Nonviral Vectors for Gene Therapy. Methods and Protocols

Citation preview

M E T H O D S I N M O L E C U L A R M E D I C I N E TM

Nonviral Vectors for Gene Therapy Methods and Protocols Edited by

Mark A. Findeis

Humana Press

Synthesis of Polyampholyte Comb-Type Copolymer

1

1 Synthesis of Polyampholyte Comb-Type Copolymers Consisting of Poly(L-lysine) Backbone and Hyaluronic Acid Side Chains for DNA Carrier Atsushi Maruyama and Yoshiyuki Takei 1. Introduction Polycations have been used as nonviral gene carriers because the polycations and DNAs form stable complexes in a noncovalent manner (1–3). The polycations, e.g., poly(L-lysine) (PLL), are reported to be conjugated with several ligands for targeted gene delivery (4–8). The physicochemical properties of the DNA complexes have been described as factors that influence transfection activity (9–11). The authors have reported (12,13) several comb-type copolymers consisting of a PLL backbone and hydrophilic dextran side chains for controling the assembling structure of DNA–copolymer complexes. The dextran chains grafted onto PLL are found to reduce aggregation of the resulting complexes and to increase the solubility of the complexes. Furthermore, the grafting degree of the copolymer affects the DNA conformation in the complex, allowing regulation of DNA compaction. The comb-type copolymers with a higher degree of grafting induce little compaction of DNA and stabilize DNA duplexes and triplexes by shielding the repulsion between phosphate anions of DNA. Moreover, the grafted chains reduce the nonspecific interaction of the PLL backbone with proteins (14). The comb-type copolymers therefore fulfill several requirements for the cellspecific carrier of DNA, if the copolymers are provided with cell-specific ligands. Hyaluronic acid (HA) is an unbranched high-molecular-weight polysaccharide consisting of alternating N-acetyl-`-D-glucosamine and `-D-glucuronic acid residues linked at the 1.3 and 1.4 positions, respectively (15). Liver sinusoidal endothelial cells possess the receptors that recognize and internalize HA (16,17). More than 90% of HAs in the blood stream are known to be taken up and metaboFrom: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

1

2

Maruyama and Takei

lized by SECs. The authors are therefore interested in HA as the ligand to deliver the DNA to the SEC. The authors’ recent study (18) shows that the complexes between PLL–HA conjugates and reporter genes were distributed exclusively in SECs, leading to gene expression in vivo. This chapter describes preparation of PLL-graft-HA (PLL-g-HA) comb-type copolymers. For the synthesis of the comb-type copolymers, high-molecularweight HA was hydrolyzed, then the HAs were covalently coupled with ¡-amino groups of PLL at their reducing end by reductive amination reaction. 2. Materials 2.1. Enzymatic Hydrolysis of HA 1. High-molecular-weight HA (5.9 × 102 kDa), obtained as its sodium salt (sodium hyaluronate), was a gift from Denki Kagaku Kogyo (Tokyo, Japan). 2. Bovine testicular hyaluronidase (EC 3.2.1.35; Type I-S, Sigma, St. Louis, MO). 3. Syringe filters, 0.45 µm (New Steradisc 25, Kurabo, Osaka, Japan).

2.2. Synthesis of PLL-g-HA Comb-Type Copolymers 1. PLL, obtained as its chloride or bromide salt, was purchased from Peptide Institute (Osaka, Japan). 2. Sodium borate buffer-NaCl: 0.1 M, pH 8.5, 0.4–1 M NaCl. 3. Sodium cyanoborohydride (NaBH3CN). 4. NaCl solution (0.5 M). 5. Dialysis membrane (Spectra/Por 7, mol wt cut-off 25,000). 6. Distilled water.

2.3. Size Exclusion Chromatography (SEC)–Multiangle Laser Light Scattering (MALLS) Apparatus 1. 2. 3. 4.

Chromatography pumping system operating at 1.0 mL/min. Size exclusion chromatography column(s). NaNO3 (0.1 M) containing 5 mM sodium phosphate buffer, pH 8.0. Na2SO4 (0.2 M) containing 5 mM sodium phosphate buffer, pH 8.0.

3. Methods 3.1. Enzymatic Hydrolysis of HA 1. Dissolve hyaluronic acid (1 g) in 120 mL water. 2. Add 20 mg bovine testicular hyaluronidase and stir at 50°C. 3. Use a small portion of the reaction to trace HA molecular weight change by SEC–MALLS. 4. When the desired molecular weight of the HA is reached, boil the mixture for 5 min to terminate the reaction. 5. After cooling down to room temperature, filter the mixture through a 0.45-µm filter to remove the denatured enzyme. The resulting HA fragments were obtained by freeze-drying.

Synthesis of Polyampholyte Comb-Type Copolymer

3

(mL) Fig. 1. Time-course of the hydrolysis of HA by hyaluronidase detected by SEC– MALLS. HA (5.9 × 102 kDa; 1 g) was hydrolyzed by hyaluronidase (20 mg) at 50°C.

Figure 1 shows the time course of the HA hydrolysis determined by a SEC– MALLS apparatus (see Subheadings 2.3. and 3.3.). The rate of hydrolysis depends on enzyme activity, so that preliminary experiment on a small scale is recommended to estimate hydrolysis rate before a large-scale reaction. The rate of hydrolysis can be regulated by changing enzyme concentration. For graft copolymer synthesis, a molecular weight ranging from 3000 to 10,000 is favorable. Because the hydrolyzed product has a large distribution in molecular weight, it is also recommended to fractionate fragments by dialysis or ultrafiltration.

3.2. Synthesis of PLL-g-HA Comb-Type Copolymers The obtained HA fragments were conjugated to PLL by reductive amination using NaBH3CN as a reducing agent (Scheme 1). The reaction proceeded through two steps. First is the Schiff’s base (-CH=N-) formation between a reductive (aldehyde) end of HA and primary amino groups of PLL. Second is reduction of the Schiff’s base to form secondary amino groups (-CH2-NH). Although the Schiff’s base is unstable and reversible, its reduced product is an irreversible covalent product. 1. Dissolve the PLL (60–120 mg) in 15 mL sodium borate buffer (0.1 M, pH 8.5) containing 0.4–1 M NaCl. 2. Add the HA fragment (100–300 mg) to the solution. If turbidity or precipitation was observed, increase NaCl concentration. PLL and HA may form an interpolyelectrolyte complex, which is unfavorable for graft copolymer synthesis. The complex formation can be avoided by increasing NaCl concentration. 3. Stir the mixture at 40°C for a few hours for Schiff’s base formation.

4

Maruyama and Takei

Scheme 1. Synthesis of PLL-g-HA comb-type copolymers. Reprinted with permission from ref. 19. Copyright 1998, American Chemical Society.

4. Add NaBH3CN to the mixture and allow to stand at 40°C for 2 d. Approximately 10 molar excess of NaBH3CN to HA is recommended. 5. Sample the solution and trace the reaction with SEC–MALLS (see Subheading 3.3. for SEC–MALLS procedure). 6. Purify the mixture by dialysis against 0.5 M NaCl aqueous solution using a Spectra/Por 7 membrane (mol wt cut-off = 25,000). 7. Desalt the sample by dialysis against distilled water, the resulting copolymer is obtained by freeze-drying. The resulting copolymer would be precipitated during the dialysis.

Figure 2 shows the time-course of the coupling reaction between PLL and HA traced by SEC–MALLS. The reaction can be detected as a decrease in peak area of free HA, increase in peak of the copolymer and in molecular weight of the copolymer. The coupling was almost completed within a few days of incubation. Note that the free HA is almost eliminated after the reaction.

Synthesis of Polyampholyte Comb-Type Copolymer

5

Fig. 2. Time-course of coupling reaction between PLL and HA detected by SEC– MALLS.

3.3. SEC–MALLS SEC was carried out using a JASCO 880-PU pumping system (Tokyo, Japan) at the flow rate of 1.0 mL/min at 25°C, with Ultrahydrogel series (Japan Waters, Tokyo, Japan) or Shodex OH pack SB-series (Showa Denko, Tokyo, Japan). A suitable combination of mobile phase and columns must be chosen, because polyelectrolytes including HA and PLL are liable to interact with the column packings, leading to delay in elution volume. In such case, molecular weight determination using the calibration curve based on molecular weight standard samples such as polyethyleneglycol and pullulan is not reliable. The choice of the mobile-phase rely on gel permeation chromatography (GPC) columns. It is highly recommended to provide a light-scattering (LS) detector system such as MALLS (Multiangle laser light scattering detector, Dawn-DSP, Wyatt Technology, Santa Barbara, CA). By using a LS detector, a direct estimation of the molecular weight is possible. The mobile phases we used are 0.1 M NaNO3 for HA fragment analysis and 0.2 M Na2SO4 containing 5 mM sodium phosphate buffer (pH 8.0) for copolymer analysis. For typical analysis, 200 µL of each sample was picked up from the reaction mixture and injected into the columns. Eluate was detected by a refractive index (RI) detector (830-RI, JASCO) and a MALLS detector. RI and LS signals were transferred to a computer to calculate number-average and weight-average molecular weight according to the instruction manual (Wyatt Technology) for Dawn-DSP.

3.4. 1H Nuclear Magnetic Research (NMR) Spectroscopic Analyses Each copolymer was dissolved in D2O (Deuterium content: 99.95% Merck, Darmstadt, Germany) containing 0.35 M NaCl. 1H-NMR spectra (400 MHz) were

6

Maruyama and Takei

Fig. 3. 1H NMR spectra of PLL (A), HA (B), and PLL-g-HA (C) in D2O. For the PLL-g-HA, D2O containing 0.35 M NaCl was used. Reprinted with permission from ref. 19. Copyright 1998, American Chemical Society.

obtained by a Varian Unity 400plus spectrometer (Palo Alto, CA), at a probe temperature of 298 K. The chemical shifts are expressed as parts/million using internal HDO molecules (b = 4.7 ppm in D2O) as a reference. As shown in Fig. 3, the 1H-NMR spectra of the comb-type copolymer showed the characteristic signals of both PLL and HA moieties: PLL, b 1.4–1.8 (`, a, b-CH2), 3.0 (¡-CH2), 4.3 (_-CH); HA, b 2.0 (NAc-CH3), 3.3–3.9 (H-2,3,4,5,6), 4.4–4.6 (H-1). From the signal ratio of methyl protons (2.0 ppm) of the N-acetyl groups of the HA-grafts to ¡-methylene protons (3.0 ppm) of the PLL backbone, the content (wt % and grafting-%) of HA in the copolymer was determined. The results of the synthesis of PLL-g-HA comb-type copolymers are summarized in Table 1. Coupling efficiency was more than about 70%. Consequently, the authors have easily prepared the various PLL–HA conjugates with a well-defined combtype structure by combining enzymatic hydrolysis and the reductive amination.

In feed

Copolymer

PLL Sample

7

1 2 3 4 5 6 7 8

Mol wtb

HA

Mnb/104

mg

Mnb/103

mg

wt%

Mn/104

Mw/Mn

4.2 4.2 7.2 7.2 7.2 4.2 7.2 7.2

61.2 61.2 122 61.2 61.2 61.2 122 61.2

2.3 2.3 3.8 3.8 3.8 1.6 1.6 1.6

94.3 189 189 189 283 94.3 94.3 189

61 76 61 76 82 61 44 76

8.5 11 15 23 31 9.3 12 24

1.5 1.4 1.4 1.6 1.5 1.4 1.5 1.4

Coupling

HA Contentc

efficiencyd Yield

Charge wt% Grafting-% ratio 50 63 53 69 77 55 38 71

5.7 9.4 3.8 7.6 11 9.8 4.9 19

0.34 0.56 0.40 0.83 1.3 0.49 0.22 1.0

%

%

66 55 73 73 71 80 80 77

67 57 55 83 57 38 51 85

aReducing reagent, 0.3 M NaBH CN; reaction temperature, 40°C; reaction time, 56 h (samples 1 and 2), 80 h (samples 3–5), 75 h (samples 6–8); 3 solvent, 0.1 M sodium borate buffer (pH 8.5) containing 0.4 M NaCl (samples 1 and 2) or 1 M NaCl (samples 3–8). b Molecular weight and its distribution (M /M ) were determined by SEC–MALLS. w n cDetermined by 1H-NMR; grafting-% = (mol fraction of the lysine residues grafted with HA) × 100%; charge ratio = [carboxyl group] / HA [amino group]PLL in copolymer. d [HA] copolymer/[HA]in feed × 100%. Reprinted from ref. 19. Copyright 1998, American Chemical Society.

Synthesis of Polyampholyte Comb-Type Copolymer

Table 1 Synthesis of PLL-g-HA Comb-Type Copolymersa

7

8

Maruyama and Takei

References 1. Kabanov, A. V., Astafyeva, I. V., Chikindas, M. L., Rosenblat, G. F., Kiselev, V. I., Severin, E. S., and Kabanov, V. A. (1991) DNA interpolyelectrolyte complexes as a tool for efficient cell transformation. Biopolymers 31, 1437–1443. 2. Boussif, O., Lezoualc’h, F., Zanta, M. A., Mergny, M. D., Scherman, D., Demeneix, B., and Behr, J. P. (1995) A versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo: Polyethylenimine. Proc. Natl. Acad. Sci. USA 92, 7297–7301. 3. Page, R. L., Butler, S. P., Subramanian, A., Gwazdauskas, F. C., Johnson, J. L., and Velander, W. H. (1995) Transgenesis in mice by cytoplasmic injection of polylysine/DNA mixtures. Transgenic Res. 4, 353–360. 4. Wu, G. Y. and Wu, C. H. (1987) Receptor-mediated in vitro gene transformation by a soluble DNA carrier system. J. Biol. Chem. 262, 4429–4432. 5. Wagner, E., Cotten, M., Foisner, R., and Birnstiel, M. L. (1991) Transferrinpolycation-DNA complexes: the effect of polycations on the structure of the complex and DNA delivery to cells. Proc. Natl. Acad. Sci. USA 88, 4255–4259. 6. Huckett, B., Ariatti, M., and Hawtrey, A. O. (1990) Evidence for targeted gene transfer by receptor-mediated endocytosis: stable expression following insulindirected entry of neo into HepG2 cells. Biochem. Pharmacol. 40, 253–263. 7. Trubetskoy, V. S., Torchilin, V. P., Kennel, S. J., and Huang, L. (1992) Use of N-terminal modified poly(L-lysine)-antibody conjugate as a carrier for targeted gene delivery in mouse lung endothelial cells. Bioconjugate Chem. 3, 323–327. 8. Martinez-Fong, D, Mullersman, J. E., Purchio, A. F., Armendariz-Borunda, J., and Martinez-Hernandez, A. (1994) Nonenzymatic glycosylation of poly-L-lysine: a new tool for targeted gene delivery. Hepatology 20, 1602–1608. 9. Perales, J. C., Grossmann, G. A., Molas, M., Liu, G., Ferkol, T., Harpst, J., Oda, H., and Hanson, R. W. (1997) Biochemical and functional characterization of DNA complexes capable of targeting genes to hepatocytes via the asialoglycoprotein receptor. J. Biol. Chem. 272, 7398–7407. 10. Wolfert, M. A., Schacht, E. H., Toncheva, V., Ulbrich, K., Nazarova, O., and Seymour, L. W. (1996) Characterization of vectors for gene therapy formed by self-assembly of DNA with synthetic block co-polymers. Hum. Gene Ther. 7, 2123–2133. 11. Kabanov, A. V. and Kabanov, V. A. (1995) DNA complexes with polycations for the delivery of genetic material into cells. Bioconjugate Chem. 6, 7–20. 12. Maruyama, A., Katoh, M., Ishihara, T., and Akaike, T. (1997) Comb-type polycations effectively stabilize DNA triplex. Bioconjugate Chem. 8, 3–6. 13. Maruyama, A., Watanabe, H., Ferdous, A., Katoh, M., Ishihara, T., and Akaike, T. (1998) Characterization of interpolyelectrolyte complexes between doublestranded DNA and polylysine comb-type copolymers having hydrophilic side chains. Bioconjugate Chem. 9, 292–299. 14. Maruyama, A., Ishihara, T., Kim, J. S., Kim, S. W., and Akaike, T. (1997) Nanoparticle DNA carrier with poly(L-lysine) grafted polysaccharide copolymer and poly(D,L-lactic acid). Bioconjugate Chem. 8, 735–742.

Synthesis of Polyampholyte Comb-Type Copolymer

9

15. Balazs, E. A., Laurent, T. C., and Jeanloz, R. W. (1986) Nomenclature of hyaluronic acid. Biochem. J. 235, 903. 16. Forsberg, N. and Gustafson, S. (1991) Characterization and purification of the hyaluronan-receptor on liver endothelial cells. Biochim. Biophys. Acta 1078, 12–18. 17. Yannariello-Brown, J., Frost, S. J., and Weigel, P. H. (1992) Identification of the Ca2+-independent endocytic hyaluronan receptor in rat liver sinusoidal endothelial cells using a photoaffinity cross-linking reagent. J. Biol. Chem. 267, 20,451–20,456. 18. Takei, Y., Maruyama, A., Nogawa, M., Asayama, S., Ikejima, K., Hirose, M., et al. (1999) A novel gene delivery system for genetic manipulation of sinusoidal )endothelial cells by triplex DNA technology: evaluation of targetability and ability to stabilize triplex formation. Hepatology 30, 298A. 19. Asayama, S., Nogawa, M., Takei, Y., Akaika, T., Maruyama, A. (1998) Synthesis of novel polyampholyle comb-type copolymers consisting of poly (L-lysine) backbone and hyaluronic acid side chains for a DNA carrier. Bioconjugate Chem. 9, 476–481.

Cationic _-Helical Peptides

11

2 Cationic _-Helical Peptides for Gene Delivery into Cells Takuro Niidome and Haruhiko Aoyagi 1. Introduction Development of nonviral gene transfer techniques has progressed, particularly the use of several kinds of cationic lipids and cationic polymers such as polylysine derivatives, polyethyleneimines, polyamidoamine dendrimers, and so on, which electrostatically form a complex with the negatively charged DNA, which can be taken up by the cells. Furthermore, targeted gene transfer has also been realized by modification of the gene carriers using cell-targeting ligands such as asialoorosomucoid, transferrin, insulin, or galactose. Recently, novel gene transfer techniques have been reported, in which an amphiphilic _-helical peptide, containing cationic amino acids is used as a gene carrier into cells. Wyman et al. (1) employed a peptide, KALA (WEAKLAKA-LAKA-LAKH-LAKA-LAKA-LKAC-EA), which is derived from the sequence of the N-terminal segment of the HA-2 subunit of the influenza virus hemagglutinin involved in the fusion of the viral envelope with the endosomal membrane. This peptide showed several functions in the transfection process, e.g., condensing DNA and causing an endosome-membrane perturbation, which enables it to deliver the incorporated DNA to the cytosol, which is essential for efficient transfection. Similarly, the authors also found the transfection technique, which is mediated by some amphiphilic _-helical peptides (e.g. Ac-LARL-LARL-LARL-LRAL-LRAL-LRAL-NHCH3 [46] and KLLK-LLLK-LWKK-LLKL-LK [Hel]) as shown in Table 1 (2–5). After that, for the purpose of refining of the peptide structure, we investigated the influence of the peptide chain length on gene transfer ability. As a result, 16 and 17 amino acid residues were sufficient to form aggregates with the DNA, From: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

11

12

Niidome and Aoyagi

Table 1 Structures of Amphiphilic _-Helical Peptides Peptide

Sequence

46 Ac-LARL-LARL-LARL-LRAL-LRAL-LRAL-NH2 4668 Ac-LARL-LRAL-LRAL-LRAL-NH2 Hel KLLK-LLLK-LWKK-LLKL-LK Hel61 LLK-LLLK-LWKK-LLKL-LK

Chain length

Cationic charge

24 16 18 17

6 4 7 6

and transfer the DNA into the cells in the deletion series of 46 and Hel, respectively (Table 1; 6). In addition, the authors succeeded in constructing a multiantennary galactose-modified peptide containing four galactose residues that serve for efficient binding to the asialoglycoprotein receptor on hepatoma cells (7). This chapter focuses on synthesis of the peptides and a method of gene transfer using them. As is well known, a peptide is readily synthesized because of the development of an automatic peptide synthesis apparatus and reagents for synthesis. From this point of view, it is expected that the gene transfer method mediated by the peptide is easily accepted by many researchers taking part in the gene therapy study. 2. Materials 2.1. Peptide Synthesis 1. Peptide synthesis apparatus (ABI 431A, PE Biosystems). 2. Fmoc-Lys(Boc) preloaded Wang resin, Rink amide resin (100–200 mesh) (Calbiochem-Novabiochem, CA). 3. Fmoc protected amino acids, 2-(1H-benzotriazole-1-yl)-1,1,3,3,-tetramethyluronium hexafluorophosphate (HBTU), N,N,-diisopropylethylamine, NMP, dichloromethane (DCM), piperidine (Watanabe Chemical, Hiroshima, Japan). 4. Thioanisole, m-Cresol, ethandithiol, trifluoracetic acid (TFA), acetonitrile (Wako Chemicals, Osaka, Japan). 5. High-performance liquid chromatography (HPLC) apparatus (Hitachi L7100 System, Tokyo, Japan). 6. Reverse-phase (RP)-HPLC column (YMC-Pack C4, q10 × 150 mm, Kyoto, Japan). 7. Matrix-assisted Laser Desorption Ionization-Time of Flight-Mass Spectra (MALDI TOF-MS) apparatus (Voyager DE STR, PE Biosystems).

2.2. Preparation of Plasmid DNA 1. Plasmid DNA, which contains a luciferase gene and SV40 promoter (PicaGene control vector, PGV-C), was purchased from Toyo Ink (Tokyo, Japan). 2. Plasmid DNA (pCMVluc), containing a luciferase gene under control of cytomegalovirus enhancer/promoter, was prepared by removing the BglII and

Cationic _-Helical Peptides

13

HindIII insert of the plasmid PGV-C (Toyo Ink), and ligating with the BglII and HindIII fragment from the pRc/CMV (Invitrogen), which contains cytomegalovirus promoter. 3. Closed circular plasmid DNA was purified by ultracentrifugation in cesium chloride gradients. The plasmid preparations showed a major band of closed circular DNA and minor amount ( 300 kDa) are able to condense DNA effectively (particle size, 150–200 nm). In contrast, when plasmid is incubated with low-molecular-weight pDMAEMA, large complexes are formed (size 0.5–1.0 µm). Using optimal conditions, the degree of transfection in vitro (approx 30% of the treated cells are transfected) is higher than that achieved with commercially available cationic lipids such as 1,2-dioleoyl3-trimethylammonium-propane (DOTAP) and Lipofectamine (12). Cells grown in vivo can be transfected ex vivo with pDMAEMA-based polyplexes with an overall transfection efficiency of ~1–2% (13). To further improve transfection performance, random copolymers were synthesized. Random copolymers of DMAEMA with ethoxytriethylene glycol methacrylate (triEGMA), N-vinylpyrrolidone (NVP), methyl methacrylate (MMA), and methacrylic acid of different molecular weights and compositions (comonomer fraction up to 66 mol%) are able to bind DNA, yielding polyplexes (14,15; and Bos and Hennink, unpublished data). However, for random copolymers of DMAEMA with triEGMA, NVP, and MMA, the polymer:plasmid ratio at which small complexes (size 200 nm) are formed increases with the mol fraction of the comonomer (14,15). A copolymer with 20 mol% MMA shows a reduced transfection efficiency and a substantial increased cytotoxicity compared with a homopolymer of the same molecular weight (14). On the other hand, for triEGMA, NVP, and methacrylic acid, the cytotoxicity of the copolymers, either complexed with DNA or in the free form, is inversely proportional to the mol fraction of these comonomers. This reduction is even more than what can be expected based on the DMAEMA mol fraction in the copolymer (14 and Bos and Hennink, unpublished data). NVPDMAEMA copolymers synthesized by polymerization to high conversion show excellent DNA binding, condensing characteristics and transfection capabilities. This is ascribed to a synergistic effect of DMAEMA-rich copolymers and NVP-rich copolymers present in this system on the complex formation with plasmid DNA (14).

Cationic Methacrylate Polymers/DNA Complexes

37

Temperature-sensitive copolymers of DMAEMA and N-isopropylacryl amide (NIPAAm) of various monomer ratios and molecular weights have been evaluated as carrier systems for DNA delivery (16). All copolymers, even with a low DMAEMA content of 15 mol%, were able to bind to DNA at 25°C. Light-scattering measurements indicate that complexation is accompanied by precipitation of the copolymer in the complex caused by a drop of the lower critical solution temperature of the copolymer. The copolymer:plasmid ratio, at which complexes with a size of approx 200 nm are formed, shows a positive correlation with the NIPAAm content of the copolymer and is independent of molecular weight of the copolymer. Complexes containing copolymers of low molecular weight or high NIPAAm content prepared at 25°C aggregate rapidly when the temperature is raised to 37°C. On the other hand, complexes containing copolymers of high molecular weight or lower NIPAAm content are stable at 37°C. The cytotoxicity of the complexes decreases with increasing NIPAAm content and is independent of molecular weight of the copolymer. The transfection efficiency as a function of the copolymer:plasmid ratio shows a bell-shaped curve. The copolymer:plasmid ratio at which the transfection efficiency is maximal rises with increasing NIPAAm content; the maximum transfection efficiency drops with increasing NIPAAm content of the copolymer (16). Besides pDMAEMA, a number of structural analogs, differing in the side chain groups, have been evaluated as transfectant (17,18). Almost all studied cationic methacrylate/methacrylamide polymers are able to condense the structure of pDNA, yielding small polyplexes (100–300 nm) and a slightly positive c potential. However, the transfection efficiency and the cytotoxicity of the polymers differ widely: the highest transfection efficiency and cytotoxicity are observed for pDMAEMA itself. Assuming that polyplexes enter cells via endocytosis, pDMAEMA apparently has advantageous properties to escape the endosome. A possible explanation is that, because of its average pKa value of 7.5, pDMAEMA is partially protonated at physiological pH and behaves as a proton sponge. This might cause a disruption of the endosome, which results in the release of both the polyplexes and cytotoxic endosomal/lysosomal enzymes into the cytosol. In contrast, the analogs of pDMAEMA studied have a higher average pKa value and have, consequently, a higher degree of protonation and a lower buffering capacity, which might be associated with a lower tendency to destabilize the endosome, resulting in both lower transfection efficiency and a lower cytotoxicity (17,18). Furthermore, structural analysis by molecular modeling techniques suggests that, of all studied polymers, pDMAEMA has the lowest number of interactions with DNA. The authors therefore hypothesize that the superior transfection efficiency of pDMAEMA-containing polyplexes can be ascribed to an intrinsic property of pDMAEMA to destabilize endosomes combined with an easy dissociation of the polyplex once present in the cytosol and/or the nucleus (17,18).

38

Bos, Crommelin, and Hennink

Functionalization of pDMAEMA, e.g., by coupling of an antibody (-fragment) or another ligand such as poly(ethylene glycol) is feasible by using a random copolymers of DMAEMA with aminoethyl methacrylate (AEMA) (19). The percentage of incorporated primary amino groups can be controlled by the feed ratio of AEMA:DMAEMA, and is usually below 10 mol%. The ligands can be coupled to the amine groups directly or, for example, via a thiol group. In this case, following the synthesis of the copolymer, protected thiol groups are introduced in a derivatization step with N-succinimidyl 3-(2-pyridyldithio) propionate and subsequent treatment with dithiothreitol. The obtained thiolated p(DMAEMA-co-AEMA) can be conjugated to antibodies and other ligands, e.g., the nuclear localization signal decapeptide Gly-Pro-Lys-Lys-Lys-Arg-Lys-ValGlu-Asp-NH2, via a disulfide linkage. In general, the coupling efficiencies are high (>90%) (19). The thiolated polymers can also be used to determine the apparent kinetic rate constants between plasmid DNA and the nonviral carrier polymers using surface plasmon resonance spectrometry (20). In this case, the polymers are attached to the gold layer through the thiol groups. Freeze-drying of these gene delivery systems can be performed using a controlled two-step drying process and sucrose as lyoprotectant (21). Freezedrying is shown to be an excellent method to preserve the size and transfection potential of pDMAEMA–plasmid complexes (8,22), even after aging at 40°C (23). The concentration of the sugars is an important factor affecting both the size and transfection capability of the complexes after freeze-drying and freezethawing. However, the type of lyoprotectant (sugar) used is of minor importance (24). The DNA topology has been shown to affect pDMAEMAmediated transfection: Circular forms of DNA (supercoiled and open-circular) show higher transfection activity than linear forms (25). Recently, the possibilities and limitations of autoclaving, filtration, and a combination of both methods for sterilization of pDMAEMA-based gene transfer complexes have been assessed (26). Agarose gel electrophoresis and circular dichroism spectroscopy shows that filtration of polyplexes does not change the topology and integrity of the DNA. Moreover, a full preservation of the transfection potential of the filtered polyplexes was observed. Precoating of the filter with polyplexes reduces the material loss and the loss of transfectivity. In contrast, autoclaving dramatically affects physical characteristics of polyplexes, resulting in a complete loss of transfection potential. Addition of sucrose to the preparation protects DNA present in pDMAEMA– DNA complexes, to some extent, from degradation during autoclaving, but the transfection potential is not retained. Filtration or autoclaving of polymer alone does not result in substantial loss of polymer integrity and material, or in decreased transfection potential. Naked DNA can easily be sterilized by filtration as well, although some DNA may be lost. Therefore, separate

Cationic Methacrylate Polymers/DNA Complexes

39

sterilization of polymer and DNA stock solutions, followed by aseptic formation and handling of polyplexes, may be an acceptable and preferred alternative to filtration of polyplexes (26). In this chapter, protocols are provided for the synthesis of pDMAEMA homo- and copolymers in water or toluene, gel permeation chromatography (GPC) analysis of the synthesized polymers, the preparation of pDMAEMA–DNA polyplexes, and subsequent determination of the plasmid integrity after formulation, sterilization of pDMAEMA-based polyplexes, a standard transfection protocol, and determination of cell viability and of transfection efficiency. 2. Materials 2.1. Common 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

12. 13. 14.

