Effect of Albumin and Polyanion on the Structure of DNA Complexes

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Bioconjugate Chem. 1999, 10, 764−772

Effect of Albumin and Polyanion on the Structure of DNA Complexes with Polycation Containing Hydrophilic Nonionic Block David Oupicky´,*,† C ˇ estmı´r Konˇa´k,† Philip R. Dash,‡ Leonard W. Seymour,‡ and Karel Ulbrich† Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Heyrovsky Sq. 2, 162 06 Prague 6, Czech Republic, and CRC Institute for Cancer Studies, University of Birmingham, Birmingham B15 2TA, U.K. Received January 15, 1999; Revised Manuscript Received April 29, 1999

Self-assembling systems based on ionic complexes of DNA with block copolymer of N-(2-hydroxypropyl)methacrylamide with 2-(trimethylammonio)ethyl methacrylate were studied as systems suitable for gene delivery. In this study, the influence of albumin and polyanion on parameters of the DNA polyelectrolyte complexes in aqueous solutions was investigated. Static and dynamic light-scattering methods were used as a main tool for characterizing these interactions. It was found that albumin is not able to release free DNA, but it can rather bind to the complexes forming ternary DNA-polycationalbumin complexes with increased hydrodynamic radii of about 10 nm. Polyanion tested, sodium poly(styrenesulfonate), was able to release free DNA in the presence of a low-molecular-weight electrolyte. In the absence of a low-molecular-weight electrolyte, only formation of ternary complexes and no DNA release was observed. The in vivo biodistribution analysis of DNA complexes showed no effect of the presence of hydrophilic nonionic poly(HPMA) on the circulatory time or organ distribution. The interaction of DNA complexes with albumin and other plasma proteins was suggested to be a major reason for the short circulatory times.

INTRODUCTION

The concept of using diblock or graft copolymers of cationic and hydrophilic nonionic monomers has been introduced as a potential way in development of nonviral gene delivery vectors (1, 2). So far, several different polycations have been used in combination with hydrophilic nonionic polymers to produce complexes with DNA. These include block copolymers of poly(ethylene glycol) (PEG) with poly(L-lysine) (PLL) (3, 4) (studied most widely) and with spermine (5), poly[N-(2-hydroxypropyl)methacrylamide] [poly(HPMA)] with poly[2-(trimethylammonio)ethyl methacrylate chloride] [poly(TMAEM)] (2, 6) or graft copolymers of PEG with PLL, dextran with PLL, poly(HPMA) with PLL and poly(TMAEM) (7, 8). Most of the above-mentioned block and graft copolymers have shown improved solution properties of their complexes with DNA (2-4). Block and graft copolymers offer the possibility of improving the solubility of the DNA complexes due to solubilization by the hydrophilic blocks. It has been shown in our previous papers (6, 8, 9) that using block or graft copolymers of N-(2-hydroxypropyl)methacrylamide (HPMA) with 2-(trimethylammonio)ethyl methacrylate chloride (TMAEM) offered such improvement in solubility of stoichiometric complexes with DNA. There was a clear relation between the content of poly(HPMA) in the copolymers and the molecular weight and size of their complexes with DNA. High contents of poly(HPMA) suppressed more efficiently aggregation of the complexed DNA through hydrophobic interactions and thus the complexes were smaller and had lower molecular weight, approaching that of the complexes containing only one DNA molecule. The effect of the * To whom correspondence should be addressed. Phone: 4202-20403212. Fax: 420-2-367981. E-mail: [email protected]. † Institute of Macromolecular Chemistry. ‡ University of Birmingham.

presence of poly(HPMA) was even more pronounced if the complexes were formed by slow titration of polycation into DNA solution, in contrast to the complexes formed by rapid titration. While DNA complexes of poly(TMAEM) formed in water by slow titration have precipitated from the solution when the charge ratio close to unity was reached, the complexes formed by the same method using both the block and the graft copolymers of TMAEM were fully soluble. Although DNA complexes of poly(TMAEM) formed in water by the rapid titration were soluble at all the charge ratios studied (0-2), the complexes formed at charge ratios close to unity showed high degrees of aggregation. We have assumed that the solubility of the DNA-poly(TMAEM) complexes formed by the rapid titration was caused by imperfect compensation of DNA phosphate groups; the slow titration method facilitates their better compensation and thus formation of more hydrophobic and insoluble complexes. It has also been shown (8) that the presence of NaCl considerably influences solution properties of the formed complexes. The complexes formed in 0.15 M NaCl solution exhibited significantly higher sizes and molecular weights than those formed in water. The formation of the complexes in distilled water resulted in small complexes, which were relatively stable when transferred into 0.15 M NaCl solution. The study showed that a relatively high content of HPMA in block or graft copolymers with TMAEM was a necessary prerequisite for achieving good stability in 0.15 M NaCl. Complexes of DNA with poly(TMAEM) and block copolymers with low contents of HPMA precipitated shortly after NaCl addition, thus, not fulfilling basic requirements for gene delivery vector intended for the in vivo use. Whereas the stability in NaCl solution and parameters of the DNA complexes (molecular weight, hydrodynamic radius) were strongly affected by the presence of hydrophilic blocks in copolymers, the ability of the polycationic part of block

