), 4.05–4.25 (1H, m, OCH 2), 4.48 (1H, m, CH), 5.81 and 5.86 (1H, s, (RO)3-CH), 6.59 and 7.49 (1H, br s, NH); MS (D/ CI) m/z calculated for (M NH4+) C24H44F3NO4 485, found 485.

2.3.4. 2,2,2-Trifluoro-N-(2-cholesteryloxy-[1,3]dioxolan-4-ylmethyl)-acetamide (4) Cholesterol (3.9 g, 10 mmol) was reacted with an equimolar amount of 2 under the same conditions as described above. Column purification gave 4 as a mixture of diastereomers (4 g, 68%). 1H NMR (CDCl3, 400 MHz): mixture of two diastereomers 50/50: d (ppm) 0.69 (3H, s, CH 3), 0.88 (6H, m, CH 3), 0.90–1.65 and 1.86–2.00 (26H, m, CH 2), 0.94 (3H, d, J 7 Hz, CH 3), 1.02 (3H, s, CH 3), 2.33 (2H, m, CH 2), 3.30–3.85 (4H, m, CH 2), 4.00 and 4.21 (1H, m, CH 2), 4.48 (1H, m, CH 2), 5.37 (1H, m, CH), 5.93 and 5.98 (1H, s, (RO)3-CH), 6.56 and 7.55 (1H, br s, NH); MS (D/CI) m/z calculated for (M+NH4+) C33H52F3NO4 601, found 601. 2.3.5. C-(2-octadecyloxy-[1,3]dioxolan-4-yl)-methylamine (5) The trifluoroacetamide 3 (6.12 g, 13 mmol) was dissolved in THF (20 ml), the solution was cooled to 0 8C, and 4% sodium hydroxide (30 ml) was added. The mixture was vigorously stirred at room temperature for 4 h, extracted with diethyl ether (3 200 ml), dried over potassium carbonate, filtered, and evaporated. The residue was purified by column chromatography with dichloromethane/methanol (9:1) as the eluant. Compound 5 was obtained as a white powder (4.38 g, 90%). 1H NMR (CDCl3, 300 MHz): mixture of two diastereomers 50/50: d (ppm) 0.89 (3H, t, J 7 Hz, CH 3), 1.2–1.45 and 1.58 (32H, m, CH 2), 2.75–3.0 (2H, m, N-CH 2), 3.53 (2H, m, OCH 2), 3.65–3.85 (1H, m, CH), 4.0–4.40 (2H, m, OCH 2), 5.81 and 5.84 (1H, s, (RO)3-CH); MS (D/CI) m/z calculated for (M+H+) C22H45NO3 372, found 372. 2.3.6. C-(2-cholesteryloxy-[1,3]dioxolan-4-yl)-methylamine (6) In a similar way as for 5, compound 4 was reacted with aqueous sodium hydroxide (15 ml, 4%) in THF (10 ml) for 4 h. Identical work-up and purification gave 6 as a white powder (2.7 g, 92%). 1H NMR (CDCl3, 300 MHz): mixture of two diastereomers 50/ 50: d (ppm) 0.69 (3H, s, CH 3), 0.85–1.75 and 1.86– 2.00 (26H, m, CH 2), 0.88 (6H, m, CH 3), 0.93 (3H, d, J 7 Hz, CH 3), 1.01 (3H, s, CH 3), 2.33 (2H, m, CH 2), 2.75–3.00 (2H, m, CH 2), 3.50 (1H, m, CH), 3.71, 3.85, 4.05, and 4.16 (2H, m, CH 2), 4.20 and 4.35 (1H,

