Effect of Pendant Group Structure on the Hydrolytic Stability of

Apr 3, 2012 - *E-mail: [email protected]; [email protected]. ... The DET pendant group is positively charged at pH 7, but in ... Citat...
0 downloads 0 Views 3MB Size
Download PDF
Article pubs.acs.org/Biomac

Effect of Pendant Group Structure on the Hydrolytic Stability of Polyaspartamide Polymers under Physiological Conditions Yijie Lu,† Mokit Chau,† A. J. Boyle,‡ Peng Liu,† Ansgar Niehoff,†,§ Dirk Weinrich,†,∥ Raymond M. Reilly,*,‡,§ and Mitchell A. Winnik*,† †

Department of Chemistry, University of Toronto, 80 St. George Street, Toronto, Ontario, Canada M5S 3H6 Leslie Dan Faculty of Pharmacy, University of Toronto, 144 College Street, Toronto, Ontario, Canada M5S 3M2 § Toronto General Research Institute, University Health Network, Toronto, ON, Canada M5G 2C4 ‡

S Supporting Information *

ABSTRACT: We describe the synthesis of metal chelating polymers based on polyaspartamide and polyglutamide backbones as carriers for 111In in radioimmunoconjugates. These polymers [PAsp(DTPA), PGlu(DTPA)] have a biotin end group and diethylenetriaminepentaacetic acid (DTPA) chelators attached to the primary amines of the diethylenetriamine (DET) pendant groups of biotin-poly{N′-[N-(2-aminoethyl)-2-aminoethyl]aspartamide} [PAsp(DET)] and of biotin-poly{N′-[N-(2-aminoethyl)-2-aminoethyl]glutamide} [PGlu(DET)]. Like Asn-containing proteins and polypeptides, polyaspartamides undergo uncatalyzed degradation under model physiological conditions (10 mM phosphate buffer, pH 7.4, 150 mM NaCl). We studied the uncatalyzed degradation of the polyaspartamide polymers by size exclusion chromatography and found that the degradation rate was sensitive to the nature of the pendant groups. The metal-free polymer underwent somewhat slower degradation than the corresponding polymers in which the DTPA groups were saturated with Eu3+ or In3+, but even after 14 days, substantial fractions of the polymers survived. We conclude that these polymers undergo negligible degradation on the time scale (24−48 h) of radioimmunotherapy treatment of tumors with 111In. From a mechanistic perspective, we note that these degradation rates are on the order of the deamidation rates reported [J. Peptide Res. 2004, 63, 426] for Asn-containing pentapeptides, with half-times on the order of 10 days, but much slower than the rapid decay (hours) reported recently [Biomaterials 2010, 31, 3707] for poly{N′[N-(2-aminoethyl)-2-aminoethyl]aspartamide} itself. This variation in degradation rate can be explained in terms of the influence of positive charges on the pendant group enhancing the acidity of the side-chain amide nitrogen of the aspartamide repeat unit. The DET pendant group is positively charged at pH 7, but in indium-loaded PAsp(DTPA) this charge is offset by the net negative charge of the DTPA-In complex.

■

INTRODUCTION We are interested in developing metal-chelating polymers (MCPs) as carriers for radionuclides in targeted delivery to cancer cells, particularly HER2+ breast cancer cells. The targeted delivery of subcellular range Auger electron-emitting radionuclides such as 111In to tumor cells is an attractive option in cancer therapy because it restricts the killing to individual cancer cells, leaving normal cells unharmed. The release of Auger electrons in the nucleus causes the double strands of DNA to break,1 resulting in cell death. For patients with HER2 amplified breast cancer, HER2 receptors are overexpressed on the surface of the cancer cells. The antibody trastuzumab (Herceptin) targets HER2 receptors and is itself used as a therapeutic agent, alone or as an adjuvant.2 Costantini et al.3 have shown that trastuzumab labeled with a diethylenetriaminepentaacetic acid (DTPA) as a metal chelator and with several copies of a nuclear localization peptide sequence (NLS) can inhibit the growth of HER2 amplified breast cancer tumors in © 2012 American Chemical Society

mice. Our approach is to develop end-functional MCPs that can carry multiple copies of 111In for delivery to a cancer cell, thereby greatly increasing the effective dose, while the endfunctional group is used for attachment to antibodies such as trastuzumab. For this purpose, we envision polymers with multiple copies of a metal-chelating group, such as DTPA as pendant groups. We have previously reported syntheses of MCPs based on a polyacrylamide backbone with pendant DTPA or DOTA (1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid) groups, and end-functionality for covalent attachment to antibodies.4,5 These polymers were employed in multiplexed immunoassays based on metal detection by inductively coupled plasma mass spectrometry. Here, for radioimmunotherapy Received: December 21, 2011 Revised: February 2, 2012 Published: April 3, 2012 1296

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

Figure 1. Strategy for rapid assembly of metal-chelating polymer (MCP) conjugates with the Fab fragment of trastuzumab (Herceptin). Biotin-endcapped polymers form a tight complex with a streptavidin (SAv) covalently linked to the Fab fragment. This conjugate is subsequently incubated with 111In to bind the radionuclide. SAv has four binding sites for biotin. Thus, more than one polymer may bind to one Fab-Sav conjugate. This strategy allows for rapid screening of different polymers for radioimmunotherapy treatment of Her2+ breast cancer tumors in animal models.