10.1021/bc990007+ CCC: $18.00 © 1999 American Chemical Society Published on Web 07/27/1999

Interaction of Albumin with DNA Complexes

or graft copolymer to interact with DNA was unaffected (8). Analysis of the complexes by atomic force microscopy revealed that whereas most block copolymers produce common spherical complexes, PEG-PLL complexes showed a mixture of extended toroidal and linear strands and spherical structures (10). It has been shown that complexation of DNA with PLL can protect it from degradation by nucleases (11); however, in the presence of mung bean nucleases, Dash (3) found partial degradation of DNA-PLL complexes, while DNA-PEG-b-PLL complexes remained completely intact. The ability to transfect cells in vitro was shown for most of the above-mentioned block and graft copolymers, although the transfection efficiency could vary considerably depending on the type of the copolymer used. DNA-PEG-b-PLL complexes showed an impressive ability to transfect 293 cells, the effect thought to result from the ability of PEG to dehydrate cell membranes (2). Presumably, the clusters of PEG exposed in closed juxtaposition can cause local permeability of the cell membrane of 293 cells. On the other hand, DNA complexes formed with poly(HPMA)-b-poly(TMAEM) mediated only limited transfection efficiency compared with that of DNA-PEG-b-PLL complexes (2). The low level of transfection in the case of DNA complexes with poly(HPMA)-b-poly(TMAEM) suggests that they do not enter the cell in such a nonspecific manner as PEG-based systems and thus this type of copolymer could be a good candidate for in vivo applicable targeted gene delivery systems after attachment of specific targeting moieties into the poly(HPMA) blocks. The complexes of DNA, similarly to other colloidal particles (like liposomes), show very low stability in the bloodstream (12, 13). They are rapidly removed from the bloodstream, which decreases their potential for specific targeting in vivo. It has been proposed that interaction of the complexes with plasma proteins (either adsorption or breaking of the complexes by proteins) can be the major obstacle for their successful in vivo use. In analogy with STEALTH liposomes (14), it was suggested that incorporation of hydrophilic, nonionic polymer onto the surface of the complexes can help to solve this problem (2, 7). The concept of using block or graft copolymers, having been developed in our laboratories for several years, seems to offer that possibility. It was suggested that the complex formation may proceed with a degree of orientation, leaving the hydrophilic blocks on the surface of the complex free to provide a protective layer (7). In this work, we have studied the interactions of the polyelectrolyte complexes of DNA with poly(TMAEM) containing hydrophilic poly(HPMA) block with albumin and synthetic polyanion. The interactions were studied by combination of several methods, including dynamic and static light-scattering and measurements of ζ-potential. The ability of albumin and polyanions to release free DNA from the complexes was tested using dynamic light scattering and agarose gel electrophoresis. Finally, the body distribution analysis was performed in order to obtain more information on the effect of the hydrophilic poly(HPMA) block on biological properties of complexes. EXPERIMENTAL PROCEDURES

Chemicals. Calf-thymus DNA and bovine serum albumin were purchased from Sigma. Sodium poly(styrenesulfonate) (PSS) (Mw ) 74 000) was from PolySciences. All other chemicals were of analytical grade.

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Synthesis of Polymers. Block copolymer was prepared by the reaction of active succinimidyl esters of carboxylic end groups in semitelechelic poly(HPMA) with semitelechelic poly(TMAEM) containing terminal amino group. Synthesis of Poly(HPMA) with Carboxylic Acid End Group [Poly(HPMA)-COOH]. Precipitation polymerization of HPMA in acetone was carried out in the presence of a chain transfer agent 3-sulfanylpropionic acid (SPA) at 50 °C for 24 h. The concentrations were as follows: initiator 2,2′-azobisisobutytonitrile (AIBN) 3 × 10-3 M, HPMA 0.8 M and SPA 7.6 × 10-3. The polymer was purified by double precipitation from methanol solution into 20-fold excess of a mixture of acetone and diethyl ether (3:1). Yield 45%. Weight (Mw)- and numberaverage (Mn) molecular weights determined by sizeexclusion chromatography (SEC) were Mw ) 14 700 and Mn ) 8400. Molecular weight calculated from the content of end carboxylic acid group was Mn ) 8500. Synthesis of Poly(TMAEM) with Amino End Group [Poly(TMAEM)-NH2]. The polymer was prepared by a solution radical polymerization in methanol in the presence of 2-aminoethanethiol hydrochloride (AET.HCl) at 50 °C for 24 h. The concentrations were as follows: initiator AIBN, 4.8 × 10-3 M; TMAEM, 0.6 M; and AET‚HCl, 8.5 × 10-2 M. The polymer poly(TMAEM)NH2 was isolated by precipitation into acetone, purified by dialysis (dialyzing tubing with molecular weight cutoff 3500), and freeze-dried. Yield 35%. Molecular weights determined by SEC using poly(HPMA) calibration were: Mw ) 10 600; Mn ) 7470. Molecular weight calculated from the content of end amino groups was Mn ) 5000. Synthesis of Active Succinimidyl Ester of Poly(HPMA)-COOH. The end carboxylic acid group was converted into succinimidyl ester by the reaction common in peptide synthesis. Poly(HPMA)-COOH (0.17 mmol) and N-hydroxysuccinimide (HOSu) (1.70 mmol) were dissolved in 6 mL of dimethylformamide (DMF) (dried with P2O5, freshly distilled before use) and cooled to -20 °C; to this solution was added a solution of N,N′dicyclohexylcarbodiimide (1.70 mmol) and 4-(dimethylamino)pyridine (0.33 mmol) in 5 mL of DMF. The solution was left overnight at 0 °C. The N,N′-dicyclohexylurea precipitated during the reaction was removed by filtration and polymer was isolated by precipitation into 15fold excess of a mixture of dry acetone and diethyl ether (3:1). The succinimidyl ester of poly(HPMA)-COOH [poly(HPMA)-COOSu] in quantitative yield was dried and used directly in the subsequent reaction. Synthesis of Block Copolymer [Poly(HPMA)block-poly(TMAEM)]. Poly(HPMA)-COOSu was dissolved in dimethyl sulfoxide (10 wt % solution) and added to a solution of poly(TMAEM)-NH2. Molar ratio of polymer end groups COOSu:NH2 was 1.2:1. The mixture was stirred at room temperature for 8 h and polymer was isolated by precipitation into 40-fold volume excess of acetone. The polymer (BLOCK-1) was dried in vacuo, excess of poly(HPMA) was removed from dry copolymer by extraction with DMF, and the product was washed with acetone and dried in vacuo. Molecular weight of the copolymer BLOCK-1 determined by SEC using poly(HPMA) calibration was Mw ) 21 000. The content of HPMA block determined by elemental analysis was 63 wt %. Formation of Polyelectrolyte Complexes. The complexes were formed in deionized water by titration of DNA solution (concentration 20 µg/mL, concentration of phosphate groups 0.0615 mM) with polycation stock