Water purified by reversed osmosis (RO water). pH-meter or pH-indicator strips. 70% Ethanol. Sodium chloride, pro analysi (pa). Sodium hydroxide (4.0, 1.0, and 0.1 N). Phosphate buffered saline (PBS): 3.6 mM KH2PO4, 6.4 mM Na2HPO4, and 145 mM NaCl, pH 7.2. 4-(2-Hydroxy-ethyl)-1-piperazine ethane sulfonic acid (HEPES). 0.22-µm Filters. Vortex mixer. Eppendorf tubes sterilized by autoclaving. Cell culture equipment: Biohazard safety cabinet or laminar airflow cabinet, equipped with burner and suction device, CO2 incubator 37°C, Bürker-Türk or Bürker counting chamber (bright lined), microscope (e.g., ×25 objective, ×10 ocular), water bath at 37°C, centrifuge. P-20, P-100, P-200, and P-1000 Gilson pipets with sterile tips. 5- and 10-mL pipets. 15- and 50-mL centrifuge tubes.

2.2. Synthesis of pDMAEMA Homo- and Copolymers in Water 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

DMAEMA stabilized with 0.15% tert-butyl-hydroxytoluene (Fluka). Ammonium peroxodisulphate (APS) (Fluka). 37% Hydrochloric acid, pa. 100 mL Infusion bottles with plastic lids and silicon rubber septa (e.g., Emergo). Needles. Nitrogen–vacuum exchange system. Water bath at 60°C. Vacuum distillation equipment. 1–10 mL Glass syringes, glass pipets. Dialysis tubing, mol wt cut-off 12–14 kDa (e.g., Medicell). Freeze-dryer.

40

Bos, Crommelin, and Hennink

2.3. Synthesis of pDMAEMA in Toluene 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

DMAEMA stabilized with 0.15% tert-butyl-hydroxytoluene (Fluka). _,_'-Azoisobutyronitril (AIBN) (Fluka). Toluene, pa. Petroleum ether, boiling range 40–60°C, technical quality. Dichloromethane. 100-mL Infusion bottles with plastic lids and silicon rubber septa (e.g., Emergo). Needles. Nitrogen–vacuum exchange system. Water bath at 60°C. Vacuum stove with temperature control at 40–60°C. Vacuum distillation equipment. Glass syringes, glass pipets.

2.4. GPC of Water-Soluble pDMAEMA Homo- and Copolymers 1. 2. 3. 4. 5. 6.

7. 8. 9. 10. 11.

Tris-(hydroxymethyl)-aminomethane (Tris). NaNO3, pa. 60% HNO3, pa. 0.22-µm Filter (e.g., Millipore GVWP04700). Degassing setup consisting of vacuum pump and ultrasound bath. High-performance liquid chromatography (HPLC) system, consisting of a pump, an autoinjector, a refractive index detector, and software with GPC option and GPC column set option. GPC column set (e.g., Shodex K80P, KB-80M, and KB802); for molecular weight higher than 10,000 g/mol, the KB802 column can be skipped. 1-mL Poly(propylene) (PP) shell vial with snap cap (Alltech). 2-mL Syringes. Nylon 13-mm syringe filters, 0.45 µm (Alltech). 15-mL PP tubes with cap.

2.5. Amplification and Purification of Plasmid 1. Bacterial strain producing an appropriate marker gene. 2. Plasmid purification kit (e.g., Giga-kit from Qiagen).

2.6. Preparation of pDMAEMA–DNA Polyplexes with Low DNA Concentration 1. Plasmid in TE-buffer (see Subheading 3.4.). 2. pDMAEMA (see Subheadings 3.1. and 3.2.).

2.7. Preparation of pDMAEMA–DNA Polyplexes with High DNA Concentration 1. 2. 3. 4.

Glacial acetic acid, pa. Sucrose. Plasmid in TE-buffer (see Subheading 3.4.). pDMAEMA (see Subheadings 3.1. and 3.2.).

Cationic Methacrylate Polymers/DNA Complexes

41

2.8. Integrity of Plasmid in Polyplexes 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Pronase. Tris-(hydroxymethyl)-aminomethane (Tris). Glacial acetic acid, pa. Ethylenediamine tetraacetic acid (EDTA). Ethidium bromide (EtBr) or SYBR Green I nucleic acid stain (Molecular Probes). Bromophenol blue. Glycerol. Marker DNA (e.g., h DNA [Gibco] digested with restriction enzyme PstI). Polyaspartic acid (pAsp) (Sigma, mol wt 50,000). Electrophoresis apparatus. UV detection system. Conical flask. Microwave oven.

2.9. Sterilization of pDMAEMA-Based Gene Transfer Complexes 1. 0.22-µm Filters (e.g., cellulose acetate) (Schleicher & Schull GmbH, Dassel, Germany).

2.10. Standard Transfection Protocol 1. 2. 3. 4. 5. 6. 7.

OVCAR 3, COS 7 or other cells. Completed culture medium, depending on the particular cell line. 10X Trypsin–EDTA (Gibco). Trypan blue for vital staining (Sigma). 96-Well plates. Centrifuge. Eight-Channel pipet.

2.11. Determination of Cell Viability 1. Sodium 3'-[1-(phenylaminocarbonyl)-3,4-tetrazolium]-bis(4-methoxy-6-nitro) benzene sulfonic acid hydrate (XTT) (Gibco). 2. N-methyl dibenzopyrazine methylsulfate (PMS) (Gibco). 3. Microplate reader.

2.12. Determination of Transfection Efficiency 1. 2. 3. 4. 5. 6. 7. 8. 9.

Tris(hydroxymethyl)-aminomethane (Tris). 37% Hydrochloric acid, pa. Triton X-100. MgCl2·6H2O. NaH2PO4·H2O. Na2HPO4·2H2O. o-Nitrophenyl-`-D-galactopyranoside (ONPG) (Sigma). `-Galactosidase (Sigma). 0.25% Glutaraldehyde solution.

42 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21.

Bos, Crommelin, and Hennink NaH2PO 4·HO. Na2HPO4·2H2O. K4Fe(CN)6·3H2O. K3Fe(CN)6. MgCl2·6H2O. 5-Bromo-4-chloro-3-indolyl-`-D-galactopyranoside (X-Gal, Gibco). Dimethylsulfoxide (DMSO): 99.9% spectrophotometric grade. Microplate reader. Gelatin powder. 85% Glycerol. Phenol. Microwave oven.

3. Methods

3.1 Synthesis of pDMAEMA Homo- and Copolymers in Water Synthesis of pDMAEMA (Fig. 1) is achieved by radical polymerization (9,10), and it can be performed in either an acidified aqueous solution (this procedure) or in toluene (see Subheading 3.2.). Because the ester bond of DMAEMA can be chemically hydrolyzed in water (27), synthesis of pDMAEMA in toluene may be preferred. However, pDMAEMA is routinely synthesized in water, which is especially useful for the synthesis of copolymers of DMAEMA and tolueneinsoluble monomers. After synthesis, the polymers are characterized by nuclear magnetic resonance (NMR) to determine if all monomer has reacted, and in the case of copolymers, also the copolymer composition. Furthermore, the polymers are characterized by GPC to determine the average molecular weight and the molecular weight distribution (see Subheading 3.3.). 1. The monomer DMAEMA should be vacuum-distilled shortly before synthesis to remove the radical scavenger tert-butylhydroxytoluene. After distillation, allow nitrogen into the distillation equipment. In some cases, the comonomer should be vacuum-distilled as well (see Note 1). 2. Dissolve 1.0 g initiator APS in 15 mL water in a 50-mL flask. 3. An initial DMAEMA concentration of 20% (v/v) for the synthesis is recommended. The molecular weight of the polymer will depend on the ratio of monomer and initiator (M:I ratio) used. Several typical examples of mixtures are given in Table 1. Add in 100-mL infusion bottles with septum appropriate amounts of water, 37% HCl and DMAEMA (in that order). Use a glass syringe to transfer DMAEMA into the flask. When copolymers are synthesized, the molar quantities of DMAEMA and the comonomer should match the quantity of DMAEMA given in the Table 1. Adjust the pH of the solution to 5.0 to prevent hydrolysis of the esterbond of DMAEMA. In this stage, chain transfer agents such as mercaptoethanol can be added to obtain low-molecular weight polymers with a functional endgroup. Add the appropriate amount of APS solution (see Table 1).

Cationic Methacrylate Polymers/DNA Complexes

43

Table 1 Reaction Components and Expected Molecular Weight 37% HCl (mL)

Water (mL)

DMAEMA (mL)

APS (mL)

M:I ratio

Mpa (expected) (106 g/mol)

4 4 4 4

34 35 35.5 35.8

10 10 10 10

2 1 0.5 0.2

100 200 400 1000

0.5 ± 0.1 1.5 ± 0.2 2.6 ± 0.4 5.0 ± 1.0

aM p

is the molecular weight at the peak of the GPC curve and is therefore independent of the peak integration. Therefore, it is useful to compare different batches of polymer.

4. 5.

6. 7.

8.

Degas the solution in the bottle with a needle through the septum with the use of a nitrogen–vacuum exchange system until first signs of evaporation of water can be seen (typically, approx 15 s), then flush nitrogen into the flask. Repeat this twice to make sure that the air atmosphere is completely replaced by nitrogen atmosphere. Incubate the flasks under shaking conditions in a water bath at 60°C for 20–22 h. Cool down the viscous solution and transfer it into dialysis tubes. If necessary, dilute the solution with water. Beware osmotic pressure: The volume of the solution may increase threefold during dialysis. Remove the buffer by dialysis for several days at 4°C using at least 5 L of water in the external compartment. Freeze-dry the polymer solution. Determine the weight of empty 500-mL or 1-L round-bottomed flasks and transfer the dialyzed polymer solution into these flasks. Do not fill a flask with more than one-third of its volume. Freeze the polymers in a film on the inner wall of the flask, while rotating the flask in liquid nitrogen. Freeze-dry the polymers overnight to remove the water. Determine the weight of the flasks with the polymer and subtract the weight of the empty flask to estimate the yield of the polymerization (usually 80–90%). Characterize the polymer by GPC (see Subheading 3.3.), and by NMR (see also Notes 2 and 3). 1H-NMR: Dissolve about 20 mg (p)DMAEMA in 1 mL CDCl . Chemical shifts: 3 a. Monomer (DMAEMA): 6.09 (s, 1H, =CH), 5.54 (s, 1H, =CH), 4.23 (t, 2H, OCH2), 2.60 (t, 2H, NCH2), 2.28 (s, 6H, N(CH3)2), 1.93 (s, 3H, C=C-CH3). b. Polymer (pDMAEMA): 4.05 ppm (b, 2H, OCH2), 2.55 ppm (b, 2H, NCH2), 2.28 ppm (b, 6H, N(CH3)2), 2.18–2.30 ppm (bm, 2H, C-CH2), 0.75–2.30 ppm (bm, 3H, C-C-CH3).

3.2. Synthesis of pDMAEMA in Toluene 1. The monomer DMAEMA should be vacuum-distilled 1 d before synthesis, as described in Subheading 3.1.1. 2. Dissolve 0.2428 g initiator AIBN in 5 mL toluene (296 mM). 3. An initial DMAEMA concentration of 20% (v/v) is recommended for the synthesis. The molecular weight of the polymer will depend on the M:I mol ratio used. Several typical examples of mixtures are given in Table 2.

44

Bos, Crommelin, and Hennink Table 2 Reaction Components and Expected Molecular Weight DMAEMA (mL)

Toluene (mL)

AIBN (mL)

M:I ratio

10 10 10 10

38 39 39.5 39.8

2 1 0.5 0.2

100 200 400 1000

Mpa (expected) (104 g/mol) 4±1 6±1 12 ± 2 22 ± 4

aM is the molecular weight at the peak of the GPC curve, and is therefore p independent of the peak integration. Therefore, it is useful to compare different batches of polymer.

4. In one flask, add appropriate amounts of DMAEMA and toluene. Add the appropriate amount of AIBN solution using a glass syringe. Degas the solution in the bottle as described in Subheading 3.1.3. 5. Incubate the flask under shaking conditions in a water bath at 60°C for 20–22 h. 6. After incubation, cool down the viscous solution and slowly add the solution drop-wise (e.g., by gravity force through a Pasteur pipet) under stirring conditions to a 10-fold bigger volume of petroleum ether to precipitate the polymer. 7. Discard the petroleum ether/toluene, wash the precipitate twice with 50–100 mL petroleum ether, and transfer the polymer into a suitable storage can or flask. Dissolve the last part of the polymer in the flask with dichloromethane and transfer the solution into the can or flask. Allow remnants of the organic solvents to evaporate in the hood overnight. 8. Dry the polymers inside a vacuum stove at 40–60°C until no toluene can be smelled and the weight of the flask with the polymer does not further decrease. If complete removal of toluene is essential, the polymer can be dissolved in water and subsequently dialyzed and freeze-dried as described in Subheadings 3.1.5–3.1.7. 9. Characterize the polymers by GPC (see Subheading 3.3.), and by NMR. For 1H-NMR assignments, see Subheading 3.1.8. and Notes 2 and 3.

3.3. GPC of pDMAEMA Homo- and Copolymers This instruction is intended for the molecular weight determination of pDMAEMA and its copolymers. 1. Prepare eluent: 0.7 M NaNO3, 0.1 M Tris-HCl, pH 7.2. Dissolve 119 g NaNO3 and 24.2 g Tris in 1.8 L RO water. Adjust pH to 7.2 with 60% HNO3, and fill up to 2 L with RO water. Filter through 0.22-µm filter, Millipore, GVWP04700. Alternative eluents (check specs of columns for compatibility) are 10 mM NaCl; 5 mM NH4+CH3COO–, pH 5.5; 0.8 M NaNO3. 2. Set HPLC system to appropriate specs, e.g., for the series of Shodex columns: flow 0.5 mL/min; detector sensitivity 256 (for concentration range of 0.5–2.0 mg/mL); detector temperature, 35°C; column oven temperature 30°C; run time 40 min; injection vol 100 µL.

Cationic Methacrylate Polymers/DNA Complexes

45

Table 3 Mp-Values of Dextran Standards for GPC Measurement of pDMAEMA Stnd. Dex1

Dex2

Dex3

Dex4

Dex5

Dex6

Mr~ a 180 991 50,000 750,000 342 1000 80,000 1,800,000 504 5000 150,000 667 12,000 270,000 82 25,000 670,000 M w=4,900,000 M p=4,500,000 M n=1,500,000

Mpb

Fluka no.

180 991 43,500 560,000 342 1080 66,700 1,450,000 504 4440 123,600 667 9890 196,000 829 21,400 401,300

aM ~ is number on bottle. r bUse M values in the weight-loading p

63416 31420 31426 63418 31416 31421 31427 63430 31417 31422 63422 31418 31423 63417 31419 31425 31428

Name Sucrose Maltohexaose Dextran 50,000 Dextran 750,000 D(+)-Maltose Dextran 1000 Dextran 80,000 Dextran 800,000 Maltotriose Dextran 5000 Dextran 150,000 Maltotetraose Dextran 12,000 Dextran 270,000 Maltopentose Dextran 25,000 Dextran 670,000 Dextran 4,900,000

table of your GPC software.

3. Calibrate the GPC columns: a. For every mixture (see Table 3) dissolve 5 mg of each dextran in 5 mL eluent. Allow dextran to dissolve overnight at room temperature. Store the solutions at 4°C. b. Measure and calculate a calibration curve. 4. For new types of copolymers, prepare a stock solution by adding 50 mg polymer to 10 mL eluent in a 15-mL PP tube. Put the tube on a rotating wheel or roller bank at 4°C for 4 d to dissolve completely. Filter the sample through a 0.45-µm syringe filter. 5. Prepare solutions with concentrations of 5, 4, 3, 2, 1, and 0.5 mg/mL by pipeting 1.0, 0.8, 0.6, 0.4, 0.2, and 0.1 mL sample respectively, in a PP vial and add eluent to obtain a final volume of 1.0 mL. 6. Inject 100 µL sample onto the GPC system and determine the molecular weight and area of the different concentrations using the software. Plot area vs concentration. This should give a linear relationship. Furthermore, Mw and M n should be independent of the concentration. 7. For validated polymers, routine analysis can be performed at 1.0 mg/mL.

46

Bos, Crommelin, and Hennink

3.4. Amplification and Purification of Plasmid Plasmids are prepared from bacterial cultures grown in the presence of a selective agent such as an antibiotic, using for instance the standard Luria Bertani medium (27). The plasmid is isolated from the culture using, e.g., an Endofree Qiagen Giga Plasmid Kit (Qiagen) according to the instructions of the manufacturer. After amplification and purification, the plasmid is collected in a Tris-EDTA buffer (TE buffer, 10 mM Tris-HCl, 1 mM EDTA, pH 8.0) to obtain a plasmid stock solution, which can be stored at 4°C for at least 1 yr; the concentration of the DNA (OD280 nm) should be checked every 3 mo. However, high EDTA concentrations may induce aggregation of the pDMAEMA–plasmid complexes at high DNA concentration. Therefore, for preparation of polyplexes with a high DNA concentration (see Subheading 3.6.), after purification, the plasmid should be taken up by a low amount of TE buffer to avoid high EDTA concentrations in the final sample. The plasmid concentration of the plasmid stock solution should therefore be higher than 3 µg/µL (see also Note 4).

3.5. Preparation of pDMAEMA–DNA Polyplexes with Low DNA Concentration Small and stable pDMAEMA–plasmid complexes (polyplexes) of low concentrations can be prepared either in HEPES buffer or in HEPES-buffered saline (HBS) (see Fig. 2 and Note 5). This procedure describes the preparation of 3:1 pDMAEMA–plasmid polyplex formulation (250 µL, containing 2.5 µg plasmid and 7.5 µg pDMAEMA). For polyplexes using a different polymer:DNA ratio, the concentration of the polymer solution should be varied. Preparation of polyplexes with higher DNA concentrations is described in Subheading 3.6. (see also Note 6). 1. Switch on the biohazard or laminar airflow safety cabinet 30 min before use and clean the surface with 70% ethanol. After opening and before closing tubes, bottles, and so on, quickly put cap and neck through a burner’s blue flame. 2. Prepare the following solutions: a. HEPES (HBS) containing 20 mM HEPES (pH 7.4) and 150 mM NaCl: Add 2.383 g HEPES and 4.383 g NaCl to 480 mL water, and adjust the pH to 7.4 with approx 1 mL 4 M NaOH. Adjust the total volume to 500 mL with water. Sterilize the buffer through a filter with 0.22-µm pore size (autoclaving causes some precipitation) (see Note 7). b. HEPES buffer (pH 7.4). Add 2.383 g HEPES to 480 mL water and adjust the pH to 7.4 with NaOH. Adjust the total volume to 500 mL with water. Sterilize the buffer through a filter with 0.22-µm pore size (see Note 7). c. Plasmid stock solution. After amplification and purification, the plasmid is collected in a Tris-EDTA buffer (see Subheading 3.4.). The plasmid stock solution can be stored at 4°C for at least 1 yr; the concentration of the DNA (OD280 nm) should be checked every 3 mo.

Cationic Methacrylate Polymers/DNA Complexes

47

Fig. 2. Preparation of polyplexes (see Subheading 3.5.).

3. 4. 5. 6. 7. 8. 9.

d. Stock solution of pDMAEMA, 5 mg/mL in HEPES buffer. Dissolve 50 mg pDMAEMA in 10 mL HEPES buffer (see step 2b). Allow the pDMAEMA to dissolve for 3–4 d at 4°C, preferably using a rotating device. Sterilize the solution through filters with 0.22-µm pore sizes. Dilute the plasmid stock solution with HBS or HEPES to obtain 200 µL containing 10 µg plasmid (50 µg/mL). Dilute the pDMAEMA stock solution with HBS or HEPES to obtain 800 µL containing 30 µg polymer (37.5 µg/mL). Add 200 µL diluted pDMAEMA solution to 50 µL diluted plasmid solution. A 250-µL sample containing 2.5 µg plasmid and 7.5 µg pDMAEMA is obtained. Immediately vortex the sample for 5 s. Repeat steps 5 and 6 to obtain three batches of polyplexes. After 30 min, the complexes are ready for use. Characterize the complexes by dynamic light scattering (DLS, particle size determination). Polyplexes prepared from newly synthesized polymers should be measured at least threefold. To characterize the polyplexes by electrophoretic mobility (c-potential), they should be prepared using HEPES buffer to dilute the plasmid and pDMAEMA stock solutions instead of HBS, and at least 1 mL should be prepared (see Note 8).

3.6. Preparation of pDMAEMA–DNA Polyplexes with High DNA Concentration pDMAEMA–plasmid complex dispersions (polyplexes) of high concentrations can be prepared without aggregation (29). The complexes are prepared at low pH (20 mM acetate buffer, pH 5.7), low ionic strength, and high viscosity (20 w/v% sucrose solution). This procedure describes the preparation of 3:1 pDMAEMA:plasmid complex dispersion (1 mL, containing 150 µg plasmid and 450 µg pDMAEMA). For polyplexes using a different polymer/DNA ratio, the concentration of the polymer solution should be varied.

48

Bos, Crommelin, and Hennink

1. Switch on the biohazard or laminar airflow safety cabinet 30 min before use and clean the surface with 70% ethanol. After opening and before closing tubes, bottles, and so on, put cap and neck through a burner’s blue flame. 2. Prepare the following solutions: a. HBS as in Subheading 3.5., step 2a. b. Acetate buffer (20 mM acetate). Add 0.57 mL glacial acetic acid to 480 mL water, and adjust the pH to 5.7 with 1.0 N and 0.1 N NaOH. Adjust the total volume to 500 mL with water. Sterilize the buffers through filters with 0.22-µm pore sizes. Opened bottles should be stored in the refrigerator and used within 1 mo. c. Sucrose–acetate buffer (40 w/v% sucrose in 20 mM acetate). Dissolve 4.00 g sucrose (2.6g) in 5 mL acetate buffer (3.2g). Add acetate buffer until the volume is 10 mL. Sterilize the buffers through filters with 0.22-µm pore sizes. d. Plasmid stock solution, as in Subheading 3.5., step 2c. However, high EDTA concentrations induce aggregation of the pDMAEMA–plasmid complexes at high DNA concentration. Therefore, after purification, the plasmid should be taken up by a low amount of TE buffer to avoid high EDTA concentrations in the final sample. The plasmid concentration of the plasmid stock solution should therefore be higher than 3 µg/µL (see Note 4). e. Stock solution of pDMAEMA as in Subheading 3.5., step 2d, using HBS instead of HEPES. 3. Dilute the plasmid stock solution with acetate buffer to obtain 100 µL containing 150 µg plasmid. 4. Add 500 µL sucrose–acetate buffer to the 100 µL diluted plasmid solution. 5. Add 310 µL acetate buffer to 90 µL pDMAEMA stock solution. A solution of 450 µg pDMAEMA in 400 µL buffer is obtained. 6. Add the 400 µL diluted pDMAEMA solution to the 600 µL diluted plasmid solution. A 1-mL sample containing 150 µg plasmid and 450 µg pDMAEMA is obtained. 7. Immediately vortex the sample gently for 5 s. 8. After 30 min, the complexes are ready for use. Alternatively, the polyplexes can be lyophilized. 9. Characterize the complexes by DLS (particle size determination) and by electrophoretic mobility (c-potential) (see Notes 9 and 10). Polyplexes prepared from newly synthesized polymers should be measured at least threefold.

3.7. Integrity of Plasmid in Polyplexes Electrophoresis through agarose is the standard method used to separate and identify DNA fragments. In this procedure, a 0.7% (w/v) agarose gel concentration is used, but different gel concentrations can be used for a broad size range of DNA molecules (see ref. 27). In order to run the plasmid on an agarose gel, the DNA needs to be dissociated from the polymer. This can be done using polyaspartic acid (see Fig. 3) (18,30). 1. Prepare the following solutions: a. 50X TAE buffer: Dissolve 242 g Tris-base, 57.1 mL glacial acetic acid, 100 mL 0.5 M EDTA (pH 8.0) in RO water to make 1 L.

Cationic Methacrylate Polymers/DNA Complexes

49

Fig. 3. Agarose gel of pCMVlacZ (lane 1), pDMAEMA–pCMVlacZ polyplexes before (lane 2) and after (lane 3) dissociation with a 100-fold excess of polyaspartic acid (see Subheading 3.7.).

2.

3.

4.

5. 6.

b. Working buffer: 1X TAE buffer: Dilute 50X TAE buffer 50-fold in RO water. c. Ethidium bromide in water (10 mg/mL). The stock solution should be stored in a bottle wrapped in aluminum foil at room temperature. d. 10X Sample buffer: 0.4% bromophenol blue, 10 mM EDTA, 50% glycerol in water. e. Dissociation buffer: Dissolve 100 mg pAsp in 5 mL PBS. Add dissociation buffer to the polyplex samples, using a 100-fold (w/w) excess of pAsp to DNA. Vortex and incubate 30 min at ambient temperature for freshly prepared polyplexes (within 2–3 h after preparation) or between 1 and 5 d at 4°C for older polyplexes. Add pronase to 1X TAE in a conical flask; e.g., for a small size gel 0.35-g pronase in 50 mL 1X TAE. Heat in a microwave until the agarose dissolves. Cool the solution to about 60°C and, when desired, add ethidium bromide to a final concentration of 0.5 µg/mL. Position combs and pour agarose solution into the mold, check for air bubbles. Leave gel to set (30–45 min at ambient temperature), carefully remove the combs and mount the gel in the electrophoresis tank. Add working buffer 1X TAE to cover the gel. Mix the samples of DNA with the sample buffer. The maximum amount of DNA that can be applied to a slot depends on the number of fragments in the sample and their sizes (15). Marker DNA (1–2 µg) can be loaded as well. Run the gel (e.g., 100 V, current unlimited) until the bromophenol blue has migrated ±6 cm through the gel. If ethidium bromide was present in the gel, examine the gel directly by UV light, or after destaining for 30 min in water. Otherwise, stain the gel by soaking it for 30–45 min in a solution of ethidium bromide (0.5 µg/mL) in water or SYBR Green I in water and destain the gel for 30–45 min in water before examining the gel.

3.8. Sterilization of pDMAEMA-Based Gene Transfer Complexes To sterilize polyplexes, filter at least 6 mL polyplex preparation through 0.22-µm filters and discard the first 3 mL of sample (26). Alternatively, filter

50

Bos, Crommelin, and Hennink

Fig. 4. Transfection protocol (see Subheading 3.9.).

at least 5 mL polymer solution (5 mg/mL) and DNA solution (10 µg/mL) separately. Before preparation of polyplexes from these sterilized solutions, the polymer and DNA concentration should be determined using optical densities 225 and 260, respectively, as measure for the concentration.

3.9. Standard Transfection Protocol This procedure describes the in vitro transfection of OVCAR-3 cells (see Note 11) and COS-7 cells (see Note 12), using cationic carriers, such as the polymer, pDMAEMA, as a vector (Fig. 4; see refs. 8–10 for typical results). The protocol can be used for all adherent cell types. For suspension type cells, cells need to be spun down before changing the medium at a speed suitable for the cell type to be transfected. 1. Switch on the biohazard or laminar airflow safety cabinet 30 min before use and clean the surface with 70% ethanol. After opening and before closing tubes, bottles, and so on, put cap and neck through a burner’s blue flame. 2. Prepare the following solutions: a. HBS as in Subheading 3.5., step 2a. b. Fetal bovine serum (FBS) (Integro), sterile and heat-inactivated (30 min at 56°C). Store at –20°C. c. Plasmid stock solution as obtained in Subheading 3.4. d. Stock solutions of carriers, e.g., 1–5 mg/mL in HBS, filtered through 0.22-µm filters, and stored at –20°C (see Subheading 3.5., step 2d). e. Completed Dulbecco's modified Eagle's medium (DMEM) culture medium: Prepare completed medium normally used for culturing the cells to be transfected. Serum can be present in the medium. f. OVCAR-3 or COS-7 cell suspension, or, alternatively, other cell suspensions. g. 0.05% Trypsin–0.02% EDTA solution (1X). Defrost the 10X trypsin–EDTA in a 37°C water bath. Add 10 mL 10X trypsin–EDTA to 90 mL PBS and store at 4°C. Tenable for 3 mo when stored at 4°C.

Cationic Methacrylate Polymers/DNA Complexes

3. 4.

5.

6.

7. 8. 9.

10. 11. 12.

51

h. 0.5% Trypan blue solution. Weigh 0.5 g trypan blue in a biohazard safety cabinet. Dissolve trypan blue in 100 mL PBS and filtrate through filtration paper to remove possible crystals. Divide in portions and store at –20°C. At room temperature, the solution is tenable for about 1 mo. Passage OVCAR-3 or COS-7 cells or other cell types 3–4 d before the transfection experiment. Detach the cells with trypsin–EDTA solution, and determine the cell number and cell viability using trypan blue. Pipet 50 µL cell suspension into an Eppendorf tube and add 50 µL 0.5% trypan blue solution. Bring some cell–trypan blue mixture into a counting chamber and count (microscope) in a number of squares the number of uncolored (vital) and blue (dead) cells in duplicate. Count at least 100 cells. When more than 10 cells are counted per square, dilute the cell suspension and count again. Determine the number of vital cells. Add 100 µL 1.1 × 105 cells/mL/well of a 96-well plate and incubate for 24 h at 37°C (the cells should be approx 60% confluent). Perform each transfection in at least six wells on two different 96-well plates and use one 96-well plate for estimation of the transfection efficiency and the other for estimation of the cell number and viability. Next day, prepare dilute solutions of the carriers and plasmids to be tested from their stock solutions in HBS. For instance: a. pDMAEMA: Prepare 0.6 mL 0.6 mg/mL from the stock solution, and make of this a series of twofold dilutions in HBS (see first column in Table 4). b. DNA: Dilute the plasmid stock solution to a final concentration of 50 µg/mL in HBS. Prepare polyplexes in threefold by pipeting 50 µL 50 µg/mL plasmid solution in a well of a 96-well plate, followed by 200 µL carrier solution. As reference, add 3× 200 µL 37.5 µg/mL pDMAEMA in HBS to 50 µL 50 µg/mL DNA in wells of a 96-well plate using the same batch of polymer for all experiments. Incubate the polyplexes for 30–60 min at room temperature. Meanwhile, aspirate culture medium from the cells (by pipeting or decanting) and add 100 µL of warm (37°C) completed DMEM. Use medium with at least 10% serum in this step. Overlay the cells (in a 96-well plate) with 100-µL polyplexes (1 µg plasmid/ well), vortex 15 s carefully, and incubate for 1 h at 37°C in the incubator. Aspirate transfection media, and add 100 µL warmed, completed DMEM into each well, and culture the cells in the incubator for 44 ± 2 h at 37°C. Estimate the transfection results. The number and viability of the cells can be estimated by the XTT colorimetric assay (see Subheading 3.10.). Using a lacZ reporter gene, the transfection efficiency can be estimated by the `-galactosidase content with the use of the substrate ONPG, or alternatively, an X-Galbased histochemical color assay can be used to estimate the number of transfected cells (see Subheading 3.11.). Express the data relative to the transfection results mediated by the reference carrier solution of pDMAEMA. The incubation times with polyplexes and the subsequent culture time can be adjusted for different cell types.