(RIT) applications, where issues such as pharmacokinetics and tissue distribution in vivo are important, we have designed a different strategy that allows for rapid scanning of a broad family of polymers of different lengths or with different pendant groups. We envision the synthesis of polymers with a terminal biotin group that can be rapidly attached to a streptavidinmodified Fab fragment of trastuzumab (SAv-Fab) via a biotinstreptavidin linkage. As a carrier of 111In, these MCP-Sav-Fab conjugates can be employed in various binding assays or pharmacokinetic studies. A cartoon depicting this strategy is shown in Figure 1. Experiments describing the synthesis and characterization of the Sav-Fab conjugate and the application of this strategy to a small library of polymers will be the subject of a separate publication.6 There may be an advantage to designing radioimmunoconjugate MCPs based on a polypeptide backbone. These polypeptides may be biodegradable or undergo spontaneous degradation under physiological conditions. Biodegradable polymers have many biomedical applications in regenerative medicine7 and in controlled drug release.8 For radioimmunotherapy, the requirements are different. Here the delivery vehicle should be stable over the time scale of tumor uptake (typically 24 to 48 h) or on the time scale of radioactive decay (t1/2 = 2.8 days for 111In). Degradation on longer time scales would facilitate elimination of the delivery vehicle components from the body. In this paper, we describe the synthesis and stability under model physiological conditions (37 °C, 10 mM phosphate buffer, pH 7.4, 150 mM NaCl) of biotin-polyaspartamides and biotin-polyglutamides with DTPA pendant groups, in the presence and absence of trivalent metal ions such as In3+. Our interest in these polymers was stimulated by a report from the Kataoka group9 that a particular polyaspartamide, poly{N′-[N(2-aminoethyl)-2-aminoethyl]aspartamide} PAsp(DET) was effective at gene transfection without the commonly encountered toxicity of cationic polymers.10 PAsp(DET) formed a tight polyion complex with plasmid DNA, and after uptake by living cells, pH sensitive protonation of the DET pendant groups caused endosomal release via the proton sponge effect and/or direct perturbation of endosomal membranes.11 Unlike other cationic polymers that have high cytotoxicity, PAsp(DET) polymers undergo spontaneous degradation under physiological conditions via anchimeric assistance (neighboring group participation), which alleviates their cumulative toxicity.10 This type of anchimeric assistance is well-known in the degradation of asparagine-containing proteins and peptides.12 This nonenzymatic degradation of Asn and the much slower degradation of Gln residues are thought to be nature’s “molecular clock” for establishing the useful lifetime of proteins in living systems.12

A surprising aspect of the results obtained by Kataoka and co-workers is the rapid time scale (hours) on which the PAsp(DET) polymer undergoes spontaneous degradation in aqueous buffer at pH 7.13 Polymers that degrade so quickly would not be useful for RIT applications. This rate is also much faster than the rate of nonenzymatic degradation of Asncontaining proteins and polypeptides. Pentapeptides with a central Asn that are thought to mimic the behavior of proteins in solution. Hundreds of these pentapeptides have been studied.12,14 The prominent reaction observed at pH 7 was the deamidation of Asn to aspartic acid. The fundamental conclusion drawn from these studies is that cleavage of the peptide (which we will refer to as “backbone degradation”) is much slower than deamidation and that deamidation rates have half-times commonly on the order of tens of days. Only when Asn is flanked by glycine are the deamidation half-times reduced to about 1 day, with much longer half-times found for all other amino acids. For this reason, we considered the idea that the rapid backbone degradation rates found to be useful by the Kataoka group were related in some important way to the nature of the pendant group on their polyaspartamide polymers. In the sections below we describe the synthesis of MCPs based upon pendant group transformation of biotin-poly(βbenzylaspartate) and biotin-poly(γ-benzylglutamate), followed by studies based on size-exclusion chromatography measurements of the spontaneous degradation at 37 °C in 10 mM phosphate buffer at pH 7.4 in the presence of 150 mM NaCl (model physiological conditions). We found, as expected, that the polyglutamide polymer, with and without trivalent metal ions, was stable to degradation over the 14 days of our study. In contrast, we found that the MCPs with the polyaspartamide backbone underwent hydrolytic degradation under these conditions, but with rates that were very sensitive to the nature of the pendant groups along the backbone as well as the charge density of these pendant groups. These rates were much slower than those reported for PAsp(DET).13 Encouragingly, we found that very little degradation occurred over a two-day period for the biotin-polyaspartamide polymer with a side chain consisting of a DET-DTPA-In in every repeat unit of the polymer. For targeted delivery of 111In (with a half-life of 2.8 days), one is primarily interested in a time scale of 24 to 48 h, typically required for tumor uptake of radioimmunoconjugates. The modest degradation rates determined here should enable the use of such polymers for the targeted delivery of 111In in vivo.