m, CH), 5.36 (1H, m, CH), 5.93 and 5.96 (1H, s, (RO)3-CH); MS (D/CI) m/z calculated for (M+H+) C31H53NO3 488, found 488. 2.3.7. a-Methoxy-x-{N-(2-octadecyloxy-[1,3]dioxolan-4-yl)methylamido}-polyethyleneglycol110 (7) MeO–PEG110–COOH (1.2 g, average M W 5000, 0.24 mmol) and compound 5 (0.1 g, 1.34 mmol) were dissolved in dichloromethane (5 ml). Triethylamine (0.188 ml, 1.34 mmol) and BOP reagent (0.143 g, 0.32 mmol) were added and the mixture was stirred at room temperature for 1 h. The reaction medium was precipitated with diethyl ether (60 ml), centrifuged, and washed with ether (4 20 ml). The white powder obtained was purified by preparative high-performance liquid chromatography (HPLC) to give 7 (0.41 g, 32%). 1H NMR (CDCl3, 300 MHz) mixture of two diastereomers 50/50: d (ppm) 0.90 (3H, t, J 7 Hz, CH 3), 1.20–1.40 and 1.50–1.75 (32H, m, CH 2), 3.39 (3H, s, O-CH 3), 3.40–3.95 (448H, m, O-CH 2-CH 2O), 4.02 and 4.03 (2H, s, CH 2), 4.00–4.25 (3H, m, CH, CH 2), 5.80 and 5.84 (1H, s, NH); MS (ES) m/z calculated for (M5+) C247H493NO116 1066.4, found 1068.9. 2.3.8. a-Methoxy-x-{N-(2-cholesteryloxy-[1,3]dioxolan-4-yl)methylamido}-polyethyleneglycol110 (8) In a similar procedure as above, MeO–PEG110– COOH (1 g, average M W 5000, 0.2 mmol) and compound 6 (0.097 g, 0.2 mmol) were coupled with BOP reagent (0.12 g, 0.27 mmol) in the presence of triethylamine (0.14 ml, 1 mmol ) to give 8, which was purified by preparative HPLC (0.38 g, 35%). 1H NMR (CDCl3, 400 MHz): mixture of two diastereomers 50/ 50: d (ppm) 0.68 (3H, s, CH 3), 0.85–1.75 and 1.85– 2.00 (26H, m, CH 2), 0.87 (6H, m, 2CH 3), 0.92 (3H, d, J 7 Hz, CH 3), 1.00 (3H, s, CH 3), 2.33 (2H, m, CH 2), 3.39 (3H, s, CH 3), 3.40–3.90 (448H, m, OCH 2-CH 2-O), 4.00–4.20 (1H, m, CH), 4.01 and 4.04 (2H, s, CH 2), 4.28 and 4.46 (1H, m, CH), 5.35 (1H, m, CH), 5.90 and 5.95 (1H, s, NH); MS (ES) m/z calculated for (M4+) C256H501NO116 1361.8, found 1364.3. 2.3.9. 2,2,2-Trifluoro-N-(2-hydroxy-1-hydroxymethylethyl)-acetamide (9) Ethyl trifluoroacetate (5.8 ml, 48 mmol) was added dropwise to a solution of serinol (4 g, 43 mmol) in