52

Bos, Crommelin, and Hennink Table 4 Ratios of Carrier:DNA in Polyplex Preparation Carrier solution (µg/mL)

DNA solution (µg/mL)

600 300 150 75 37.5 18.75 9.38 4.69

50 50 50 50 50 50 50 50

Carrier/DNA (w/w) 48 24 12 6 3 1.5 0.75 0.38

3.10. Determination of Cell Viability To determine the influence of carrier–plasmid complexes on cell viability and proliferation, the number of viable cells can be measured using an XTT colorimetric assay (33; see also Note 13). The assay is based on the cleavage of the yellow tetrazolium salt, XTT, to form an orange formazan dye by dehydrogenase activity in active mitochondria. Therefore, this conversion only occurs in living cells. The formazan dye formed is soluble in aqueous solutions and is directly quantified using a microplate reader. This procedure can also be applied for suspension type cells. 1. Switch on the biohazard or laminar flow safety cabinet 30 min before use and clean the surface with 70% ethanol. After opening and before closing tubes, bottles, and so on, put cap and neck through a burner’s blue flame. 2. Prepare the following solutions: a. XTT stock solution, 1 mg/mL: Dissolve XTT in plain RPMI 1640 at 1 mg/mL. Fill out in 10-mL portions and store in the dark at –20°C. b. Electron-coupling reagent: Dissolve PMS in PBS at 0.383 mg/mL (1.25 mM). Fill out in 250-µL portions and store in the dark at –20°C. c. XTT-solution: Freshly prepared each experiment. Thaw XTT stock solution and electron-coupling reagent in a water bath at 37°C. Mix each vial to obtain a clear solution. For one 96-well plate, add 100 µL electron-coupling reagent to 5 mL XTT stock solution. Mix by vortexing. 3. Make a calibration curve with living cells: Detach adherent cells with trypsin– EDTA solution, determine the cell number and cell viability using trypan blue, according to the procedure described in Subheading 3.9.4. Dilute the cells to 1 × 106 cells/mL (cell stock) with completed culture medium. Make a calibration curve in a 96-well plate, e.g., as in Table 5. 4. Incubate the cells for 2 h at 37°C in the CO2-incubator. 5. Add 50 µL XTT-solution (step 2c) per well. 6. Incubate for 1 h at 37°C in the CO2-incubator.

Cationic Methacrylate Polymers/DNA Complexes

53

Table 5 Cell Viability Calibration Curve Cells/well A B C D E F G H

100 µL cell stock 100 µL culture medium + 100 µL cell stock 100 µL culture medium + 100 µL B 100 µL culture medium + 100 µL C 100 µL culture medium + 100 µL D 100 µL culture medium + 100 µL E 100 µL culture medium + 100 µL F, mix and pipet 100 µL out µL culture medium

100 × 103 50 × 103 25 × 103 12.5 × 103 6.25 × 103 3.13 × 103 1.56 × 103 0

7. Measure the absorbance at 490 nm with a reference wavelength of 655 nm. 8. With the calibration curve of living cells, the number of viable cells can be calculated.

3.11. Determination of Transfection Efficiency The enzyme activity of `-galactosidase, expressed upon transfection of OVCAR-3 cells, COS-7 cells, or other cell types with lacZ reporter gene, can be determined using the substrate o-nitrophenyl-`-D -galactopyranoside (ONPG; 28). For suspension type cells, cells need to be spun down before changing the medium, at a speed suitable for the cell type to be transfected. 1. Switch on the biohazard or laminar airflow cabinet 30 min before use and clean the surface with 70% ethanol. 2. Prepare the following solutions: a. Lysis buffer, containing 50 mM Tris–HCl buffer (pH 8.0), 150 mM NaCl, and 1% Triton X-100. For 100 mL, dissolve 0.6057 g Tris, 0.8766 g NaCl in 80 mL water, add 1.0 mL Triton X-100, and adjust the pH to 8.0 with (diluted in water) hydrochloric acid. Fill up to 100 mL with water. b. 100X MgCl2-solution (0.1 M): Dissolve 0.2033 g MgCl2·6H2O in 10 mL water. c. 0.2 M NaH2PO4: Dissolve 5.52 g NaH2PO4·H2O in 200 mL water. The solution is tenable for 3 mo at room temperature. d. 0.2 M Na2HPO4: Dissolve 17.80 g Na2HPO4·2H2O in 500 mL water. The solution is tenable for 3 mo at room temperature. e. 0.1 M sodium phosphate buffer (pH 7.4): Mix 9.5 mL 0.2 M NaH2PO4 (step 2c) with 40.5 mL 0.2 M Na2HPO4 (step 2d) and 50 mL water. f. 10 mg/mL ONPG solution: Dissolve 1000 mg ONPG in 100.0 mL 0.1 M sodium phosphate buffer, pH 7.5 (step 2e). Fill out in 5-mL aliquots in 5- or 10-mL plastic tubes and store in the dark at –20°C. g. 1000 U/mL `-galactosidase stock solution: Add 1.0 mL lysis buffer to 1000 U `-galactosidase. Fill out in 20-µL aliquots in Eppendorf cups and store in the dark at –20°C. Use immediately after thawing and do not freeze and store more than once.

54

Bos, Crommelin, and Hennink

Table 6 Galactosidase Calibration Curve Sample A B C D E F G H

3. 4. 5.

6. 7.

8.

Lysis buffer (µL) 100 100 100 100 100 100 100

Enzyme solution 200 µL 5 U/mL 100 µL A 100 µL B 100 µL C 100 µL D 100 µL E 100 µL F

`-galactosidase (mU/well) 100 50 25 12.5 6.25 3.13 1.56 0

h. ONPG-staining solution: For estimating enzyme activity in all wells of a 96well plate, mix 18.5 mL 0.1 M sodium phosphate buffer, pH 7.5, 200 µL 100X Mg solution, and 1.35 mL 10 mg/mL ONPG solution. Aspirate the media from the cells in the well plate and wash once with 100 µL cold PBS (4°C). Add 20 µL cold lysis buffer (4°C) to the cells in a well, and incubate for 20 min at 4°C. Meanwhile, make the different dilutions for the calibration curve of `-galactosidase from the 1000 U/mL stock solution (2 g). Add 10 µL 1000 U/mL to 90 µL lysis buffer and an aliquot of 50 µL of this mixture to 950 µL lysis buffer, to obtain an enzyme solution of 5 U/mL. Make 7× twofold dilutions of this final solution in a 96-well plate to obtain standard enzyme solutions, according to the scheme in Table 6. Add 20 µL of the standard enzyme solutions A–H to empty wells in the 96-well plate, which is used for, e.g., transfection. Add 180 µL ONPG-staining solution (2 h) to each well (both standards and samples). Incubate the well plate(s) at 37°C and measure the absorbance at 415 nm in a microplate reader, using the absorbance at 655 nm as a reference, until the highest standard shows an absorbance of 2 or more (approx 30–45 min). Calculate the `-galactosidase amount of a sample in mU/well by comparison with the linear standard curve.

Alternatively, cells containing `-galactosidase upon transfection of cells with a lacZ reporter gene can be histochemically stained in vitro (Fig. 5). For suspension type cells, cells need to be spun down before changing the medium at a speed suitable for the cell type to be transfected. 1. Switch on the biohazard or laminar airflow cabinet 30 min before use and clean the surface with 70% ethanol. 2. Prepare the following solutions:

Cationic Methacrylate Polymers/DNA Complexes

55

Fig. 5. X-Gal stained OVCAR3 cells (see Subheading 3.10.). The blue dots are nuclei of cells transfected with pCMVlacZ. a. Working fixative (0.25% glutaraldehyde solution): Made fresh each time by diluting a 25% glutaraldehyde solution 100-fold with PBS. b. 0.2 M NaH2PO4: Dissolve 5.52 g NaH2PO4·H2O in 200 mL RO water. Stored at room temperature, the stock solution is tenable for 3 mo. c. 0.2 M Na2HPO4: Dissolve 17.80 g (0.1 mol) Na2HPO4·2 H2O in 500 mL RO water. Stored at room temperature, the stock solution is tenable for 3 mo. d. 0.1 M Sodium phosphate buffer (pH 7.4): For 100 mL, mix 9.5 mL 0.2 M NaH2PO4 (step 2b) with 40.5 mL 0.2 M Na2HPO4 (step 2c) and 50 mL RO water. Stored at 4°C, it is tenable for 3 mo. e. 0.5 M Potassium hexacyanoferrat(II): Dissolve 0.564 g K4Fe(CN)6·3H2O in 2.67 mL water in a 15-mL centrifuge tube. Protected from light (e.g., by aluminum foil) and stored at 4°C, the stock solution is stable for 3 mo. f. 0.5 M Potassium hexacyanoferrat(III): Dissolve 0.415 g K3Fe(CN)6 in 2.52 mL water in a 15-mL centrifuge tube. Protected from light and stored at 4°C, the stock solution is stable for at least 3 mo. g. 1.0 M MgCl2: Dissolve 2.03 g MgCl2·6H2O in 10 mL water in a 15-mL centrifuge tube. Stored at room temperature, it is tenable for 3 mo. h. 40 mg/mL X-Gal in DMSO: Dissolve 400 mg X-Gal in 10 mL DMSO (99.9% spectrophotometric grade). Fill out in 500-µL portions (in Eppendorf tubes or glass vials, not in polycarbonate or polystyrene ones) and store in the dark at –20°C. After using part of it, freeze the remainder for use again (may become slightly yellow). Do not repeat thawing and freezing more than 5×. i. X-Gal stain solution, freshly made each time. Required volume is approx 0.1 mL/cm2: 1 mL/well of 6-well plate (9.4 cm2), 0.1 mL/well of 24 well

56

3. 4. 5. 6. 7. 8. 9.

10.

Bos, Crommelin, and Hennink plate (2 cm2), 50 µL/well of a 96-well plate (0.38 cm 2). For 10 mL staining solution, mix: 100 µL 0.5 M (K4Fe(CN) 6·3H2O) (step 2e), 100 µL 0.5 M (K3Fe[CN]6) (step 2f), 20 µL 1 M MgCl2 (step 2g), and 250 µL X-Gal solution (step 2h). Bring the volume to 10 mL with 9.53 mL 0.1 M sodium phosphate buffer (step 2d). Aspirate media from monolayers of cells to be assayed. Rinse gently but thoroughly with PBS (for a 96 well plate, 100 µL per well). After aspirating PBS, overlay the cells with fixative (step 2a) and incubate at 4°C for 5 min (for a 96-well plate, 75 µL/well). Aspirate the fixative and rinse gently twice with PBS at room temperature (for a 96-well plate, 100 µL/well). Aspirate the PBS and overlay the cells with X-Gal stain solution (step 2i) (for a 96-well plate, 50 µL/well). Replace the microplate lids and incubate overnight at 37°C. View cells under microscope. Cells expressing `-galactosidase are detectable by their overall blue color or their blue colored nuclei (when the lacZ-plasmid used has an NLS-signal). Count the number of blue cells/well. After washing with PBS, the well plates can be stored at 4°C for 2 d. The plates can be stored for a longer period of time (several years at room temperature) after preservation (see Subheading 3.12.).

3.12. Conservation of X-Gal Colored Cells After performing a color reaction with cells, the cells can be conserved. At elevated temperatures, the conservation solution is a transparent fluid. It congeals when cooled to room temperature. Overlaid with this substance, cells are covered with a jelly transparent film at room temperature. Therefore, they are not susceptible to bacteria, and can be stored at room temperature for infinite time. It is still possible to study them under a microscope. 1. 2. 3. 4. 5. 6. 7. 8.

Suspend 17.4 g gelatin powder in 71 mL water. Add 128 mL 85% glycerol and mix. Add 2.17 g phenol and dissolve it by stirring. Heat at 60°C (or in a microwave oven) until the mixture is a clear, transparent, and air-bubble-free fluid. Fill out in 50-mL portions and store the tubes at room temperature. Prior to use heat a portion in the microwave oven until the substance is fluid. Wash the cells once with 100 µL PBS/well. Add mixture to the wells with the help of a pipette (just enough to cover the cells). Shortly after adding the fluid, it turns into a jelly transparent film protecting the cells.

4. Notes 1. If necessary, the distilled solution can be stored at –20°C for at least 1 wk. 2. High molecular weight pDMAEMA synthesized in water must be dissolved in

Cationic Methacrylate Polymers/DNA Complexes

3. 4. 5.

6.

7.

8.

9. 10. 11. 12. 13.

57

D2O (20 mg/mL) because it does not dissolve in CDCl3. Chemical shifts are much broader than in CDCl 3: a. Monomer (DMAEMA): 5.95 (s, 1H, =CH), 5.52 (s, 1H, =CH), 4.08 (t, 2H, OCH2), 2.58 (t, 2H, NCH2), 2.09 (s, 6H, N[CH3]2), 1.72 (s, 3H, C=C-CH3). b. Polymer (pDMAEMA): 3.95–4.35 ppm (b, 2H, OCH2), 3.00–3.60 ppm (b, 2H, NCH2), 2.50–2.90 ppm (b, 6H, N[CH3]2), 1.30–2.10 ppm (bm, 2H, CCH2), 0.50–1.10 ppm (bm, 3H, C-C-CH3) (see Note 3). The monomer peaks at 6.09 (5.95 in D 2O) ppm and 5.54 (5.52 in D2O) ppm must be absent in polymer samples. If the concentration of the plasmid intended for use is too low, the plasmid can be precipitated in 70% ethanol (28) and redissolved at higher concentration. HBS (step 2a) should be used for in vitro transfection, whereas HEPES (step 2b) is to be preferred for determining the c-potential. Therefore, stock solutions of pDMAEMA can best be prepared in HEPES buffer. The size of polyplexes is influenced by the quantities of DNA and polymer solutions that are mixed, as well as by the speed at which the polymer is added to the DNA solution. Also, the buffer (HEPES or HBS) influences the size of the polyplexes. Closed bottles with sterilized solutions can be stored at room temperature and may be used up to 1 yr after preparation. Opened bottles should be stored in the refrigerator and used within 2 mo. The obtained particle size and c-potential of the pDMAEMA–plasmid complexes prepared in HEPES buffer are approx 100 nm and 30 mV, respectively; the obtained particle size of the pDMAEMA–plasmid complexes prepared in HBS is approx 160 nm. For the refractive index and viscosity, values of 1.363 and 1.546, respectively, must be used (when measured at 25°C). The obtained particle size and c-potential of the pDMAEMA–plasmid complexes are approx 175 nm and 28 mV, respectively. OVCAR-3 cells are human ovarian carcinomas (adenocarcinoma) and originate from American Type Culture Collection (ATCC) (31,32). COS-7 cells are CV-1 cell lines derived from African green monkey kidney cells. The cell line originates from ATCC. Alternatively, cell numbers can be counted using a Bürker counting chamber.

Acknowledgments The authors wish to thank Dr. Jong-Yuh Cherng, Dr. Petra Van de Wetering, Dr. Herre Talsma, Nancy M. E. Schuurmans-Nieuwenbroek, Dr. Wouter L. Hinrichs, and OctoPlus B.V. for the use of their data and protocols. The European Union (grant PL 970002) funds G. W. Bos. References 1. Behr, J. P. (1993) Synthetic gene transfer vectors. Acc. Chem. Res. 26, 274–278. 2. Ledley, F. D. (1997) Pharmaceutical approach to somatic gene therapy. Pharm. Res. 13, 1595–1614.

58

Bos, Crommelin, and Hennink

3. Godbey, W. T., Wu, K. K., and Mikos, A. G. (1999) Poly(ethylenimine) and its role in gene delivery. J. Control. Release 60, 149–160. 4. De Smedt, S. C., Demeester, J., and Hennink, W. E. (2000) Cationic polymer based gene delivery systems. Pharm. Res. 17(2), 113–126. 5. Kawai, S. and Nishizawa, M. (1984) New procedure for DNA transfection with polycation and dimethyl sulfoxide. Mol. Cell. Biol. 4, 1172–1174. 6. Curiel, D. T., Wagner, E., Cotten M., Birnstiel, M. L., Agarwal, S., Li, C. M., Loechel, S., and Hu, P. C. (1992) High-efficiency gene transfer mediated by adenovirus coupled to DNA- complexes. Hum. Gene Ther. 3, 147–154. 7. Boussif, O., Lezoualc’h, F., Zanta, M. A., Mergny, M. D., Scherman, D., Demeneix, B., and Behr, J. P. (1995) A versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo: polyethylenimine. Proc. Nat. Acad. Sci. USA 92, 7297–7301. 8. Cherng, J. Y., Van de Wetering, P., Talsma, H., Crommelin, D. J. A., and Hennink, W. E. (1996) Effect of size and serum proteins on transfection efficiency of poly([2-dimethylamino] ethyl methacrylate)-plasmid nanoparticles. Pharm. Res. 13, 1038–1042. 9. Van de Wetering, P., Cherng, J. Y., Talsma, H., and Hennink, W. E. (1997) Relation between transfection efficiency and cytotoxicity of poly(2-(dimethylamino)ethyl methacrylate)/plasmid complexes. J. Control. Release 49, 59–69. 10. Hennink, W. E. and Van de Wetering, P. (1997) Cationic polyacrylates and poly(alkyl) acrylates and acrylamides for use as carriers of nucleic acid in the transformation of animal cells. PCT Int. Appl. Patent no. WO9715680A1, p. 37. 11. Felgner, P. L., Barenholz, Y., Behr, J. P., Cheng, S. H., Cullis, P., Huang L., et al. (1997) Nomenclature of synthetic gene delivery systems. Human Gene Ther. 8, 511–512. 12. Fonseca, M. J., Storm, G., Hennink, W. E., Gerritsen, W. R., and Haisma, H. J. (1999) Cationic polymeric gene delivery of `-glucuronidase for doxorubicin prodrug therapy. J. Gene Med. 1, 404–417. 13. Van de Wetering, P., Schuurmans-Nieuwenbroek, N. M. E., Hennink, W. E., and Storm, G. (1999) Comparative transfection studies of human ovarian carcinoma cells in vitro, ex vivo and in vivo with poly(2-(dimethylamino)ethyl methacrylate)-based polyplexes. J. Gene Med. 1, 156–165. 14. Van de Wetering, P., Schuurmans-Nieuwenbroek, N. M., Van Steenbergen, M. J., Crommelin, D. J., and Hennink, W. E. (2000) Co-polymers of 2(dimethylamino)ethyl methacrylate with ethoxytriethylene glycol methacrylate or N-vinyl-pyrrolidone as gene transfer agents. J. Control. Release 64, 193–203. 15. Van de Wetering, P., Cherng, J. Y., Talsma, H., Crommelin, D. J., and Hennink, W. E. (1998) 2-(Dimethylamino) ethyl methacrylate based (co) polymers as gene transfer agents. J. Control. Release 53, 145–153. 16. Hinrichs, W. L., Schuurmans-Nieuwenbroek, N. M., Van de Wetering, P., and Hennink, W. E. (1999) Thermosensitive polymers as carriers for DNA delivery. J. Control. Release 60, 249–259.

Cationic Methacrylate Polymers/DNA Complexes

59

17. Van de Wetering, P., Moret, E. E., Schuurmans-Nieuwenbroek, N. M., Van Steenbergen, M. J., and Hennink, W. E. (1999) Structure-activity relationships of water-soluble cationic methacrylate/methacrylamide polymers for nonviral gene delivery. Bioconjugate Chem. 10, 589–597. 18. Arigita, C., Zuidam, N. J., Crommelin, D. J., and Hennink, W. E. (1999) Association and dissociation characteristics of polymer/DNA complexes used for gene delivery. Pharm. Res. 16, 1534–1541. 19. Van Dijk-Wolthuis, W. N., van de Wetering, P., Hinrichs, W. L., Hofmeyer, L. J., Liskamp, R. M., Crommelin, D. J., and Hennink, W. E. (1999) A versatile method for the conjugation of proteins and peptides to poly(2-[dimethylamino]ethyl methacrylate). Bioconjugate Chem. 10, 687–692. 20. Wink, T., De Beer, J., Hennink, W. E., Bult, A., and Van Bennekom, W. P. (1999) Interaction between plasmid DNA and cationic polymers studied by surface plasmon resonance spectrometry. Anal. Chem. 71, 801–805. 21. Talsma, H., Cherng, J., Lehrmann, H., Kursa, M., Ogris, M., Hennink, W. E., Cotten, M., and Wagner, E. (1997) Stabilization of gene delivery systems by freeze-drying. Int. J. Pharm. 157, 233–238. 22. Cherng, J. Y., Van de Wetering, P., Talsma, H., Crommelin, D. J., and Hennink, W. E. (1997) Freeze-drying of poly([2-dimethylamino]ethyl methacrylate)-based gene delivery systems. Pharm. Res. 14, 1838–1841. 23. Cherng, J. Y., Talsma, H., Crommelin, D. J. A., and Hennink, W. E. (1999) Long term stability of poly([2-dimethylamino]ethyl methacrylate)-based gene delivery systems. Pharm. Res. 16, 1417–1423. 24. Cherng, J. Y., Van de Wetering, P., Talsma, H., Crommelin, D. J., and Hennink, W. E. (1999) Stabilization of polymer-based gene delivery systems. Int. J. Pharm. 183, 25–28. 25. Cherng, J. Y., Schuurmans-Nieuwenbroek, N. M., Jiskoot, W., Talsma, H., Zuidam, N. J., Hennink, W. E., and Crommelin, D. J. (1999) Effect of DNA topology on the transfection efficiency of poly([2-dimethylamino]ethyl methacrylate)plasmid complexes. J. Control. Release 60, 343–353. 26. Bos, G. W., Trullas-Jimeno, A., Jiskoot, W., Crommelin, D. J. A., and Hennink, W. E. (2000) Sterilization of poly(dimethylamino) ethyl methacrylate-based gene transfer complexes. Int. J. Pharm. 15, 211(1–2), 79–88. 27. Van de Wetering, P., Zuidam, N. J., Van steenbergen, M. J., Van der Houwen, O. A. G. J., Underberg, W. J. M., and Hennink, W. E. (1998) A mechanistic study of the hydrolytic stability of poly(2-[dimethyl]aminoethyl methacrylate). Macromolecules 31, 8063–8068. 28. Sambrook, J., Fritsch, E. F., Maniatis, T., eds. (1989) Molecular Cloning, A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 29. Cherng, J. Y., Talsma, H., Verrijk, R., Crommelin, D. J., and Hennink, W. E. (1999) Effect of formulation parameters on the size of poly-([2dimethylamino]ethyl methacrylate)-plasmid complexes. Eur. J. Pharm. Biopharm. 47, 215–224.

60

Bos, Crommelin, and Hennink

30. Katayose S. and Kataoka K. (1997) Water-soluble polyion complex associates of DNA and poly(ethyleneglycol)-poly(L-lysine) block co-polymer. Bioconjugate Chem. 8, 702–707. 31. Hamilton, T. C., Young, R. C., McKoy, W. M., Grotzinger, K. R., Green, J. A., Chu, E. W., et al. (1983) Characterization of a human ovarian carcinomal cell line (NIH:OVCAR-3) with androgen and estrogen receptors. Cancer Res. 43, 5379–5389. 32. Hamilton, T. C., Foster, B. J., Grotzinger, K. R., McKoy, W. M., Young, R. C., and Ozols, R. F. (1983) Development of drug sensitive and resistant human ovarian cancer cell lines. A model system for investigating new drugs and mechanisms of resistance. Proc. Am. Assoc. Cancer Res. 24, 313. 33. Scudiero, D. A., Shoemaker, R. H., Paull, K. D., Monks, A., Tierney, S., Nofziger, T. H., et al. (1988) Evaluation of a soluble tetrazolium/formazan assay for cell growth and drug sensitivity in culture using human and other tumor cell lines. Cancer Res. 48, 4827–4833.

Steric Stabilization of DNA Complexes

61

5 Stabilization of Polycation–DNA Complexes by Surface Modification with Hydrophilic Polymers David Oupicky, Martin L. Read, and Thierry Bettinger 1. Introduction Polycation–DNA complexes represent promising synthetic vectors for gene delivery, showing good transfection activities in vitro and safety in vivo. However, simple polycation–DNA complexes suffer from several disadvantages that limit their potential usefulness in vivo. Advances in this field thus rely on better control of the structure, colloidal, and surface properties of condensed DNA particles. Physicochemical stability of simple polycation–DNA complexes is limited. The complexes are usually stable in water or in low-concentration buffers, because of their positive charge; in physiological saline, however, this charge is suppressed by the screening effect of salt and, as a result, the complexes aggregate. Their positive charge (beneficial for transfection efficiency in vitro [1]) also means that they are prone to interaction with various proteins, which represents another major drawback for their use in vivo. When distribution kinetics on nonviral vectors based on polycations (or lipids) have previously been examined in vivo, they were invariably cleared quickly from the bloodstream (2). This is thought to result from binding of serum proteins, which promotes the uptake of complexes by the liver. Systemic application of these complexes would require (1) formation of small particles (~100 nm) facilitating diffusion, extravasation through vascular fenestration, and cellular uptake (endocytosis), (2) formation of neutral particles to minimize nonspecific interactions with proteins and negatively charged surfaces of cell membranes. Apart from reducing nonspecific interactions in vivo, possible specific uptake of the complexes by components of reticuloendothelial system should also be avoided. (3) Cell-specific binding and internalization are also required, and are achievable by their enhanced circulation times in the blood. From: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

61

62

Oupicky, Read, and Bettinger

In order to overcome the previously mentioned problems of simple polycation– DNA complexes, a range of block and graft copolymers of polycations and hydrophilic nonionic polymers, (such as polyethylene glycol (PEG), dextran, and poly(N-[2-hydroxypropyl]methacrylamide) (PHPMA) (3–7) have been synthesized and tested. Complexes formed with these polymers, indeed, showed improved stability against aggregation and interaction with proteins, compared to simple polycation–DNA complexes, but no improvement of in vivo properties (blood circulation times) was observed (3,7,8). Using block copolymers in constructing nonviral gene delivery vectors is discussed in Chapter 3 by Choi and Park. An alternative approach, inspired by experiences with liposomes and nanoparticles in the field of drug delivery systems, has been adopted recently into the field of nonviral gene delivery vectors. The approach relies on surface modification of preformed complexes with hydrophilic polymers. The rationale behind this approach, also called “steric stabilization,” is to provide the complexes with a protecting surface polymer layer, which significantly affects interparticle forces. First, it influences van der Waals attractive forces; and second, it can give rise to repulsion between the particles. The magnitude of repulsion arising from the presence of the layer depends on the density with which it covers the surface; the more thinly it is spread, the smaller its effectiveness in preventing the particles from approaching one another (9). The concept of steric stabilization has been exploited in the field of drug delivery as a way of preventing the particles from aggregation, but also as a way of reducing their interactions with opsonins and components of the reticuloendothelial system, which effectively increased circulation times of these particles (10). Steric stabilization of particles or liposomes generally increases biocompatibility, reduces immune response, increases in vivo stability, and delays clearance by the reticuloendothelial system. Modification of DNA complexes (as with liposomes, proteins) can thus provide many benefits for both in vivo and in vitro applications. Covalent coupling of hydrophilic polymers to complexes can alter their surface and solubility properties, effectively masking the intrinsic character of the surface. Although a range of suitable polymers is available, DNA complexes have so far been modified (coated) with the most frequently used polymer, PEG, and also by PHPMA (11–16). The purpose of this chapter is to describe basic methods used for surface coating of DNA complexes with hydrophilic polymers and their subsequent characterization. 2. Materials 1. 2. 3. 4.

Plasmid DNA. 10 mM HEPES buffer, pH 7.4. Poly-L-lysine (PLL). Polyethylenimine (PEI).

Steric Stabilization of DNA Complexes 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

63

mPEG(5000)-SPA. N-(2-hydroxypropyl) methacrylamide (HPMA). _,_'-Azoisobutyronitrile (AIBN). 3-Mercaptopropionic acid (3-MPA). Acetone. Ether. Methanol. Dimethylformamide (DMF). N,N'-Dicyclohexylcarbodiimide (DCCD). 4-Dimethylaminopyridine (DMAP).

3. Methods 3.1. Coating of DNA Complexes An important prerequisite for coating DNA complexes is the availability of free primary (or secondary) amino groups. Polycations based on tertiary and quaternary amino groups have been successfully used for gene delivery in vitro, but they cannot be as easily chemically modified (17,18). Among the described polycations containing primary or secondary amino groups, the most widely used are PLL and branched PEI.

3.2. DNA Complexes Suitable for Coating and Their Formation In order to prepare small complexes (~100 nm) it is vital to keep the concentration of DNA and salts low during formation. If possible, the complexes should be made in water or in low-concentration buffers (10–50 mM). PLL, if obtained from Sigma, is supplied as a hydrobromide salt and complexes can be prepared directly in water. PEI, however, is usually not supplied as a salt, and needs to be protonated first by dissolving in appropriate buffer (HEPES, pH 7.4). PLL complexes with amine:phosphate (N:P) ratios higher than 1.2 and PEI complexes with N:P higher than 3 are generally suitable for coating reactions as DNA is fully condensed, and the complexes already show positive c potential, indicating the presence of free amino groups on the surface of the complexes (12,19,20). In order to increase the size stability of the complexes, however, it is advisable to work at higher N:P ratios (~2 for PLL and 4–5 for PEI), because complexes around neutrality (N:P ~1 for PLL:DNA and N:P ~2–3 for PEI) tend to aggregate more easily. The following protocol describes a standard method of preparing PLL and PEI complexes in the authors’ laboratories. 1. Prepare solution of DNA in 10 mM HEPES, pH 7.4, at a of concentration 20 µg/mL. (All methods described in this chapter have been performed with plasmid DNA– circular 5 kb expression vector containing a cytomegalovirus promoter-driven luciferase reporter and ampicillin resistance gene.)