■

EXPERIMENTAL SECTION

Materials and Methods. All reagents and solvents, including diethylenetriaminepentaacetic acid (DTPA) (98%, Aldrich), tetrahydrofuran (THF) (anhydrous, Aldrich), and other compounds, were 1297

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

NaOH was added to dissolve the DTPA and bring the solution pH to 8.5 (monitored by a pH meter). DMTMM (200 mg, 7 equiv per polymeric primary amine group) was dissolved in 4 mL of water with sonication and added quickly dropwise with stirring to the first solution. This solution was given 5 min to prereact. Then a solution of PAsp(DET) (20 mg in 4 mL water) was added quickly but dropwise with stirring. After another 10 min, the reaction solution was then transferred to a 15 mL 3 kDa MWCO Millipore Amicon spin filter and washed with water (9 × 15 mL). Finally, the aqueous solution was freeze-dried to yield the sodium salt of polyaspartate with the DET pendent groups fully conjugated to DTPA (PAsp(DTPA)-1). Yield: 41 mg (80%). SEC: Mn = 11 kDa, Mw/ Mn = 1.13. 1 H NMR (D2O, 60 °C): PAsp(DTPA)-1: δ(ppm, integrated peak areas reported are based on methine protons of the aspartamide repeat unit as a reference): 4.6−4.5 ppm (broad, 1H/Asp, integration =1.00), 3.9−2.6 (broad, 2H/Asp (−CH2−CO) plus 8H per DET (−NH− CH2CH2-NHCH2CH2-NH−) plus 18H (9 CH2) groups per DTPA, integration =28.57) and biotin end group signals 2.3−2.2 (broad, 2H per polymer, −CH2CH2CH2CH2CO−, integration =0.060), 1.75− 1.45 (broad, 4H per polymer, −CH2CH2CH2CH2CO−, integration =0.120), 1.45−1.30 (broad, 2H per polymer,−CH2CH2CH2CH2CO−, integration =0.060). The degree of polymerization (DPn = 33) was calculated by comparing the integration of the 1H NMR signals at 2.3−2.2 ppm and 1.7−1.3 (the biotin end group) to that at 4.6−4.5 ppm (the methine proton on the backbone). PAsp(DTPA)-2: Experiments to obtain a polymer (PAsp(DTPA)2) in which only a fraction of the pendant −NH2 groups were converted to a DTPA monoamide began with a different sample of the PAsp(DET) precursor, PAsp(DET)-2 (cf., Table 2, below, and Supporting Information). For this reaction, 1.6 equiv of DMTMM per primary polymeric amine group was used. Briefly, DMTMM (46 mg) was added into the DTPA solution (adjusted with 5 M aqueous NaOH to pH 8.5), and the solution was stirred for 5 min. Then PAsp(DET)-2 (20 mg in 4 mL of DI water) was added into the solution and stirred for 10 min. The reaction solution was then transferred to a 15 mL 3 kDa MWCO Millipore Amicon spin filter and washed with water (9 × 15 mL). Finally, the aqueous solution was freeze-dried to yield the sodium salt of polyaspartate with DET pendant groups partially conjugated to DTPA (PAsp(DTPA)-2). Yield: 26 mg (71%). 1 H NMR (D2O, 60 °C): PAsp(DTPA)-2: δ(ppm): 4.6−4.4 (broad singlet, 1H per monomer, backbone methine, integration =1.00, reference), 3.6−2.8 (broad, 2H/Asp (−CH2−CO) plus 8H per DET plus 18H (9 CH2) groups per DTPA, integration =19.43), and biotin end group signals 2.3−2.2 (broad, 2H per polymer, −CH2CH2CH2CH2CO−), integration =0.024), 1.75−1.45 (broad, 4H per polymer, −CH2CH2CH2CH2CO−, integration =0.054), 1.45− 1.30 (broad, 2H per polymer, −CH2CH2CH2CH2CO−, integration =0.025). The degree of polymerization (DPn = 78) was calculated by comparing the integration of the 1H NMR signals at 2.3−2.2 ppm and 1.7−1.3 (the biotin end group) to that at 4.6−4.5 ppm (the methine proton on the backbone). The fraction of repeat units carrying a DTPA group, expressed as the % DTPA functionality, was calculated from the NMR integration values using eq 1.