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THF (20 ml) at 0 8C. After 2 h, the reaction mixture was concentrated, the residue dissolved in ethyl acetate (25 ml), and this solution washed twice with aqueous potassium hydrogen sulfate (0.5 M, 5 ml) and brine (5 ml). It was dried over anhydrous MgSO4, filtered, and evaporated to give a white powder (8.1 g, 99%). 1H NMR (CDCl3, 300 MHz): d (ppm) 3.65 (4H, mt, 2 CH 2), 3.9 (1H, m, CH), 4.3 (2H, t, 2 OH), 7.93 (1H, m, NH); MS (D/CI) m/z calculated for (M+H+) C5H8F3NO3 188, found 188. 2.3.10. 2,2,2-Trifluoro-N -(2-methoxy-2-methyl[1,3]dioxan-5-yl)-acetamide (10) The diol 3 (7.9 g, 42 mmol) was dissolved in dichloromethane (30 ml); trimethyl orthoacetate (16.1 ml, 127 mmol) and PTSA (0.073 g, 0.4 mmol) were then added and the solution stirred at room temperature for 3 h. The solution was then diluted with dichloromethane (100 ml), washed successively with saturated aqueous sodium hydrogen carbonate (3 40 ml) and brine (40 ml), and dried over anhydrous potassium carbonate. It was filtered and concentrated to a white powder (9.9 g, 96%). 1H NMR (CDCl3, 300 MHz): mixture of two diastereomers 75/25: d (ppm) 1.49 and 1.50 (3H, s, CH 3), 3.34 and 3.35 (3H, s, O-CH 3), 3.66 (1H, dm, J 12 Hz, O-CH 2), 3.82 (1H, dd, J 11 Hz, J 8, OCH 2), 3.9–4.0 and 4.20–4.35 (1H, m, CH), 3.97 (1H, dd, J 11 Hz, J 5, CH 2), 4.33 (1H, dm, J 12 Hz, CH 2), 6.38 and 7.04 (1H, br s, NH); MS (D/CI) m/z calculated for (M+H+) C8H12F3NO4 244, found 244. 2.3.11. 2,2,2-Trifluoro-N-(2-octadecyloxy-2-methyl[1,3]dioxan-5-yl)-acetamide (11) The cyclic orthoester 10 (3 g, 12.3 mmol) and octadecyl alcohol (3 g, 11.1 mmol) were melted together at 80 8C during 2 h. The fused reaction mixture was then poured into cold acetonitrile (50 ml), where it precipitated. Compound 11 was obtained by recrystallization from acetonitrile (2.85 g, 53%). 1H NMR (CDCl3, 400 MHz): d (ppm) 0.9 (3H, t, J 7 Hz, CH 3), 1.20–1.50 (30H, m, CH 2), 1.52 (3H, s, CH 3), 1.64 (2H, m, CH 2), 3.48 (2H, t, J 7 Hz, CH 2), 3.67 (2H, dd, J 1.5 Hz, J 12, CH 2), 3.94 (1H, dm, J 8.5 Hz, CH), 4.36 (2H, dd, J 1 Hz, J 12 Hz, CH 2), 7.14 (1H, br d, J 8 Hz, NH); MS (D/CI)

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m/z calculated for (M+H+) C25H46F3NO4 482, found 482. 2.3.12. 2,2,2-Trifluoro-N-{2-(3-hydroxypropyl-cholesterylcarbamate)-2-methyl-[1,3]dioxan-5-yl}-acetamide (12) A mixture of 10 (0.5 g, 2 mmol) and (3hydroxypropyl)-cholesterylcarbamate (1 g, 2 mmol) in toluene was refluxed for 4 h in the presence of catalytic pyridinium p-toluene sulfonate (5 mg, 0.02 mmol). The solution was then evaporated and the crude residue purified by column chromatography (cyclohexane/ethylacetate, 1:1) to give 12 (0.98 g, 70%). 1H NMR (CDCl3, 300 MHz): d (ppm) 0.70 (3H, s, CH 3), 0.88 (6H, d, CH 3), 0.94 (3H, d, CH 3), 1.04 (3H, s, CH 3), 2.12 (3H, s, CH 3), 2.32 (2H, m, CH 2), 3.38 (2H, bq, CH 2NH), 3.75 (3H, m, CH 2O and CHNH), 4.30 (4H, m, 2 CH 2O), 4.52 (1H, m, CH), 4.86 (1H, bs, NH), 5.40 (1H, bd, CH), 6.88 (1H, bs, NH); MS (ES) m/z calculated for (M+Na+) C38H61F3N2O6 721, found 721. 2.3.13. 2-Octadecyloxy-2-methyl-(1,3)dioxan-5ylamine (13) Compound 11 (1 g, 2 mmol) was dissolved in a mixture of THF (10 ml) and NaOH (1 N, 10 ml), and stirred for 4 h at room temperature. The partly concentrated solution was extracted with diethyl ether (3 100 ml) to give 13 as a white powder after evaporation (0.81 g, quantitative). 1H NMR (CDCl3, 400 MHz): d (ppm) 0.89 (3H, t, J 7 Hz, CH 3), 1.20–1.50 (30H, m, 15 CH 2), 1.49 (3H, s, CH 3), 1.62 (2H, m, CH 2), 2.71 (1H, m, CH), 3.46 (2H, t, J 7 Hz, CH 2-O), 3.54 (2H, dd, J 10 Hz, CH 2O), 4.30 (2H, dd, J 10 Hz; 1.5, CH 2-O); MS (D/CI) m/z calculated for (M+H+) C23H47NO3 386, found 386. 2.3.14. 2-(3-Hydroxypropyl-cholesterylcarbamate)-2methyl-[1,3]dioxan-5-ylamine (14) In a similar procedure as above, compound 12 (0.1 g, 0.14 mmol) was treated overnight at room temperature by a mixture of NaOH (1 N, 1 ml) and isopropanol (1 ml). It was then partially evaporated and extracted with dichloromethane. The organic phase was evaporated to give crude 14, which was purified by column chromatography and eluted with dichloromethane/methanol (85:15), buf-