64

Oupicky, Read, and Bettinger

2. Add PLL solution (2.5 mg/mL in water) in a single addition to achieve the desired N:P ratio and mix solution quickly by inverting several times. 3. Alternatively, prepare solution of DNA at 40 µg/mL in 10 mM HEPES, pH 7.4, and mix with equal volume of PEI in 10 mM HEPES, pH 7.4, containing the calculated amount of polycation. 4. Leave the complexes for at least 30 min prior to use.

3.3. Coating with Monofunctional Polymers 3.3.1. PEG-Coated Complexes The presence of primary amino groups in the PLL–DNA and PEI–DNA complexes predetermines the choice of the chemistry useful for coating reactions. As mentioned in the introduction the most widely used polymer in modification of particulate drug delivery systems is PEG. Amino-reactive analogs of PEG suitable for coating reactions are available in a wide range of molecular weights and types of reactive groups. They can be obtained from Shearwater Polymers (www.swpolymers.com), who specialize in synthesis of various PEG derivatives. The most commonly used reagents are those based on the active esters of carboxylic acid groups and carbonates. Carboxyl groups activated as N-hydroxysuccinimidyl (NHS) esters are highly reactive toward amine nucleophiles, forming an amide bond and releasing free NHS. The reaction of NHS esters with thiol or hydroxyl groups is possible, but does not yield stable conjugates. Several types of mPEG-NHS are available, but we prefer to use succinimidyl esters of methoxy-PEG propionic acid (mPEG-SPA) because of their increased hydrolytic stability compared to other derivatives. mPEG-SPA, with mol wt 2000, 5000, and 20,000, can be obtained from Shearwater Polymers; PEG(5000)-SPA is also available from Fluka (no. 85969). If a limited amount of accessible amino groups is available, fork-like (PEG)2-NHS containing two mPEG molecules attached to amino groups of lysine, is the reagent of choice to increase PEG density on the coated complexes (also available from Shearwater Polymers). The reaction should be performed in buffers of pH 7.0–9.0 (hydrolysis halflife for mPEG-SPA at pH 8.0 at 25°C is 16.5 min). We have found pH 7.4–8.0 suitable. Increasing pH above 8.0 (together with higher salt concentration) can lead to aggregation of the complexes because of charge screening. The following protocol describes coating of DNA complexes (Fig. 1) using monofunctional mPEG(5000)-SPA (for structure, see Fig. 2). 1. Prepare PLL–DNA complexes at N:P ratio 2, or PEI–DNA complexes at N:P ratio 3–5, in 10–50 mM HEPES, pH 7.4–7.8 (final DNA concentration, 20 µg/mL). 2. Dissolve mPEG(5000)-SPA in water at concentration of 20 mg/mL prior to use; alternatively, make stock solution in dry dimethylsulfoxide (DMSO) and store at –80°C.

Steric Stabilization of DNA Complexes

65

Fig. 1. Schematic presentation of coating (and attachment of targeting ligands to) DNA complexes with multifunctional (PHPMA-ONp) and monofunctional polymers (PHPMA-NHS and mPEG[5000]-SPA). 3. Add the PEG solution to complex solution at desired concentration (0.05–1 mg/mL final concentration is usually optimal). 4. Allow the reaction to proceed at room temperature for at least 3 h: Complexes are then ready to use.

3.3.2. Complexes Coated with Semitelechelic PHPMA We have used PHPMA as an alternative monofunctional (semitelechelic) hydrophilic polymer to PEG. PHPMA is known to be a nontoxic and nonimmunogenic polymer that has previously been used in the field of drug delivery as a carrier of low-molecular-weight drugs (21), and also in

66

Oupicky, Read, and Bettinger

Fig. 2. Chemical structures of polymers used for coating of DNA complexes.

combination with cationic polymers for the formation of DNA complexes (7). The properties of this polymer are in many ways similar to those of PEG. However, it has significant advantages in terms of its structural versatility, enabling easy incorporation of various functionalities. Monofunctional PHPMA can be prepared by radical polymerization in the presence of chain transfer agents, for introducing carboxylic acid endgroup 3-MPA (7) is suitable. The following protocol describes, in detail, synthesis of PHPMA containing one carboxylic endgroup and its activation to NHS reactive ester (Fig. 2). 3.3.2.1. SYNTHESIS OF PHPMA CONTAINING REACTIVE SUCCINIMIDYL ESTER GROUP (PHPMA-NHS) 1. Dissolve 1.2 g HPMA (Polysciences, no. 08242), 5 mg AIBN (initiator of radical polymerization) (Fluka, no. 11630), and 10 mg 3-MPA (Fluka, no. 63770) in 10 mL methanol.

Steric Stabilization of DNA Complexes

67

2. Bubble nitrogen through the solution for 10 min to remove dissolved oxygen. 3. Polymerize in sealed ampule at 50°C for 24 h. 4. Precipitate the solution into 200 mL acetone/ether (3:1 v/v) and isolate the polymer by filtration through sintered glass filter. 5. Purify the polymer by redissolving in methanol and precipitating into acetone/ ether (3:1). 6. Dry polymer in vacuum and determine number of end carboxylic groups by titration with 0.05 M NaOH (molecular weight should be 8000–10,000 g/mol). 7. Convert the carboxylic acid end-group into succinimidyl active ester by dissolving 0.2 g of the obtained polymer in 1 mL dry DMF; add 25 mg NHS. 8. Cool the solution to –20°C, and add a solution of 52 mg DCCD (Fluka, no. 36650) and 2 mg DMAP (Fluka, no. 39405) in 0.3 mL DMF. 9. Leave the solution overnight at 0°C. 10. Remove the precipitated N,N'-dicyclohexylurea by filtration, and precipitate the polymer into 20 mL mixture of dry acetone–diethyl ether (3:1). 11. Isolate the precipitated polymer by filtration and dry it under vacuum at room temperature. 12. The polymer is now ready for use in the coating reaction; however, it can be stored for a limited period of time at –20°C under argon.

3.3.2.2. COATING PROCEDURE 1. Prepare PLL–DNA complexes, at N:P ratio 2 or PEI:DNA complexes at N:P ratio 3–5, in 10 mM HEPES, pH 7.4 2. Dissolve PHPMA–NHS in water at a concentration 15 mg/mL. 3. Add the PHPMA–NHS solution to complex solution at desired concentration (0.2–1 mg/mL final concentration is usually optimal). 4. Let the reaction proceed at room temperature for at least 3 h: Complexes are then ready to use.

3.4. Coating with Multifunctional Polymers Coating with multifunctional polymers was recently developed in this laboratory as an alternative approach to the more common coating with monofunctional PEG. An advantage of coating with multifunctional polymer is that it provides complexes not only with better stability, because of increased hydrophilic character of their surface, but also with lateral stabilization, which was found to be an important factor influencing their biological properties (15). We have used a statistical copolymer of HPMA with methacryloylated tetrapeptide, GlyPheLeuGly, having terminal carboxyl group activated as reactive 4-nitrophenyl ester (PHPMA-ONp) (Fig. 2). Multifunctional PHPMA is not commercially available, and needs to be synthesized. The synthesis is well described by Ulbrich (22), and its description here would be beyond the scope of this chapter. The coating procedure using multifunctional PHPMA itself is similar to that with monofunctional polymers. Schematic representation of the differences are

68

Oupicky, Read, and Bettinger

illustrated in Fig. 1. The polymer of typical mol wt 20,000 contains on average 12 reactive 4-nitrophenyl groups/molecule, and this enables multivalent attachment to the surface of the complex, which is assumed to be cooperative in nature. The coating then stabilizes the complex by crosslinking its surface, introducing another degree of stabilization compared to monofunctional polymers. The following protocol describes a procedure used to coat DNA complexes with multifunctional PHPMA-ONp. 1. Dissolve PHPMA-ONp in pure water (5 mg/mL) and add to complexes (prepared according to the standard protocol in 10 mM HEPES, pH 7.4), at a final concentration of 200 µg/mL (i.e., 40 µL/mL). 2. Accelerate the reaction by addition of HEPES buffer, pH 7.8, 0.5 M to 50 mM concentration. 3. Incubate for at least 3 h at ambient temperature or overnight at 4°C. 4. Reaction can be monitored by measuring the disappearance of 4-nitrophenyl esters at 274 nm (¡274 = 10,500 L/[mol/cm]). The PHPMA-ONp hydrolysis halflife at pH 8.0 at 25°C is about 30 min. 5. Before use, the remaining 4-nitrophenyl groups can be aminolyzed by addition of aminoethanol solution 2 µL/mL of complexes to a final aminoethanol concentration of 0.0004% v/v.

3.5. Attaching of Targeting Agents Simple polycation–DNA complexes show good transfection activity in vitro, mostly because of their nonspecific uptake by adsorptive endocytosis. Coating the complexes with hydrophilic polymers changes their surface properties, namely, the positive charge responsible for cellular uptake. As a result, the complexes show decreased uptake into cells and a reduction in transfection activity (15,23). This is in fact desirable because it enables retargeting of complexes to selected cells or tissues. In order to achieve targeted delivery, it is necessary to attach targeting ligands to the coated complexes. Both types of described polymers can in principle be used for attaching targeting ligands to the surface of the complexes. However, only the use of multifunctional PHPMA-ONp has been published so far. The principle of attachment of targeting ligands onto DNA complexes by means of multifunctional PHPMA-ONp is schematically shown in Fig. 1.

3.5.1. Attachment of Targeting Agents via Multifunctional PHPMA Polymers with multivalent reactive sites can be used to couple numerous smaller molecules to complexes, which can serve as targeting ligands for binding sites on cells. PHPMA contains several reactive groups, part of which can be used for the coating reaction and the rest for attaching targeting ligands (Fig. 1). Using multifunctional PHPMA for attaching targeting ligands to the

Steric Stabilization of DNA Complexes

69

complexes is easy, and the attachment is realized through amino groups. Targeting ligands, if of a protein nature, usually contain amino groups in sufficient amount. The disadvantage is the limited control on the specificity of binding, and also a possibility of crosslinking the complexes via the multifunctional polymer. For efficient attachment of targeting ligands, the coating reaction has to be stopped by addition of the targeting ligand at a suitable time in order to preserve sufficient amounts of reactive 4-nitrophenyl groups, and at the same time allow proper coating of the complexes (15). The following protocol describes the coating of PLL-DNA complexes and attachment of the targeting ligand transferrin. 1. Dissolve PHPMA-ONp in pure water at the concentration of 5 mg/mL and add to complexes at a final concentration of 200 µg/mL (i.e., 40 µL/mL). 2. Add HEPES buffer (pH 7.8, 0.5 M) to 50 mM concentration to accelerate the reaction. 3. Leave the reaction mixture for 2 h at 4°C. 4. Add 10 µL of transferrin solution (10 mg/mL) to the complex solution and incubate overnight at 4°C. 5. Other targeting ligands may require different timing of addition to the coated complexes and different concentration of ligand.

3.5.2. Attachment of Targeting Agents via Heterobifunctional PEG Another possibility for attaching targeting agents to coated complexes would be the use of heterobifunctional PEG. An advantage of using heterobifunctional PEG could be better control over the product structure, as reactive groups toward two different functionalities are used. This approach has been successfully used for attaching targeting ligands (antibodies) to liposomes (24). However, to the authors’ knowledge, no paper describing the use of this approach for targeting of polycation–DNA complexes has yet been published.

3.6. Properties and Analysis of Coated Complexes 3.6.1. Colorimetric Determination of Degree of Modification The efficiency of the coating reaction can be estimated by measuring the decrease in the number of amino groups in complexes. A wide range of reagents is available for this purpose including 2,4,6-trinitrobenzenesulfonic acid (TNBS) (25), ninhydrin (13), and fluorescamine (12). The advantage of using TNBS is that the assay is performed in fully aqueous solution (0.05 M borate buffer); both fluorescamine and ninhydrin assays are performed in aqueous–organic solutions, which raises the question of possible influence of the presence of organic solvents on the complex properties. Fluorescence-based fluorescamine assay, on the other hand, is at least 10-fold more sensitive and can be performed more quickly than absorption–based TNBS and ninhydrin assays.

70

Oupicky, Read, and Bettinger

The following two protocols are for determination of amino groups in DNA complexes. The fluorescamine assay is described in Chapter 11 by Read et al. 3.6.1.1. TNBS ASSAY 1. Mix 0.4 mL of complexes (made at 20 µg/mL DNA concentration) with 0.4 mL 0.1 M Na2B4O7·10H2O, and add 0.02 mL TNBS solution diluted from 1 M stock solution (Fluka, no. 92822) to 15 mg/mL with water. 2. Incubate for 45 min at room temperature and measure absorbance at 420 nm (absorbance of the solution made from PLL (20,000)–DNA N:P = 2 is usually about 0.35). 3. As a blank, add 0.02 mL TNBS to 0.8 mL borate buffer. 4. Compare absorbances of the coated complexes with those of the original complexes and calculate % of modified amino groups.

3.6.1.2. NINHYDRIN ASSAY 1. Prepare solution A (2.5 g ninhydrin in 50 mL ethanol) and solution B (1.3 mg potassium cyanide in 2 mL water and 98 mL pyridine). 2. Dilute sample containing ~10 µg polycation (PLL, PEI) with 0.1 mL solution B and 0.075 mL solution A. 3. Stop the reaction after heating for 10 min at 95°C by addition of 0.75 mL 60% ethanol and measure the absorption of the resulting blue solution at 570 nm (the assay gives identical results with free or DNA-bound PEI).

3.6.2. Size of Coated Complexes The introduction of a surface layer of hydrophilic polymer causes an increase in the size of the complexes that can be easily measured by dynamic light scattering (photon correlation spectroscopy [PCS]). Under the previously described conditions used for coating reactions, the authors have usually observed an increase in diameter following modification with mPEG(5000)-SPA of about 15–20 nm and slightly higher (about 20–25 nm) in the case of multifunctional PHPMA-ONp (mol wt 20,000). The thickness of the coating polymer layer is related to a reduction in phagocytic uptake, as demonstrated on polystyrene nanoparticles. For particles of the size of typical DNA complexes (~100 nm), a layer thickness of approx 10 nm should be adequate to confer particle nonrecognition in vivo (10). The typical changes in size of PLL–DNA and PEI–DNA complexes, following their modification with mPEG(5000)-SPA and multifunctional PHPMA-ONp, are shown in Fig. 3. The increase of size mostly results from the presence of the surface polymer layer, but could also partly reflect crosslinking of complexes when using multifunctional PHPMA-ONp. According to Davis (10), the thickness (L) of the PEG layer on the surface of modified nanoparticles is linearly related to the number of ethylene oxide groups (EO): L (nm) = 0.125 EO

Steric Stabilization of DNA Complexes

71

Fig. 3. Effect of coating PLL (20,000)–DNA and PEI (25,000)–DNA complexes with mPEG(5000)-SPA and multifunctional PHPMA-ONp on the size of the complexes (measured by PCS).

This equation gives 14-nm thickness for PEG(5000) (i.e., increase in diameter of 28 nm), which is slightly more than the observed increase of the size of the complexes as documented in Fig. 3. Details about measuring sizes by PCS are described in Chapter 11 by Read et al. and Chapter 22 by Wiethoff et al.

3.6.3. Altered c Potential Using excess polycation for complex formation results in generating positively charged particles. Attachment of hydrophilic polymer (PEG) to the surface of the complexes screens (masks) the surface charge and as a result, a decreased c potential is usually observed. In the case of multifunctional PHPMA-ONp, recharging is observed because some reactive 4-nitrophenyl ester groups on PHPMA-ONp do not react with amino groups on the surface of the complex, but hydrolyze leaving a free carboxylic acid group, giving the complexes a negative c potential. The influence of coating on c potential of DNA complexes is shown in Fig. 4. The protocol for measuring c potential of DNA complexes is in Chapter 11 by Read et al.

3.6.4. Increased Stability Against Salt-Induced Aggregation The size of polycation–DNA complexes depends on their N:P ratio, and on the ionic strength of the solution. At high N:P ratios, simple polycation:DNA complexes are usually well-stabilized in low-ionic-strength solutions by

72

Oupicky, Read, and Bettinger

Fig. 4. Effect of coating PLL (20,000)–DNA and PEI (25,000)–DNA with mPEG(5000)-SPA and multifunctional PHPMA-ONp on the c potential of the complexes.

electrostatic repulsion. With increasing ionic strength of the solution, the attractive van der Waals forces become stronger than the electrostatic repulsion, because of the screening effect of salt suppressing the charged layer surrounding the particle. As a result, complexes aggregate with increasing salt concentrations. Several techniques can be used to study the stability of the coated complexes against salt-induced aggregation. The simple one suitable for routine testing is to follow changes in turbidity of the complex solution after addition of salt. For this purpose, a fluorimeter with excitation and emission wavelengths set to the same value (600 nm) is the easiest option. The uncoated polycation–DNA complexes usually precipitate very fast, forming large aggregates shortly after the addition of NaCl, so that the quality of coating is easily checked. Properly coated complexes should be stable in 0.15 M NaCl for an extended period of time (at least 24 h with only minimal change in size). However, distinct differences between properly coated and uncoated complexes are observed after several minutes in 0.15 M NaCl solution. Better information about the changes in the properties of the complexes in salt solution can be obtained by dynamic light scattering or a combination of static and dynamic light scattering techniques (7).

3.6.5. Morphology of Coated Complexes To obtain the full picture about the influence of the coating on the properties of the complexes, it is advisable to use several analytical techniques. Light scattering techniques (PCS) are powerful tools, but it is not always easy to

Steric Stabilization of DNA Complexes

73

obtain information about morphology of the complexes. Electron microscopy, on the other hand, gives direct image of the complexes (limitations of this technique are described in Chapters 11 and 12). Special attention needs to be paid to the possibility that coating of the complexes will disrupt their condensed structure. As a result, extended open structures can be obtained. Various extended and open structures were observed for DNA complexes of PEG-PLL and PEG-PEI graft and block copolymers (3,23). The presence of such structures has not been reported for coated complexes (PEG, PHPMA-ONp, PHPMA-NHS), but the possibility of disrupting and consequently decreasing the stability of the complexes as a direct result of coating cannot be completely ruled out.

3.6.6. Albumin-Induced Turbidity Assay Simple polycation–DNA complexes are cleared quickly from the bloodstream after intravenous administration (2). This is thought to result from binding of serum proteins. Reducing the amount of proteins binding to the complexes is thus believed to be essential for successful systemic gene delivery. The addition of albumin to simple polycation–DNA complexes in water results in significant turbidity, which may be utilized as a convenient and simple measure of the stability of the complexes toward albumin (or other proteins) interaction. Albumin is able to bind to simple polycation–DNA complexes forming ternary complexes (2). At certain ratios, the ternary complexes are hydrophobic and excessive aggregation can be observed. However, at high concentrations, albumin is able to reduce the aggregation of PLL–DNA complexes, acting as a surface stabilizer (1,11), which suggests that, although the simple PLL–DNA complexes show only low size stability in saline solution or in low concentrations of albumin, their size could be well stabilized in the presence of higher concentrations of proteins in the blood. However, the adsorbed protein still represents a problem for successful systemic delivery. Although the addition of low concentrations of albumin (90%).

3.6.9. Freeze-Drying of Coated Complexes It is practical to store all pharmaceutical formulations in solid state. Successful freeze-drying of the simple polycation–DNA complexes requires the presence of cryoprotectants, such as sucrose, in order to preserve original size and biological properties (27). Multifunctional PHPMA-ONp-coated complexes can be easily freeze-dried and stored in solid state even without the presence of cryoprotectants during freeze-drying. We have successfully freeze-dried both concentrated PHPMA-coated and diluted complexes. The presence of 0.5% lactose in the solution has only a minimal effect on the sizes of reconstituted complexes.

Steric Stabilization of DNA Complexes

77

References 1. Ogris, M., Steinlein, P., Kursa, M., Mechtler, K., Kircheis, R., and Wagner, E. (1998) Size of DNA/transferrin-PEI complexes is an important factor for gene expression in cultured cells. Gene Ther. 5, 1425–1433. 2. Dash, P. R., Read, M. L., Barrett, L. B., Wolfert, M. A., and Seymour, L. W. (1999) Factors affecting blood clearance and in vivo distribution of polyeletrolyte complexes for gene delivery. Gene Ther. 6, 643–650. 3. Wolfert, M. A., Schacht, E. H., Toncheva, V., Ulbrich, K., Nazarova, O., and Seymour, L. W. (1996) Characterization of vectors for gene therapy formed by self-assembly of DNA with synthetic block copolymers. Hum. Gene Ther. 7, 2123–2133. 4. Kabanov, A. V. and Kabanov, V. A. (1995) DNA complexes with polycations for the delivery of genetic material into cells. Bioconjugate Chem. 6, 7–20. 5. Katayose, S. and Kataoka, K. (1997) Water-soluble polyion complex associates of DNA and poly(ethylene glycol)-poly(L-lysine) block copolymer. Bioconjugate Chem. 8, 702–707. 6. Seymour, L.W., Kataoka, K., and Kabanov, A. V. (1998) Cationic block copolymers as self-assembling vectors for gene delivery, in Self-assembling Complexes for Gene Delivery: From Laboratory to Clinical Trials (Kabanov, A. V., Felgner, P. L., and Seymour, L. W., eds.), Wiley, New York, pp. 219–239. 7. Oupicky, D., Konak, C., Dash, P. R., Seymour, L. W., and Ulbrich, K. (1999) Effect of albumin and polyanion on the structure of DNA complexes with polycation containing hydrophilic nonionic block. Bioconjugate Chem. 10, 764–772. 8. Toncheva, V., Wolfert, M. A., Dash, P. R., Oupicky, D., Ulbrich, K., Seymour, L. W., and Schacht, E. H. (1998) Novel vectors for gene delivery formed by self-assembly of DNA with poly(L-lysine) grafted with hydrophilic polymers. Biochim. Biophys. Acta 1380, 354–368. 9. Everett, D. H. (1988) Basic Principles of Colloid Science. The Royal Society of Chemistry, London. 10. Davis, S. S. and Illum, L. (1988) Polymeric microspheres as drug carriers. Biomaterials 9, 111–115. 11. Oupicky, D., Howard, K. A., Konak, C., Dash, P. R., Ulbrich, K., and Seymour, L. W. (2000) Steric stabilization of poly-L-lysine/DNA complexes by covalent attachment of semitelechelic poly[N-(2-hydroxypropl)methacrylamide]. Bioconjugate Chem. 11(4), 492–501. 12. Read, M. L., Etrych, T., Ulbrich, K., and Seymour, L. W. (1999) Characterisation of the binding interaction between poly( L-lysine) and DNA using the fluorescamine assay in the formation of non-viral gene delivery vectors. FEBS Lett. 461, 96–100. 13. Ogris, M., Brunner, S., Schuller, S., Kircheis, R., and Wagner, E. (1999) PEGylated DNA/transferrin-PEI complexes: reduced interaction with blood components, extended circulation in blood and potential for systemic gene delivery. Gene Ther. 6, 595–605.

78

Oupicky, Read, and Bettinger

14. Kircheis, R., Schuller, S., Brunner, S., Ogris, M., Heider, K.-H., Zauner, W., and Wagner, E. (1999) Polycation-based DNA complexes for tumor-targeted gene delivery. J. Gene Med. 1, 111–120. 15. Dash, P .R., Read, M. L., Fisher, K., Howard, K. A., Wolfert, M., Oupicky, D., et al. (2000) Decreased binding to proteins and cells of polymeric gene delivery vectors surface modified with a multivalent hydrophilic polymer and retargeting through attachment of transferrin. J. Biol. Chem. 275, 3793–3802. 16. Plank, C., Mechtler, K., Szoka, F. C., and Wagner, E. (1996) Activation of the complement system by synthetic DNA complexes: a potential barrier for intravenous gene delivery. Hum. Gene Ther. 7, 1437–1446. 17. Wolfert, M. A., Dash, P. R., Nazarova, O., Oupicky, D., Seymour, L. W., Smart, S., et al. (1999) Polyelectrolyte vectors for gene delivery: influence of cationic polymer on biophysical properties of complexes formed by self-assembly with DNA. Bioconjugate Chem. 10, 993–1004. 18. van de Wetering, P., Moret, E. E., Schuurmans-Nieuwenbroek, M. E., van Steenbergen, M. J., and Hennink, W. E. (1999) Structure-activity relationship of water-soluble cationic methacrylate/methacrylamide polymers for nonviral gene delivery. Bioconjugate Chem. 10, 589–597. 19. Bettinger, T., Remy, J.-S., and Erbacher, P. (1999) Size reduction of galactosylated PEI/DNA complexes improves lectin-mediated gene transfer into hepatocytes. Bioconjugate Chem. 10, 558–561. 20. Zou, S.-M., Erbacher, P., Remy, J.-S., and Behr, J.-P. (2000) Systemic linear polyethylenimine (L-PEI)-mediated gene delivery in the mouse. J. Gene Med. 2(2), 128–134. 21. Duncan, R. and Ulbrich, K. (1993) Development of N-(2-hydroxypropl)methacrylamide copolymer conjugates for delivery of cancer chemotherapy. Macromol. Chem. Macromol. Symp. 70/71, 157–162. 22. Ulbrich, K., Subr, V., Strohalm, J., Plocova, D., Jelinkova, M., and Rihova, B. (2000) Polymeric drugs based on conjugates of synthetic and natural macromolecules. I. Synthesis and physico-chemical characterisation. J. Control. Release 64, 63–79. 23. Erbacher, P., Bettinger, T., Belguise-Valladier, P., Zou, S., Coll, J.-L., Behr, J.-P., and Remy, J.-S. (1999) Transfection and physical properties of various saccharide, poly(ethylene glycol), and antibody-derivatized polyethylenimines (PEI). J. Gene Med. 1, 210–222. 24. Mastrobattista, E., Koning, G. A., and Storm, G. (1999) Immunoliposomes for the targeted delivery of antitumor drugs. Adv. Drug Delivery Rev. 40, 103–127. 25. Snyder, S. L. and Sobocinski, P. Z. (1975) An improved 2,4,6-trinitrobenzenesulfonic acid method for the determination of amines. Anal. Biochem. 64, 284–288. 26. Lasic, D. D., Martin, F. J., Gabizon, A., Huang, S. K., and Papahadjopoulos, D. (1991) Sterically stabilized liposomes: a hypothesis on the molecular origin of the extended circulation times. Biochim. Biophys. Acta 1070, 187–192. 27. Cherng, J .Y., Van De Wetering, P., Talsma, H., Crommelin, D. J. A., and Hennink, W. E. (1997) Freeze-drying of poly([2-dimethylamino]ethyl methacrylate)-based gene delivery systems. Pharm. Res. 14, 1838–1841.

Disulfide Cationic Lipids

79

6 Use of Disulfide Cationic Lipids in Plasmid DNA Delivery Fuxing Tang and Jeffrey A. Hughes 1. Introduction Gene therapy provides a paradigm of the treatment of human diseases. The ultimate goal of gene therapy is to cure both inherited and acquired disorders by removing the original causes, i.e., adding, blocking, correcting, or replacing genes. Although gene therapy trials have been initiated worldwide for more than two decades, little has been achieved in clinically curing diseases. One of the major hurdles for gene therapy is the lack of an efficient gene delivery system. An ideal gene delivery system should be specifically targeting, biodegradable, nontoxic, nonimmunogenic, and stable for storage. Cationic liposomes are the most extensively investigated nonviral vectors. It is generally believed that DNA–liposome complexes enter cells via endocytosis, although other pathways such as membrane fusion may exist (1,2). The barriers involved in the transfection process in vitro generally include the following aspects (3): 1. 2. 3. 4. 5. 6.

Formation of the liposome–DNA complexes. Entry of complexes into cell. Escape of DNA from the endosomes. Dissociation of DNA from liposomes. Entry of DNA into nucleus. DNA transcription.