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solution (positive charge concentration 0.012 M). Two modes of preparation were used in this study. (1) Rapid titration: a polycation stock solution was added as quickly as possible (typically in 1 s) to 3 mL of a stirred DNA solution. (2) Slow titration: a polycation stock solution was slowly added using a programmable syringe pump to 3 mL of a stirred DNA solution, the rate of addition being 30 µL/h (full compensation of negative DNA phosphate groups was achieved in 30 min). Calf-thymus DNA was used in all experiments, unless otherwise stated. Characterization of Polymers. Molecular weights of poly(HPMA)-COOH and poly(TMAEM)-NH2 were determined by size-exclusion chromatography (SEC) in 0.05 M Tris buffer (pH 8.1) on a Superose 12 column (Pharmacia). Number-average molecular weights (Mn) were also calculated from the content of the end functional carboxylic or amino groups, assuming one functional end group per macromolecule. The content of the end carboxylic groups of poly(HPMA)-COOH was determined by titration with 0.05 M NaOH using automatic titrator (Radiometer). The amount of amino end groups of poly(TMAEM)-NH2 was determined by the 2,4,6-trinitrobenzenesulfonic acid assay using molar absorption coefficient 10 500 L/(mol cm). Agarose Gel Electrophoresis. The ability of albumin or polyanion to release free DNA from the complexes was also determined by agarose gel electrophoresis. DNA complexes in water in the presence of various concentrations of albumin or polyanion were electrophoresed on agarose gel (0.5%) in 0.05 M Tris buffer, pH 7.5, for 30 min at 90 V. Ethidium bromide (1 µg/mL of gel) was incorporated in the gel to show the location of DNA using UV detection (254 nm). Analysis of ζ-Potential. The charge of the calfthymus DNA complexes was determined by using a Zetamaster system (Malvern Instruments). The system was calibrated using a -55 mV standard. Solutions of the complexes (5 mL) were measured five times for 30 s at 1000 Hz with zero field correction. Static Light Scattering (SLS). Static light-scattering measurements were performed with a commercial Fica 40 apparatus in vertically polarized light at wavelength λ0 ) 632.8 nm, angle θ ) 90°, and temperature 25 °C. The apparatus was calibrated with benzene at θ ) 90°. The data are expressed as the scattered light intensity at 90 ° normalized with intensity scattered by benzene. Dynamic Light Scattering (DLS). Polarized DLS measurements were made in the angular range 30-130° using a light-scattering apparatus equipped with an HeNe (632.8 nm) and Ar-ion laser (514.5 nm) and an ALV 5000, multibit, multi-tau autocorrelator covering approximately 10 decades in delay time τ. The time autocorrelation functions were fitted assuming the Pearson distribution of characteristic relaxation times, τc:

z(τc) ) τ po pτc-p-1 exp(-τc/τo)/Γ(p)

(1)

where τo and p are parameters, and Γ(p) is the gamma function of parameter p. The Pearson distribution was chosen for the simplicity of its mathematical treatment. The apparent hydrodynamic radius, Rh, was calculated from the zero angle limit of apparent collective diffusion coefficient, Da, using the Stokes-Einstein equation:

