Method for producing a polyglycerol nanogel for the encapsulation and release of biologically active substances

Abstract

A method for producing a polyglycerol nanogel is disclosed, the method comprising the following steps: Mixing an aqueous solution of first polyglycerol macromonomers, which are modified with a first reactive group, with an aqueous solution of second polyglycerol macromonomers, which are modified with a second reactive group, wherein the first reactive group and the second reactive group can react with each other forming a chemical bond; transferring the mixture into an organic non-solvent; and precipitation of a polyglycerol nanogel consisting of first polyglycerol macromonomers and second polyglycerol macromonomers which are covalently bound to each other. According to an aspect of the invention, the method is characterized in that the organic non-solvent is miscible with water and in that the method is carried out without adding surface-active substances.

Claims

1. A method for producing a polyglycerol nanogel, comprising the following steps: mixing an aqueous solution of first polyglycerol macromonomers, which are modified with a first reactive group, with an aqueous solution of second polyglycerol macromonomers, which are modified with a second reactive group, wherein the first reactive group and the second reactive group can react with each other forming a covalent bond, transferring the mixture of both aqueous solutions into an organic non-solvent being miscible with water, allowing diffusion of water from the aqueous solutions into the organic non-solvent, thereby increasing the concentration of the first polyglycerol macromonomers and the second polyglycerol macromonomers, precipitation of a hydrophilic polyglycerol nanogel consisting of first polyglycerol macromonomers and second polyglycerol macromonomers which are covalently bound to each other, wherein the covalent bond between the first polyglycerol macromonomers and the second polyglycerol macromonomers is established by a reaction of the first reactive group and the second reactive group, which takes place spontaneously only as a consequence of transferring the mixture of both aqueous solutions into the organic non-solvent and increasing the concentration of the first polyglycerol macromonomers and the second polyglycerol macromonomers in the aqueous solutions, wherein the method is carried out at a temperature of between 0° C. to 25° C. without adding a surface-active substance.

2. The method according to claim 1, wherein method is carried out without using ultrasound.

3. The method according to claim 1, wherein the first polyglycerol macromonomers are present in a first concentration and the second polyglycerol macromonomers are present in a second concentration, wherein the first and the second concentration lie in a range of 0.1 to 30 mg/ml independently of each other.

4. The method according to claim 1, wherein the polyglycerol nanogel is transferred into an aqueous phase after precipitation.

5. The method according to claim 1, wherein the precipitation is carried out in the presence of a labile substance.

6. The method according to claim 1, wherein the first polyglycerol macromonomers and/or the second polyglycerol macromonomers contain a pH-labile group, which is still present in the polyglycerol nanogel formed.

7. The method according to claim 6, wherein the pH-labile group is selected from the group consisting of acetals, ketals, enol ethers, esters, amides, hydrazones, hydrazides, oximes, maleic acid derivatives, carbamates, hydroxylamine imines, iminium compounds, enamines, silyl ethers and silyl enol ethers.

8. The method according to claim 1, wherein the first polyglycerol macromonomers and/or the second polyglycerol macromonomers have a terminal modification of the type —R—R′, which is covalently bound to a linear or branched polyglycerol structure of the first polyglycerol macromonomers and/or of the second polyglycerol macromonomers, wherein R is a pH-labile group and R′ is a bioorthogonal terminal group that can undergo a reaction according to click chemistry.

9. The method according to claim 1, wherein the first reactive group is an alkyne group and the second reactive group is an azide group.

10. The method according to claim 1, wherein the precipitation of the polyglycerol nanogel takes place without adding a compound containing copper.

11. The method according to claim 5, wherein the labile substance is a peptide, a protein, DNA, RNA and/or a hormone.

12. The method according to claim 1, wherein the precipitation is carried out in the presence of a biologically active substance.