The strategy for overcoming any of the above barriers should increase transgene expression, and the formulations, which overcome the major barriers, will result in greater transgene expression. Toxicity is one of the major barriers that limits the application of cationic lipids in clinical trials. The use of ester, amide, and carbamate linkages to tether From: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

79

80

Tang and Hughes

polar and hydrophobic domains of cationic lipid is a common strategy to lower toxicity. Ester bonds are biodegradable, but the introduction of the ester bond may also decrease the stability of liposomes in systemic circulation, when the liposomes are used in clinical trials. For example, Aberle et al. (4) constructed a tetraester cationic lipid and the liposome demonstrated lower toxicity in NIH 3T3 cells than did 3`(N-[N',N' dimethyl-aminoethane] carbamoyl) cholesterol (DC-Chol). However, this liposome must be used within 2 h of preparation. Carbamate bond is biodegradable and more stable than an ester bond in aqueous solution. Huang’s group (5) first introduced carbamate in the cationic lipid DC-Chol. DC-Chol was the first lipid used in clinical trials because of its combined properties of transfection efficiency, stability, and low toxicity (2). Another barrier for cationic lipid-mediated plasmid DNA (pDNA) delivery is the low transfection efficiency compared to the viral system. To solve these problems, the authors designed a class of disulfide cationic lipids that can enhance transfection activity and decrease the toxicity, but not sacrifice the stability of liposomes in aqueous solution (6,7). The strategy was to take advantage of the high intracellular reductive environment to use a disulfide linkercontaining cationic lipid that is selectively stable outside cells; it can be reduced in cells by intracellular reductive substances such as glutathione. The reduction of the disulfide linker resulted in the collapse of complexes, thus decreasing toxicity (Fig. 1) and enhancing the release of DNA from DNA–liposome complexes (Fig. 2). Dissociation of DNA from DNA–liposome complexes is one of the major barriers for cationic liposome-mediated gene transfection (1,3,8). The enhancement of the release of DNA was expected to increase transgene expression. Introduction of disulfide linker into cationic lipids enhanced transgene expression and decreased the toxicity of cationic lipid-mediated plasmid delivery (Figs. 3 and 4; 6,7,9,10). Disulfide conjugate techniques have been widely used in drug delivery to achieve high delivery efficiency (11–13). The normal method used in bioconjugation involves crosslinking or modification reactions using disulfide exchange processes to form disulfide linkage with sulfhydryl-containing molecules such as 3-(2-pyridyldithio)propionic acid n-hydroxy-succinimide ester (SPDP) (11–13). This method results in more stable disulfide compounds than the original compound, and thus is not suitable for the purpose of decreasing toxicity. The authors proposed a direct conjugation method to introduce disulfide lipid in cationic lipids-1', 2'-dioleoyl-sn-glycero-3'-succinyl-2-hydroxyetheyl disulfide ornithine conjugate (DOGSDSO) and cholesteryl hemidithiodiglycolyl Tris(aminoethyl)amine conjugate (CHDTAEA) (6,7). This method applied routine chemical synthesis, which is more economical, compared with the disulfide exchange method. For example, the price of SPDP, which is the key compound used in the classic disulfide exchange method, is thousands of times more expensive than dithioglycolic acid, which is used to introduce a disulfide

Disulfide Cationic Lipids

81

Fig. 1. Cytotoxicity of the liposome–DNA complexes was studied in CHO and SKnSH cells. A fixed dose of 1 µg/well pDNA was mixed with increasing amounts of cationic liposomes and used in the toxicity assay. Cell viability was calculated as percentage of survival cells as stated in Subheading 3. (A) Toxicity in CHO cells. (B) Toxicity in SKnSH cells. 䉬, CHDTAEA–DOPE, 䊏, CHSTAEA–DOPE; 䉱, DCChol–DOPE. Data is shown as mean ± SD (n = 3).

linker in the authors’ methods. This economical method will demonstrate an advantage in scale-up clinical trials and in pharmaceutical industry applications. Another advantage of this method is that disulfide bonds with different stability can be chosen for various applications. 2. Materials 1. 2-Hydroxyethyl disulfide (Aldrich, Milwaukee, WI), dithiodiglycolic acid (Pfaltz & Bauer, Waterbury, CT). 2. 1,2-Dioleoyl-sn-glycero-3-succinate, cholesterol (Avanti Polar Lipids, Ala– baster, AL).

82

Tang and Hughes

Fig. 2. Gel electrophoretic analysis of the release of pDNA from cationic liposomes–DNA complexes in 10 mM glutathione in phosphate-buffered saline. (A) DNA; (B) CHDTAEA–DOPE + DNA (2:1,w/w); (C) CHSTAEA–DOPE + DNA (2:1,w/w).

Fig. 3. Transgene expression in primary rat neuronal cultures transfected with a fixed amount of pDNA (3 µg) and various liposomal formulations. Number of replicates, n = 4. Each experiment was repeated at least 3×. The error bars represent mean ± SD.

3.

L-Ornithine,

2-(tert-butoxycarbonyloxyimino)-2-phenylacetonitrile (BOC-ON), triethylamine (TEA), 1,3-dicyclohexylcarbodiimide (DCCD), 1,4-dioxane, trifluoroacetic acid, 4-pyrrolidinopyridine, ethyl acetate, dichloromethane and methanol (Aldrich). 4. Chromatographic silica gel (200–425 mesh) (Fisher, Fair Lawn, NJ). 5. 1,2-Dioleoyl phosphatidylethanolamine (DOPE) (Avanti Polar Lipids).

Disulfide Cationic Lipids

83

Fig. 4. Comparison of the transfection of pGL3-luciferase plasmid by CHDTAEA–DOPE, CHSTAEA–DOPE, and DC-Chol in CHO and SKnSH cells. A fixed dose of 2 µg/well pGL3 DNA was mixed with increasing weight ratios of cationic liposomes (calculation based on cationic lipid) and used for transfection. (A) Transfection in CHO cells; (B) Transfection in SKnSH cells. 䉬, CHDTAEA– DOPE, 䊏, CHSTAEA–DOPE; 䉱, DC-Chol–DOPE. Data is shown as mean ± SD (n = 3). 6. 7. 8. 9.

Rotary evaporator Buchi 011(Buchi, Switzerland). Kontes ChromaflexTM chromatography column (Kontes,Vineland, NJ). Sonic Dismembrator 60 (Fisher Scientific). LiposoFastTM (Avestin, Ottawa, Canada).

84

Tang and Hughes

3. Methods 3.1. Synthesis of DOGSDSO (see Fig. 5) 1. Dissolve a total of 0.5 g A (ornithine) (0.0030 mol) and TEA in 6 mL water and 4 mL dioxane. 2. Add a total of 1.6 g BOC-ON (0.0066 mol) to the solution and stir the mixture at room temperature overnight. 3. Add 6 mL saturated sodium bicarbonate solution to the reaction mixture and extract the solution 3× with 20 mL ethyl acetate. Product B should be in aqueous layer at this step. 4. To the aqueous layer, add 20 mL 5% citric acid solution. Extract product B with 3× 20 mL ethyl acetate. 5. Collect the organic layers and dry over anhydrous sodium sulfate. Evaporate the organic solvent on a rotary evaporator. Pure product B is a colorless oil. 6. 0.388 g B (1.167 mmol), 0.360 g 2-hydroxyethyl disulfide (2.334 mmol), and 0.0173 g 4-pyrrolinopridine are dissolved in 20 mL methylene chloride under nitrogen (see Note 1). 7. To the solution, 1.284 mL 1 M DCC in dichloromethane is added via syringe under nitrogen. The mixture is stirred at room temperature under nitrogen overnight. 8. The white precipitate N,N-dicyclohexyl urea is filtered and the solvent is evaporated. 9. Product C is purified by chromatography on silica gel with hexane:ethyl acetate (1:1, v/v). Rf = 0.25. 10. 0.075 g 1,2-dioleoyl-sn-glycero-3-succinate (D) (0.104 mmol), 0.0788 g C (0.156 mmol), and 0.002 g 4-pyrrolidinopridine (0.0156 mmol) are mixed in dichloromethane under nitrogen. 11. 0.156 mL 1 M DCCD in dichloromethane is added to the mixture via syringe under nitrogen. The reaction mixture is stirred at room temperature under nitrogen overnight. 12. White precipitate N,N-dicyclohexyl urea is filtered and product E is purified by chromatography on silica gel with an eluant of hexane:ethyl acetate (3:1). Rf = 0.25. 13. To remove the BOC group, 2 mg E is dissolved in dichloromethane in a 50-mL round bottomed flask. The solvent is evaporated on a rotary evaporator and the flask is cooled on ice for 10 min to 0°C. 14. 5 mL trifluoroacetic acid is added to the flask at 0°C and the flask is incubated at room temperature for 5 min. Excess trifluoroacetic acid is evaporated on a rotary evaporator and evaporated further using a stream of nitrogen for 30 min. The product DOSGDSO is ready for preparation of liposomes (see Note 2).

3.2. Synthesis of CHDTAEA (see Fig. 6) The strategy of synthesis of CHDTAEA is similar to that of DOGSDSO with some modification. The electron-withdrawing properties of two _-carboxyl groups at the `-position of disulfide bond weaken the disulfide linker in CHDTAEA and make DNA easily releasable by reductive substances.

Disulfide Cationic Lipids

85

Fig. 5. Scheme of synthesis of 1', 2'-dioleoyl-sn-glycero-3'-succinyl-2-hydroxyethyl disulfide ornithine conjugate (DOGSDSO). 1. A total of 2.0 g (6 mmol) cholesterol and 1.9 g dithiodiglycolic acid (12 mmol) are dissolved in 30 mL ethyl acetate under nitrogen (see Note 3). 2. A total of 12 mmol DCCD, 0.19 g 4-pyrrolidinopyridine (1.2 mmol), and 1.43 mL TEA are added to the solution at 0°C. 3. The mixture is warmed to room temperature by standing and stirred under nitrogen overnight.

86

Tang and Hughes

Fig. 6. Scheme of synthesis of cholesteryl hemidithiodiglycolyl Tris(aminoethyl)amine (CHDTAEA). 4. The reaction mixture is filtered and washed by a 5% solution of citric acid, followed by brine, 3×. 5. The organic layer is collected and dried over sodium sulfate. 6. The organic solvent was evaporated on rotary evaporator. 7. The product cholesteryl hemidithiodiglycolate is purified on silica gel with an eluant of hexane:ethyl acetate:acetic acid (30:10:1). Rf = 0.3. 8. N,N-BOC2-Tris(2-aminoethyl) amine is prepared by treating Tris(2-aminoethyl) amine with two equivalents of BOC-ON in wet tetrahydrofuran. 9. A total of 0.3 g cholesteryl hemidithiodiglycolate (0.5 mmol), 0.24 g N,NBOC2-Tris(2-aminoethyl) amine (0.65 mmol), and 0.007 g 4-pyrrolidinopridine (0.005 mmol) is dissolved in 20 mL of dichloromethane. 10. A total of 0.6 mmol DCCD in dichloromethane and 0.06 mL TEA is added dropwise at 0°C. 11. The mixture is stirred at room temperature under nitrogen overnight. 12. The product N,N-BOC2–CHDTAEA is separated and purified on silica gel with a developer of dicholomethane:methanol (10:1). Rf = 0.35. 13. To remove BOC group, 2 mg N,N-BOC2 –CHDTAEA is dissolved in dichloromethane in a 50-mL round -bottomed flask. The solvent is evaporated on a rotary evaporator and the flask is cooled on ice for 10 min to 0°C. 5 mL TFA is added to the flask at 0°C and the flask is incubated at room temperature for 5 min. Excess TFA is evaporated on a rotary evaporator and evaporated further using a stream of nitrogen for 30 min. The product CHDTAEA is ready for preparation of liposomes.

3.3. Liposome Preparation 1. 5 mg Disulfide lipid DOGSDSO or CHDTAEA is dissolved in chloroform in a 250-mL round-bottomed flask.

Disulfide Cationic Lipids

87

2. One molar equivalent of DOPE is mixed with DOGSDSO or CHDTAEA in a flask. 3. Chloroform is evaporated on a rotary evaporator at room temperature and residual solvent is dried using a stream of nitrogen for 10 min (see Note 4). 4. Sterile water is added to the flask to hydrate the lipid film (see Note 5). 5. The flask is shaken at 30°C for 30 min. 6. The liposomes are hydrated at 4°C overnight. 7. The liposome suspension is sonicated using a Sonic Dismembrator for 2 min at 4°C to form homogenized liposomes (see Note 6). 8. When certain sizes of liposomes are required, the liposome suspension is passed through a membrane of specific sizes using LiposoFastTM (Avestin). 9. The liposomes are stored at 4°C until used for plasmid delivery. The liposomes do not lose activity for at least 1 yr.

4. Notes 1. The esterifcation reaction is hard to complete at room temperature. 4-pyrrolidinopyridine is a catalyst that can assist the reaction to be completed. 2. Disulfide lipid is not stable on chromatography column via separation. Because this step was reported to be quantitative (14), no further purification was attempted. 3. Ethyl acetate is a solvent with moderate polarity. Ethyl acetate was a better solvent than N,N-dimethylformide in this special case. 4. Temperature should be controlled to not exceed 30°C in considering the stability of disulfide bond. 5. The amount of water added was decided by the concentration required in the experiments. The concentration used in this laboratory is 1 mg/mL for in vitro or 5 mg/mL for in vivo experiments. In in vivo experiments, 5% dextrose solution is recommended to keep isotonic. Liposomes can still be made in sterile water (more stable) at a high concentration and diluted to concentration required in experiments, by dextrose solution of high concentration to reach a final concentration of 5% dextrose. 6. When the disulfide liposomes were made using sonicator, the size of liposomes was in a range of 130 ± 20 nm.

References 1. Rolland, A. P. (1998) From genes to gene medicines: recent advances in nonviral gene delivery. Crit. Rev. Ther. Drug Carrier Syst. 15, 143–198. 2. Gao, X. and Huang, L. (1995) Cationic liposome-mediated gene transfer. Gene Ther. 2, 710–722. 3. Zabner, J., Fasbender, A. J., Moninger, T., Poellinger, K. A., and Welsh, M. J. (1995) Cellular and molecular barriers to gene transfer by a cationic lipid. J. Biol. Chem. 270, 18,997–19,007. 4. Aberle, A. M., Tablin, F., Zhu, J., Walker, N. J., Gruenert, D. C., and Nantz, M. H. (1998) A novel tetraester construct that reduces cationic lipid-associated cytotoxicity. Implications for the onset of cytotoxicity. Biochemistry 37, 6533–6540. 5. Gao, X. and Huang, L. (1991) A novel cationic liposome reagent for efficient transfection of mammalian cells. Biochem. Biophys. Res. Commun. 179, 280–285.

88

Tang and Hughes

6. Tang, F. and Hughes, J. A. (1999) Use of dithiodiglycolic acid as a tether for cationic lipids decreases the cytotoxicity and increases transgene expression of plasmid DNA in vitro. Bioconjug. Chem. 10, 791–796. 7. Tang, F. and Hughes, J. A. (1998) Introduction of a disulfide bond into a cationic lipid enhances transgene expression of plasmid DNA. Biochem. Biophys. Res. Commun. 242, 141–145. 8. Escriou, V., Ciolina, C., Helbling-Leclerc, A., Wils, P., and Scherman, D. (1998) Cationic lipid-mediated gene transfer: analysis of cellular uptake and nuclear import of plasmid DNA. Cell. Biol. Toxicol. 14, 95–104. 9. Tang, F., Wang, W., and Hughes, J. A. (1999) Cationic liposomes containing disulfide bonds in delivery of plasmid DNA. J. Liposome Res. 9, 331–347. 10. Ajmani, P. S., Tang, F., Krishnaswami, S., Meyer, E. M., Sumners, C., and Hughes, J. A. (1999) Enhanced transgene expression in rat brain cell cultures with a disulfide-containing cationic lipid. Neurosci. Lett. 277, 141–144. 11. Boutorine, A. S. and Kostina, E. V. (1993) Reversible covalent attachment of cholesterol to oligodeoxyribonucleotides for studies of the mechanisms of their penetration into eucaryotic cells. Biochimie 75, 35–41. 12. Legendre, J. Y., Trzeciak, A., Bohrmann, B., Deuschle, U., Kitas, E., and Supersaxo, A. (1997) Dioleoylmelittin as a novel serum-insensitive reagent for efficient transfection of mammalian cells. Bioconjug. Chem. 8, 57–63. 13. Trail, P. A., Willner, D., Knipe, J., Henderson, A. J., Lasch, S. J., Zoeckler, M. E., et al. (1997) Effect of linker variation on the stability, potency, and efficacy of carcinoma-reactive BR64-doxorubicin immunoconjugates. Cancer Res. 57, 100–105. 14. 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. USA 86, 6982–6986.

Lipid–ODN Interaction with LDL

89

7 Interactions of Lipid–Oligonucleotide Conjugates with Low-Density Lipoprotein Erik T. Rump, Erik A. L. Biessen, Theo J. C. van Berkel, and Martin K. Bijsterbosch 1. Introduction The ability of antisense oligonucleotides to interdict, sequence-specifically, the expression of pathogenic genes affords an exciting new strategy for therapeutic intervention (1–3). Oligonucleotides with physiological phosphodiester internucleotide bonds are rapidly degraded, predominantly by exonucleases. Numerous oligonucleotide analogs have therefore been synthesized to confer resistance toward nuclease activity (3). The phosphorothioate analog is the most extensively studied, and phosphorothioate oligodeoxynucleotides have been shown to be potent inhibitors of the expression of their target genes in vitro and in vivo (1,3). However, phosphorothioate oligodeoxynucleotides also bind avidly and nonspecifically to proteins, thus provoking a variety of non-antisense effects (4). Oligonucleotide analogs that do not bind to proteins are therefore expected to display less nonantisense side effects. However, protein binding also affects the in vivo disposition of oligonucleotides. Nonphosphorothioate oligonucelotide analogs generally do not bind to serum proteins, and are therefore rapidly cleared from the circulation, protein-bound phosphorothioate oligodeoxynucelotides circulate much longer (5,6). The authors’ aim is to prolong the circulation time of nonphosphorothioate oligonucleotides, in order to increase the exposure of target cells to the oligonucleotides. The approach is to conjugate oligonucleotides with lipids, with the objective of inducing association of the oligonucleotide with longcirculating lipid particles (lipoproteins and lipoprotein-like particles). The biological fate of the particle-associated oligonucleotide will then be determined by the lipid carrier. The authors’ studies focused on low-density From: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

89

90

Rump et al.

lipoprotein (LDL), the main cholesterol-transporting vehicle in human circulation. LDL is a spherical particle (diameter 23 nm), consisting of an apolar core of cholesteryl esters and triglycerides, which is surrounded by a shell of cholesterol and phospholipids (7). A large part of the surface is covered by apoprotein B100, which is recognized by specific LDL receptors. LDL is slowly cleared from the circulation (7,8), which makes it suitable for prolonging the circulation of associated oligonucleotides. A second reason to choose LDL as carrier is that it has been shown that a variety of tumor cell types (e.g., leukemic cells) internalize large amounts of LDL via the LDL receptor (9). Association of an oncogene-specific antisense oligonucleotide with LDL may lead to a higher uptake of the oligonucleotide by tumors, and consequently higher therapeutic efficacy. This chapter describes the conjugation of various lipids to a c-myb-directed oligonucleotide. The association of the lipid–oligonucleotide conjugates (lipid– ODNs) with LDL is characterized, as well as the stability of the lipid-ODN–LDL complexes in vitro in rat plasma and in vivo in rats. 2. Materials 2.1. Synthesis of Activated Lipid Structure 1. 2. 3. 4. 5. 6. 7. 8.

Lithocholic acid. Oleic acid. 3_-(oleoyloxy)-5`-cholanic acid. 3_,7_-bis(oleoyloxy)-5`-cholanic acid. 3`-(oleoyloxy)-5-cholenic acid. Pentafluorophenol. Dicyclohexylcarbodiimide. Cholesterol chloroformate.

2.2. Synthesis and Purification of 3'-Lipid-ODN 1. An 18-mer antisense oligonucleotide complementary to c-myb (10) (5'-GTG CCG GGG TCT TCG GGC-3') was from Eurogentec (Seraing, Belgium) and had a phosphodiester backbone. The antisense oligonucleotide had three phosphorothioate linkages at the 5' end and a C7-amino linker at the 3'-end. 2. [3H]-Labeled 3'-amine 18-mer antisense oligonucleotide, radiolabeled with 3H by heat-catalyzed exchange at the C8 positions of the purine nucleotides (see Note 1 and refs. 11,12). 3. [3H]ISIS-9388, a 3'-cholesteryl-conjugated phosphorothioate oligodeoxynucleotide specific for murine intercellular adhesion molecule-1 (13) was kindly provided by Dr. M. Manoharan (ISIS Pharmaceuticals, Carlsbad, CA). 4. Na125I (carrier-free) and 3H2O were from Amersham (Amersham, UK). 5. Low-melting multipurpose agarose and Agarase from Pseudomonas atlantica were obtained from Boehringer (Mannheim, Germany). 6. Tween-20 was from Merck (Darmstadt, Germany).

Lipid–ODN Interaction with LDL 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17.

91

Lithium perchlorate (LiClO4). Acetone. Dimethylformamide. Dioxane. N,N-Diisopropylethylamine (DIPEA). Dichloromethane. 50 mM Triethylamine ammonium acetate, pH 7.0. Agarase. 30 mM Bis-Tris, containing 10 mM EDTA, pH 6.5. Acetonitrile. 45 mM Tris-borate buffer, containing 0.1 mM EDTA, pH 8.4.

2.3. Determination of Melting Temperatures 1. A 28-mer sense oligonucleotide (5'-CCA TGG CCC GAA GAC CCC GGC ACA GCA T-3') was from Eurogentec and had a phosphodiester backbone. 2. Phosphate-buffered saline (PBS): 10 mM sodium phosphate buffer, pH 7.4, containing 0.15 M NaCl.

2.4. LDL and Lipid-ODNs 1. Human LDL: density 1.024–1.063 g/mL. 2. Hionic Fluor TM scintillation cocktail. 3. Superose 6 Precision chromatography column, 3.2 × 300 mm (Pharmacia).

2.5. Plasma Clearance and Liver Association 1. Male Wistar rats (180–230 g). 2. Ketamine hydrochloride was from Eurovet (Bladel, the Netherlands). 3. HypnormTM (0.315 mg/mL fentanyl citrate and 10 mg/mL fluanisone) was from Janssen-Cilag (Sauderton, UK). 4. Thalomonal TM (0.05 mg/mL fentanyl and 2.5 mg/mL droperidol) was from Janssen-Cilag. 5. Hionic Fluor and Emulsifier SafeTM scintillation cocktails and Soluene-350TM were from Packard (Downers Grove, IL). 6. Heparinized collection tubes.

3. Methods 3.1. Conjugation of Oligonucleotides with Lipid Structures The authors synthesized a series of conjugates of lipids with a 18-mer antisense oligonucleotide complementary to the c-myb proto-oncogene. The oligonucleotide had a phosphodiester backbone with three phosphorothioate linkages at the 5' end as protection against 5'-exonucleases. At the 3' end, the oligonucleotide was provided with an amino group to enable conjugation. Figure 1 shows the structures of the lipids that were used for conjugation. To enable coupling to the 3'-amino-tailed oligonucleotide, the lipids were

92

Rump et al.

Fig. 1. Lipid structures of lipid-ODNs.

activated. Except cholesterol (ODN-4), all lipids were activated at their carboxylic acid functionalities with pentafluorophenol (see ref. 11 for full details of the synthesis of the activated lipid structures). Cholesterol chloroformate was used for the conjugation of cholesterol. Conjugation of the

Lipid–ODN Interaction with LDL

93

amino-terminated oligonucleotide with the activated lipids was performed in a mixture of water and the aprotic solvents, dimethylformamide and 1,4-dioxane (1:4:4; v/v/v), in which the organic base DIPEA was present to create mild basic conditions. After conjugation, the lipid-ODNs were purified. ODN-2, ODN-3, and ODN-4 (conjugates of oligonucleotide with litocholic acid, oleic acid, and cholesterol, respectively) were purified by reversed-phase high-performance liquid chromatography (RP-HPLC). Purification of conjugates of the oligonucleotide with the oleoyl steroid ester structures (ODN-5, ODN-6, and ODN-7) was not possible by RP-HPLC; no conjugated products were recovered. These lipid-ODNs were separated from unconjugated oligonucleotide by electophoresis in a 1% agarose gel, containing 0.1% Tween-20. Separation is accomplished by the formation of micelles of lipid-ODNs and Tween-20. The lipid-ODNs were retrieved from the gel by melting the gel, followed by digestion of the gel material by Agarase.

3.1.1. Synthesis of Activated Lipid Structure 5`-Cholanic acid 3_-ol pentafluorophenyl ester, oleic acid pentafluorphenyl ester, 3_-(oleoyloxy)-5`-cholanic acid pentaflorophenyl ester, 3_,7`-bis(oleoyloxy)-5`-cholanic acid pentaflorophenyl ester, and 3`(oleoyloxy)-5-cholenic acid pentaflorophenyl ester were synthesized as described in full detail earlier by activating lithocholic acid, oleic acid, 3_(oleoyloxy)-5`-cholanic acid, 3_,7`-bis(oleoyloxy)-5`-cholanic acid, and 3`-(oleoyloxy)-5-cholenic acid, respectively, with pentafluorophenol (11). The identity of all compounds was verified by mass spectrometry and nuclear magnetic resonance.

3.1.2. Synthesis and Purification of 3'-Lipid-ODN 1. The amino-terminated antisense oligonucleotide is precipitated as a Li-salt with 10 vol of 3% LiClO4 in acetone. 2. The oligonucleotide is subsequently dissolved in H2O and precipitated again with 10 vol acetone to remove final traces of LiClO4. 3. In a typical derivatization experiment, 15 nmol of oligonucleotide is dissolved in 350 µL H2O/dimethylformamide/dioxane (1:4:2, v/v/v). Subsequently, 1 µmol activated lipid, dissolved in 100 µL dioxane, and 35 µmol DIPEA are added. For the preparation of ODN-2, ODN-3, and ODN-4, the mixtures were incubated for 48 h at 37°C, and, for the preparation of ODN-5, ODN-6, and ODN-7, the mixtures were incubated for 48 h at 56°C. 4. Solvents are removed in a Speed-Vac concentrator, and the residue is taken up in 200 µL dichloromethane and 200 µL H2O. The layers are separated by centrifugation, and the organic layer is washed twice with 200 µL H2O. 5. The aqueous phases are combined and freeze-dried. 6. Derivatization of (3H)oligonucleotide is performed as described in steps 1–5 with 2 nmol labeled oligonucleotide and 4 nmol unlabeled oligonucleotide.

94

Rump et al.

7. Oligonucleotides conjugated with lithocholic acid, oleic acid or cholesterol are purified by RP-HPLC on a Waters C4 column (5 µm, 300 A, 300 × 3.9 mm) by applying a gradient of 1% CH3CN/min in 50 mM triethyl ammonium acetate (pH 7.0) at a flow rate of 1 mL/min. The gradient (5–50%) is started after elution for 5 min at 5% CH3CN. Oligonucleotides are detected at 260 nm. 8. All other lipid-conjugated oligonucleotides are purified by gel electrophoresis in a 1% (w/v) low-melting multi purpose agarose gel, containing 0.1% Tween-20, at pH 8.4 (45 mM Tris-borate buffer, containing 0.1 mM EDTA). Gel slices containing lipid-ODNs are melted for 5 min at 65°C. The agarose is digested for 2 h with Agarase (40 U/mL gel) at 45°C in 30 mM Bis-Tris, containing 10 mM EDTA. The lipidODNs are precipitated with 10 vol of acetone. To remove traces of undigested agarose, the precipitate is taken up in 200 µL H2O and passed over a filter paper (no. 589, Schleicher and Schüll). Lipid-ODNs are isolated in a yield of 35–75%.

3.2. Interaction of Lipid–ODNs with Their Target Sequences: Determination of Melting Temperatures To ascertain that the attached lipid structures do not interfere with the interaction of the antisense oligonucleotide with the target sequence, the authors determined the melting temperatures of antisense–sense hybrids. A 28-mer sense oligonucleotide was utilized with five overhang nucleotides at both the 3'- and 5'-end, to assess a possible interaction of the lipid with the singlestranded part of the hybrid. The melting temperatures of the hybrids with the conjugated ODNs differed not appreciably (maximally 2°C) from that of the hybrid with the unconjugated ODN-1 (Table 1). Thus, the bulky steroid lipids (particularly the oleoyl steroid esters) do not significantly interfere with the overhang nucleotides of the target sequence. These findings are consistent with reports of other lipid-ODNs (14). Melting temperatures of hybrids of the antisense lipid-ODNs with an 28-mer sense oligonucleotide were determined using a Perkin-Elmer spectrophotometer equipped with a PTP-6 thermal programmer. 1. Equimolar amounts of both oligonucleotides are dissolved to a concentration of 3.6 µM in PBS. 2. The mixtures are placed for 2 min at 96°C and subsequently slowly cooled to room temperature to allow annealing. 3. Then, the temperature is adjusted to 35°C and hybrids are melted by increasing the temperature to 95°C at a rate of 0.5°C/min (see Note 2).

3.3. Interaction of Lipid-ODNs with LDL To examine which of the conjugated lipid structures is able to associate the oligonucleotide with LDL, radiolabeled lipid-ODNs were incubated for 2 h at 37°C in PBS with equimolar amounts of radioiodinated LDL. Aliquots of the incubation mixtures were analyzed by agarose gel electrophoresis at pH 8.8 in

Lipid–ODN Interaction with LDL

95

Table 1 Melting Temperatures of Hybrids of Antisense Lipid-ONDs and Sense Oligonucleotide Oligonucleotide ODN-1 ODN-2 ODN-3 ODN-4 ODN-5 ODN-6 ODN-7

Melting temperature (°C) 71 72 72 73 70 69 70

Equimolar amounts of antisense lipid-ODNs and 28-mer sense oligonucleotide, dissolved in PBS, were heated for 2 min at 96°C, and subsequently slowly cooled to room temperature to allow annealing. Then, the temperature was adjusted to 35°C and hybrids were melted by increasing the temperature to 95°C at a rate of 0.5°C/min. Differences in melting temperatures measured in duplicate runs were 95% of the [3H]ODN-4, [3H]ODN-5, and [3H]ODN-7 comigrated in the gel with radioiodinated LDL (Rf, 0.2). Thus, conjugation of the oligonucleotide with cholesterol or the oleoyl esters of lithocholic acid or cholenic acid induces spontaneous and almost complete association of the oligonucleotide with LDL. These steroids meet the structural requirements for LDL anchors as defined by Firestone et al. (15; see Note 3). When lipid-ODNs were associated at higher molar ratios, the increase in the electrophoretic mobility was more clear (data not shown). Figure 2 also shows that conjugation of lithocholic acid and oleic acid (ODN-2 and ODN-3) did not induce association of the oligonucleotide with LDL at all. Only a proportion of the [3H]oligonucleotide conjugated to the bis-oleoyl steroid ester (ODN-6) comigrated with LDL. A substantial proportion of 3H-radioactivity migrated at Rf 0.2–1.0, which suggests that the lipid-ODN does associate with LDL, but that the complex slowly dissociates during electrophoresis. The lipid moiety of ODN-6 probably does not partition with its complete steroid structure into the lipids of LDL, but only with the two oleoyl chains. The complete bisoleoyl steroid ester structure may be too bulky to associate spontaneously with LDL. However, ODN-6 may well be incorporated in the recently developed artificial LDL-like carrier systems (16,17; see Note 4).

Fig. 2. Association of lipid-ODNs with LDL; analysis by agarose gel electrophoresis. Equimolar amounts of 125I-LDL and lipid(1.7 µM), dissolved in PBS + 1 mM EDTA, pH 7.4, were incubated for 2 h at 37°C. Aliquots of the incubation mixtures were subjected to gel electrophoresis in a 0.75% (w/v) agarose gel in 75 mM Tris-hippuric acid buffer (pH 8.8). After electrophoresis, the gel was cut into slices that were assayed for 3H-radioactivity (䊊), and 125I-radioactivity (䊉). The results are expressed as percentages of the recovered radioactivities. Recoveries were >95%.