fered by 0.1% of triethylamine (80 mg, 92%). 1H NMR (CDCl3, 300 MHz): d (ppm) 0.69 (3H, s, CH 3), 0.88 (6H, d, CH 3), 0.94 (3H, d, CH 3), 1.04 (3H, s, CH 3), 1.54 (3H, s, CH 3), 2.34 (2H, m, CH 2), 2.82 (1H, m, N-CH), 3.35 (2H, m, N-CH 2), 3.6 (4H, m, O-CH 2), 4.28 (2H, m, O-CH 2), 4.52 (1H, m, CH), 4.82 (1H, bs, NH), 5.38 (1H, m, CH); MS (ES) m/z calculated for (M+H+) C36H62N2O5 603, found 603. 2.3.15. a-Methoxy-x-{N-(2-methyl-2-octadecyloxy[1,3]dioxan-5-yl)-amido}-polyethyleneglycol110 (15) To a solution of MeO–PEG110–COOH (1.15 g, 0.23 mmol) in dichloromethane (5 ml) were added successively triethylamine (0.18 ml, 0.13 mmol), 13 (0.1 g, 0.26 mmol), and BOP reagent (0.172 g, 0.39 mmol). The mixture was stirred for 1 h at room temperature, then the desired compound was precipitated by the addition of diethyl ether (60 ml) and centrifuged. The precipitate was washed several times with ether, dried over anhydrous MgSO4, and purified by preparative HPLC to give pure 15 (0.42 g, 34%). 1H NMR (CDCl3, 400 MHz): d (ppm) 0.89 (3H, t, J 7 Hz, CH 3), 1.20–1.50 (30H, m, CH 2), 1.48 (3H, s, CH 3), 1.55–1.75 (2H, m, CH 2), 3.39 (3H, s, O-CH 3), 3.40–3.90 (448H, m, O-CH 2), 3.89 (1H, m, CH), 4.05 (2H, s, O-CH 2), 4.30 (2H, m, OCH 2), 7.57 (1H, d, J 8.5 Hz, NH); MS (ES) m/z calculated for (M+Na)4+ C246H491NO115 1331, found 1335.2. 2.3.16. a-Methoxy-x-{N-2-(3-hydroxypropyl-cholesterylcarbamate)-2-methyl-[1,3]dioxan-5-yl-amido}polyethyleneglycol110 (16) To a solution of compound 14 (0.12 g, 0.2 mmol) and MeO–PEG 110 –COOH (1 g, 0.2 mmol) in dichloromethane (5 ml) were added triethylamine (0.16 ml, 1.1 mmol) and BOP (0.12 g, 0.27 mmol). After 1 h at room temperature, diethyl ether was added (60 ml) and the precipitate was centrifuged and washed several times with ether. It was dried and purified by HPLC to give 16 as a white powder (0.44 g, 40%). 1H NMR (CDCl3, 500 MHz): d (ppm) 0.68 (3H, s, CH 3), 0.88 (6H, d, CH 3), 0.93 (3H, d, CH 3), 1.03 (3H, s, CH 3), 2.08 (3H, s, CH 3), 2.32 (2H, m, CH 2), 3.34 (2H, q, N-CH 2), 3.38 (3H, s, O-CH 3), 3.5–3.8 (448H, m, O-CH 2), 4.02 (2H, m , O-CH 2), 4.25 (4H, m, 2 O-CH 2), 4.52 (1H, m, CH), 4.90