Rh ) kT/6πηDa

(2)

where k is the Boltzmann constant, T is absolute temperature, and η (0.894 cP) is the viscosity of water at 25 °C. The experimental error of the aggregate size determination was typically about 3%. Radiolabeling of DNA. The linearized plasmid pCMVβ-Gal was radiolabeled using the Ready-to-Go DNA Labeling Kit (-dCTP) from Pharmacia Biotech. The DNA was purified from unincorporated nucleotides using MicroSpin DNA purification columns from Pharmacia Biotech. Body Distribution Analysis. Radiolabeled DNA (50 ng/mL) was added to a calf-thymus DNA solution (20 µg/ mL), and the complexes were formed using different polycations and various charge ratios. The complex solutions were diluted with PBS buffer to balance the solution osmotically prior to injecting 100 µL into the tail vein of BALB/c mice. After 30 min, the animals were killed and their liver, kidneys, intestines, heart, and lungs were removed. The blood and tissue samples were analyzed for radioactivity in a Packard Scintillation Counter. The total amount of radioactivity in blood was determined by assuming that the volume of blood in a mouse is 8.64 mL/100 g. In all cases, the tails of the mice were removed prior to dissolution and were not assayed for radioactivity. The percentage of radioactivity in each organ is given as a percentage of the recovered dose. RESULTS

Good candidate for in vivo delivery of DNA should exhibit prolonged circulation times in the bloodstream in order to accomplish targeting. In analogy with other particulate systems (liposomes, nanoparticles), the use of cationic block copolymers containing hydrophilic blocks is thought to reduce the removal of polyelectrolyte DNA complexes by the immune system. Since hydrophobic forces are thought to be the main driving force in phagocytosis of nanoparticles and liposomes, the presence of hydrophilic polymers decreases hydrophobic nature of these delivery systems. Particles containing hydrophilic polymers usually display reduced protein adsorption and consequently increased circulation times and altered body distribution (15). We have evaluated several block and graft copolymers of HPMA and TMAEM (8, 9). We have chosen the block copolymer BLOCK-1 as a polymer that can form polyelectrolyte complexes with DNA that are relatively small and are sufficiently stable in saline solution. Before testing the biodistribution of its complexes with DNA, we have studied the interaction of these complexes with albumin. Proteins can interact with DNA complexes among others by electrostatic and hydrophobic interactions. To investigate electrostatic interactions, it is important to measure the charge of the complexes, and in order to suppress these interactions, it is reasonable to attempt to form them charge-neutral. The charge of the complexes was evaluated by measuring their ζ-potentials. Figure 1A shows the dependencies of ζ on the ratio of positive charges of polycation and negative charges of DNA (φ) for poly(TMAEM)-NH2 and block copolymer BLOCK-1 used for the formation of DNA complexes by the rapid titration method. In both cases, ζ monotonically increases and crosses zero between charge ratios 0.7 and 0.8. The complexes gain positive charge at higher charge ratios (φ > 0.8). Since the ζ-potential is related to the sum of all charges of complexes, the shift of complete neutralization of charges to φ values lower than 1 might be due to



Figure 2. Dependence of ζ-potential on the ratio between positive charges of polycation and negative charges of DNA (φ) for DNA-BLOCK-1 complexes. The complexes were formed at φ ) 0.65 by rapid titration and the aliquots of BLOCK-1 solution were added up to φ ) 0.9.

Figure 1. Dependence of ζ-potential on the ratio between positive charges of polycation and negative charges of DNA (φ). (A) complexes formed by rapid titration using poly(TMAEM)NH2 (b) and BLOCK-1 (O); (B) complexes formed by slow titration using BLOCK-1.

positive counterions entrapped inside the complexes by electrostatic interactions with DNA. It has been shown by Manning (16) that when DNA is neutralized by polycations to the critical φ (0.88), the hydrophobicity of DNA is so high that it cannot stay in its extended form and collapses. The collapse then may cause entrapping of remaining free phosphate groups inside the complex. The incorporation of excess block copolymer to the complexes at φ > 0.8 manifested in positive ζ-potential values is probably caused by partly neutralized copolymer molecules on the surface of the complexes. Similar behavior was already observed by several other authors (2, 7, 17). On the contrary, formation of DNA complexes with almost neutral charge was reported by Kataoka using PEG-b-PLL (4). The point of zero ζ-potential around φ ≈ 0.8 also well corresponds to a maximum in the size and molecular weight dependencies observed in our previous paper (9). This confirms that these maxima are caused by the hydrophobically driven aggregation, which most easily proceeds if the complexes are uncharged (ζ ) 0). To our surprise, there was no significant difference in ζ-potential between complexes formed by slow or rapid titration (Figure 1B). On the basis of our previous data, we have intuitively anticipated that the slow titration method facilitates formation of complexes with better compensated DNA negative charges. In such case, the recharging of the complexes after reaching zero ζ-potential could be avoided, and as a consequence, neutral complexes could be easily obtained. Since this assumption was not justified, we have attempted to obtain neutral complexes by finding the exact charge ratio of neutrality (Figure 2). The change from the negatively to positively charged complexes occurs suddenly in a very narrow interval of charge ratios and thus,