Description

BRIEF DESCRIPTION OF THE DRAWINGS

(1) FIG. 1 shows a schematic illustration of an exemplary embodiment of a method for producing a polyglycerol nanogel;

(2) FIG. 2A shows a graphic illustration of the decomposition of a pH-labile polyglycerol nanogel at different pH values;

(3) FIG. 2B shows a graphic illustration of the L-asparaginase-II release from a pH-labile polyglycerol nanogel at different pH values;

(4) FIG. 3 shows a schematic illustration of an exemplary embodiment of a method for producing a polyglycerol nanogel with the simultaneous embedding of a protein;

(5) FIG. 4A shows second derivatives of absorption spectra of the model protein L-asparaginase II in the amide-I region after encapsulation and degradation of the nanogels;

(6) FIG. 4B shows second derivatives of absorption spectra of the model protein L-asparaginase II in the amide-II region after encapsulation and degradation of the nanogels;

(7) FIG. 5 shows a graphic illustration of the enzyme activity of the model protein L-asparaginase II after encapsulation and release from the nanogels and

(8) FIG. 6 shows a flow chart to illustrate the individual steps of the method of an exemplary embodiment.

DETAILED DESCRIPTION

(9) FIG. 1 shows a schematic illustration of a method for producing a polyglycerol nanogel. First polyglycerol macromonomers (1) are here mixed with second polyglycerol macromonomers (2) in low concentration. The first macromonomers carry in each case five propargyloxybenzacetal groups (10) as first reactive group or alkyne group. The second macromonomers carry in each case five azide groups (20) as second reactive groups or azide groups. The alkyne unit of the propargyloxybenzacetal groups (10) here is terminally located. This alkyne unit as such is stable. The azide groups (20) of the second macromonomers (2) are likewise terminally located. The mixture of first macromonomers (1) and second macromonomers (2) is present in an aqueous phase and is introduced into acetone (4) as organic non-solvent. Subsequently, there is a diffusion (5) of the water from the aqueous phase (3) into the acetone (4). Thereby, the concentration of the first macromonomers (1) and the second macromonomers (2) in the aqueous phase (3) increases. Thereby, the concentration of the macromonomers (1, 2) critical for initiating the reaction is reached, so that there is a reaction between the alkyne groups (10) and the azide groups (20). Covalent bonds form between the first macromonomers (1) and the second macromonomers (2), resulting in a formation of nanoparticles (6). In this manner, a polyglycerol nanogel 6 forms by swelling. This reaction, schematically illustrated in FIG. 1, will now be illustrated in more detail with the help of a specific exemplary embodiment.

(10) Producing p-propargyloxy-benzaldehyde

(11) 2.00 g 4-hydroxybenzaldehyde (16.38 mmol) were dissolved in 50 ml acetone. 15.15 g potassium carbonate (109.62 mmol) were added, and the suspension was stirred for 30 minutes under reflux. After the solution had cooled down to room temperature, 2.12 ml propargylic bromide (19.10 mmol) were added over a period of 2.5 hours. Thereafter, the suspension was heated for 1.5 hours under reflux. The suspension was then filtered and the solvent of the filtrate was evaporated in vacuum. 50 ml dichloromethane were added and the organic phase thereafter was washed twice with 20 ml 1 M caustic soda and once with 20 ml water. The organic phase was dried over magnesium sulfate. The raw product was purified by means of Kugelrohr distillation. A white crystalline solid was obtained. As the results of an analysis show, the reaction was complete with a yield of about 80%.

(12) Producing p-propargyloxy-benzdimethylacetal

(13) 1 g p-propargyloxy-benzaldehyde (6.29 mmol) was dissolved in 10 ml trimethyl orthoformate. Thereafter, 108 mg dry p-toluenesulfonic acid (0.629 mmol) were added. The reaction mixture was heated to 65° C. for one hour and thereafter quenched by adding 25 ml saturated sodium hydrogen carbonate solution. The mixture was extracted three times with, in each case, 25 ml ethyl acetate, and the fractions were merged and dried over sodium sulfate. The ethyl acetate was evaporated and p-propargyloxy-benzdimethylacetal was obtained as a yellowish oil at a yield of 90%.