(1H, bt, NH), 5.38 (1H, m, CH), 7.48 (1H, d, NH); MS (ES) m/z calculated for (M4+) C259H506N2O117 1379.5, found 1382. 2.4. Degradation studies The stability of the PEG lipid products under acidic conditions was evaluated at different pH values in 0.1 M acetate buffer. The percentage of degradation was deduced from the integration of the peaks corresponding either to the starting material or to the methoxy-PEG product of hydrolysis, using an HPLC (Merck Hitachi) coupled to an Evaporative Light Scattering Detector (ELSD Eurosep). An XTerra RP-8 column (Waters) was used, eluted by a water/acetonitrile linear gradient (1 ml/min, 0–15 min, 20–100% CH3CN), buffered with 0.01% HPLC-grade triethylamine. A stock solution of the PEG lipid (10 mg/ml in 0.1 M triethylammonium acetate buffer, pH 7.5) was made and 0.1-ml aliquots were diluted in 1 ml of the desired buffer (pH 4 and 5), giving the initial time of the reaction. The pH of a sample was measured to ascertain that this buffer mixture did not noticeably affect the final pH. Final concentration of the PEG lipid was in the range of 10 4 M. Measures at close intervals could be achieved by quenching aliquots of the reaction medium with excess triethylamine; other points were obtained through the use of the autosampler timing of the HPLC equipment. All the experiments were carried out at room temperature.

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mM NaCl) was added dropwise under vortex conditions to obtain a ratio of cationic lipid over DNA of 3 nmol/Ag, at which the neutrality of lipid cationic charges over DNA anionic charges was obtained. The size of the complexes was measured as a function of the percentage of PEG lipid by dynamic light scattering with a Coulter N4+ particle sizer at 632 nm. 2.6. Influence of PEG lipid percentage on DNA compaction Compaction of DNA in the liposomes containing different percentages of PEG lipid was verified by addition of ethidium bromide (3 Al of a 1 g/l solution for 8 Ag of DNA) and measurement of the fluorescence (k em 590 nm), which decreases when DNA is compacted. The DNA compaction was also evidenced by the loss of DNA electrophoretic mobility on agarose gel. 2.7. Stability of the complexes at different pH values Lipid/PEG lipid/DNA complexes, prepared as described above, were subjected to different pH values (6.0 and 5.0) in 0.1 M sodium citrate buffer at 37 8C. Size increase was measured as a function of time. The hydrolysis of the pH-sensitive PEG lipids induced a deshielding of the lipoplexes and aggregation of the neutral particles. 2.8. Lipoplex formulation for in vitro experiments

2.5. Lipoplex stabilization Mixing cationic lipids and DNA at a charge ratio lipid/DNA of 1 resulted in neutral particles that rapidly formed aggregates of micrometer size. Two aspects were evaluated: (1) the ability of PEG lipids to stabilize the cationic lipids/DNA neutral complexes and their influence on DNA compaction; and (2) the influence of pH on the lipoplex shielded by the PEG lipids. Complexes were prepared by mixing liposomes of RPR120535/DOPE (1/1) (400 Al, 60 AM, 10 mM Hepes buffer, pH 7.4) with different amounts of PEG lipid (corresponding to 5–30% mol of PEG lipid/lipid in 10 mM Hepes, pH 7.4). After 30 min at room temperature, the DNA solution (0.02 g/l, 400 Al, 300