from the practical point of view, charge-neutral complexes cannot be easily obtained. Since the recharging of the complexes coincides with the maxima of molecular weight and size of the complexes and with sudden repulsion of intercalated ethidium bromide from DNA (8), the dramatic conformation change (coil-globule transition) of DNA structure (collapse) is assumed to occur in this charge ratio interval. To induce collapse of the extended DNA structure, it is necessary to use charge ratios higher than 0.8. Then, the positive charge of the complexes (Figures 1 and 2) has to be considered when evaluating their interactions with proteins. After interaction with proteins, positive charges of the complexes could promote binding of negatively charged proteins. In certain cases, negatively charged protein could possibly replace DNA in the complex, and free DNA may be released. Since the major protein with negative net charge occurring in blood plasma in high concentrations is albumin, we have used it in this study. A possible release of free DNA from the complexes was assessed by agarose gel electrophoresis. We have shown by electrophoresis that albumin is not able to destroy the complexes and release free DNA, but it could probably interact with positively charged complexes and form ternary albumin-DNA-polycation complexes. In that case, some parameters of the DNA complexes, such as charge, molecular weight, and hydrodynamic radius, have to be changed. To verify this possibility, we have used static and dynamic lightscattering and ζ-potential techniques. DNA complexes with BLOCK-1 were first formed in water at φ ) 0.9, and then increments of albumin solution were added. The dependence of ζ-potential of the complexes on the concentration of albumin is shown in Figure 3A. The increasing concentration of albumin causes gradual change in ζ from positive to negative values. Reaching constant negative ζ above 1 mg/mL of albumin suggests possible saturation of the complexes with albumin. Almost all the positively charged complexes of BLOCK-1 formed at different charge ratios altered their ζ-potential to negative in the presence of 1 mg/mL of albumin (Figure 3B). The complexes formed with high excess of polycation (φ ) 6) are an exception. In this case, substantial quantity of polycation is free in solution and forms additional complexes with albumin having hydrodynamic radii in the range 10-100 nm and ζ varying from negative to positive values depending on the polycation/ albumin ratio. In consequence, polycation-albumin complexes may interfere with DNA complexes and distort the


Oupicky´ et al.

Figure 4. Effect of the addition of 1 mg/mL of albumin to DNA-BLOCK-1 complexes formed at different φ on the scattering intensity at 90°. The complexes were formed in water, scattering intensity was measured (O), then albumin was added and scattering intensity of the solution was again measured (b). The scattering intensity of the DNA-BLOCK-1 complexes in the presence of 1 mg/mL of albumin on addition of 0.15 M NaCl was finally measured (×). Figure 3. Effect of albumin on ζ-potential of DNA complexes. (A) DNA-BLOCK-1 complexes were formed at φ ) 0.9 and effect of increasing albumin concentration on ζ-potential was monitored. (B) DNA-BLOCK-1 complexes were formed at φ ) 0.7, 0.9, 1.1, 2, and 6, their ζ-potential was measured (9) and then effect of 1 mg/mL of added albumin on ζ-potential was monitored (0).

results of ζ-potential measurements as in the case of the complexes formed at φ ) 6 (Figure 3B). Polycationalbumin complexes are positively charged at high φ values and most albumin is probably consumed in the formation of the complexes with free polycation. As a consequence, the overall measured ζ-potential remains positive. Light-scattering techniques can contribute to better understanding of the effect of albumin on the DNA complexes. Figure 4 demonstrates the effect of an addition of albumin (1 mg/mL) on the normalized intensity of scattered light (the scattered light intensity at 90 ° normalized with intensity scattered by benzene) of complexes formed at different charge ratios in water. Scattering intensity is used instead of customary molecular weights because the concentration of albumin built in DNA complexes is unknown. A minor increase in the scattering intensity is observed after albumin addition to the complexes at φ ) 0.9. This corresponds well with the observed positive ζ of the complexes formed at φ ) 0.9 (Figure 1). The increase in scattering intensity suggests an increase in molecular weight of the complexes due to their interaction with albumin. In addition, an abrupt break above the charge ratio 1.1 is observed. We suppose that the abrupt break is provoked mostly by the interaction of albumin with free polycation. The polycation-albumin complexes bring additional scattering to that of DNA complexes. The scattering intensity is again decreased if NaCl is added (Figure 4) to the complexes in the presence of albumin. A markedly dramatic decrease is observed for the complexes formed

at φ > 1.2. We assume that this decrease is mainly caused by break-up of the polycation-albumin complexes. The polycation-albumin complexes are not stable in the presence of a low-molecular-weight electrolyte (NaCl) due to only weak interactions. This is in contrast to the complexes of DNA with polycations, where a strong cooperative effect induces high stability. The decrease in scattering intensity after NaCl addition is significant but yet it has not reached the scattering intensity of DNABLOCK-1 complexes. This observation suggests that some albumin could be possibly entrapped inside the DNA complex (by hydrophobic interactions) increasing thus molecular weight of the complex (manifested by increased scattering intensity). The static light-scattering methods require precise knowledge of particle concentrations in solution to calculate their molecular weights. Dynamic light scattering, on the contrary, offers the possibility of determining directly the hydrodynamic radii of the complexes without precise knowledge of the particle concentration. The effect of increasing concentration of albumin on the distribution of hydrodynamic radii [A(Rh)] of the complexes (DNABLOCK-1) formed in water by slow titration (φ ) 0.9) is demonstrated in Figure 5. An addition of albumin causes a shift of radii distribution to higher values. Saturation of the complexes with albumin was observed above 0.7 mg/mL of albumin where the hydrodynamic radius of the complexes was found to be almost independent of albumin concentration. We assume that the shift of the size distribution is caused predominantly by albumin binding on the surface of the complexes forming there a “protein layer” (hydrodynamic radius of albumin ≈ 4 nm). The average hydrodynamic radius of the complexes in the presence of 1 mg/mL of albumin increased by about 10 nm compared with the original complex. The complexes formed at φ ) 2 exhibited considerably higher shifts of the hydrodynamic radii distributions after