(14) Producing 3-azidopropyl 4-toluenesulphonate

(15) In a two-necked flask, equipped with a dropping funnel and a stir bar, 3-azidopropanol (3.187 g, 31.52 mmol) and triethylamine (9.65 ml, 69.3 mmol, 2.2 molar equivalents (eq.)) were dissolved in dichloromethane (30 ml). After cooling down to 4° C. by means of an aqueous ice bath, a solution of tosyl chloride (6.61 g, 34.7 mmol, 1.1 eq.) in dichloromethane (30 ml) was added in drops within 10 min. The reaction was carried out further for 6 h at room temperature and the reaction process was monitored by IR-spectroscopy and thin-layer chromatography. After filtering out the formed salt, the filtrate was concentrated, taken up in dichloromethane (50 ml) and washed with a 1M NH.sub.4Cl-solution (20 ml). The organic phase was dried over MgSO.sub.4 and concentrated. The substance was ultimately purified by column chromatography on silica gel (hexane/ethyl acetate, 3:1) in order to obtain 3-azidopropyl 4-toluenesulphonate (6.44 g, 80%) as a yellow liquid. R.sub.f: 0.64 (hexane/ethyl acetate, 3:1).

(16) Producing 4-(3-azidopropoxy)-2-methoxybenzaldehyde

(17) K.sub.2CO.sub.3 (16.50 g, 119.5 mmol, 5 eq.) was added into a solution of 4-hydroxy-2-methoxybenzaldehyde (3.66 g, 23.9 mmol, 1.2 eq.) and 3-azidopropyl 4-toluenesulphonate (5.08 g, 19.91 mmol, 1 eq.) in acetone (70 ml). The reaction was carried out for 16 h under reflux and, after cooling down to room temperature, salt was filtered out and the filtrate was concentrated. After the uptake of the residue in dichloromethane (20 ml), the organic phase was washed with water (20 ml), dried over MgSO.sub.4 and then concentrated. The substance was ultimately purified by column chromatography on silica gel (hexane/ethyl acetate, 3:1) in order to obtain 4-(3-azidopropoxy)-2-methoxybenzaldehyde (3.98 g, 85%) as a transparent liquid. R.sub.f: 0.54 (hexane/ethyl acetate, 3:1).

(18) Producing 4-(3-azidopropoxyl)benzaldehyde

(19) K.sub.2CO.sub.3 (13.55 g, 98.05 mmol, 5 eq.) was added into a solution of 4-hydroxybenzaldehyde (2.39 g, 19.61 mmol, 1.2 eq.) and 3-azidopropyl 4-toluenesulphonate (5.08 g, 19.91 mmol, 1 eq.) in acetone (70 ml). The reaction was carried out for 16 h under reflux and, after cooling down to room temperature, salt was filtered out and the filtrate was concentrated. After the uptake of the residue in dichloromethane (20 ml), the organic phase was washed with water (20 ml), dried over MgSO.sub.4 and then concentrated. The substance was ultimately purified by column chromatography on silica gel (hexane/ethyl acetate, 2:1) in order to obtain 4-(3-azidopropoxyl)benzaldehyde (3.98 g, 91%) as transparent liquid. R.sub.f: 0.67 (hexane/ethyl acetate, 2:1).

(20) Producing 4-(3-azidopropoxy)-2-methoxybenzaldehyde-dimethylacetal

(21) Trimethyl orthoformate (5.20 ml, 49.02 mmol, 5 eq.) and PTSA without water (169 mg, 0.98 mmol) were added into a solution of 4-azidopropoxy-2-methoxybenzaldehyde (2.00 g, 9.80 mmol) in degassed methanol (20 ml). The reaction was carried out for 20 h under reflux and, after cooling down, quenched with aqueous ammonia (0.5 ml). After concentrating the solution, ethyl acetate (50 ml) was added and the solution was washed with water (50 ml). After drying the organic phase over MgSO.sub.4, the organic phase was concentrated in order to obtain 4-(3-azidopropoxy)-2-methoxybenzaldehyde dimethylacetal (2.32 g, 95%) as a yellow liquid. R.sub.f. 0.84 (hexane/ethyl acetate, 3:1).

(22) Producing 4-(3-azidopropoxyl)benzaldehyde dimethylacetal

(23) Trimethyl orthoformate (4.64 ml, 43.72 mmol, 5 eq.) and PTSA without water (169 mg, 0.98 mmol) were added to a solution of 4-azidopropoxybenzaldehyde (2.00 g, 8.51 mmol) in degassed methanol (20 ml). The reaction was carried out for 20 h under reflux and, after cooling down, quenched with aqueous ammonia (0.5 ml). After concentrating the solution, ethyl acetate (50 ml) was added and the solution was washed with water (50 ml). After drying the organic phase over MgSO.sub.4, the organic phase was concentrated in order to obtain 4-(3-azidopropoxy)-2-methoxybenzaldehyde dimethylacetal (2.38 g, 97%) as a yellow liquid. R.sub.f: 0.57 (hexane/ethyl acetate, 4:1).