Lipoplexes were prepared in a Hepes/mes/pipes buffer at pH=8.0, charge ratio (+/ )=10 from liposomes described above (RPR120535/DOPE 1:1). Plasmid DNA used contained the luciferase (Luc) reporter gene under the cytomegalovirus (CMV) promoter. Brij 700 and pH-sensitive alkyl derivatives were added to the preformed lipoplexes at 10% polymer/total lipids ratio, and chol-PEG and pHsensitive cholesteryl derivatives were inserted at 20% ratio. 2.9. Cell culture In vitro experiments were performed on HeLa cells. The cells were grown in minimum essential

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medium (MEM; Gibco), supplemented with 2 mM glutamine, penicillin/streptomycin (100 U/ml and 100 Ag/ml, respectively), and 10% (vol/vol) FBS. Cultures were maintained at 37 8C in a 5% CO2/air incubator. 2.10. Transfection procedure Gene delivery efficiency was evaluated using a plasmid carrying a luciferase reporter gene. HeLa cells were seeded in 24-well plates (3104 cells/well) 1 day before transfection. Immediately before lipofection, cells were washed twice with fresh medium without serum. Then, lipoplexes were added to the cells. Ten percent of serum was added to the cell culture 2 h after lipofection. The cells were incubated for 24 h at 37 8C in the presence of 5% CO2.

The reaction pathway is represented in Scheme 3: orthoester synthons were prepared either by reaction of N-protected 1-amino-propanediol with trimethyl orthoformate or N-protected serinol with trimethyl orthoacetate, under PTSA catalysis to give 2 and 10, respectively. These orthoesters were then reacted with octadecanol, cholesterol, or (3-hydroxypropyl)-cholesteryl carbamate in the presence of a trace of PTSA in refluxing toluene, to give products 3, 4, 11 and 12, respectively. The TFA-protecting groups were removed by the action of aqueous sodium hydroxide in THF and led to compounds 5, 6, 13, and 14, which were condensed with a-carboethoxy-N-methoxy-PEG [18] in the presence of BOP reagent to give final products 7, 8, 15, and 16. 3.2. Studies of PEG lipid hydrolysis

2.11. Luciferase assay Cells were washed with PBS and lysed with 200 Al of cell culture lysis reagent (Promega). Luciferase expression was quantified on 5 Al of centrifuged lysate supernatant using a luciferase assay kit (Promega). Light emission was measured using a luminometer (Multilabel counter 1420 Victor 2 ; EG&G Wallac), equipped with a coinjector that delivered 80 Al of luciferase substrate into 40 Al of cell extracts. Relative light units (RLU) were calculated versus background activity. Light emission was normalized to the protein concentration of cell extracts, determined using the Bradford protein assay kit (Bio-Rad).

3. Results and discussion 3.1. Synthesis of PEG lipids In a preliminary approach, four pH-sensitive compounds were prepared with different substitutions and structures (Scheme 2). Two compounds (7, 15) were obtained with an octadecyl alkyl chain, and two others (8, 16) with a cholesteryl group. Both types comprised an orthoester group with either a five- or six-membered cycle, respectively. The six-membered cycle in compounds 15 and 16 was substituted with a methyl group introduced by the use of the trimethyl orthoacetate reagent.

The rate of hydrolysis of the acid-sensitive PEG compounds 7, 8, 5, and 16 was evaluated at pH 4.0 and 5.0 in 0.1 M acetate buffer (Fig. 1). The percentage of degradation was determined with HPLC by calculating the ratio between the peak area corresponding to the product of hydrolysis and the starting material, respectively. This calculation method was used according to the nonlinear response of ELSD, which excluded accurate standardization between runs. After 30 min, monoalkyl compound 7 was 30% degraded at pH 4.0, and only 10% at pH 5.0, whereas its six-membered ring analog 15 was completely hydrolyzed at pH 4.0, and 30% degraded at pH 5.0. Changing the monocatenar chain (7) for a cholesterol moiety (8) did not significantly alter this behavior. The acidsensitive cholesterol derivative 8 exhibited a slightly better stability, and was stable over 1 h at pH 5.0 and only 25% was degraded at pH 4.0. It was hypothesized that this difference could be due to the greater lipophilicity surrounding the orthoester group in the case of the cholesterol-PEG 8. This could impair the penetration of water molecules at this site, thus slowing down the hydrolysis. Accordingly, compound 16, which includes a spacer between the orthoester group and the cholesterol moiety, was hydrolyzed much faster than its monoalkyl counterpart 15, with a complete degradation of 16 being obtained in 1 h at pH 5.0 and within 10 min at pH 4.0.