used without further purification unless otherwise noted. Water was purified through a Milli-Q water purification system (18 MΩ cm). Sodium bicarbonate/carbonate buffer (pH 9.4, 200 mM) was purchased from Pierce Biotechnology. All other buffers were prepared in our laboratory. The Spectra/Pro dialysis membranes (MWCO 1 kDa and MWCO 3.5 kDa) were purchased from Spectrum Laboratories, Inc. Millipore Amicon spin filters (15 mL, 3 kDa MWCO) were purchased from Fisher Scientific, Canada. Poly(sodium methacrylate), Mn = 340 kDa, Mw/Mn = 1.10 was obtained from Polymer Standards Service Mainz. 111InCl3 (>7.4 GBq/mL) was purchased from Nordion (Kanata, ON, Canada). 4-(4,6-Dimethoxy1,3,5-triazin-2-yl)-4-methylmorpholinium chloride (DMTMM, Acros Organics, 99+%, from Fisher Scientific, Canada), L-glutamic acid γbenzyl ester (BLG), β-benzyl-L-aspartate (BLA), N,N-dimethylformamide (DMF) anhydrous, N-methylpyrrolidone (NMP), dichloridemethane (DCM) anhydrous, diethylenetriamine (DET), phosgene solution (∼20% in toluene), indium chloride, and europium chloride hexahydrate were purchased from Sigma Aldrich. (+)-Biotinyl-3,6dioxaoctanediamine was synthesized in our lab,6 although it is also commercially available from Pierce Biotechnology.15 Instrumentation. The nominal molecular weights and polydispersity indices (PDI = Mw/Mn) of all water-soluble samples were measured with a Viscotek size exclusion chromatograph (SEC) equipped with a Viscotek VE3210 UV/vis detector, a VE3580 refractive index detector, and Viscotek ViscoGEL G4000PWXL and G2500PWXL columns (kept at 30 °C). The flow rate was maintained at 1.0 mL/min using a Viscotek VE1122 Solvent Delivery System and a VE7510 GPC Degasser. The eluent used was 0.2 M KNO3, 25 mM pH 8.5 phosphate buffer, and 200 ppm NaN3 in water, and the system was calibrated with poly(methacrylic acid) standards. All samples were dissolved in sodium bicarbonate/carbonate buffer (pH 9.4, 200 mM) prior to injection. The polydispersity indices of N-methylpyrrolidone (NMP)-soluble polymers were measured with a Viscotek GPC Max gel-permeation chromatograph (GPC) equipped with a VE3580 refractive index detector. The eluent, NMP, was kept at a flow rate of 0.6 mL/min at 80 °C. The column was calibrated with poly(methyl methacrylate) standards. Size-exclusion high performance liquid chromatography (SEHPLC) measurements were performed on a Biosep S-4000 column (300 × 4.6 mm) eluted with 0.1 M NaH2PO4 (pH 7.0) at a flow rate of 0.35 mL/min using a Series 200 pump (PerkinElmer, Wellesley, MA, USA) interfaced with a flow scintillation analyzer; Radiomatic 610TR (PerkinElmer). 1 H NMR (400 MHz) spectra were recorded on a Varian Mercury 400 spectrometer. Chemical shifts were referenced to either tetramethylsilane (TMS) or a residual solvent peak. All spectra were collected as 64 transients with a delay time of 10 s. Thermogravimetric analysis (TGA) measurements were carried out with a TA Instruments model SDT Q600 instrument. Data were collected from 25 to 600 °C in two isothermal steps. First, the temperature was ramped up to 100 °C at a heating rate of 10 °C/min and kept at 100 °C for 200 min. Then the temperature was ramped up to 600 °C at a speed of 10 °C/min and kept at 600 °C for 200 min. All analyses were carried out under a stream of air. Polymer Synthesis and Pendant Group Transformations. Experimental details for the syntheses of β-benzyl-L-aspartate Ncarboxyanhydride (BLA-NCA), γ-benzyl-L-glutamate N-carboxyanhydride (BLG-NCA), and the (+)-biotinyl-3,6-dioxaoctanediamineinitiated poly(β-benzyl-L-aspartate) (PBLA) and poly(γ-benzyl-Lglutamate) (PBLG) and the reactions of these polymers with diethylenetriamine to form the corresponding biotin-poly{N′-[N-(2aminoethyl)-2-aminoethyl]aspartamide} PAsp(DET) and glutamide PGlu(DET) are presented in the Supporting Information. Representative 1H NMR spectra of these monomers and polymers are presented in Figures S1−S4 and S6−S8 of the Supporting Information. Synthesis of PAsp(DTPA)-1 and PAsp(DTPA)-2. PAsp(DTPA)-1: An aqueous solution containing a large excess of DTPA was prepared by adding DTPA (3.0 g, 80 equiv per polymeric primary amine group) and 4 mL of H2O to a 100 mL round-bottom flask. Next, 5 M aqueous

DTPA functionality % = 100(B/ A − 10)/(28 − 10) (polyaspartamide)

(1)

Here, B refers to the integral of the signal at 3.6−2.5 ppm (B, due to the CH2 groups of the DTPA groups (18H/DTPA), the −CH2−C O protons of the aspartamide (2H/Asp), and the CH2 groups of the DET pendant chains (8H/DET)). A refers to the well-resolved signal at 4.6−4.5 ppm (d in Figure 1, below) from the backbone methine (1H/Asp). A more detailed explanation of eq 1 is presented in the Supporting Information. In this way, we determined that PAsp(DTPA)-1 was fully functionalized (within NMR error) with DTPA groups. In contrast, in PAsp(DTPA)-2, 53% of the repeat units contained a DTPA group, and 47% were unreacted DET groups. 1298

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

Scheme 1. Synthesis of Metal Chelating Polymers Based upon Biotin-Poly(β-benzylaspartate) and Biotin-Poly(γbenzylglutamate) Precursors