Figure 5. Effect of increasing concentration of albumin on the distribution of hydrodynamic radii of DNA-BLOCK-1 complexes. The complexes were formed at φ ) 0.9 (O), and then the distribution of hydrodynamic radii of DNA-BLOCK-1 complexes was measured after albumin addition [final concentrations of albumin: 0.4 (b), 0.7 (×), and 1 mg/mL albumin (9)].

Figure 7. Effect of PSS on the distribution of hydrodynamic radii of DNA-BLOCK-1 complexes. (A) The complexes were formed at φ ) 1.1 in distilled water (O), effect of an addition of PSS is shown (b). (B) The complexes were formed at φ ) 1.1 in 0.15 M NaCl (O), effect of an addition of PSS is shown 30 min (b) and 3 days (×) after addition.

Figure 6. Effect of 1 mg/mL albumin and 0.15 M NaCl on the distribution of hydrodynamic radii of DNA-BLOCK-1 complexes. The complexes were formed at φ ) 2 (O). Effect of albumin (b) and subsequent addition of NaCl (×) on the distribution of hydrodynamic radii of the complexes is shown.

albumin addition (1 mg/mL) (Figure 6) in water, compared with the complexes formed at φ ) 0.9 (Figure 5). A significant increase in average hydrodynamic radius and polydispersity was observed after albumin addition due to the formation of polycation/albumin complexes. In agreement with our previous results, large complexes formed mostly between albumin and free polycation have broken after addition of NaCl to their solutions (final concentration 0.15 M) (Figure 6). The solution then contained DNA-BLOCK-1 complex particles similar to those prior to the albumin addition. Corresponding experiment with the complex formed at the 0.9 charge ratio showed that a slight increase in the complex size only slightly decreased in 0.15 M NaCl, suggesting that the interaction of albumin with the complexes is relatively more stable than that with free polycations. This is in agreement with the static light-scattering experiments (Figure 4), which indicate that some albumin could be bound to the complexes even in 0.15 M NaCl. The complexes in the presence of albumin were stable over a period of hours, but after 24 h, the starting aggregation was observed. From the practical standpoint, the com-

plexes should remain in circulation until they reach their target site; as the required half-life in the bloodstream is several hours (13), the observed stability could be sufficient. It has been shown that certain polyanions (polyaspartic acid, heparin) can release free DNA from the complexes (3). This reaction can also possibly account for the low stability of the complexes in the bloodstream. We have studied the ability of synthetic polyanion sodium poly(styrenesulfonate) (PSS) to release free DNA from its complexes with BLOCK-1 by agarose gel electrophoresis and dynamic light scattering. To compare the effect of PSS on DNA complexes in water and in 0.15 M NaCl, we have used a high concentration of PSS. The ratio of positive charges of BLOCK-1 used for complex formation and negative charges of PSS was 1:25. The use of a high excess of PSS should ensure that in the case of exchange polyelectrolyte reaction between DNA and PSS in DNA complexes with BLOCK-1, the possibly formed complexes PSS-BLOCK-1 have relatively small size and thus do not interfere with the DNA complexes. Figure 7 shows the distribution of hydrodynamic radii of the DNA-BLOCK-1 complexes prior to and after addition of PSS. The complexes in water (Figure 7A) showed only a small increase in the hydrodynamic radius. Similarly to the interaction of complexes with albumin, a shift of the size distribution to higher values was observed. The hydrodynamic radius of these complexes virtually did not change during the 3 days of observation. The same experiment in 0.15 M NaCl (graph B) showed a similar increase in Rh and shift of the Rh


Oupicky´ et al.

Figure 8. Body distribution of radiolabeled DNA (9) and of the complexes of radiolabeled DNA with poly(TMAEM)-NH2 (0) or BLOCK-1. The latter complexes were formed in water at φ ) 0.7 (0), 0.9 (0), and 1.1 (0). The complex or DNA solutions were diluted with PBS buffer prior to injecting. The mice were killed after 30 min, and the blood and tissue samples were analyzed for radioactivity.