(24) Producing hPG.sub.7,7, which is functionalized with 7 p-azidopropoxy-methoxybenzacetal units (hPG.sub.7,7-7-p-azidopropoxy-benzacetal)

(25) 1 g hPG.sub.7,7 (0.13 mmol) and 365.3 mg of 4-(3-azidopropoxy)-2-methoxybenzaldehyde dimethylacetal (1.3 mmol) were dissolved in 4 ml n-methyl-2-pyrrolidone, and 22 mg p-toluenesulfonic acid without water (0.13 mmol) were added. The reaction mixture was held at room temperature for three hours and the condensed methanol was removed from the reaction mixture by cryo-distillation. The reaction was quenched by adding 1 ml aqueous ammonia. The n-methyl-2-pyrrolidone was evaporated by cryo-distillation, and the remaining residue was again dissolved in basified water (basified water contains 0.05 wt % aqueous ammonia). The solution was dialyzed in basified water for five days, wherein the dialysate was changed every three hours. After freeze-drying, hPG.sub.7,7 functionalized with seven p-azidopropoxy-methoxybenzacetal units was obtained as a viscous wax. The reaction took place with a conversion of 71% and a yield of 78%.

(26) Producing hPG.sub.7,7, which is functionalized with 7 p-azidopropoxybenzacetal units (hPG.sub.7,7-7-p-azidopropoxy-benzacetal)

(27) 1 g hPG.sub.7,7 (0.13 mmol) and 326.3 mg of 4-(3-azidopropoxyl)benzaldehyde dimethylacetal (1.3 mmol) were dissolved in 4 ml n-methyl-2-pyrrolidone, and 22 mg p-toluenesulfonic acid without water (0.13 mmol) were added. The reaction mixture was held at room temperature for three hours and the condensed methanol was removed from the reaction mixture by cryo-distillation. The reaction was quenched by adding 1 ml aqueous ammonia. The n-methyl-2-pyrrolidone was evaporated by cryo-distillation, and the remaining residue was again dissolved in basified water (basified water contains 0.05 wt % aqueous ammonia). The solution was dialyzed in basified water for five days, wherein the dialysate was changed every three hours. After freeze-drying, hPG.sub.7,7 functionalized with seven p-azidopropoxy-methoxybenzacetal units was obtained as a viscous wax. The reaction took place with a conversion of 69% and a yield of 83%.

(28) Producing hPG7,7, which is functionalized with 7p-propargyloxy-benzacetal units (hPG.sub.7,7-7-p-propargyloxy-benzacetal)

(29) 1 g hPG.sub.7,7 (0.13 mmol) and 250 mg p-propargyloxy-benzdimethylacetal (1.3 mmol) were dissolved in 4 ml n-methyl-2-pyrrolidone, and 22 mg p-toluenesulfonic acid without water (0.13 mmol) were added. The reaction mixture was heated to 120° C. for three hours and the condensed methanol was removed from the reaction mixture by cryo-distillation. After cooling down to room temperature, the reaction was quenched by adding 1 ml aqueous ammonia. The n-methyl-2-pyrrolidone was evaporated by cryo-distillation, and the remaining residue was again dissolved in basified water (basified water contains 0.05 wt % aqueous ammonia). The solution was dialyzed in basified water for two hours, wherein the dialysate was changed every three hours. After freeze-drying, hPG.sub.7,7 functionalized with seven p-propargyloxy-benzacetal units was obtained as a viscous wax. The reaction took place with a conversion of 70% and a yield of 80%.