Synthesis of PGlu(DTPA). A large excess of DTPA (3.0 g, 80 equiv per primary polymeric amine group) and 4 mL of H2O were added to a 100 mL round-bottom flask. Next, 5 M aqueous NaOH was added to form a solution of pH of 8.5. DMTMM (200 mg, 7 equiv per primary polymeric amine group) was dissolved in 4 mL of water with sonication and added quickly dropwise with stirring to the first solution. This solution was given 5 min to prereact. Then a solution of PGlu(DET) (20 mg) in 4 mL of water was added quickly dropwise with stirring. After another 10 min, the reaction solution was then transferred to a 15 mL 3 kDa MWCO Millipore Amicon spin filter and washed with water (9 × 15 mL). Finally, the aqueous solution was

freeze-dried to yield the sodium salt of polyglutamate with DET pendant groups fully conjugated to DTPA (PGlu(DTPA)). The ratio of the DTPA/DET group of the polymer and DPn were characterized by 1H NMR. Yield: 39 mg (76%). SEC (aqueous, RI) Mn = 11.6 kDa, Mw/ Mn = 1.16. PGlu(DTPA): 1H NMR (D2O, 25 °C): δ (ppm, integrated peak areas reported were based on the methine protons on the backbone (1H as the reference), 4.4−4.0 (broad, 1H per monomer, backbone methine, integration =1.00), 3.6−2.8 (broad, 18H per DTPA plus 8H per DET, integration =25.74), 2.2−1.8 (broad, 4H per monomer, −CH2CH2−CO, integration =3.98), and biotin end group signals 1299

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

1.6−1.4 (broad, 4H per polymer −CH2CH2CH2CH2−CONH−, integration =0.138), 1.4−1.2 (broad, 2H per polymer −CH2CH2CH2CH2−CONH−, integration =0.079). The degree of polymerization (DPn = 28) was calculated by comparing the integration of the 1H NMR signals at 2.3−2.2 ppm and 1.7−1.3 (the biotin end group) to that at 4.6−4.5 ppm (the methine proton on the backbone). The functionality of the DTPA groups was calculated by eq 2. Within NMR error, the polymer was found to be fully functionalized with DTPA groups.

in assessing their utility in radioimmunotherapy applications. We begin with a description of the synthesis and characterization of the polymers. This is followed by studies by size exclusion chromatography of the degradation kinetics. Finally, we interpret our results in terms of how the charges on the pendant group influence the mechanism of the degradation reaction. Polymer Synthesis. Samples of poly(β-benzyl-L-aspartate) (PBLA) and poly(γ-benzyl-L-glutamate) (PBLG) were synthesized by ring-opening polymerization of the respective Ncarboxyanhydrides using the amino-biotin derivative 1 as the initiator as shown in Scheme 1. Details of the synthesis of these samples are provided in the Supporting Information. The degrees of polymerization of the polymers were determined by 1 H NMR by comparing the integration of the end group protons on the biotin to the pendant group signals. For example, for PBLA-1, a value of DPn = 30 was calculated by comparing the integral of the 1H NMR signals at 1.4−2.2 ppm (the biotin end-group) to that at 7.2−7.6 ppm for the −C6H5 groups of the benzyl esters. Analysis of the polymer by size exclusion chromatography (SEC, N-methylpyrrolidone eluent) indicated that the polymerization reactions were successful and that polymers PBLG and PBLA-1 had narrow chain length distributions. In the synthesis of a second sample of PBLA (PBLA-2), we obtained a polymer with a higher mean degree of polymerization and a broader molecular weight distribution. As seen in the data summarized in Table 1, the values of DPn

DTPA functionality % = 100(B/ A − 8)/(26 − 8) (polyglutamide)

(2)