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Scheme 3. (i) Trimethyl orthoformate, PTSA, room temperature. (ii) Trimethyl orthoacetate, PTSA, room temperature. (iii) Octadecanol (3 and 11) or cholesterol (4) or (3-hydroxypropyl)-cholesteryl carbamate (12), PTSA, toluene, reflux. (iv) NaOH, H2O/THF (50/50). (v) a-Carboethoxy-N-methoxy-PEG, BOP, Et3N.

As expected from the chemical properties of the orthoester group, compounds bearing a six-membered cycle were more sensitive to hydrolysis than their five-membered cycle analogs. Moreover, in the case of the six-membered ring derivatives 15 and 16, the introduction of a methyl group, providing a greater stability for the intermediate carbocation, further enhanced the hydrolysis reaction [15]. On the other hand, stock solutions in pH 7.5 buffer of the different pH-sensitive lipid and cholesterol PEGs that were used in these experiments did not show noticeable degradation after several days at room temperature and several weeks at 4 8C.

As shown in Fig. 2, the colloidal stabilization of isoelectric lipid/DNA complexes was obtained for a PEG lipid/total lipids ratio of 4–5% with compounds 7, 15, and chol-PEG. Compaction of DNA was not noticeably influenced by the amount of PEG lipid added (Fig. 3). Increasing the percentage of lipid– polymer insertion from 5% to 30% resulted in a 5– 10% fluorescence increase, indicating a better DNA accessibility for ethidium bromide, thus DNA release from the lipoplexes. However, in the range of the amount of PEG lipid necessary for complex stabilization, no significant DNA release was noticed. 3.4. Influence of pH on lipoplex stability

3.3. Stabilization of neutral lipoplexes and DNA compaction Mixing cationic lipids and DNA at a charge ratio lipid/DNA of 1 led to the formation of micrometer-size aggregates. Insertion of an appropriate amount of PEG lipid in lipoplex bilayer inhibited this aggregation [19].

The colloidal stability of the lipoplexes under isoelectric lipid/DNA conditions and coated with the different pH-sensitive PEG lipids was evaluated at two different pH values and compared to those obtained with the non-pH-sensitive chol-PEG at a ratio PEG lipid/total lipid of 3%. It was important to

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Fig. 3. Influence of the PEG lipid amount on DNA compaction.

Fig. 1. Hydrolysis of compounds 7, 8, 15, and 16 in 0.1 M acetate buffer at pH 4 and 5 as a function of time.

4). The best acid-labile PEG lipid candidate was identified as the cholesteryl six-membered ring orthoester derivative 16, for which destabilization was obtained after only 30 min at pH 5.0. The relative instability obtained with the monoalkyl compounds 7 and 15 even at pH 6.0 as compared to 16 may reflect a weaker interaction of the monoalkyl-PEG with the lipoplex, compared to the cholesterol derivatives (8 and 16). In this case, the lower stability of the PEG lipid insertion in the liposome bilayer should contribute to lipoplex destabilization. Thus, the observed aggregation behavior at higher pH (i.e., pH 6.0) for the monoalkyl compounds 7 and 15 might not be completely attributed to the hydrolysis of the orthoester group.

set the percentage of PEG lipid to a minimum by ensuring stabilization of the particles, since the sensitivity of the complexes to acidic medium would depend upon the number of hydrolyzed molecules required to induce destabilization and aggregation. Size increase corresponding to PEG lipid degradation and neutral lipoplex aggregation was measured (Fig.