(30) Producing homobifunctional 1PG.sub.5-biscyclooctyne

(31) P(EEGE).sub.5-Br (4 g, 0.8 mmol) was dissolved in tetrahydrofuran (20 ml) and the solution was cooled down to 4° C. by means of an ice bath. After the addition of triethylamine (2.23 ml, 16 mmol) and mesyl chloride (0.62 ml, 8 mmol), the reaction was carried out for one day at room temperature. After salt filtration, the polymer was purified by means of dialysis in THF. Subsequently, the polymer (2 g, 0.4 mmol) was taken up in DMF (20 ml) and caused to react with NaN.sub.3 (520 mg, 8 mmol) at 80° C. for three days, the salt was filtered out, the protective groups were deprotected by means of ethanolic HCL (1 vol. %) and then purified by means of a three-day dialysis. Thereafter, the azided polymer (1.8 g, 0.36 mmol) was reduced for three days in a water-THF mixture (10 ml, 1:1) by triphenylphosphine (377.3 mg, 1.44 mmol). The diamine formed (1 g, 0.2 mmol) was ultimately with BCN (138.6 mg, 0.44 mmol) in dichloromethane (10 ml) with triethylamine (0.88 mmol, 123 μL) as base. The polymer was processed by means of a three-day dialysis in a water-acetone mixture (1:1) in order to obtain 1PG.sub.5-biscyclooctyne.

(32) Producing a Polyglycerol Nanogel by Nanoprecipitation

(33) 5 mg hPG.sub.7,7-7-p-propargyloxy-benzacetal (0.6 μmol) and 7 mg hPG.sub.7,7[N.sub.3].sub.7 (0.9 μmol) were dissolved in 0.5 ml purified deionized water, independently of one another. Tris(3-hydroxypropyltriazolylmethyl) (THPTA), copper sulfate and sodium ascorbate were added to the hPG.sub.7,7-7-p-propargyloxy-benzacetal solution in precisely that order. The solutions were cooled down to 4° C. The solutions were then mixed with each other and quickly added to 20 ml acetone, which was stirred by a magnetic stirrer. This now led to the precipitation of polyglycerol nanoparticles, which were visible as bluish-appearing dispersions. The particle size was detected by means of dynamic light scattering (DLS). After three hours, the gel formation reaction was quenched by adding an excess of 50 mg azidoglycerol (427 μmol). After 12 hours, 20 ml purified deionized water were added, and the acetone was evaporated in order to obtain a bluish-shimmering nanogel dispersion in water. The nanogel was separated from the aqueous phase by centrifugation at 4000 rpm and washed five times with purified deionized water. The nanogel was thereafter characterized by means of DLS, optical microscopy and transmission electron microscopy.

(34) Embedding Proteins, Including an L-Asparaginase II, a Bovine Serum Albumin, the Antibody IgG and a Lysozyme, in the Nanogel

(35) 2 mg hPG.sub.7,7-7-p-propargyloxy-benzacetal (0.2 μmol) and 3 mg hPG.sub.7,7[N.sub.3].sub.7 (0.3 μmol) were dissolved in 0.5 ml purified deionized water, independently of one another. THPTA and copper acetate were added to the hPG.sub.7,7-7-p-propargyloxy-benzacetal solution. Furthermore, the protein was added to the hPG.sub.7,7[N.sub.3].sub.7 solution. The solutions were cooled down to 4° C. Thereafter, the solutions were mixed and quickly added to 20 ml acetone, which was stirred on a magnetic stirrer. After three hours, the gel formation reaction was quenched by adding an excess of 50 mg azidoglycerol (427 μmol). After 12 hours, the nanogel was separated from the liquid phase by means of centrifugation at 4000 rpm and washed five times with purified deionized water.

(36) Embedding Proteins, Including an L-Asparaginase II, a Bovine Serum Albumin, the Antibody IgG and a Lysozyme, in the Nanogel by Copper-Free Click Chemistry

(37) 2 mg hPG.sub.7,7-7-p-azidopropoxy-benzacetal (0.2 μmol) and 4 mg 1PG.sub.5-biscyclooctyne (0.6 μmol) were dissolved in 0.5 ml purified deionized water, independently of one another. Furthermore, the protein was added to the hPG.sub.7,7-7-p-azidopropoxy-benzacetal solution. The solutions were cooled down to 4° C. Thereafter, the solutions were mixed and quickly added to 20 ml acetone, which was stirred on a magnetic stirrer. After three hours, the gel formation reaction was quenched by adding an excess of 50 mg azidoglycerol (427 μmol). After 12 hours, the nanogel was separated from the liquid phase by means of centrifugation at 4000 rpm and washed five times with purified deionized water.