Metal Loading Experiments. A DTPA polymer solution was prepared by dissolving a sample of polymer (5 mg, PAsp(DTPA)-1, PAsp(DTPA)-2, or PGlu(DTPA)) in ammonium acetate buffer (2.4 mL, 20 mM, pH 6.0). To prepare a europium-containing polymer, a EuCl3 solution (0.25 mL, 10 mg/mL) was prepared added to the polymer solution (1.3−1.4 equiv of Eu3+ per DTPA units of the polymer). After 30 min at 37 °C, the solution was transferred to a 4 mL 3 kDa MWCO Millipore Amicon spin filter and washed three times with Tris buffer (Tris 25 mM, NaCl 150 mM, KCl 2 mM, pH 7.4) and then water. Then the final solution was filtered with a 200 nm syringe filter and freeze-dried. Other Eu-loaded polymers were prepared in a similar manner. To prepare an indium-containing polymer, a solution of InCl3 (200 μL, 45 mmol/mL, ca. 1.3 equiv per primary polymeric DTPA group) in citric acid solution (pH 5.0, 100 mM) was added to a solution containing 5 mg of polymer sample (PAsp(DTPA)-1, PAsp(DTPA)-2, or PGlu(DTPA)) in citric acid solution (1.0 mL, pH 5.0, 100 mM). After 1 h, the solution was transferred to a 4 mL 3 kDa MWCO Millipore Amicon spin filter and washed with aqueous sodium acetate/ acetic acid (pH 5.5, 100 mM) twice and DI water twice. Finally, the solution was filtered with a 200 nm syringe filter and freeze-dried, and the products, PAsp(DTPA)-1:In, PAsp(DTPA)-2:In, or PGlu(DTPA):In were obtained depending on the precursor used. Degradation Kinetics Experiments. Solutions of PAsp(DTPA)1, PAsp(DTPA)-2, and PGlu(DTPA) and their metal-containing derivatives for degradation kinetics experiments were prepared as follows: Each polymer sample was dissolved (2.0 mg/mL) in phosphate buffer (pH 7.4, 10 mM containing 150 mM NaCl) in a 2 dram vial and incubated at 37 °C, controlled by a water bath. Aliquots (100 μL) were withdrawn periodically over 14 days and stored at −18 °C to arrest further degradation. Before SEC analysis, these aliquots were mixed with the SEC eluent (0.2 M KNO3, 200 ppm NaN3, and 25 mM, pH 8.5 phosphate buffer) and an internal standard (poly(sodium methacrylate), Mn = 340 kDa, Mw/Mn = 1.10). Each sample for SEC analysis contained 0.5 mg/mL of polymer and 0.2 mg/ mL of the internal standard. Degradation Kinetics Experiments of PAsp(DTPA)-1 Loaded with 111In. PAsp(DTPA)-1 (100 μg) were radiolabeled by incubation in 100 mM sodium acetate (pH 6.0) with 4 MBq of 111InCl3 for 1 h at RT. The polymer was then saturated with stable In3+ by adding a 50fold molar excess of InCl3 compared to total DTPA and stirring the mixture with the radiolabeled polymer for 1 h at RT in 100 mM sodium acetate (pH 6.0). Excess InCl3 was removed by ultrafiltration on an Amicon-Ultra device (3 kDa MWCO). The polymer loaded with 111In, PAsp(DTPA):111In, was then transferred into a vial, and phosphate buffer (pH 7.4, 10 mM containing 150 mM NaCl) was added such that the final concentration of polymer was 0.5 mg/mL. The solution was incubated at 37 °C. Aliquots (20 μL) were withdrawn periodically (0, 1, 24, 48, 72, and 100 h) and analyzed by size-exclusion HPLC.

Table 1. Characteristics of the Polymers Synthesized by Ring-Opening Polymerization polymer sample d

PBLA-1 PBLA-2d PBLG

yield %

[M]/[I]a

DPnb

Mw/Mnc

81 74 79

27 45 25

30 51 26

1.15 1.32 1.13

a Ratio of the monomer concentration [M] to the initiator concentration [I]. bFrom 1H NMR end group analysis. cFrom SEC measurements in NMP with refractive index (RI) detection. dPBLA-1 and PBLA-2 refer to two separate syntheses of biotin-poly(βbenzylaspartate). Note the broader polydispersity of the PBLA-2 sample.

obtained by NMR for the various polymers were consistently in good agreement with the [monomer]/[initiator] feed ratios.16 Since all polymers reported here (e.g., PBLA-1) contain a biotin end group at the C-terminal of the polypeptide, we omit mention of the biotin in our notation for the polymers. The aminolysis reaction conditions needed to convert the benzyl ester groups to N-substituted polyamide derivatives were different for PBLA and PBLG. For the reaction of PBLA with diethylenetriamine (DET), we employed conditions similar to those reported by Kataoka et al.,11 who found that the benzyl ester groups of PBLA underwent a quantitative reaction with primary amines under very mild conditions. In the 13C NMR spectra of the product, they found a split of the carbonyl and methylene groups of aspartamide units, suggesting that the aminolysis of PBLA was accompanied by intramolecular isomerization of aspartamide units to form β-aspartamide units. We carried out the aminolysis of PBLA with a large excess of DET in NMP as solvent at 0 °C for 2 h. After 20% aqueous acetic acid was added into the reaction mixture in an ice bath, the polymer was separated from excess amine by dialysis. The NMR spectrum of the PAsp(DET) in Figure S4

■

RESULTS AND DISCUSSION The focus of this paper is on the spontaneous degradation kinetics of metal chelating polymers based on a polyaspartamide and a polyglutamide backbone as an important first step 1300