Fig. 2. Stabilization of neutral lipoplexes by different amounts of PEG lipids.

Fig. 4. Colloidal stability of the lipoplexes in 0.1 M sodium citrate buffer at pH 5 and 6 as a function of time.

3.5. Transfection results Cationic lipoplexes bearing pH-sensitive monoalkyl-PEG (7 and 15) or cholesteryl-PEG (8) compounds have been tested in vitro and compared to lipoplexes bearing the non-pH-sensitive analogs (Fig. 5). In the absence of serum, the lipoplexes without PEG lipid gave a high transfection level, as previously shown with our cationic lipoplexes [8]. When lipoplexes were coated with either 10% Brij or 20% chol-PEG, the level of transfection was significantly reduced by a factor 30 and 300, respectively, as was similarly reported [20]. Insertion of the least pHsensitive monoalkyl compound 7 also inhibited the transfection efficacy. Conversely, cationic lipoplexes including either the most pH-sensitive alkyl-PEG derivative 15 or the pH-sensitive cholesteryl derivative 8 increased the transfection level by a factor of 10 and 25, respectively, as compared to the corresponding non-pH-sensitive formulations obtained with Brij 700 or chol-PEG. It should be noted, however, that the general level of transfection observed with lipoplexes containing pH-sensitive PEG lipid was still lower than that observed with uncoated cationic lipoplexes, possibly because of partial DNA release from the complexes due to the insertion of a large amount of lipid PEG (10–20%) (see Fig. 3). This observation might also be due to incomplete hydrolysis of pH-sensitive PEG lipid, thus leading to only partial release of DNA in the

Fig. 5. HeLa cell transfection in the absence of serum using RPR120535/DOPE/DNA lipoplexes including Brij 700, PEG lipids 7 and 15 (10%), and chol-PEG and PEG lipid 8 (20%) as compared to the cationic lipoplexes.

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cytoplasm. We have shown that the presence of an acid-sensitive linker in PEG lipid restored the transfection as compared to non-pH-sensitive PEG lipidcoated lipoplexes. Since the lipoplexes were prepared at pH=8.0, and that the pH of the cellular medium was maintained at pH 7.4, no prior degradation of the pH-sensitive polymers could have occurred as shown by our stability studies. The complexes might have been destabilized by low pH in the endosomal compartment, thus releasing DNA, preferentially for the pH-sensitive formulation as compared to the nonpH-sensitive one. These results are in good agreement with the chemical and physico-chemical studies. Indeed, the highest level of transfection in the presence of PEG lipid was obtained with the six-membered cycle PEG lipid 15, which was also found to be one of the most pH-sensitive molecules in our degradation studies (Fig. 1).

4. Conclusion We have described the synthesis of new pHsensitive PEG conjugates. The sensitivity of these compounds to different pH values is related to the structure of the orthoester linkage and can therefore be adjusted accordingly. The destabilization at slightly acidic pH of globally neutral lipoplexes at the isoelectric point, coated with these sensitive PEG lipids, correlated well with the chemical degradation of the molecules alone. This confirmed that the hydrolysis of the orthoester group could proceed, despite the insertion of the PEG lipid in the lipoplex lipid bilayer. Moreover, the use of pHsensitive PEG lipid in the formulation of lipoplexes gave a higher level of transfection than with nonpH-sensitive analogs on HeLa cells culture at pH=7.4. It is not excluded, however, that degradation of these complexes might have occurred in the endosomal compartment, thus favoring transfection. These observations make these types of conjugates promising candidates for the formulation of pHsensitive liposomes delivering genes or drugs to low pH tissues, such as solid tumors and inflammatory sites. Work is in progress to validate the use of these new compounds for gene delivery to tumors in vivo.

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Acknowledgements This work was supported by grants from Gencell, the Ministe`re de l’Enseignement et de la Recherche, and the Association Franc¸aise pour les Myopathies. We also thank M. Vuilhorgne and his staff (Structural Analysis Department of Aventis).

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