(38) Determining the Size of the Nanogel Particles

(39) As the subsequent Table 1 shows, the size of the polyglycerol nanogels obtained depends on the starting concentration of the macromonomers employed.

(40) TABLE-US-00001 TABLE 1 Dependency of the size of the formed polyglycerol nanoparticles on the starting concentration of the macromonomers employed c (macromonomer)/ d/nm PDI d/nm PDI (mg/ml) (in acetone) (in acetone) (in water) (in water) 12 580 0.03 820 0.07 6 440 0.02 610 0.03 3 310 0.06 430 0.08 1.5 102 0.04 145 0.07

(41) The lower the starting concentration of the macromonomers, the smaller the diameter of the nanogels formed. Here, in Table 1, c is the concentration, d the diameter and PDI the polydispersity. Whereas with a macromonomer concentration of 1.5 mg/ml, polyglycerol nanogels having a diameter of about 100 nm in acetone were obtained, this diameter increased to just under 600 nm at a starting concentration of 12 mg/ml macromonomers. After transferring the nanogels into water, there was a further swelling of the nanogels due to the integration of water molecules. Thereby, the measured diameter of the nanogels also increased.

(42) Polydispersity is a measure for the scattering of the particle sizes and indicates that the nanogels have a very narrow size distribution. When the particles are transferred from acetone into water, the particle sizes increase. This suggests the swelling of the particles.

(43) Determining the Nanogel Degradation Kinetics

(44) Nanogel dispersions were incubated at 37° C. and at different pH values. After different times of incubation, the nanogels were cooled down to 4° C., neutralized and separated from degraded fragments by means of a 5-minute centrifugation at 4000 rpm. Thereafter, the UV-absorption of the degraded fragments located in the supernatant solution was observed at 350 nm. During the degradation, more and more degraded fragments go into solution, causing the absorption to rise. The corresponding result of this experiment is illustrated in FIG. 2A. One can see well that at a pH value of 7.4 the integrity of the polyglycerol nanoparticles is not affected. Rather, the nanoparticles remain stable at this pH value. Only when the pH value is lowered, there is a degradation of the polyglycerol nanoparticles, for then the benzacetal compound contained in the nanoparticles formed is broken up. As is evident from FIG. 2A, the degradation of the polyglycerol nanoparticles furnished with the benzacetal groups goes faster, the lower the set pH value. At a pH value of 4, the polyglycerol nanoparticles are completely degraded in less than five hours.

(45) The complete nanogel degradation was confirmed by means of DLS-size measurements and .sup.1H-NMR-spectroscopic measurements.

(46) Controlled Release of Asparaginase Initiated by pH-Dependent Nanogel Degradation

(47) The polyglycerol nanogel loaded with L-asparaginase II according to the protocol explained above (10 mg/ml nanogel and 0.5 mg/ml L-asparaginase II) was acidified with hydrochloric acid to pH 4 or pH 5, respectively. The samples were incubated at room temperature (25±2° C.) under a slight motion (300 rpm). Individual samples were collected over the course of three days and, thereafter, analyzed by means of size exclusion high-performance chromatography (SEC-HPLC). In order to stop the nanogel degradation, the samples were neutralized with 0.1 M caustic potash prior to the SEC-HPLC. For the SEC-HPLC, 50 μl of the neutralized samples were injected into a HPLC equipped with a TSKgel G40000 PWXL column (300×7.8 mm, 10 μm particle size). An isocratic elution with a buffer of 20 mM NaHPO4, 150 mM NaCl and 0.003 mM NaN3 (pH 7.4) at a flow rate of 0.4 ml/min took place. The concentration of the L-asparaginase II was determined by means of UV-absorption at 280 nm and fluorescence detection (excitation with 295 nm and emission at 348 nm).

(48) FIG. 2B shows the corresponding results of this examination. The obtained HPLC-chromatograms show that L-asparaginase II can be detected by means of the intrinsic tryptophan fluorescence without being impaired by polyglycerol or polyglycerol fragments. In the intact polyglycerol nanogel loaded with L-asparaginase II, no free L-asparaginase could be detected. Hence, the loading or embedding efficiency was at roughly 100%. By means of the chromatograms the percentage of the released L-asparaginase II was determined and plotted against time in FIG. 2B. As expected, the L-asparaginase II was released faster at pH 4 (square data points) than at pH 5 (round data points), since at a lower pH value, a faster nanogel degradation takes place.