(3H)ODNs

96 Rump et al.

96

Lipid–ODN Interaction with LDL

97

3.3.1. Isolation and Radioiodination of LDL Human LDL (density 1.024–1.063 g/mL) was isolated from the serum of fasted volunteers by density gradient ultracentrifugation (18) and were dialyzed against PBS containing 1 mM EDTA. Radioiodination was performed at pH 10.0 with carrier-free 125I as described by McFarlane (19). Protein concentrations of the lipoproteins were determined by the method of Lowry et al. (20) with bovine serum albumin as standard.

3.3.2. Determination of Association of Lipid–ODNs with LDL 1. Equimolar amounts of 125I-labeled LDL and 3H-labeled lipid-ODNs (1.7 µM), dissolved in PBS containing 1 mM EDTA, are incubated for 2 h at 37°C (see Note 5). 2. Aliquots of the incubation mixtures are subjected to gel electrophoresis in a 0.75% (w/v) agarose gel in 75 mM Tris-hippuric acid buffer, pH 8.8. 3. After electrophoresis, the gel is cut into slices, and the 125I radioactivity is counted after addition of 0.5 mL Soluene-350. 4. The gel slices are allowed to dissolve for 24 h at room temperature. 5. Then, 3 mL Hionic Fluor is added and samples are counted for 3H-radioactivity. The measured values of 3H-radioactivity are corrected for the contribution of 125I-radioactivity.

3.4. Exchange of Lipid–ODNs from LDL to Plasma Proteins The preceding subheading demonstrated that only the conjugates of the oligonucleotide with cholesterol or oleoyl steroid esters (ODN-4, ODN-5, and ODN-7) associate quantitatively with LDL. To examine the stability of the complexes of the lipid-ODNs with LDL in a biological matrix, the authors studied the exchange of the lipid-ODNs from preformed lipid-ODN–LDL complexes to components in rat plasma. Lipid-[3H]ODN–LDL complexes were incubated for 5 or 25 min with rat plasma and subsequently subjected to size-exclusion chromatography. The fractions were monitored for 3H-radioactivity, and the results are depicted in Fig. 3. Although HDL is the main lipoprotein present in rat plasma, no significant redistribution of ODN-4, ODN-5 and ODN-7 from LDL to HDL (elution vol 1.57 mL) was seen. The lipid-ODNs redistributed to some extent to triglyceride-rich lipoproteins (elution vol 0.9 mL). The oleoyl steroid ester-conjugated oligonucleotides (ODN-5 and ODN-7) (Fig. 3B,D) appeared to be most stably complexed with LDL, because after 25 min of incubation, >70% of the radioactivity still was associated with the LDL fractions. At that time, only approx 50% of the cholesteryl-conjugated ODN (ODN-4) (Fig. 3A) was found to be associated with LDL. The lipid-ODNs, ODN-4, ODN-5, and ODN-7, have a partially phosphorothioate (PS)-modified backbone (17% PS linkages). Many studies have found that PS linkages in oligonucleotides induce binding to serum proteins (22,23). Therefore, the authors also examined the stability of complexes of LDL and [3H]ISIS-9388, a full

98

Rump et al.

Fig. 3. Chromatographic profiles after incubation of lipid-ODN–LDL complexes with rat plasma. Preformed lipid-ODN/LDL complexes were incubated with rat plasma for 5 min (䊊) or for 25 min (䊉) at 37°C. The samples were subsequently injected onto a Superose 6 column (Pharmacia) and the fractions were assayed for 3H-radioactivity. The 3H-radioactivity in the fractions are expressed as percentage of recovered radioactivity (recoveries were >95%). The gray zones indicate the fractions containing 90% of the lipid-ODN–LDL complex at t = 0 min. (A) ODN-4; (B) ODN-5; (C) ODN-7; (D) ISIS-9388.

PS oligonucleotide that is conjugated at the 3' end with cholesterol. Upon incubation with rat plasma, ISIS-9388 dissociated more rapidly from a preformed complex with LDL than the partially PS-modified lipid-ODNs. After 25 min incubation, less then 20% of ISIS-9388 was found to be associated with LDL (Fig. 3D; see Note 6).

3.5. Determination of Exchange of Lipid–ODNs from LDL to Proteins in Plasma 1. Equimolar amounts of 125I-labeled LDL and 3H-labeled lipid-ODNs (200 pmol), dissolved in 100 µL of PBS containing 1 mM EDTA, are incubated for 2 h at 37°C. 2. Subsequently, 50 µL of the mixtures are injected onto a Superose 6 Precision Column (3.2 × 300 mm) (Smart System, Pharmacia) and eluted at a flow rate of 50 µL/min with PBS.

Lipid–ODN Interaction with LDL

99

3. 50 µL Fractions are collected and the three main fractions containing the lipidODN–LDL complexes are pooled. 4. 50 µL Aliquots of the pooled fractions are subsequently incubated at 37°C with 40 µL citrated rat plasma. 5. After incubation, the mixtures are injected onto the Superose 6 column and the column is eluted as described previously. The fractions are assayed for 3H-radioactivity.

3.6. Behavior of Lipid–ODN–LDL Complexes In Vivo To establish which of the lipid steroid structures is the most effective LDL anchor in vivo, the authors examined in the rat the plasma clearance of complexes of LDL and lipid-(3H)ODNs, ODN-4, ODN-5, and ODN-7. When injected without LDL, these lipid-ODNs were rapidly cleared from the circulation (Fig. 4A,C,E). At 5 min after injection, more than 95% of the dose was cleared, and a significant proportion of the radioactivity (15–40% of the dose) was recovered in the liver (Fig. 4B,D,F). The plasma clearance of the lipid-ODNs complexed with LDL was studied utilizing double-labeled complexes (lipid-(3H)ODN/(125I)LDL), which allows monitoring of both the lipid-ODN and the LDL carrier. Fig. 4A,C,E show that (125I)LDL was slowly cleared from the circulation, with a concomitant low liver uptake. These findings are consistent with the previously reported half-life of 5–6 h for LDL in the rat (8). The lipid-ODNs in the lipid-[3H]ODN/[125I]LDL complexes were cleared much slower than the noncomplexed lipid-ODNs (Fig. 4A,C,E). At 5 min after injection of the LDLcompexed lipid-ODNs, only 51, 39, and 24% of ODN-4, ODN-7, and ODN-5, respectively, had been cleared from the circulation (vs >95% for the noncomplexed lipid-ODNs). Thus, when complexed with LDL, both oleoyl steroid ester-conjugated oligonucleotides (ODN-5 and ODN-7) were more slowly cleared than the cholesteryl-conjugated ODN-4. Accordingly, the plasma area under the curve (AUC) of the LDL-complexed ODN-4 (1.49 ± 0.37 µg/min/mL) was significantly lower that the LDL-associated ODN-5 and ODN-7 (6.82 ± 1.34 µg/min/mL and 4.61 ± 0.38 µg/min/mL, respectively). The noncomplexed lipidODNs had plasma AUCs < 0.4 µg/min/mL. Compared to the free lipid-ODNs, the liver uptake of LDL-complexed lipid-ODNs was reduced (Fig. 4B,D,F). However, the lipid-ODN–LDL complexes were not completely stable, since the clearance of the lipid-ODNs did not completely resemble the clearance of LDL (Fig. 4A,C,E). The reduction of the clearance rate of the lipid-ODNs, achieved by complexation with LDL, was most evident for the oleoyl steroid ester-conjugated ODNs (ODN5 and ODN-7). The cholesteryl-conjugated ODN (ODN-4) displayed the highest leakage from LDL. 3.6.1. Determination of Plasma Clearance and Liver Association 1. Male Wistar rats are anesthetized by subcutaneous injection of a cocktail of ketamine-HCl, fentanyl, droperidol, and fluanisone (75, 0.04, 1.1, and 0.75 mg/kg, respectively).

100

Rump et al.

Fig. 4. Plasma clearance and liver association of lipid-ODNs and lipid-ODN–LDL complexes. Rats were injected with the free lipid-[3H]ODNs (䉱) or lipid-(3H)ODN/ (125I)LDL complexes (3H,䊊; 125I, 䊉). At the indicated times, the amounts of radioactivity in plasma (A,C,E) and liver (B,D,F) were determined. Values are means ± SEM of two (free lipid-ODNs) or three (ODN–LDL complexes) separate experiments. A and B: ODN-4; C and D: ODN-5; E and F: ODN-7. 2. The abdomen is opened. Free lipid-[3H]ODNs (4 µM in PBS), or complexes of lipid-[3H]ODNs with (125I)LDL (4 µM, complexes prepared as described above), are injected via the vena penis at a dose of 5 µg lipid-ODN/kg body wt.

Lipid–ODN Interaction with LDL

101

3. At the indicated times, 250 µL blood samples are taken from the inferior vena cava and collected in heparinized tubes. The blood samples are centrifuged for 10 min at 500g, and the plasma is assayed for radioactivity. 4. The total amounts of radioactivity in plasma are calculated using the equation: Plasma vol (mL) = [0.0291 × body wt (g)] + 2.54 (24) 5. At the indicated times, liver lobules are tied off and excised, and at the end of the experiment, the remainder of the liver is removed. 6. The amount of radioactivity in the liver at each time-point is calculated from the radioactivities and weights of the liver samples and is corrected for radioactivity in plasma present in the tissue at the time of sampling (85 µL/g fresh weight). 7. The plasma concentration-time AUC is calculated by computerized nonlinear fitting (Graphpad Prism, Graphpad Software, San Diego, CA).

3.6.2. Determination of Radioactivity Samples containing 3H are counted in a Packard Tri-Carb 1500 liquid scintillation counter. Liquid samples are counted without further processing using Emulsifier Safe or Hionic Fluor scintillation cocktails. Agarose gel slices are first dissolved in Soluene-350. Tissue samples are processed using a Packard 306 Sample Oxidizer. Samples containing both 125I and 3H, are first assayed for 125I-radioactivity using a Packard Auto-Gamma 5000 counter. The 3H-radioactivity is subsequently measured as described above, and corrected for the contribution of 125I-radioactivity.

3.7. Discussion This chapter describes the conjugation of an amino-terminated oligonucleotide with several lipid structures in solution phase. The conjugated lipid structures did not affect the association of the oligonucleotide with its target sequence, as judged by the lack of effects on the melting temperatures of the antisense–sense hybrids. Several of the lipid-ODNs, namely those conjugated with cholesterol or oleoyl steroid ester moieties, associate readily with LDL. The stability of the complexes of these lipid-oligonucleotides with LDL was investigated in vitro by incubation with rat plasma and in vivo in rats. Detailed reports of the experiments presented here were published earlier (11,25). In vitro, the authors examined the exchange of the lipid-oligonucleotides from preformed lipid-ODN–LDL complexes to components in rat plasma. The more lipidic oleoyl steroid ester structures appeared to be better LDL-anchors than the cholesteryl moiety. ISIS-9388 (a 3'-cholesteryl-conjugated phosphorothioate oligonucleotide) redistributed to a much higher extent from LDL to plasma proteins than the partially phosphorothioate-modified lipid-ODNs (ODN-4, ODN-5, and ODN-7). This is likely to be the result of the high affinity of the full phosphorothioate oligonucleotide for plasma proteins (22,23).

102

Rump et al.

When the lipid-ODNs were injected in rats without LDL, they were rapidly cleared from the circulation, meaning that lipid-conjugation alone is not sufficient to achieve a prolonged half-life in the circulation. A substantial amount of the lipidODNs was recovered in the liver, which may be ascribed to recognition by scavenger receptors on liver cells (13,26). When the lipid-ODNs were administered as complexes with LDL, the plasma clearance of the lipid-ODNs was considerably delayed and their liver uptake reduced. The oligonucleotides containing oleoyl steroid ester structures (ODN-5 and ODN-7) were more slowly cleared than the cholesteryl-conjugated ODN-4. The clearance and liver uptake of the LDL particles in the complexes was not altered, which indicates that complexation with the lipidODNs does not affect the integrity of the particle. Taken together, of all the steroid structures tested, the lithocholic acid-3_oleate structure (present in ODN-5) most effectively reduced clearance of the oligonucleotide in vivo and the exchange from LDL in vitro. The strong association of this steroid ester lipid anchor with the lipids of LDL is probably primarily responsible for the prolonged circulation.The backbone chemistry of the oligonucleotide is also important. A modification with low affinity for plasma proteins (e.g., oligoncucleotide with little phosphorothioate linkages or nonphosphorothioate oligonucleotides, such as morpholino or peptide nucleic acid oligomers [27,28]), will reduce dissociation of lipid-conjugated oligonucleotides from lipoproteins. Oligonucleotides conjugated with the oleoyl steroid esters remain associated to LDL and the retarded plasma clearance results in a higher exposure to target cells. Oligonucleotides associated with LDL (or LDL-like particles [16,17]) may be taken up along with the particles via LDL receptors that are overexpressed on various types of tumor cells, e.g., leukemic cells (9). The firm association of lipid-ODNs with LDL or LDL-like particles thus has the additional advantage that enhanced exposure is accompanied by enhanced rate of uptake by LDL receptor-overexpressing tumor cells. 4. Notes 1. Radiolabeling of oligoncucleotides. The 3'-amine antisense oligonucleotide was radiolabeled with 3H by heat-catalyzed exchange at the C8 positions of the purine nucleotides as described previously (11,12). The radiolabeled oligonucleotide was stored at –20°C. No loss of radioactivity from the oligonucleotide was detected during 6 mo of storage at –20°C. The specific activity of the radiolabeled oligonucleotide was 80 × 106 dpm/mg. 2. Differences in melting temperatures measured in duplicate runs were log (I/I0) = C¡(h)l

(1)

where A(h) is the absorbance at the specified wavelength, I and Io are the final and initial intensities of light transmitted, C represents sample concentration, ¡(h) the molar extinction coefficient at the absorbance wavelength, and l the pathlength of the sample cell (5). In cases in which ¡(h) is not known, the following approximations for natural nucleic acids are usually adequate: 1 A260 U double-stranded DNA (dsDNA) = 50 µg/mL; 1 A260 U single-stranded DNA (ssDNA) = 37 µg/mL; 1 A260 U ssRNA = 40 µg/mL.

UV Absorption and CD Spectroscopy

257

Note that 1 A260 U is defined here as the quantity of polynucleotide having an absorbance of 1.0 at 260 nm, when dissolved in 1 mL buffer and measured in a 1 cm cuvet at 20°C. A typical buffer might contain 20 mM sodium phosphate at pH 7.0 and 0.1 M NaCl. However, the actual absorbance of nucleic acids is dependent on both pH and ionic strength. As the pH changes, the bases undergo protonation and deprotonation reactions, producing shifts and alterations in their absorbance spectra (5), and the extinction coefficient of DNA is decreased in the presence of salts, compared to pure water (6). For example, in 10 mM Tris and 1 mM EDTA (pH 7.4), 1 A260 corresponds to a dsDNA concentration of 45 µg/mL; in highly purified water 1 A260 U corresponds to only 38 µg/mL of the same polynucleotide (7). This ionic strength dependence necessitates careful consideration in any quantitative work. Nucleic acid preparations may contain trace amounts of protein impurities after isolation and extraction from culture, which must be quantitated and/or removed from the samples. The common method of quantitation is to measure the ratio of absorbance values at 260 and 280 nm (A260:A280). A sample that is considered free of protein contaminants has a typical ratio of 1.8 (DNA) to 2.0 (RNA) (8,9). Additionally, a measure of phenol contamination has been established using A260:A270, with a ratio of *1.2 indicating a phenol-free preparation (10). There are problems with this approach, however, as pointed out by Glasel and others (11–13), because the extinction coefficients originally used in determining acceptable ratios have been recalculated in the intervening years, leading to a possible over- or underestimation of purity. Furthermore, as described above, solution conditions such as pH and ionic strength may also affect the absorbance values (9). Nevertheless, when used with care, the A260:A280 ratio is still of some value under certain conditions. For example, ratios of less than 1.7 are indicative of impurities in the preparation, and, as the ratio decreases, the amount of UV-absorbing contaminants (i.e., proteins or phenol) are presumably increasing (13). Another approach used to estimate the degree of purity of nucleic acid preparations is the use of second-derivative spectroscopy, which is described below. In either method, purity estimates should be supplemented by electrophoretic and chromatographic analysis.

2.2.2. Application: Analysis of Thermal Disruption of Polynucleotide Secondary Structure (DNA “Melting”) Double stranded DNA undergoes a highly cooperative helix-to-coil transition at increasing temperatures. These melting curves arise from the large amount of hypochromism present in the absorption spectrum of dsDNA. The absorbance of double-stranded nucleic acids is approx 40% lower in intensity than that predicted from the base composition (3; see Subheading 2.2.1.). This hypochromic effect results from the presence of nondegenerate interactions

258

Braun, Kueltzo, and Middaugh

between the bases, leading to a loss in intensity. Degenerate interactions between the bases are also present, but these result in a splitting and redistribution of intensities, with no net intensity loss. As the strands begin to unwind and separate, the interactions between the bases decrease, increasing UV absorption to a point at which the helix is fully unwound. At high temperatures when the strands behave like random coils, however, there are still some interactions between the bases, and as a result the maximum absorbance reached is still only ~88% of the weighted average of the monomers (1). From a plot of A260 (or similar wavelengths) vs temperature, the midpoint of the melting transition can be determined and used to define the melting temperature (T m). Derivative curves of the thermal transitions (plots of dnA/dTn vs T) often exhibit fine structure indicative of a stepwise melting process (14). Differences in the stability of regions of the helix, caused by base pair composition or chemical modification of the bases (e.g., methylation or crosslinking) are reflected in the fine structure. Changes in the derivative profile may also provide information regarding the origin of these more complex structural transitions. The wavelength dependence of the individual derivative transitions, as well as the midpoint of the general melting curve, can be used to obtain information about the base composition of the regions being perturbed, or of the polynucleotide as a whole. By following the melting of DNA simultaneously at 260 and 282 nm, the fractional GC content of the melting region, and of the entire molecule, can be obtained, since the change in absorbance at 282 nm for a G-C bp greatly exceeds that of an A-T bp; the opposite is observed at 260 nm (15). Additionally, a linear correlation between base composition and Tm has been established (3,15–17). The deconvolution approach may also be employed to determine the effects of specific base modifications and/or solvent perturbation on the temperature stability of nucleic acids (18). DNA tertiary structural features, such as triplexes, hairpins, and pseudoknots, as well as properties of DNA–drug complexes (17), are all potentially detectable by absorbance-monitored melting (19). The formation of cruciform structures from inverted repeat sequences has been detected through the comparison of predicted and experimental Tm values (20). Transitions that are not accompanied by hyperchromism, as shown by Davis et al. (19), may be undetectable by derivative UV spectroscopy. CD, however, may be able to detect such transitions, and is often used as a complementary technique (see Subheading 3. for a detailed discussion). Changes in the thermal melting curve of the DNA, especially the finer transitions observed in the derivative, may provide useful information for the structural characterization of polymer:DNA complexes used in gene therapy. Melting curves have been used extensively in the evaluation of drug-nucleic acid interactions, enabling determination of 6H°, 6S°, 6G°, as well as association constants and stoichiometries of the binding reactions (14,17).

UV Absorption and CD Spectroscopy

259

The same technique has been applied to the interaction of cationic polymers with DNA. For example, the interaction of linear DNA with polylysine (PL) has been extensively characterized by this approach (21,22). More recently, Katayose and Kataoka were able to distinguish between free linear DNA and DNA complexed to a polyethylene glycol-poly(L)lysine block copolymer using the biphasic character of the unfolding transition (23). Unfortunately, the high intrinsic melting point of supercoiled plasmid DNA (Tm > 90°C) may limit the application of such analyses, since many instruments are unable to collect absorbance spectra over the entire Tm range, making the analysis incomplete or even impossible. However, open-circular and linear DNA melt at lower temperatures and are readily analyzed in complexed form (see the discussion of differential scanning calorimetry in Chapter 21 by Lobo et al.). Additionally, even though the Tm of the helix–coil transition is linearly dependent on the log of salt concentration, at high salt concentrations, the Tm actually begins to decrease (17). Thus, the Tm of supercoiled DNA can be lowered with high salt concentration or by the addition of chaotropic agents, such as NaClO4 (7.2 M), to permit thermal transitions to be detected (14,24). One must take into consideration, however, the effect of changes in ionic strength on the interaction of the DNA with the polymer when employing such methods. Changes from high to low (90% supercoiled and assume that any observed properties are primarily those of the biologically active supercoiled form. Determination of molar ellipticity (i.e., normalized or intrinsic CD spectra) requires an accurate measurement of nucleic acid concentration. The absorption of nucleotides is strongly dependent on the pH and ionic strength of the buffer (7,9), as described in Subheading 2.2.1.; therefore, one needs to carefully calibrate concentration measurements if CD spectra of nucleic acids are to be quantitatively comparable. Furthermore, one needs to be careful that spectral changes, produced directly by altering the solvent environment are not confused with structural changes. In the CD experiments described here, the solution concentrations of the nucleic acid solutions were determined by their absorbance at 260 nm (A260) and calibration curves were employed as needed. Investigators must know the molarity of nucleic acid solutions to prepare complexes at specified charge ratios, an important parameter in gene delivery

UV Absorption and CD Spectroscopy

273

Table 1 Molecular Masses of Nucleic Acids Deoxynucleotide base 324.5 g/mol Ribonucleotide base 340.5 g/mol 1 kb dsDNA (Na salt) 6.5 × 105 g/mol 1 kb ssDNA (Na salt) 3.3 × 105 g/mol 1 kb ssRNA (Na salt) 3.4 × 105 g/mol h DNA (48,502 bp) 3.1 × 107 g/mol qX-174 DNA (5386 b) 3.6 × 106 g/mol E. coli DNA (4 × 106 bp) 3.1 × 109 g/mol

efficiency. Furthermore, to aid comparison among CD spectra the data are usually reported normalized as molar ellipticity. Therefore, estimated molecular weights of various commonly used nucleic acids are summarized in Table 1. These values are approximations based on the average weight of the four nucleotides in either RNA or DNA. A wide variety of cationic polymers and lipids are currently being used as vehicles for nonviral gene delivery. Some general classes of materials in use include peptide-based polymers (e.g., PL), peptoids (N-substituted polyglycines), branched-chain polymers (e.g., polyethylenimine and polyamidoamine dendrimers), and cationic lipids. These materials complex with DNA primarily by electrostatic interactions of positively charged functional groups with the negatively charged phosphates of the DNA backbone. Most of the materials used to enhance the delivery of DNA for gene therapy applications are not optically active, and therefore allow observation of the DNA CD without interfering spectral contributions. An important exception to this is peptide-and protein-based polymers. The bulk of their stronger CD contribution, however, occurs below 250 nm so that changes in the 275-nm band can often be detected with minimal interference. Although polypeptide aromatic side chains have CD peaks in this region, their intensity is usually low. If the CD spectra of these materials are of low intensity compared to the nucleic acids, their spectra can be subtracted from the complex spectra with standard software analysis tools, although this involves the tenuous assumption that their CD spectra are not altered during the complexation process.

3.3.2. Methods: DNA Complexes One of the most effective ways to use CD to investigate gene delivery complexes is comparison of CD spectra of complexes of different polymer compositions. This method is most simply performed as a titration of the gene delivery agent into a known amount of DNA. This results in a series of spectra illustrating the state of DNA in a known native secondary structure (e.g.,

274

Braun, Kueltzo, and Middaugh

solution B form) and spectra representative of DNA within the gene delivery complex at various ratios of cationic polymers to DNA. Most nonviral gene delivery vehicles are heterogeneous, noncovalent complexes between DNA and the cationic delivery agent. Therefore, consistency in the method used to form the complexes is critical for reproducibility of spectra. The following preparation procedure for cationic lipid–DNA complexes demonstrates some of the important parameters, but these will vary, depending on the polymer employed. Cationic lipids are effective delivery agents that have already been employed to deliver genes in a number of human clinical trials. CD studies of two cationic lipids, dimethyldioctadecylammonium bromide (DDAB) and dioleoyl-1,2-diacyl3-trimethylammonium propane (DOTAP), are presented here as examples. Cationic lipids interact at least partially with DNA through an electrostatic interaction between their positively charged headgroups and the negatively charged phosphate backbone of the DNA. Complexes are typically prepared at charge ratios (+/–) of 0.25–2. These charge ratios are calculated by assuming a single positive charge for each cationic lipid. Lipids were prepared as liposomes in 10 mM Tris-HCl buffer at pH 7.4. The lipid, solubilized in chloroform, was added to a glass vial, and dried with nitrogen to a thin film. At this point, weights were recorded, to provide a measure of the amount of lipid for future calculations. The lipids were then dispersed in buffer and extruded 10× through a 0.22-µm filter. Temperatures were maintained above the glass transition of the liposome. Supercoiled (>95%) DNA was prepared by diluting a stock solution to the required concentration, typically 10–100 µg/mL. An optimal signal:noise ratio can be achieved at 20–100 µg/mL in a 1-mm pathlength cell. The DNA solution is assayed by absorbance at 260 nm after dilution, and the experimental solution concentration calculated from the result. The DNA concentration is usually calculated in terms of the molar concentration of deoxynucleotide bases, employing an average molecular weight of 324.5 g/ mol (see Table 1). Complexes are individually prepared by adding equal volumes of liposome and DNA solution at the appropriate concentrations. The lower concentration solution is always added to the larger, to avoid passing through charge neutrality, which can result in precipitation of the complexes. At some charge ratios, the complexes are physically unstable, and are therefore prepared immediately prior to spectroscopic analysis.

3.3.3. Results The formation of lipoplexes, consisting of the cationic lipid DDAB, and a DNA plasmid, results in significant changes in the native CD spectra of DNA. As Fig. 10 illustrates, with increasing charge ratio, a peak intensity shift occurs in the 210, 220, 245, and 275 nm bands. In addition, shifts in peak position also occur. Up to a 0.75 charge ratio (+/–), the DNA manifests a continuous change in the 275-nm CD

UV Absorption and CD Spectroscopy

275

Fig. 10. Plasmid DNA (9.1 kb) complexed to increasing amounts of DDAB liposomes in 10 mM Tris buffer at pH 7.4 and at charge ratios of 0, 0.25, 0.50, 0.75, and 1.0.

peak with the magnitude decreasing as the charge ratio increases. The shifts occurring in the 210, 220, and 245 nm bands, however, do not occur directly in parallel with the 275-nm peak changes, although they do seem to follow a trend. With increasing charge ratio, the intensity decreases in the 245- and 210-nm troughs, and increases in the 220-nm peak. The exception is the sample that precipitated at a charge ratio of 1. In this case, the 245- and 275-nm peaks follow the established trends, but the 210- and 220-nm peaks decrease to near zero. Commonly, the spectra of precipitated materials dramatically decrease in intensity. That sample material is grossly aggregated or precipitated may not, however, be immediately apparent from the shape of the observed spectra. Another important observation is appearance of the red shift that occurs in the 245 and 275 nm bands, but is absent in the 210- and 220-nm peaks. As discussed previously, a characteristic feature of absorption flattening is a red shift. Experiments described in Subheading 3.5., however, suggest that absorption flattening does not significantly contribute to these spectra. The addition of neutral helper lipids often provides a significant enhancement in gene transfer efficiency of cationic lipids (47). Figure 11 illustrates the CD spectrum of lipoplexes consisting of DOTAP present in an equal molar ratio with the helper lipid L-_-dioleoyl phosphatidylethanolamine (DOPE). Note that the actual concentration of lipid is twice the previous example at any

276

Braun, Kueltzo, and Middaugh

Fig. 11. Plasmid DNA (9.1 kb) complexed to increasing amounts of DOTAP:DOPE liposomes in 10 mM Tris buffer at pH 7.4 and at charge ratios (+/–) of 0, 0.25, 0.50. and 0.75.

particular charge ratio. The spectra are similar to the previous lipoplexes with conservation of the native DNA bands at 210, 220, 245, and 275 nm. The spectral changes in the 275-nm peak are similar to those described in the DDAB lipoplexes with a decrease in magnitude correlating with increasing charge ratio. In fact, this feature seems conserved in the CD spectra of lipoplexes containing various combinations of cationic and helper lipids. A distinguishing feature, in contrast to the DDAB lipoplexes, is the 245-nm trough, which increases in magnitude with increasing charge ratio. This characteristic can be assigned primarily to the influence of the helper lipid, DOPE. The remaining bands, at 210 and 220 nm, manifest increasing and decreasing magnitudes, respectively, as described in the previous example. The utility of CD in such studies is evident from the finding that lipoplexes of similar charge ratios, but differing in lipid composition are both similar (i.e., 275-nm peak features) and different (i.e., 245 nm trough features) in their CD spectral features.

3.4. Applications: DNA–Cationic Polymer Complexes The other major class of nonviral gene delivery agents is the cationic polymers. This class contains a wide variety of compounds, but PEI and polylysine have been frequently used, and are discussed as representatives of this class. Polyethylenimine is a linear or branched-chain polymer containing both primary and secondary amines. The molecular weight may vary considerably, with nonviral vehicles being produced from PEI of 2–800 kDa, although a mol

UV Absorption and CD Spectroscopy

277

wt )2 kDa requires formulation with an endosomolytic agent, to ensure efficacy (48). The higher-mol-wt forms of PEI have been found to be more efficient delivery vehicles, but this highly branched polymer has been shown to have some cytotoxicity (49). In contrast, low-mol-wt PEI (e.g., ~12 kDa) has significantly diminished cytotoxicity. The generally accepted charge ratios used for most cationic nonviral delivery agents are less applicable to PEI because of its multiple pKa values, which are altered by nucleic acid binding (48). Complexes are therefore formed based on a PEI nitrogen:DNA phosphate ratio (N:P). Figure 12 illustrates the spectra of small (2 kDa) linear PEI complexed to plasmid DNA at various N:P ratios. Even at the lowest N:P ratio examined (0.5), the 275-nm native DNA peak decreases and red shifts (~280 nm), producing strong negative ellipticity at ~260 nm. This new trough reaches a maximum at 2.0 N:P ratio. At an N:P ratio of 6.7, the 260-nm trough is reversed, decreasing in magnitude, with a corresponding increase in the 285–290 nm peak. The CD spectrum of the 10- and 20-N:P ratio complexes overlay one another, implying the presence of a stable form. The large peak shifts and changes in intensity are more characteristic of psi-type CD spectra than those seen in cationic lipid complexes, although not as dramatic as those that arise in polymer- and salt-condensed DNA (29,44). The polylysines are another frequently used class of cationic polymer. These linear peptides have been extensively studied both as simple models of DNA:protein interactions and for their DNA condensing properties. The CD spectra of polylysine–DNA complexes (Fig. 13) show a dramatic effect on the bands of uncomplexed plasmid with addition of polylysine. The 275-nm peak decreases in magnitude, which seems to be a common observation as the molar ratio of various cationic delivery agents’ increase. The 258-nm crossover point of B-form DNA, however, red shifts, as seen in B A s transitions. The 210- and 220-nm bands both decrease in their negative ellipticity values, with increasing charge ratio. Weiskoff et al. (50) examined the CD of polylysine–DNA complexes by various preparation methods. At high salt concentration (1 M NaCl) and a 0.72 charge ratio, the complexes exhibited a psi-type CD spectra. At this charge ratio, the characteristic 275-nm peak of B form is lost, and a large negative psi-type trough is produced at 270 nm. At lower salt concentrations ()0.5 M NaCl), however, the polylysine–DNA complexes had more moderate spectral changes. At low salt, the major spectral feature was a decrease in the 275-nm peak similar to that seen here.