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

(Supporting Information) demonstrates that, in our reactions, the benzyl ester groups were converted completely to the corresponding amide. We were unable to analyze this reaction product directly by SEC because the amine polymers did not elute from the SEC columns on our instrument. To analyze the polyaspartamides by SEC, the amines of the polyaspartamides were succinylated with succinic anhydride. The results are presented in Table S1. The low polydispersity indices and the lack of side peaks in GPC traces of these succinylated polymers demonstrate that no significant interchain cross-linking occurred in these reactions. The corresponding aminolysis reaction with PBLG was carried out under more forcing conditions, again with a large excess of DET (10 mL for 200 mg polymer) in 10 mL of DMF in the presence of 2-pyridone as a catalyst.17 The reaction was carried out at 40 °C for 24 h, with a similar workup as described for the reaction product with PBLA. The quantitative conversion of PBLG to PGlu(DET) was verified by 1H NMR from the peak integration ratio of the methine of the polypeptide backbone to the methylene protons adjacent to the amine and amide nitrogens in the PGlu(DET) pendant groups. The absence of peak broadening in GPC traces of succinylated PGlu(DET) provides good evidence that no significant interchain cross-linking occurred in these reactions. The next step involved attachment of diethylenetriaminepentaacetic acid (DTPA) to each of the pendant primary amino groups using 4-(4,6-dimethoxy-1,3,5-triazin-2-yl)-4-methylmorpholinium chloride (DMTMM) as the coupling agent, using a protocol that we have described elsewhere.7 To convert the DET groups of the PAsp(DET) and PGlu(DET) to DETDTPA pendant groups, we pretreated a large molar excess (based on primary amine groups) of DTPA in water at pH 8.5 with a limiting amount of DMTMM to convert some of the carboxylic acid groups to an activated ester. By varying the amount of DMTMM, we were able to tune the fraction of the DET pendants groups conjugated to DTPA. To the activated DTPA solution, we added a solution of the PAsp(DET) or PGlu(DET) polymer in DI water. After 10 min, excess reagents were removed using a spin filter with a 3 kDa cutoff. The polymer was then freeze-dried. The 1H NMR spectrum of PAsp(DTPA)-1 is presented in Figure 2. Table 2 summarizes the characteristics of the DTPA polymers. The number of DTPA groups per polymer was calculated by comparing the sum of the integration of the signals in 1H NMR of DTPA and DET (signals between 3.6 and 2.8 ppm) and integration of the signals of the backbone methine as described in the Experimental Section (eq 1 for PAsp(DET); eq 2 for PGlu(DET)). For the reaction described above (for PAsp(DTPA)-1, PGlu(DTPA) in the presence of excess DMTMM coupling reagent), we found that the ratios of DET to DTPA groups in these polymers were 1:1. The diethylenetriamine pendant groups of PAsp(DET) and PGlu(DET) contain both primary and secondary amines. To explain our finding of one DTPA per DET pendant group, we assume that the activated DTPA reacted selectively with the primary amine with essentially quantitative yield and that the secondary amine shows little or no reactivity with DTPA when DMTMM was used as the amine-coupling reagent at pH 8.5. SEC traces for these DTPA polymers indicated a narrow distribution of chain lengths. These samples showed no peak asymmetry or additional higher molecular weight peaks that one would expect if interchain cross-linking by DTPA occurred during the reaction.

Figure 2. 1H NMR spectrum (D2O, 60 °C) of PAsp(DTPA)-1. End group analysis shows the degree of polymerization to be 33, by comparing the integration of the biotin end group signals a, b, and c to that of the backbone signal d. The DTPA functionality was calculated by comparing the integral of peak e (due to DTPA, DET, and −CH2CO−) to the integral of peak d due to a proton on the polymer backbone (eq 1). Within NMR error, the polymer was fully functionalized with DTPA groups.

Table 2. Characteristics of the PAsp(DTPA), PGlu(DTPA) Samples sample

DPna

DPn of the starting materialb

Mw/ Mnc

PAsp(DTPA)-1 PAsp(DTPA)-2 PGlu(DTPA)

33 76 28

30 51 26

1.13 1.27 1.16

1

H NMR molecular weightd (kDa) 19.0 32.6 16.5

a

From 1H NMR end group analysis. bValues from Table 1. cFrom SEC analysis with refractive index (RI) detection. dThese molecular weight values were calculated from DPn assuming that the DTPA groups were in their fully protonated form. For PAsp(DTPA)-2, this calculation assumes 53% DTPA pendant groups and 47% N-[N-(2aminoethyl)-2-aminoethyl] pendant groups, as characterized by 1H NMR.

To partially conjugate DET pendant groups to DTPA groups, we varied the amount of DMTMM coupling reagent added to the reaction. Here we used 1.6 equiv of DMTMM for each DET pendant group (along with 80 equiv of DTPA). For this reaction, we employed a different PAsp(DET) sample [PAsp(DET)-2] prepared from PBLA-2. This distinction is important, because both PBLA-2 and PAsp(DET)-2 have significantly broader polydispersities than the samples used for the synthesis of PAsp(DTPA)-1. The DTPA functionality of the resulting polymer was characterized by 1H NMR (cf., eq 1, Experimental Section). In this way we calculated that PAsp(DTPA)-2 had DTPA coupled to 53% of the pendant groups and 47% unreacted DET amine groups. We also found that the degree of polymerization increased from 51 (PBLA-2) to 76 when PBLA-2 was converted to PAsp(DTPA)-2, accompanied by a lowered yield. We attribute these changes to loss of the low molecular weight component of the polymer during spin filtration, and we rationalize this loss of material in terms of the broader polydispersity of the sample. We imagine that some of the lower molecular weight components of the sample were not retained by the spin filter used in the polymer purification step. 1H NMR spectra of PAsp(DTPA)-2 and 1301

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

sample were withdrawn periodically (between 0 to 13.8 days) and stored at −18 °C to arrest further hydrolytic degradation. At the end of the experiment (ca. 14 days), all samples were thawed and analyzed by SEC. The results for the three polyglutamide polymers (PGlu(DTPA), PGlu(DTPA:In), and PGlu(DTPA:Eu)) are shown in Figure 3. The SEC trace of