(49) FIG. 3 shows a schematic illustration of the previously explained embedding of a protein 7 in the formed nanoparticles or the formed nanogel 6. A joint nanoprecipitation of polyglycerol macromonomers and a protein leads to an in situ gel formation, wherein the also precipitated proteins are embedded inside of the nanogel in their native form. When the protein-loaded polyglycerol nanogel 8 is transferred into an acidic medium, it comes to the degradation and protein release 9. One can make use of this fact, because low pH values predominate in inflamed or tumor tissue of an organism. In this manner it is possible to place a nanogel loaded with a therapeutic protein into an organism, wherein the therapeutic protein is released only at its site of action (namely the inflamed tissue with a low pH value). Instead of embedding or integration, one can also speak of the encapsulation of a protein or of another active substance.

(50) As already mentioned, harsh reaction conditions are not suited to maintain a protein or another labile substance in its native and active form. In order to prove that proteins in the native form can be embedded in the formed nanogel with the method presently introduced, the secondary structure of L-asparaginase II, as an exemplarily embedded enzyme, was detected after encapsulation in and release from the nanogel. This was done by means of the Fourier transformation infrared spectroscopy (FTIR), wherein measuring was done in the form of attenuated total reflection measurements (ATR). The employed ATR cell was held at a constant temperature of 25° C. 25 μl of a sample were given onto the ATR cell under dry nitrogen and measured against PBS buffer with a pH value of 5 or against water as control. 120 scans for each experiment at a resolution of 4 cm.sup.−1 were carried out, wherein a water vapor correction took place. The second derivatives of the obtained absorption spectra were used for further data analysis.

(51) The result of these FTIR-examinations is illustrated in FIGS. 4A and 4B. Here, FIG. 4A shows absorptions observed in the region of the amide-I band, whereas absorptions in the region of the amide-II band are illustrated in FIG. 4B. The amid-I band is sensitive to C═O stretching vibrations and is well suited to determine the secondary structure of a protein.

(52) The second derivative of the spectrum of native L-asparaginase II dissolved in water (freshly prepared) is illustrated as a dashed line in FIGS. 4A and 4B. The dotted line shows the second derivative of an absorption spectrum of L-asparaginase II which was stored for seven days in PBS buffer at a pH value of 5. The continuous line finally shows the second derivative of an absorption spectrum of L-asparaginase II released from polyglycerol nanogels after seven days.

(53) As can be seen from FIG. 4A, neither the band intensities nor the wave numbers of the three different samples differ from each other significantly. Rather, merely a slight shift of the band characteristic for a α-helical secondary structure at 1660 cm.sup.−1 by 1 cm.sup.−1 to lower wave numbers can be observed. With the band characteristic for a β-sheet secondary structure at approximately 1635 cm.sup.−1 for the native protein, no shift can be observed. Altogether, however, these shifts lie within the range of error due to the measuring technique. Thus, it is to be assumed that the secondary structure is not changed by an encapsulation of the L-asparaginase II in the polyglycerol nanoparticles.

(54) This finding is also confirmed by an analysis of the amide-II band. The amide-II band provides information about the N—H bending vibrations and the C—N stretching vibrations. Herein, when storing the L-asparaginase II in water or encapsulating this enzyme in the polyglycerol nanogels, likewise no significant band shift (see FIG. 4B) can be detected.

(55) The observed absorptions in the amide-I and amide-II region are illustrated in the subsequent Table 2.

(56) TABLE-US-00002 TABLE 2 Absorptions of the L-asparaginase in the amide-I and amide-II region, determined with the help of the second derivatives of corresponding absorption spectra Absorptions in Absorptions in the region of the the region of the amide-I band/cm.sup.−1 amide-II band/cm.sup.−1 L-asparaginase II in water 1660.5 1634.4 1549.6 (freshly prepared) L-asparaginase II in PBS 1659.5 1635.4 1550.5 pH 5.0 (7 d storage) L-asparaginase II after 1659.0 1637.3 1547.6 release (7 d storage)
Determining the Asparaginase Activity

(57) The activity of the L-asparaginase II was determined by means of Neβler's ammonia quantification.