3.5. Discussion The nonviral gene delivery complexes discussed are generally large inhomogeneous structures. Therefore, at this time, it is difficult to correlate spectral changes to specific alterations in supramolecular structure. However, the following discussion may provide some insight into how the formation of

278

Braun, Kueltzo, and Middaugh

Fig. 12. Plasmid DNA (9.1 kb) complexed to PEI at N:P ratios of 0, 0.5, 1, 2, 6.7, 10, and 20.

Fig. 13. CD spectra of polylysine/DNA complexes in 2 mM sodium phosphate buffer at pH 7.4 and at charge ratios (+/–) of 0, 0.2, 0.4, 0.6, and 0.8.

supramolecular structures influences CD spectra. When DNA complexes to cationic polymers, aggregates in the size range of 50–200 nm are typically formed (see Chapter 22 by Wiethoff and Middaugh). The CD of DNA present

UV Absorption and CD Spectroscopy

279

in such structures will generally differ significantly from that of free polynucleotides. The fact that such spectra are significantly distorted has been known for some time, as the result of studies of entities such as viruses and chromosomes. In general, one finds that as the size of such complexes increases, tails begin to appear in regions outside absorption bands. Larger aggregates also often display CD bands with dramatically altered intensities (usually larger) and distorted peak shapes. As indicated previously, the latter are often referred to as psi-type effects. The origin of the tails is now well accepted to be caused by differential scattering of either right or left circularly polarized light. This phenomenon, however, is generally thought to contribute only a limited amount to the abnormal spectra seen in DNA–polymer complexes. Another possible origin of such spectral alterations could be Duysens’ flattening, which arises from a screening phenomenon among the dispersed particles. The authors have examined the various complexes discussed above over a range of concentrations and pathlengths. If absorption flattening is responsible for the spectral changes seen upon complex formation, variations in spectra should be observed as these parameters are varied, but this is not seen. The authors therefore conclude that a statistically uneven distribution of complexes does not play a major role in the CD spectra induced by polymer binding. Another possibility is that the polymers produce actual structural changes in the structure of the DNA itself (e.g., a B A C transition) (44). In general, however, the newly induced CD spectra cannot be clearly identified as any known structural form. Furthermore, X-ray diffraction studies of DNA complexes; which display typical distorted spectra, are seen to be present in the normal B form (50). Currently, the most attractive hypothesis is that large-scale chiral elements in the structure of the complex interact differently with differentially handed, circularly polarized light. These elements, or liquid crystalline phases, have been reported in both condensed DNA and cationic lipid–DNA complexes (47,51). The ordered packing is generally of two types, with nematic and hexagonal phases aligned along the long axis of neighboring molecules; second-order cholesteric phases align similarly, with the addition of a twist or helix perpendicular to the first-order alignment (52). Supramolecular helices of cholesteric phases propagate one circular polarization of light in preference to the other, in a wavelength-dependent fashion (52). The basic idea is that, when the size of any DNA-containing particle approaches that of the incident light, the overall chirality of the complex will have a significant effect in either increasing or reducing the CD signals. That chiral order within the complex can be responsible for these spectral perturbations has been directly demonstrated in a simple model system (41). When a highly concentrated solution of DNA is pressed between two quartz plates and rotated, a psi–like spectrum of

280

Braun, Kueltzo, and Middaugh

both intensity-enhanced and shape-distorted peaks is produced. If the rotation is reversed, the sign of the psi–like peak is reversed with little alteration in peak shape. If the film is twisted back and forth numerous times to randomize the large-scale chiral order, essentially natural spectra result. Thus, it is clear that in at least some cases, large-scale chiral order can produce major changes in CD spectra of DNA. These effects appear to primarily arise from electrostatic coupling between chromophores. Such long-range coupling appears to be possible when the complex is large (>h/4), has at least a certain density of chromophores (>1/nm3), and has a defined three-dimensional structure (29). The critical chromophoric interactions usually seem to be the result of a combination of static dipole, intermediate, and radiation coupling. As a consequence, structural factors, such as the size, pitch, handedness, density of chromphores, and internal order of the aggregate, all control the sign, shape, and magnitude of the altered CD spectra seen in DNA–polymer complexes. However, most cases in the literature find major increases in the intensity of the CD spectra induced by polymers. In systems containing agents as diverse as various cationic homopolypeptides (e.g., PL, polyarginine, polyhistidine), spermine, histones, polyethylene glycols, as well as high concentrations of hydroxylcontaining organic solvents and certain salts, psi-like spectra are seen in which the DNA CD peaks are dramatically enhanced. This, in fact, is not what is seen when gene delivery polymers are complexed to supercoiled DNA plasmids over the range of charge ratios used to prepare active delivery complexes. One possible explanation may be that the long-range chirality of the complexes formed is opposite to that usually observed, resulting in the observed decreases in spectral intensity. Another possibility is that the gene delivery complexes lack the high degree of chiral order seen in the other systems, thus permitting more subtle effects to dominate. For example, X-ray studies indicate that cationic lipid–linear DNA complexes exist as alternating layers of lipid bilayers and DNA (47,51). These supercoiled plasmids could in effect be stretched out (i.e., pseudo-linearized) by lipid binding, thus lowering electrostatic coupling, and thereby reducing dichroic effects. The observation that the unwinding of negatively supercoiled DNA produces decreases in the intensity of the 277-nm CD peak is consistent with this hypothesis (36). Furthermore, at the secondary structure level, in a series of papers by Chen and Hanlon (53–55), it was shown that a subtle increase in winding angle results in substantial decrease in the rotational strength above 260 nm. Clearly, DNA CD spectra are sensitive to subtle differences in the structure of the DNA component at both the tertiary and secondary level. However, the spectra observed can be considered to sensitively reflect the overall structural organization of the complex and thus provide a valuable measure of structural integrity.

UV Absorption and CD Spectroscopy

281

4. Conclusion Of the many techniques currently employed in the analysis of nucleic acids and their complexes, UV-absorption spectroscopy is rarely the first choice, except in the case of concentration determination. As argued here, however, there appear to be specific applications of the technique that can provide structural information with a few fast and simple experiments. Current instrumentation is inexpensive, user-friendly, and, most important, already present in most laboratories. It seems probable that this ready availability of equipment, accompanied by the considerations outlined above, will lead to an increased use of the technique, especially in the context of derivative analysis of gene delivery complexes. CD is also a sensitive technique for investigating nonviral gene delivery systems. CD spectral differences provide a method for differentiating between complexes constructed with different gene delivery agents. Although interpretation of the structural basis of spectral differences in these complex constructs is elusive, the spectral features are reproducible. The fact that CD seems to provide some insight into the supramolecular organization of these components is actually beneficial, since recent work suggests structural features are important in the delivery process. Thus, this technique is a potentially valuable tool in the characterization of gene delivery formulations and could be used to provide a measure of quality control. An intriguing possibility is that correlations between the CD of gene delivery complexes and their efficiency of transfection may be present, which could aid in the construction of more efficacious vehicles. References 1. van Holde, K. E., Johnson, W. C., and Ho, P. S. (1998) Principles of Physical Biochemistry. Prentice-Hall, Upper Saddle River, New Jersey. 2. Owen, T. (1996) Fundamentals of Modern UV-vVisible Spectroscopy: A Primer. Hewlett Packard Co., Germany. 3. Bloomfield, V. A., Crothers, D. M., and Ignacio Tinoco, J. (1974) Physical Chemistry of Nucleic Acids. Harper & Row, New York. 4. Fasman, G. D., ed. (1975) Handbook of Biochemistry and Molecular Biology. CRC, Cleveland, OH. 5. Cantor, C. R. and Schimmel, P. R. (1980) Biophysical Chemistry Part II: Techniques for the Study of Biological Structure and Function. W. H. Freeman, San Francisco, CA. 6. Beavan, G. H., Holiday, E. R., and Johnson, E. A. (1955) Optical properties of nucleic acids and their components, in The Nucleic Acids (Chargaff, E. and Davidson, J. N., eds.), Academic, New York, pp. 493–553. 7. McGown, E. L. (2000) UV Absorbance measurements of DNA in microplates. Biotechniques 28, 60–64. 8. Baumann, C. G. and Bloomfield, V. A. (1995) Large-scale purification of plasmid dna for biophysical and molecular biology studies. Biotechniques 19, 884–890.

282

Braun, Kueltzo, and Middaugh

9. Wilfinger, W. W. (1997) Effect of pH and ionic strength on the spectrophotometric assessment of nucleic acid purity. Biotechniques 22, 474–481. 10. Stulnig, T. M. and Amberger, A. (1994) Exposing contaminating phenol in nucleic acid preparations. Biotechniques 16, 403–404. 11. Glasel, J. A. (1995) Validity of Nucleic acid purities monitored by 260nm/280nm absorbance ratios. Biotechniques 18, 62–63. 12. Huberman, J. A. (1995) Importance of measuring nucleic acid absorbance at 240 nm as well as at 260 and 280 nm. Biotechniques 18, 636. 13. Manchester, K. L. (1995) Value of A260/A280 ratios for measurement of purity of nucleic acids. Biotechniques 19, 208–210. 14. Wada, A., Yabuki, S., and Husimi, Y. (1980) Fine structure in the thermal denaturation of DNA: high temperature-resolution spectrophotometric studies. Crit. Rev. Biochem. 9, 87–144. 15. Blake, R. D. and Hydorn, T. G. (1985) Spectral analysis for base composition of DNA undergoing melting. J. Biochem. Biophys. Meth. 11, 307–316. 16. Felsenfeld, G. (1971) Analysis of temperature-dependant absorption spectra of nucleic acids, in Procedures in Nucleic Acid Research (Cantoni, G. L. and Davies, D. R., eds.), Harper & Row, New York, pp. 233–261. 17. Wilson, W. D., Tanious, F. A., Fernandez-Saiz, M., and Rigl, C. T. (1997) Evaluation of drug-nucleic acid interactions by thermal melting curves, in Methods in Molecular Biology, vol. 90: Drug-DNA Interaction Protocols (Fox, K. R., ed.), Humana, Totowa, NJ, pp. 219–240. 18. Blackburn, G. M. and Gait, M. J., eds. (1996) Nucleic Acids in Chemistry and Biology. Oxford University Press, Oxford. 19. Davis, T. M., McFail-Isom, L., Keane, E., and Williams, L. D. (1998) Melting of a DNA hairpin without hyperchromism. Biochemistry 37, 6975–6978. 20. McCampbell, C. R., Wartell, R. M., and Plaskon, R. R. (1989) Inverted repeat sequences can influence the melting transitions of linear DNAs. Biopolymers 28, 1745–1758. 21. Carroll, D. (1972) Complexes of polylysine with polyuridylic acid and other polynucleotides. Biochemistry 11, 426–433. 22. Mandel, R. and Fasman, G. D. (1976) Chromatin models. interaction between dna and polypeptides containing L-lysine and L-valine: circular dichroism and thermal denaturation studies. Biochemistry 15, 3122–3130. 23. Katayose, S. and Kataoka, K. (1997) Water-soluble polyion complex associates of dna and poly(ethylene glycol)-poly(L-lysine) block copolymer. Bioconj. Chem. 8, 702–707. 24. Thumm, W., Seidl, A., and Hinz, H.-J. (1988) Energy–structure correlations of plasmid DNA in different topological forms. Nuc. Acids Res. 16, 11,737–11,757. 25. Mach, H., Sanyal, G., Volkin, D. B., and Middaugh, C. R. (1997) Applications of ultraviolet absorption spectroscopy to the analysis of biopharmaceuticals, in ACS Symposium Series. American Chemical Society, pp. 186–205. 26. Mach, H., Middaugh, C. R., and Lewis, R. V. (1992) Detection of proteins and phenol in dna samples with second-derivative absorption spectroscopy. Anal. Biochem. 200, 20–26.

UV Absorption and CD Spectroscopy

283

27. Kim, M.-H., Ulibarri, L., Keller, D., Maestre, M. F., and Bustamante, C. (1986) Psi-type circular dichroism of large molecular aggregates. III. Calculations. J. Chem. Phys. 84, 2981–2989. 28. Keller, D. and Bustamante, C. (1986) Theory of the interaction of light with large inhomogeneous molecular aggregates. I. Absorption. J. Chem. Phys. 84, 2961–2971. 29. Keller, D. and Bustamante, C. (1986) Theory of the interaction of light with large inhomogeneous molecular aggregates. II. Psi-type Circular Dichroism. J. Chem. Phys. 84, 2972–2980. 30. Neidle, S., ed. (1999) Oxford Handbook of Nucleic Acid Structure. Oxford University Press, New York. 31. Ivanov, V. I., Minchenkova, L. E., Schyolkina, A. K., and Poletayev, A. I. (1973) Different conformations of double-stranded nucleic acid in solution as revealed by circular dichroism. Biopolymers 12, 89–110. 32. Middaugh, C. R., Evans, R. K., Montgomery, D. L., and Casimiro, D. R. (1998) Analysis of plasmid DNA from a pharmaceutical perspective. J. Pharm. Sci. 87, 130–146. 33. Mathews, C. K. and van Holde, K. E. (1996) Biochemistry. Benjamin/Cummings, Menlo Park, California. 34. Johnson, W. C. (1994) CD of nucleic acids, in Circular Dichroism: Principles and Applications (Nakanishi, K., Berova, N. and Woody, R. W., eds.), VCH, New York, pp. 523–540. 35. Johnson, W. C. (1996) Determination of the conformation of nucleic acids by electronic CD, in Circular Dichroism and the Conformational Analysis of Biomolecules (Fasman, G. D., ed.) Plenum, New York, pp. 433–468. 36. MacDermott, A. J. and Drake, A. F. (1986) circular dichroism of postively and negatively supercoiled DNA. Stud. Biophys. 115, 59–67. 37. Glaeser, R. M. and Jap, B. K. (1985) Absorption Flattening in the circular dichroism spectra of small membrane fragments. Biochemistry 24, 6398–6401. 38. Mao, D. and Wallace, B. A. (1984) Differential light scattering and absorption flattening optical effects are minimal in the circular dichroism spectra of small unilamellar vesicles. Biochemistry 23, 2667–2673. 39. Wallace, B. A. and Teeters, C. L. (1987) Differential absorption flattening optical effects are significant in the circular dichroism spectra of large membrane fragments. Biochemistry 26, 65–70. 40. Bustamante, C., Tinoco, I., Jr., and Maestre, M. F. (1983) Circular differential scattering can be an important part of the circular dichroism of macromolecules. Proc. Natl. Acad. Sci. USA 80, 3568–3572. 41. Maestre, M. F. and Reich, C. (1980) Contribution of light scattering to the circular dichroism of deoxyribonucleic acid films, deoxyribonucleic acid-polylysine complexes, and deoxyribonucleic acid particles in ethanolic buffers. Biochemistry 19, 5214–5223. 42. Phillips, C. L., Mickols, W. E., Maestre, M. F., and Tinoco, I., Jr. (1986) Circular differential scattering and circular differential absorption of DNA-protein condensates and of dyes bound to DNA-protein condensates. Biochemistry 25, 7803–7811.

284

Braun, Kueltzo, and Middaugh

43. Reich, C., Maestre, M. F., Edmondson, S., and Gray, D. M. (1980) Circular dichroism and fluorescence-detected circular dichroism of deoxyribonucleic acid and poly[d(A-C)•d(G-T)] in ethanolic solutions: a new method for estimating circular intensity differential scattering. Biochemistry 79, 5208–5213. 44. Tinoco, I., Bustamante, C., and Maestre, M. (1980) Optical activity of nucleic acids and their aggregates. Ann. Rev. Biophys. Bioeng. 9, 107–141. 45. Tinoco, I. J. and Mickols, W. (1987) Absorption, scattering, and imaging of biomolecular structures with polarized light. Ann. Rev. Biophys. Biophys. Chem. 16, 319–349. 46. Ausubel, F. M., et al., eds. (1989) Current Protocols in Molecular Biology. Wiley, New York. 47. Koltover, I., Salditt, T., and Safinya, C. R. (1999) Phase diagram, stability, and overcharging of lamellar cationic lipid-DNA self-assembled complexes. Biophys. J. 77, 91–924. 48. Huang, L., Hung, M.-C., and Wagner, E. (1999) Non-Viral Vectors for Gene Therapy. Academic, San Diego, CA. 49. Fischer, D., Bieber, T., Li, Y., Elsasser, H.-P., and Kissel, T. (1999) Novel Nonviral vector for DNA delivery based on low molecular weight, branched polyethylenimine: effect of molecular weight on transfection efficiency and cytotoxicity. Pharm. Res. 16, 1273–1279. 50. Weiskoff, M. and Jei Li, H. (1977) Poly (L-lysine) -DNA interactions in NaCl solutions: B - C and B - psi transitions. Biopolymers 16, 669–684. 51. Koltover, I., Salditt, T., Rädler, J. O., and Safinya, C. R. (1998) An inverted hexagonal phase of cationic liposome-DNA complexes related to DNA release and delivery. Science 281, 78–81. 52. Gottarelli, G. and Spada, G. P. (1994) Application of CD to the study of some cholesteric mesophases, in Circular Dichroism: Principles and Applications (Nakanishi, K., Berova, N., and Woody, R. W., eds.), VCH, New York, pp. 105–119. 53. Chen, C., Kilkuskie, R., and Hanlon, S. (1981) Circular dichroism spectral properties of covalent complexes of deoxyribonucleic acid and n-butylamine. Biochemistry 17, 4987–4995. 54. Chen, C., Pheiffer, B. H., Zimmerman, S. B., and Hanlon, S. (1983) Conformational characteristics of deoxyribonucleic acid-butylamine complexes with C-type circular dichroism spectra 1. An X-ray fiber diffraction study. Biochemistry 22, 4746–4751. 55. Fish, S. R., Chen, C., Thomas, G. J., Jr., and Hanlon, S. (1983) Conformational characteristics of deoxyribonucleic acid-butylamine complexes with C-type circular dichroism spectra 2. A Roman spectroscopic study. Biochemistry 22, 4751–4756.

Synthetic Vectors by IR Spectroscopy

285

20 Characterization of Synthetic Gene Delivery Vectors by Infrared Spectroscopy Sirirat Choosakoonkriang, Christopher M. Wiethoff, Lisa A. Kueltzo, and C. Russell Middaugh 1. Introduction For many decades, infrared (IR) spectroscopy has been used to characterize the structure of molecules. In IR spectroscopy, absorption of light, corresponding to vibrational and rotational transitions of a molecule, is measured. For a transition to be IR-active, a change in the dipole moment of a particular bond must occur upon excitation. This vibrational energy is not only dependent on the chemical nature of the particular covalent bonds, but also on the environment of these coupled atoms and bonds. IR spectroscopy has been previously employed in the study of the structure of nucleic acids, producing not only information about the individual bases, sugars, and phosphate backbone, but also providing information about the helical conformation of polynucleotides (1–3). IR spectroscopy has also been successfully applied to the analysis of lipids, as well as to numerous other polymers (4). Thus, IR spectroscopy potentially possesses the ability to obtain structural information about all of the components of most synthetic gene delivery complexes, as well as changes in the structure of polymeric or lipid components upon complex formation. In addition to the ability to gather detailed structural information, there are also some practical advantages to the use of IR spectroscopy for the study of plasmid DNA and DNA complexes compared to other techniques, including the availability of a variety of sampling techniques, permitting the analysis of samples in a wide variety of physical states including solutions, solids, and gels. There is also no upper limit to the size of the sample molecule examined, allowing both short oligonucleotides and higher molecular weight DNA to be From: Methods in Molecular Medicine, vol. 65: Nonviral Vectors for Gene Therapy Edited by: M. A. Findeis © Humana Press Inc., Totowa, NJ

285

286

Choosakoonkriang et al.

studied. IR spectroscopy is not a destructive technique, and requires only small amounts of material, making it ideal for the analysis of valuable samples. Here are reviewed IR instrumentation, common sampling geometries in relation to the physical states of samples, and different methods of data analysis. Examples of the use of Fourier-transformed infrared (FTIR) to characterize the interaction of DNA with several polycations used in gene delivery including cationic lipids and polymers, are presented and discussed. 2. Representative IR Vibrational Modes 2.1. Characteristic IR Absorption Bands of Polynucleotides The most useful vibrational bands of DNA are observed between 1800 and 700 cm–1, with approx 35 well-defined absorption bands occurring in this region of the IR spectrum (Fig. 1). Four general aspects of DNA structure are reflected in the vibrational modes in this region. Bands between 1750 and 1500 cm–1 primarily represent vibrations of the bases. Vibrations caused by DNA basesugar entities, heavily dependent on glycosidic torsion angles are observed between 1500 and 1250 cm–1. Strong absorption occurs for the phosphate groups and deoxyribose between 1250 and 1000 cm–1 and the region below 1000 cm–1 primarily contains vibrations of the phosphodiester bond coupled to vibrations of the deoxyribose. Table 1 provides a more complete listing of the IR vibrational modes of DNA. Vibrational modes in the spectral region from 1750 to 1500 cm–1 result from carbonyl stretching and N-H bending, as well as C = C and C = N stretching vibrations of the nucleic acid bases. Each of the four bases, when present as mononucleotides or single-stranded homopolymers, has characteristic bands in this region, and it is common to observe multiple bands in this region that result from the overlap of bands from the individual bases. Unfortunately, this region contains a strong water absorption band, making peak assignments difficult. Water subtraction algorithms have greatly reduced this problem, but studies in this region are often performed in deuterium oxide (D2O), to eliminate the water interference. Therefore, the following assignments in this region are reported for D2O solutions. Assignments in other regions of the polynucleotide spectra discussed below are for samples in water solutions. Guanine has characteristic bands at 1531, 1581, and 1668 cm–1 (5). Similarly, this region contains bands at 1506, 1524, 1619, and 1652 cm–1 (cytosine), 1626 cm–1 (adenine) and 1632, 1663, and 1695 cm–1 (thymine) (5). In many cases, these characteristic vibrations of the nucleic acid bases undergo abrupt changes when involved in intermolecular base-pairing in duplex DNA. Important spectral differences upon base-pairing, as seen for the interaction of complementary homopolymers, include a shift of the carbonyl stretching vibration of guanine from 1668 to 1689 cm–1, accompanied by a strong reduction in intensity of the

Synthetic Vectors by IR Spectroscopy

287

Fig. 1. FTIR absorbance spectra of DNA (plasmid) in H2O solution (top) (top to bottom: 4.0, 3.0, 2.0, 1.0, and 0.5 mg/mL), and on polyethylene cards (bottom) (top to bottom: 50 µL of 5.4, 4.0, 3.0, 2.0, 1.0, and 0.5 mg/mL).

peak at 1581 cm–1 (5). Cytosine shows little change in its carbonyl stretching vibration at 1652 cm–1, but the band at 1524 cm–1 can no longer be observed upon base-pairing. In the case of adenine, the band at 1626 cm–1 is shifted down to 1622 cm–1 (5). The carbonyl bands for thymine (1695 and 1663 cm–1) change only with respect to their relative intensities, while the 1632 cm–1 band shifts to 1641 cm–1.

288

Choosakoonkriang et al.

Table 1 Representative IR Vibrational Modes of DNA Nucleic acid base Adenine Cytosine

Wavenumber (cm–1)a,b 1626 (1622 upon base pairing) 1652 1619 1524 (not observed upon bp) 1506 1668 (1689 upon bp) 1581 1531 1695 1663 1632 (1641 upon bp)

Guanine

Thymine

Conformationally Sensitive Bands (cm–1) Ab

Bb

Cc

Zb

1705 1418 1375 1335

1715 1425 1375 1344 1328

1710 1425 1375 1344 1328

1695 1408 1355

1275

1281

1240 1188

1225

1320

970 (triplet)

970 (singlet)

Assignmentb,c Base carbonyl indicative of base pairing Deoxyribose dGdA dA dT dC dT

1265 1230–1217 1215

Antisymmetric phosphate stretch

1069

Deoxyribose

1065 1013

968

Deoxyribose 929

898 (triplet)

897 (singlet) 840

891 833

806

Deoxyribose Deoxyribose Deoxyribose

a Assignments

are for bases in D2O. are from refs. 2 and 5. cAssignments are from ref. 9. b Assignments

The helical conformation of DNA potentially exists in multiple forms depending on factors such as salt concentration, base sequence and composition, relative humidity, and pH. The most common antiparallel, double-stranded, helical geometries of DNA are the right-handed A- and B-form and the left-handed

Synthetic Vectors by IR Spectroscopy

289

Z-form families. Two important differences in DNA helical geometries allow one to spectroscopically distinguish one form from another. Differences include departures of the signature bands for base-pairing from that of the archetypal Bform geometry and differences in the conformation of the sugars in the helix (e.g., N-type, C3' endo/anti geometry for the A-form and S-type, C2' endo/anti geometry for the B form). The IR spectra of these various forms contain tell-tale bands that permit discrimination between each of the helical geometries. Of the two right-handed geometries, the B form prevails in aqueous solutions at moderate ionic strength and neutral pH. Thus, it is this form of DNA that is thought to most commonly occur in biological systems. The A form occurs primarily under conditions of low water activity (66–47% relative humidity) or in high concentrations of organic solvents, such as 80% EtOH (2,5). Between 1750 and 1500 cm–1, a band around 1715 cm–1, caused by the carbonyl stretching of the bases reflects the intermolecular hydrogen bonding of base pairs for the B form of DNA in H2O (5). This band is shifted down to 1705 cm–1 for the A form. In the spectral region between 1500 and 1250 cm–1, several vibrational modes are characteristic of B-form DNA. A vibration caused by the deoxyribose sugar (C2' endo) occurs at ~1425 cm–1, but is shifted to approx 1418 cm–1 for A form (5). Another characteristic sign of the right-handed geometry is the torsion angle of the glycosidic bond for purine bases. Both A and B forms manifest a band near 1375 cm–1, because of the anticonformation of deoxyguanylate and deoxyadenylate (6). In comparing poly(dA)·poly(dT) (B form) to poly(rA)·poly(dT) (A form), the B-form spectra include a doublet at 1344 and 1328 cm–1 from the adenine and thymine bases, respectively, as well as a single band at 1281 cm–1, arising from the N-3 H-bending vibration of thymine (2). For the A form, the doublet bands are merged into a single peak near 1335 cm–1 and the N-3 H-bending vibration is found at about 1275 cm–1. In the region between 1250 and 1000 cm–1, information regarding the antisymmetric and symmetric phosphate- stretching vibrations is contained. The B form exhibits a strong band at 1225 cm–1 due to the antisymmetric stretching vibration of the phosphate. This band is found ~15 cm–1 higher in A-form DNA near 1240 cm–1. The position of the symmetric-stretching vibration of the phosphate is essentially independent of the helical geometry of DNA and is found at about 1089 cm–1. As mentioned above, the spectral region between 1000 and 700 cm–1 contains information about the vibrations of the phosphate esters coupled to vibrations of the deoxyribose ring. A band at 970 cm–1 is seen for the B form; a poorly resolved triplet centered at this position is observed for A form. The B form also shows two bands at 894 and 840 cm–1; the A form displays a wellresolved triplet at 898, 882, and 864 cm–1 (2,5). The puckering of the ribose ring for the A- and B-form geometries gives rise to bands at 807 cm–1 (N type, C3'

290

Choosakoonkriang et al.

endo/anti) and 840 cm–1 (S type, C2' endo/anti) for the A and B forms, respectively (5). Thus, distinct spectral bands for the A and B forms allow the two major helical geometries of DNA to be unambiguously distinguished by IR spectroscopy. Several variants of B-form DNA have been identified, including the C and D forms. The C form has been detected in the presence of specific counterions, such as Li+ and Cs+ (7), and has been shown to occur with reduced H2O activity (lower relative humidity), while specific sequences (polyd(A-T)·polyd[AT]) have been found to be present in the D form (5). Conflicting reports have indicated that the antisymmetric phosphate stretching vibration of C-form DNA occurs anywhere from 1230 (7) to 1217 cm–1 (8). Additional differences between the C and B form may be observed by a shift in the base carbonyl stretching peak from 1715 cm–1 (B form) to ~1710 cm–1 (C form) (8,9). Perhaps the most consistent difference observed between the IR spectra of B- and C-form DNA is the appearance of a more prominent band around 1069 cm–1 in the C-form spectra, compared to a weak shoulder at 1071 cm–1 seen in B-form spectra (8,9). Initial studies using circular dichroism have suggested the existence of C-DNA, when DNA is complexed with either cationic lipids (10,11) or a cationic polymer, such as polylysine (12). These results are complicated by distortions in the CD spectra of large complexes, making interpretation difficult (see Chapter 19 by Braun et al.). The IR spectrum of the left-handed double helical Z-form DNA contains several distinct differences compared to its right-handed counterparts. In the case of Z-form DNA, the band reflecting the presence of base-pairing seen at 1715 cm–1 for B form is shifted to around 1695 cm–1 (5). The phosphate antisymmetric-stretching vibration is also present at lower frequency, near 1215 cm –1. Additionally, new absorption bands around 1320 cm–1 (cytosine), 1265, 1123, 1067, 1013, and 928 cm–1 are indicative of the Z form (13). Although no information regarding IR vibrational frequencies have been reported for ^-form DNA, this highly ordered state has been proposed to exist for several polycation–DNA complexes (11,14). The many conformationally sensitive bands of DNA provide an excellent basis with which to examine complex formation in gene delivery vehicles. In particular, analysis of the shifts in position of these peaks, as polycations bind to DNA, should permit the structure of complexed DNA to be probed.