distribution immediately after a PSS addition. From the very beginning, a second population of small particles with the hydrodynamic radius around 10 nm was apparent. The proportion of these particles increased with time at the expense of the DNA complexes. Together with the alteration of the relative content of these two populations, the intensity of the scattered light of the solution at 90° decreased, indicating dissociation of the DNA complexes. This means that large, highly scattering particles slowly disappear. We suppose that, in the absence of NaCl, the exchange polyelectrolyte reaction virtually does not take place and only formation of ternary complexes DNABLOCK-1-PSS occurs, as manifested by a minor increase in the mean hydrodynamic radius. In the presence of 0.15 M NaCl, the exchange reaction can take place. This is manifested by an increasing quantity of 10 nm particles and by a decrease in intensity of scattered light. These observations agree with the results published in ref 18. They have demonstrated that the exchange polyelectrolyte reactions can take place only in the presence of a low-molecular-weight electrolyte. We suppose that the particles with Rh ) 10 nm correspond to complexes PSS-BLOCK-1. The exchange reaction is relatively slow and thus the interaction of DNA complexes in the bloodstream with possibly present polyanions does not have to be an obstacle in successful in vivo gene delivery. Agarose gel electrophoresis was used in order to observe release of free DNA from the complexes. Since DNA complexes do not migrate in the gel, the released free DNA can be easily monitored by this method. PSS was able to release free DNA from the complexes and the amount of DNA released from the complexes increases with increasing concentration of PSS in solution. We have studied the organ distribution of different DNA complexes 30 min after an i.v. tail vein injection (Figure 8). The figure compares the organ distribution of DNA-BLOCK-1 complexes formed at three different φ (0.7, 0.9, 1.1) and that of the complex DNA-poly(TMAEM)-NH2 (φ ) 0.9). To our surprise, the DNABLOCK-1 complexes have manifested almost no difference in their biodistribution compared with DNApoly(TMAEM)-NH2 complexes. Unlike STEALTH liposomes, the presence of poly(HPMA) in the complexes has apparently no effect on the prolongation of circulation

times of the complexes. Neither did exhibit the effect of varying charge ratio in the complex formation and any significant difference in the organ distribution, although the complexes formed at φ ) 0.7 had a negative ζ-potential. No explanation for such an observation was available until now. The accumulation of the complexes in the liver could be probably a consequence of both the charge and relatively high hydrodynamic radius of the complexes (19). DISCUSSION

It has been shown that DNA-PLL complexes are rapidly cleared from the bloodstream, with a half-life of less than 5 min. The interaction of the complexes with albumin was proposed to have a major impact on the circulation of the complexes in blood (13). It was suggested that the main reason could be an increase in the size of the complexes. To test this hypothesis we have studied the interaction of DNA complexes with albumin by combination of several analytical techniques. As we have shown above, albumin is able to interact with DNA-BLOCK-1 complexes and increase their hydrodynamic radii, nevertheless, the increase was not so significant to explain the observed biodistribution. The exchange reaction between DNA and polyanions which caused a release of free DNA is relatively slow, and thus the interaction of DNA complexes in the bloodstream with possibly present polyanions does not have to be an obstacle in successful in vivo gene delivery. On the basis of these promising observations, we have expected prolonged plasma circulation of DNA complexes with block copolymers. In analogy with STEALTH liposomes, we have expected the hydrophilic poly(HPMA) block to increase the circulatory times of the complexes. To explain the fact, that using block copolymers we have observed no significant improvement of the biodistribution of the complexes, it is necessary to review the analogy with STEALTH liposomes. In our previous physicochemical study (9), we have shown that increasing relative HPMA content in both the block and graft copolymers used for the formation of DNA complexes resulted in decreasing density of the complexes. In the case of a graft copolymer containing 80% of HPMA, the



Figure 9. (A) structure of brush polymer (DNA fully compensated with block copolymers). (B) Proposed structure of DNA complexes with block or graft copolymers in solution. (C) Structure of STEALTH liposome.

complexes displayed an expanded structure that was very close to that of a flexible polymer coil, as assessed by the ratio of radius of gyration and hydrodynamic radius of the complexes. Then, the complexes have “swollen and expanded” structure, where their hydrophobic parts might tend to concentrate in the central core of the complex and hydrophilic poly(HPMA) blocks on the surface, but microphase separation of the hydrophobic and hydrophilic parts is virtually impossible. The frequently suggested micelle-like structure with hydrophobic core surrounded by hydrophilic poly(HPMA) or PEG shell seems to be not fully realistic (4, 7) in case for highmolecular-weight DNA. To better explain a possible structure of the DNA complexes with block or graft copolymers, it is necessary to realize that full compensation of DNA with molecular weight of almost 7 million requires several hundreds of polycation macromolecules (molecular weight ≈ 20 000). Then the complexes have the structure of so-called brush polymers (Figure 9A). The DNA complexes are usually about 100 nm in diameter, and thus it is not possible for poly(HPMA) blocks located on the DNA parts in the center of the complex to reach the surface and they must be also located in the core of the complex, where they cause an internal swelling (Figure 9B). In STEALTH liposomes, the protecting polymer layer is located on the surface (Figure 9C), where it sterically stabilizes the liposome due to osmotic and entropic effects of an inert, nonionic, hydrophilic, and flexible polymer coating, which increases repulsive pressure above these surfaces (20). The complexes with the structure closer to STEALTH liposomes, i.e., those with hydrophobic DNA-polycation core surrounded by hydrophilic protecting surface layer, would be more stable in the bloodstream, due to the ability of the compact surface layer to sterically stabilize the particle. We could conclude that a compact surface layer of hydrophilic nonionic polymer is probably required to efficiently decrease interaction with proteins and, consequently, to achieve good stability in the bloodstream. The absence of such compact continuous layer of poly(HPMA) in the DNA complexes studied in this work results in facile binding

of albumin or other proteins to the complexes. The complexes then may have a structure of “large albumin spheres” with positive or negative surface charge dependent on the amount of bound albumin. Then, other plasma proteins (opsonins) could bind and also promote removal of the DNA complexes from the bloodstream, preferably by the liver. A potential way of obtaining the DNA complexes with poly(HPMA) chains located only on their surface might be the chemical modification of the preformed complexes containing surface reactive groups with a reactive poly(HPMA) derivative. ACKNOWLEDGMENT