(58) In order to carry out the asparaginase activity tests, 50 μl L-asparaginase II, 100 μl Tris-HCl with a pH value of 8.6 and 850 μl L-asparagine monohydrate buffer solution were incubated at 37° C. for 10 minutes. After the addition of 50 μl of a 1.5 M solution of trichloroacetic acid and subsequent centrifugation, 100 μl of the supernatant were added to Neβler's reagent. After 10 minutes, the optical density at 436 nm was determined and compared to a calibration curve as well as corrected by the total enzyme content. Calculating the enzyme activity then took place according to the following formula:

(59) $U\text{/}\mathrm{mg}=\frac{\mathrm{\mu mol}\mathrm{released}\mathrm{ammonia}}{10\min \times \mathrm{mg}\mathrm{enzyme}\mathrm{in}\mathrm{the}\mathrm{reaction}}$

(60) A unit (1 U) of the detected enzyme activity here corresponds to the released amount of ammonia in micromol per 10 minutes from asparagine as substrate.

(61) The result of this quantification showed an activity of the freshly prepared asparaginase solution of 98.6 U/mg, which coincides with the data specified by the manufacturer (98.2 U/mg). When transferring L-asparaginase II into a PBS buffer with pH 5.0, the activity decreased by 10% to 86.1 U/mg. Storing the L-asparaginase II in the buffer over 7 days, reduces the activity by another 2.5% to 86.2 U/mg. An identical value, within the limits of measurement accuracy, could be detected for the activity of the L-asparaginase II which was encapsulated in a polyglycerol nanogel and released again, after its release. The measurement results are illustrated in the subsequent Table 3, together with the respectively detected standard deviations (SD).

(62) TABLE-US-00003 TABLE 3 Enzyme activity of the L-asparaginase II Specific Specific activity/ activity/ SD/ SD/ (U/mg) % (U/mg) % L-asparaginase II in water 98.6 100 4.4 4.4 (freshly prepared) L-asparaginase II in PBS 89.1 90.8 0.1 0.1 pH 5.0 (freshly prepared) L-asparaginase II in PBS 86.1 87.3 0.7 0.8 pH 5.0 (7 d storage) L-asparaginase II after 86.2 87.5 0.9 1 release (7 d storage)

(63) A corresponding graphic illustration of the specific enzyme activity can be seen in FIG. 5. Here, the enzyme activity of the freshly prepared native L-asparaginase II in water was set to 100%. It is also evident from this graphic illustration that the L-asparaginase II, encapsulated in a polyglycerol nanogel, has an activity after its release that corresponds to the activity of L-asparaginase II dissolved in buffer. This confirms the results established by the FTIR-measurements. So, L-asparaginase II maintains its native secondary structure even after encapsulation in a polyglycerol nanogel. Furthermore, an encapsulation of L-asparaginase II in polyglycerol nanogels does not reduce the enzyme activity.

(64) FIG. 6 shows a flow chart, which serves to illustrate an exemplary embodiment of the method claimed.

(65) In a step 100 preceding the actual method itself, a synthesis of first polyglycerol macromonomers and of second polyglycerol macromonomers takes place.

(66) In a first step of the process 110, an active merging of the first and the second polyglycerol macromonomers, a labile substance (a protein or enzyme, for instance) and, if required, a catalyst, which catalyzes the click reaction taking place later, takes place.

(67) In a second step of the process 120, an active transferring of the merged substances into an organic non-solvent takes place. “Active” here means that an operator executes the corresponding steps by his or her willful actions.

(68) In a third step of the process 130, there are two spontaneous chemical reactions, so that one can also speak of a double spontaneity. On the one hand, a spontaneous precipitation of the merged substances takes place with the spontaneous formation of nano-aggregates. On the other hand, a spontaneous cross-linking of the nano-aggregates takes place by a click reaction (forming covalent bonds between the first polyglycerol macromonomers and the second polyglycerol macromonomers).

(69) Afterwards, the cross-linked nanoparticles are actively transferred into an aqueous phase in a forth step of the process 140.

(70) Thereafter, in a fifth step of the process 150, a spontaneous swelling of the cross-linked particles takes place in aqueous phase.