2.2. Major IR Bands of Cationic Lipids and Select Polycations Cationic lipids have been extensively used in nonviral gene therapy as vectors for the delivery of DNA to cells because of their low toxicity, reduced immunogenicity, and established efficacy, providing a simple system for DNA delivery. Cationic lipid systems have been shown to produce significant levels of gene expression in tissues when administered intravenously (15), with

Synthetic Vectors by IR Spectroscopy

291

Fig. 2. FTIR absorbance spectra for 1,2-dioleoyl-3-trimethylammonium-propane (DOTAP) (top, 1.2 mg/mL) and dimethyldioctadecylammonium bromide (DDAB) (bottom, 2.5 mg/mL) liposomes in solution.

activity demonstrated in multiple tissues (16–22), increasing the attractiveness of the system. A variety of such vectors is currently in human clinical trials. Based on the potential of these systems, the authors concentrate here on the IR characterization of examples of cationic lipid-DNA complexes, although, as is shown, extension to other polymeric-based delivery systems is straightforward. Cationic lipid molecules usually have several IR-active groups (Fig. 2). The main vibrational modes observed in lipids originate from the molecular vibrations of the acyl or alkyl chains and the head groups. There are two major

292

Choosakoonkriang et al.

types of acyl chain vibrational modes, the C–H vibration characteristic of the hydrophobic hydrocarbon region, and the C–O vibration characteristic of the glycerol–acyl-chain interface. These bands are easily identified (Table 2). C–H stretching bands appear in the region of 3200–2700 cm–1, primarily because of the antisymmetric (2925 cm –1 ) and symmetric (2852 cm–1 ) vibrations of the methylene groups (23). Bending vibrations are observed for methyl and methylene groups, including the symmetric CH3 deformation (umbrella-type) vibration at 1375 cm–1, a CH2 scissoring band between 1474 and 1468 cm–1, a series of CH2 wagging bands between 1190 and 1345 cm–1, and CH2 twisting/rocking peaks in the region of 720–1150 cm–1 (23). The ester carbonyl-stretching mode has a strong absorption band that appears at 1700–1750 cm–1. The position of this band is sensitive to the geometry of the packing of the acyl chains and glycerol moiety, as well as the hydration state of the lipid headgroup (23). For the cationic lipid, 1,2-dioleoyl-3-trimethylammoniumpropane (DOTAP), the ester C = O stretch appears ~1739 cm–1 in solution (24). The headgroups of membrane lipids also produce a number of characteristic IR bands. In phospholipids, for example, the antisymmetric PO2– stretching vibrations result in a strong IR band in the range of 1220–1240 cm–1, with a symmetric stretch occurring at ~1085 cm–1 (23). Unfortunately, these features significantly overlap the related vibrations in DNA, complicating spectral assignments of complexes. The trimethylammonium headgroup of many lipids has several vibrational modes. Bending modes of the C–H group appear as medium to weak bands at 1485 (antisymmetric) and 1405 (symmetric) cm–1. The N-C-bending vibration can be followed at 970 (antisymmetric) and 920 (symmetric) cm–1. These bands are weak to medium in intensity (25). Polycations such as polyethylenimine (PEI) or poly-L-lysine have IR spectra with a number of distinct bands. Polylysine possesses an NH3+ group, which has symmetric and antisymmetric bending modes at ~1520 and 1630 cm–1 (see Table 2). PEI manifests -NH2+- deformations in the frequency range of 1560–1620 cm–1 (26). Polylysine also possesses the Amide I and Amide II regions, which overlap with much of the in-plane double bond vibrations of the DNA bases and the conformationally sensitive DNA base-sugar coupled vibrations in the range of 1700–1450 cm–1. However, polylysine and PEI do not possess major absorption bands in the spectral region dominated by the deoxyribose-phosphate backbone (1250–950 cm–1), nor do they absorb in the region just above 1700 cm–1, where conformationally sensitive bands, indicative of DNA base-pairing occur. This allows one to follow changes in the double stranded helical geometries caused by changes seen in base pairing and the sugar-phosphate backbone of DNA upon formation of complexes with many polycations. A more detailed list of vibrational bands observed in lipids and polycations is shown in Table 2.

Synthetic Vectors by IR Spectroscopy

293

Table 2 IR Absorption Bands of Common Lipids and Polycations Wavenumber (cm–1) 3000–3500 3200–3500 3038 2956 2920 2870 2850 1720–1750 1630 1623 1520–1650 1520 1485 1473 1468 1460 1405 1370–1380 1200–1400 1228 1170 1085 1070 1060 970 920 720–730 aAssignments

Assignmenta N-H Stretch O-H Stretch (R-OH) CH3 Antisymmetric stretch (Choline) CH3 Antisymmetric stretch CH2 Antisymmetric stretch CH3 Symmetric stretch CH2 Symmetric stretch C=O Stretch of ester NH3+ Antisymmetric bend COO– Antisymmetric stretch N-H Bend NH3+ Symmetric bend (RNH3+) N(CH3)3 C-H Antisymmetric bend CH2 Scissoring (triclinic) CH2 Scissoring (hexagonal) CH3 Antisymmetric bend +N(CH ) C-H Symmetric bend 3 3 CH3 Symmetric bend CH2 Wagging band progression PO2– Antisymmetric stretch CO-O-C Antisymmetric stretch PO2– Symmetric stretch CO-O-C Symmetric stretch C-O-P-O-C Stretch +N(CH ) N-C Antisymmetric bending 3 3 +N(CH ) N-C Symmetric bending 3 3 CH2 Rocking

are from refs. 27, 28, and 31.

3. Instrumentation, Sampling Geometries, and Sample Preparation Over the past decade, development of IR instrumentation has focused on FTIR spectrometers. The newer FTIR instruments provide a number of advantages compared to the older grating models, including faster sampling times (typically 1 vs 20 min for a 2 cm–1 resolution spectrum), high resolution (as low as 1000:1). It is, in fact, often the case that the second derivative of high-resolution spectra of macromolecular samples having a low S:N ratio is too noisy to give accurate peak positions, and “phantom” peaks may be introduced. Switching to a slightly lower resolution often eliminates this problem. However, probably the major advantage of this technique is its ability to determine with great accuracy the positions of not only well-resolved peaks, but also weak shoulders and underlying peak components. This application is especially important when studying complex systems, such as those of interest here, which consist of a large number of overlapping peaks.

4.2. Fourier Self-Deconvolution Often, several peaks of interest lie in the same region of the spectrum and are present as a single broad peak. In some cases, such as the analysis of the complex Amide I bands of protein molecules, the underlying components

300

Choosakoonkriang et al.

cannot be detected by eye. The second-derivative spectrum can be used as indicated, with the intensities of the individual second-derivative peaks used to estimate the relative contribution of each component. Another band-narrowing technique, Fourier self-deconvolution (FSD), can also be used to deconvolute overlapping spectral features. The FSD technique is especially useful from a quantitative perspective because the relative integral areas of the peaks are conserved upon deconvolution. This type of deconvolution is a built in function in most modern data analysis programs and is quite useful. The proper use of this technique requires some care, however. The operator is required to set a single postulated bandwidth at half height for all underlying components and an enhancement parameter representing the degree to which features are resolved. Preliminary results are then calculated, and one iteratively searches for optimal parameters. This method can be subjective and a good rule of thumb is to establish the bandwidth and enhancement parameters with one sample, and, if possible, to remain consistent with them throughout all subsequent deconvolutions of a sample set. Overdeconvolution of the peak can easily occur, introducing spurious peaks. This problem can be identified by the introduction of oscillations into the baseline of empty spectral regions near the peak of interest. Iterative alteration of the FSD parameters involving increases in the bandwidth and decreases in the amplification factor can then be used to minimize the problem. Again, high S:N ratios are necessary for the use of this method, but these are usually obtainable with modern FTIR instruments. It should also be noted that zero-order spectra can be smoothed prior to secondderivative or FSD analysis, but loss of critical information may result. A good example of the application of FSD is in the analysis of the carbonyl region of phosphatidylethanolamine (PE), the IR spectrum of which contains a broad peak centered around 1738 cm–1 (23). This peak has been resolved into two separate components at 1743 and 1726 cm–1 using FSD. It was originally thought that the presence of two peaks resulted from differences in the sn-1 and sn-2 esters of the lipid. However, 13C-isotope editing of one of the ester groups has revealed that both esters contribute to each of the two peaks. It is thought that differences in hydrogen bonding of the carbonyl results in a splitting of the peak with the lower frequency being the result of a species that has more interaction with the solvent water (23). Changes in the relative contributions of the resolved peaks to the zeroorder spectra were followed as a function of temperature, and shifts in the peak position in the zero-order spectra were found to be caused more by changes in their relative intensity than changes in their vibrational energy (peak position).

4.3. Curve Fitting The quantitative analysis of poorly resolved peaks, such as those seen in macromolecules, can present many problems. Although FSD can help identify the presence and location of such bands, it cannot be used to quantitate the

Synthetic Vectors by IR Spectroscopy

301

relative area of each peak. Curve-fitting methods are therefore generally used to determine the relative amounts of previously detected underlying components. In most curve-fitting programs, Gaussian and/or Lorentzian functions are used to represent characteristic component absorption bands. Initial positions and size of the underlying peaks can be found either automatically by the software, or manually determined by the user. The second-derivative and FSD spectra are best used as a guideline for manual peak selection. Once the initial parameters are set, a nonlinear least-squares method is used to get bestfit values of peak position, height and width for each band. In the analysis of DNA and DNA complexes, this technique has not yet been fully explored, although isolated reports have appeared for DNA (9). 5. Application to Gene Delivery Vehicles: Analysis of Cationic Lipid–DNA Complexes Previously, IR spectroscopy has been employed in the characterization of lipids, carbohydrates, nucleic acids (31), and DNA–metal ion binding (32–35). Moreover, IR has been used to detect conformational changes and the interaction of DNA with drugs (36,37), natural or synthetic diamines (5), and peptides (38), as well as changes occurring under various sample conditions, including altered salt concentration (39) and humidity levels (40). Based on these previous studies, the authors have developed methodology for the study of the interaction of DNA with cationic lipids. Here is discussed the analysis of cationic lipid–DNA complexes from spectra obtained by both ATR and thinfilm transmission methods, although similar results have been obtained by the less convenient transmission methods but are not presented.

5.1. Preparation of Liposomes and Complexes The cationic lipid DOTAP was deposited from chloroform solutions on the sides of a glass vial by evaporating the solvent under a stream of nitrogen gas. The resulting film was then placed under vacuum for several hours to remove residual solvent. The lipid was dispersed in 10 mM Tris buffer, pH 7.4, at temperatures above that of the lipid gel-liquid crystalline phase transition for 0.5–1 h. Unilamellar vesicles were prepared by extruding the suspension 10× through a 100 nm pore polycarbonate membrane. All liposomes were used within 3 d of preparation. Cationic lipid–DNA complexes were prepared using various weight ratios of lipid:DNA. Aliquots of lipid and DNA to be mixed were always of equal volume, and were combined by adding the less concentrated component to the more concentrated one. A 9.1-kb plasmid containing >95% supercoiled DNA was used in all studies. The samples were stirred continuously for 10 min, and equilibrated at room temperature for at least 20 min before measurement. All complexes were used the same day as preparation and the final DNA concentration was 1 mg/mL in all cases.

302

Choosakoonkriang et al.

5.2. IR Measurements FTIR spectra were collected using a Nicolet Magna-IR 560 spectrometer equipped with a MCT detector. Spectra were obtained at 4 cm–1 resolution, by co-addition of 256 interferograms. The instrument was continuously purged with dry air to minimize water vapor and CO2 absorption. Samples were measured either by attenuated total reflectance (ATR) or a thinfilm method involving the deposition of the sample on a PE film. Background scans with minimal water vapor were collected before sampling. For ATR measurements, ~1 mL of each sample was placed in a trough ATR accessory (Thermal A.R.K., SpectraTech) equipped with a ZnSe crystal (45 degree angle of reflection, 12 bounces). The samples covered the entire crystal. The sample compartment was purged with dry air for at least 20 min before the spectrum was collected. For the transmission study, sampling cards with a PE membrane were used (3 M IR card, type 61, 19-mm aperture). For each sample, 50 µL was deposited onto the center of the PE card and dried under mild vacuum overnight, prior to measurement. After each card was placed in the transmission accessory, the sample compartment was again allowed to purge for at least 20 min before collecting spectra. All samples were prepared and analyzed in triplicate at a minimum. Because the spectral shifts of interest are often small (95% supercoiled. In general, whatever the topological form of the DNA, it should be as homogeneous as possible. Glassware should be exhaustively rinsed with distilled and deionized water that has been filtered through a 0.22-µm or smaller filter. Additionally, it may be required that the glassware be cleaned with chromic acid solutions to ensure adequate cleanliness. Regarding solution preparation, glassware with groundglass connectors such as volumetric flasks should be avoided to eliminate glass particulates. All buffers should be filtered to remove dust material. In addition to simply using a syringe-based filter system, filtration devices can be produced using peristaltic pumps to cycle buffer and samples repeatedly through a filter or series of filters. The use of these continuous filtration devices is strongly recommended for weakly scattering samples and in any situation in which reproducibility problems are encountered. If solutions containing DNA, complexing agent, or a complex of these molecules are to be filtered, however,

360

Wiethoff and Middaugh

verification of a lack of extensive filter binding of particles and an absence of structural changes induced by filtration should be performed, if possible, before exhaustive characterization of the filtered samples. Concentration can be an important parameter when making light scattering measurements. Depending on the size and shape of complexes, concentration is crucial to obtaining accurate estimates of the molecular size in static experiments. To measure the radius of gyration, the assumption of negligible interparticle interactions is made. Thus, dilute solutions are best employed (see Eqs. 9 and 12). The configuration of the instrument may dictate the range of concentrations applicable because of the power of the light source, the ability to attenuate the light source with neutral density filters and the ability to adjust the cross-sectional area of the light seen by the photomultiplier tube, relative to the coherence area by means of an aperture. In general, a useful DNA concentration range for initial evaluation is between 10 and 100 µg/mL. The DNA concentration may need to be higher if the DNA is to be evaluated in the absence of complexing agents. Ideally, samples should be prepared in a concentration range in which the intensity of scattered light is linearly dependent on the concentration, assuming the physical properties (e.g., size or shape) of the complex do not change over this concentration range. If a sample is too concentrated in DLS experiments, the results may suggest a hydrodynamic radius that is smaller than the true value. This artifact is caused by deviation from the assumption that a single scattering event occurs. In concentrated solutions, multiple scattering events (i.e., a given photon is scattered from one particle to another before it is detected) can occur, producing an apparent increase in the fluctuations in the intensity of scattered light with time. To minimize this problem, it has been suggested that the optical density (i.e., turbidity) for a 1-cm pathlength cell of 2 mg/mg DNA) are cytotoxic to endothelial cells in vitro. Brazeau et al. (47) used an in vitro system of cultured muscle tissue to assess myotoxicity of various cationic macromolecules. They found that cationic liposomes were less myotoxic compared to the dendrimers and polylysine. Myotoxicity was dependent on concentration and mol wt. The presence of plasmid DNA significantly reduced toxicity probably because of the decrease of overall positive charge of the complexes. When DNA– liposome complexes were injected into the renal pelvis of mice, there were no histological changes in the kidney or changes in renal functions (15,16). PEI appears to have low toxicity both in vivo and in vitro (17).

Renal Gene Therapy

387

Intravenous administration of some liposomal drugs can trigger immediate hypersensitivity reactions, including symptoms of cardiopulmonary distress. Szebeni et al. (48) reported that iv injection of 5-mg boluses of large multilamellar liposomes (dimyristoyl phosphatidylcholine, dimyristoyl phosphatidylglycerol, and cholesterol at 50:5:45 mol ratio) into pigs (32–48 kg) caused a significant rise in pulmonary arterial pressure, pulmonary and systemic vascular resistance, and a decline in cardiac output. These changes peaked between 1 and 5 min after injection, subsided within 10–20 min, and were lipid-dose-dependent. There was a close quantitative and temporal correlation between the hemodynamic changes and the elevations of plasma thromboxane B2 level. Those changes could be inhibited by an anti-C5a monoclonal antibody and soluble CR1. The results of that study (48) indicate that liposomes may activate complements and cause lifethreatening events. However, this complication has not been reported in the studies of liposome-mediated gene transfer by other investigators (43,44). Whether this phenomenon is associated with this particular type of liposome or particular animal model, remains to be determined.

7.4. Fusogenic Protein So far, no toxicity associated with HVJ–liposome complex in small animals has been reported. Repeated administration of the HVJ–liposome induced antibody formation, but the transfection efficiency is not significantly affected (4).

7.5. Gene Products For hereditary renal diseases, gene therapy will introduce a normal, but novel, protein to the host who has not been exposed to such a protein prior to treatment. Theoretically, after gene therapy, patients may develop antibodies against therapeutic gene products, particularly if they are circulatory or membranous proteins. Therefore, it may be advisable to measure those antibodies, and, if they are present at a high titer, immunosuppressive therapy may be added in conjunction of gene therapy at least at the initial period to inhibit antibody formation. 8. Conclusions Gene therapy for renal diseases may become a clinical reality in the near future. The most promising field is the application of gene therapy for renal transplantation. The use of gene therapy for acute tubular necrosis, acute glomerulonephritis, and renal vascular disease is also likely to be developed. The most difficult area is gene therapy for hereditary or chronic renal diseases, because long-term and cellspecific expression of transgenes is still far from achievable. New strategies are needed to overcome these obstacles in this area. Finally, as more is learned about the side effects of nonviral gene therapy, efforts should be made to explore the

388

Lien and Lai

mechanisms of these side effects, and to minimize them. With continuous progress in vector technologies and gene therapy strategies, the authors are confident that the era of gene therapy is coming soon. Acknowledgments This work was supported by NIH grant RO1DK52358 and a grant from the Dialysis Clinic, a nonprofit organization. References 1. Lien, Y. H. and Lai, L. (1997) Gene therapy for renal diseases. Kidney Int. 52(Suppl. 61), S85–S88. 2. Lipkowitz, M. S., Klotman, M. E., Bruggeman, L. A., Nicklin, P., Hanss, B., Rappaport, J., and Klotman P. E. (1996) Molecular therapy for renal diseases. Am. J. Kidney Dis. 28, 475–492. 3. Moullier, P., Salvetti, A., Champion-Arnaud, P., and Ronco, P. M. (1997) Gene transfer into the kidney: current status and limitations. Nephron 77, 139–151. 4. Imai, E. and Isaka, Y. (1998) Strategies of gene transfer to the kidney. Kidney Int. 53, 264–72. 5. Kelley, V. R. and Sukhatme, V. P. (1999) Gene transfer in the kidney. Am. J. Physiol. 276, F1–F9. 6. Isaka, Y., Akagi, Y., Kaneda, Y., and Imai, E. (1998) The HVJ liposome method. Exp. Nephrol. 6, 144–147. 7. Tomita, N., Higaki, J., Morishita, R., Kato, K., Mikami, H., Kaneda, Y., and Ogihara, T. (1992) Direct in vivo gene introduction into rat kidney. Biochem. Biophy. Res. Commun. 186, 129–134. 8. Akami, T., Arkawa, K., Okamoto, M., Akioka, K., Fujiware, I., Nakai, I., et al. (1994) Introduction and expression of human CD59 gene in the canine kidney. Transplant Proc. 26, 1315–1317. 9. Yamada, T., Horiuchi, M., Morishita, R., Zhang, L., Pratt, R. E., and Dzau, V. J. (1995) In vivo identification of a negative regulatory element in the mouse renin gene using direct gene transfer. J. Clin. Invest. 96, 1230–1237. 10. Isaka I., Fujiwara Y., Ueda, N., Kaneda, Y., Kamada, T., and Imai, E. (1993) Glomerulosclerosis induced by in vivo transfection by transforming growth factor-` or platelet-derived growth factor gene into the rat kidney. J. Clin. Invest. 92, 2597–2601. 11. Akagi, Y., Isaka, Y., Arai, M., Kaneko, T., Takenaka, M., Moriyama, T., et al. (1996) Inhibition of TGF-beta 1 expression by antisense oligonucleotides suppressed extracellular matrix accumulation in experimental glomerulonephritis. Kidney Int. 50, 148–155. 12. Maeshima Y., Kashihara, N., Yasuda, T., Sugiyama, H., Sekikawa, T., Okamoto, K., et al. (1998) Inhibition of mesangial cell proliferation by E2F decoy oligodeoxynucleotide in vitro and in vivo. J. Clin. Invest. 101, 2589–2597. 13. Ledley, F. D. (1995) Nonviral gene therapy: the promise of genes as pharmaceutical products. Hum. Gene Ther. 6, 1129–1144.

Renal Gene Therapy

389

14. Lien, Y. H. and Lai, L. (1997). Liposome-mediated gene transfer into the tubules. Exp. Nephrol. 5, 132–136. 15. Lai, L., Moeckel, G. W., and Lien, Y. H. (1997) Kidney-targeted liposome-mediated gene transfer in mice. Gene Ther. 4, 426–431. 16. Lai, L., Chan, D., Erickson, R. P., Hsu, S. J., and Lien, Y. H. (1998) Correction of renal tubular acidosis in carbonic anhydrase II deficient mice with gene therapy. J. Clin Invest. 101, 1320–1325 17. Boletta, A., Benigni, A., Lutz, J., Remuzzi, G., Soria, M. R., and Monaco, L. (1997) Nonviral gene delivery to the rat kidney by polyethylenimine. Hum. Gene Ther. 8, 1243–1251. 18. Oberbauer, R., Schreiner, G. F., and Meyer, T. W. (1995) Renal uptake of an 18mer phosphorothioate oligonucleotide. Kidney Int. 48, 1226–1232. 19. Carome, M. A., Kang, Y. H., Bohen, E. M., Nicholson, D. E., Carr, F. E., Kiandoli, L. C., et al. (1997) Distribution of the cellular uptake of phosphorothioate oligodeoxynucleotides in the rat kidney in vivo. Nephron 75, 82–87. 20. Oberbauer, R., Schreiner, G. F., Biber, J., Murer, H., and Meyer, T. W. (1996) In vivo suppression of the renal Na+/Pi cotransporter by antisense oligonucleotides. Proc. Natl. Acad. Sci. USA 93, 4903–4906. 21. Wang, Z. Q., Felder, R. A., and Carey, R. M. (1999) Selective inhibition of the renal dopamine subtype D1A receptor induces antinatriuresis in conscious rats. Hypertension 33, 504–510. 22. Noiri, E., Peresleni, T., Miller, F., and Goligorsky, M. S. (1996) In vivo targeting of inducible NO synthase with oligodeoxynucleotides protects rat kidney against ischemia. J. Clin. Invest. 97, 2377–2383. 23. Haller, H., Dragun, D., Miethke, A., Park, J. K., Weis, A., Lippoldt, A., Gross, V., and Luft, F.C. (1996) Antisense oligonucleotides for ICAM-1 attenuate reperfusion injury and renal failure in the rat. Kidney Int. 50, 473–480. 24. Dragun, D., Tullius, S. G., Park, J. K., Maasch, C., Lukitsch, I., Lippoldt, A., et al. (1998) ICAM-1 antisense oligodeoxynucleotides prevent reperfusion injury and enhance immediate graft function in renal transplantation. Kidney Int. 54, 590–602. 25. Dragun, D., Lukitsch, I., Tullius, S. G., Qun, Y., Park, J. K., Schneider, W., Luft, F. C., and Haller, H. (1998) Inhibition of intercellular adhesion molecule-1 with antisense deoxynucleotides prolongs renal isograft survival in the rat. Kidney Int. 54, 2113–2122. 26. Stepkowski, S. M., Wang, M. E., Condon, T. P., Cheng-Flournoy, S., Stecker, K., Graham, M., et al. (1998) Protection against allograft rejection with intercellular adhesion molecule-1 antisense oligodeoxynucleotides. Transplantation 66, 699–707. 27. Isaka, Y., Brees, D. K., Ikegaya, K., Kaneda, Y., Imai E., Noble, N. A., and Border, W. A. (1996) Gene therapy by skeletal muscle expression of decorin prevents fibrotic disease in rat kidney. Nature Med. 2, 418–423. 28. Isaka, Y., Akagi, Y., Ando, Y., Tsujie, M., Sudo, T., Ohno, N., et al. (1999) Gene therapy by transforming growth factor-beta receptor-IgG Fc chimera suppressed extracellular matrix accumulation in experimental glomerulonephritis. Kidney Int. 55, 465–475.

390

Lien and Lai

29. Isaka, Y., Akagi, Y., Ando, Y., and Imai, E. (1997) Application of gene therapy to diabetic nephropathy. Kidney Int. 52(Suppl. 60), S100–S103. 30. Lenkkeri, U., Mannikko, M., McCready, P., Lamerdin, J., Gribouval, O., Niaudet, P. M., et al. (1999) Structure of the gene for congenital nephrotic syndrome of the finnish type (NPHS1) and characterization of mutations. Am. J. Human Genet. 64, 51–61. 31. Scheiman, S. J., Guay-Woodford, L. M., Thakker, R. V., and Warnock, D. G. (1999) Genetic disorders of renal electrolyte transport. N. Engl. J. Med. 340, 1177–1187. 32. Lai, L. and Lien, Y. H. (1999) Homologous recombination-based gene therapy. Mini-review. Exp. Nephrol. 7, 11–14. 33. Lai, L., Chau, B., and Lien, Y. H. (1999) In vivo gene targeting in carbonic anhydrase II deficient mice by chimeric RNA/DNA oligonucleotides. Annual meeting of American Society of Gene Therapy, Washington, DC. Conference Proceedings, p. 236a. 34. Schwarting, A., Moore, K., Wada, T., Tesch, G., Yoon, H. J., and Kelley, V. R. (1998) IFN-gamma limits macrophage expansion in MRL-Fas(lpr) autoimmune interstitial nephritis: a negative regulatory pathway. J. Immunol. 160, 4074–4081. 35. Morishita, R., Gibbons, G. H., Horiuchi, M., Ellison, K. E., Nakama, M., Zhang, L., et al. (1995) A gene therapy strategy using a transcription factor decoy of the E2F binding site inhibits smooth muscle proliferation in vivo. Proc. Natl. Acad. Sci. USA 92, 5855–5859. 36. Kaneda, Y., Morishita, R., and Dzau, V. J. (1997) Prevention of restenosis by gene therapy. Ann. NY Acad. Sci. 811, 299–308. 37. Poston, R. S., Tran, K. P., Mann, M. J., Hoyt, E. G., Dzau, V. J., and Robbins, R. C. (1998) Prevention of ischemically induced neointimal hyperplasia using ex vivo antisense oligodeoxynucleotides. J. Heart Lung Transplant. 17, 349–355. 38. Isner, J. M. and Asahara, T. (1999) Angiogenesis and vasculogenesis as therapeutic strategies for postnatal neovascularization. J. Clin. Invest. 103, 1231–1236. 39. Lopez , J. J., Laham, R. J., Stamler, A., Pearlman, J. D., Bunting, S., Kaplan, A., et al. (1998) VEGF administration in chronic myocardial ischemia in pigs. Cardiovasc. Res. 40, 272–281. 40. Swenson, K. M., Ke, B., Wang, T., Markowitz, J. S., Maggard, M. A., Spear, G. S., et al. (1998) Fas ligand gene transfer to renal allografts in rats: effects on allograft survival. Transplantation 65, 155–160. 41. Olthoff, K. M., Judge, T. A., Gelman, A. E., da Shen, X., Hancock, W. W., Turka, L. A., and Shaked, A. (1998) Adenovirus-mediated gene transfer into cold-preserved liver allografts: survival pattern and unresponsiveness following transduction with CTLA4Ig. Nature Med. 4, 194–200. 42. Schwartz, D. A., Quinn, T. J., Thorne, P. S., Sayeed, S., Yi, A. K., and Krieg, A. M. (1997) CpG motifs in bacterial DNA cause inflammation in the lower respiratory tract. J. Clin. Invest. 100, 68–73. 43. Stewart, M. J., Plautz, G. E., Buono, L. D., Yang, Z. Y., Xu, L., Gao, X., et al. (1992) Gene transfer in vivo with DNA-liposome complexes: safety and acute toxicity in mice. Hum. Genet. Ther. 3, 267–275.

Renal Gene Therapy

391

44. Canonico, A. E., Plitman, J. D., Conary, J. T., Meyrick, B. O., and Brigham, K. L. (1994) No lung toxicity after repeated aerosol or intravenous delivery of plasmidcationic liposome complexes. J. Appl. Physiol. 77, 415–419. 45. Monteith, D. K., Horner, M. J., Gillett, N. A., Butler, M., Geary, R., Burckin, T., Ushiro-Watanabe, T., and Levin A. A. (1999) Evaluation of the renal effects of an antisense phosphorothioate oligodeoxynucleotide in monkeys. Toxicol. Pathol. 27, 307–317. 46. Henry, S. P., Taylor, J., Midgley, L., Levin, A. A., and Kornbrust, D. J. (1997) Evaluation of the toxicity of ISIS 2302, a phosphorothioate oligonucleotide, in a 4-week study in CD-1 mice. Antisense Nucleic Acid Drug Dev. 7, 473–481. 47. Brazeau, G. A., Attia, S., Poxon, S., and Hughes, J. A. (1998) In vitro myotoxicity of selected cationic macromolecules used in non-viral gene delivery. Pharm. Res. 15, 680–684. 48. Szebeni, J., Fontana, J. L., Wassef, N. M., Mongan, P. D., Morse, D. S., Dobbins, D. E., et al. (1999) Hemodynamic changes induced by liposomes and liposomeencapsulated hemoglobin in pigs: a model for pseudoallergic cardiopulmonary reactions to liposomes. Role of complement and inhibition by soluble CR1 and anti-C5a antibody. Circulation 99, 2302–2309.