The support of the EU by Grant ERBIC20CT97005 and of the Grant Agency of the Czech Republic by Grant 307/ 96/K226 is gratefully acknowledged. LITERATURE CITED (1) 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 (K. V. Kabanov, P. L. Felgner, and L. W. Seymour, Eds.) pp 219-239, John Wiley & Sons, New York. (2) 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, 21232133. (3) Dash, P. R., Toncheva, V., Schacht, E. H., and Seymour, L. W. (1997) Synthetic polymers for vectorial delivery of DNA: characterization of polymer-DNA complexes by photon correlation spectroscopy and stability to nuclease degradation and disruption by polyanions in vitro. J. Controlled Release 48, 269-276. (4) 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.

772 Bioconjugate Chem., Vol. 10, No. 5, 1999 (5) Kabanov, A. V., Vinogradov, S. V., Suzdaltseva, Y. G., and Alakhov, V. Y. (1995) Water-Soluble Block Polycations as Carriers for Oligonucleotide Delivery. Bioconjugate Chem. 6, 639-643. (6) Konˇa´k, C ˇ ., Mrkvicˇkova´, L., Nazarova, O., Ulbrich, K., and Seymour, L. W. (1998) Formation of DNA complexes with diblock copolymers of poly(N-(2-hydroxypropyl) methacrylamide) and polycations. Supramol. Sci. 5, 67-74. (7) 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. (8) Oupicky´, D., Konˇa´k, C ˇ ., and Ulbrich, K. (1998) DNA Complexes with Block and Graft Copolymers of N-(2-Hydroxypropyl)methacrylamide and 2-(Trimethylammonio)ethyl methacrylate. J. Biomater. Sci., Polym. Ed. (in press). (9) Oupicky´, D., Konˇa´k, C ˇ ., and Ulbrich, K. (1999) Preparation of DNA Complexes with Diblock Copolymers of Poly[N-(2Hydroxypropyl)methacrylamide] and Polycations. Mater. Sci. Eng. C (in press). (10) Wolfert, M. A., and Seymour, L. W. (1996) Atomic Force Microscopy of the influence of the molecular weight of poly(L-lysine) on the size of polyelectrolyte complexes formed with DNA. Gene Ther. 3, 269-273. (11) Chion, H. C., Tangco, M. V., Levine, S. M., Robertson, D., Karmisk, L., Wu, C. H., and Wu, G. Y. (1994) Enhanced resistance to nuclease degradation of nucleic acids complexed to asialoglycoproteins-polylysine carriers. Nucleic Acids Res. 22, 5439-5446. (12) Oja, C. D., Semple, S. C., Chonn, A., and Cullis, P. R. (1996) Influence of dose on liposome clearance: critical role of blood proteins. Biochim. Biophys. Acta 1281, 31-37.

Oupicky´ et al. (13) Dash, P. R. (1998) The Development of Polymer-based Synthetic Vectors for Use in Gene Therapy. Ph.D. Thesis, University of Birmingham. (14) Lasic, D. D., and Needham, D. (1995) The “stealth” liposome: A prototypical biomaterial. Chem. Rev. 95, 26012628. (15) Moghimi, S. M. (1995) Mechanisms regulating body distribution of nanospheres conditioned with pluronic and tetronic block copolymers. Adv. Drug Delivery Rev. 16, 183193. (16) Manning, G. S. (1969) Limiting laws and counterion condensation in polyelectrolyte solutions. I. Colligative properties. J. Chem. Phys. 51, 924-938. (17) Wolfert, M. A. Self-assembling systems based on synthetic polymers for gene delivery. (1998) Ph.D. Thesis, University of Birmingham. (18) Bakeev, K. N., Izumrudov, V. A., Kuchanov, S. I., Zezin, A. B., and Kabanov, V. A. (1992) Kinetics and Mechanism of Interpolyelectrolyte Exchange and Addition Reactions. Macromolecules 25, 4249-4254. (19) Takakura, Y., and Hashida, M. (1998) Pharmacokinetics of macromolecules and synthetic gene delivery systems. In Self-assembling complexes for gene delivery: From laboratory to clinical trial (A. V. Kabanov, P. L. Felgner, and L. W. Seymour, Eds.), pp 295-306, John Wiley & Sons, New York. (20) Lasic, D. D. (1997) Recent developments in medical applications of liposomes: sterically stabilized liposomes in cancer therapy and gene delivery in vivo. J. Controlled Release 48, 203-222.

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