PGlu(DTPA) are presented in Figures S9 and S10 in the Supporting Information. Additional information is needed to convert degrees of polymerization determined by NMR to molecular weights. To calculate the actual molecular weight of these fully and partially converted polymers, we must also account for the number of counterions present per polymer. To determine the mean number of sodium ions per polymer, we employ thermal gravimetric analysis (TGA). The polymer samples were held at 100 °C for 200 min to remove residual moisture and then at 600 °C for 300 min. A representative TGA trace is presented in Figure S11 (Supporting Information). The mass loss due to moisture at 100 °C was 7.1% for PAsp(DTPA)-1 and 4.2% for PGlu(DTPA). The final ceramic yield was 20.0% for PAsp(DTPA)-1 and 18.6% for PGlu(DTPA), corresponding to conversion of the sample to Na2CO3.20 From these data, we calculate that PAsp(DTPA)-1 contains 2.6 Na+ ions, whereas PGlu(DTPA) contains 2.3 Na+ ions per repeat unit. The sodium content of 2.6 or 2.3 Na+ ions per DTPA is consistent with the range of pKa values for DTPA itself.18 We summarize these data in Table 3, where the “adjusted molecular weight”

Figure 3. Time dependence (days) at 37 °C of the SEC profiles of PGlu(DTPA), PGlu(DTPA:In), and PGlu(DTPA:Eu) in phosphate buffer solution (10 mM, pH 7.4) containing 150 mM NaCl. The fact that the lines corresponding to different incubation times cannot be resolved indicates that the polymers do not undergo any significant degradation on these time scales.

Table 3. H2O and Na+ Content and Adjusted Molecular Weight for PAsp(DTPA)-1 and PGlu(DTPA) Calculated from TGA Analysis

sample

H2O per DTPA unit

Na+ per DTPA unit

PAsp(DTPA)-1 PGlu(DTPA)

2.7 1.6

2.6 2.3

DPna

1 H NMR molecular weightb (kDa)

adjusted molecular weightc (kDa)

33 28

19.0 16.5

22.6 18.8

PGlu(DTPA), the polymer before metal-chelation, shows a narrow and symmetrical peak centered at a retention volume (Rv) of 14.3 mL. Metal complexation of PGlu(DTPA) with Eu3+ and with In3+ caused a small shift to longer retention volumes. The metal-free polymer has a net negative charge associated with each repeat unit. This charge leads to electrostatic repulsion between pendant groups, resulting in a larger hydrodynamic volume for the metal-free polymer than for its metal-containing counterparts, which have zwitterionic pendant groups. The (−1) charge of the DTPA-Eu complex is neutralized by the positive charge on the secondary amine of the DET spacer group, as shown in Figure 4. With increasing

a

From end group analysis by 1H NMR. bCalculated from DPn, assuming fully protonated DTPA groups. cThe recalculated molecular weight that includes the mass contribution of water molecules and sodium counterions in the sample.

includes the contribution from both water and sodium ions. These values can be used to determine the moles of polymer from weighed samples. Backbone Degradation of PGlu(DTPA) and PAsp(DTPA). In this section, we examine the influence of pendant group structure and the role of DTPA-bound metal ions in the degradation of PGlu(DTPA) and PAsp(DTPA)-1 in solutions approximating physiological conditions. Sample solutions of polymer (2.0 mg/mL) were prepared in 10 mM phosphate buffer (pH 7.4) containing 150 mM NaCl. In parallel, similar samples of both polymers saturated with Eu3+ [PGlu(DTPA:Eu), PAsp(DTPA:Eu)-1] and with In3+ [PGlu(DTPA:In), PAsp(DTPA:In)-1] were prepared in the same buffer medium. We assume that the binding constant for Eu3+ with a monofunctional DTPA is similar to that of Gd3+, which is on the order of 1017 at physiological pH. The binding constant for In3+ is even stronger, on the order of 1029.19 Because of these strong binding affinities, we assume that essentially all of the DTPA units on the MCPs contain a metal ion at pH 7.4. In a previous publication, we showed that for a metal-chelating poly(N-alkylacrylamide) polymer with a DTPA chelator attached to each pendant groups, the number of Gd3+ ions bound per polymer was (within experimental error) equal to the number of DTPA pendant groups on each polymer.5 To mimic physiological conditions, each polymer sample (2.0 mg/mL) was incubated at 37 °C in 10 mM phosphate buffer (pH 7.4) containing 150 mM NaCl. Aliquots of each

Figure 4. Charge distribution and the zwitterionic properties on the DTPA polymer loaded with metal ions.

incubation time, no changes are observed in the SEC traces for all the polyglutamate derivatives. This result is consistent with the known hydrolytic stability of polyglutamides at neutral pH.13 As anticipated by Kataoka et al.,13 the hydrolytic degradation of polyaspartamides was much more rapid. Our interest is in the influence of the pendant group on the rate of this degradation. The evolution of the SEC traces with incubation 1302

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

time for the metal-free PAsp(DTPA)-1 is shown in Figure 5A. All traces are normalized to the peak of the poly(sodium

Figure 6. Time dependence of the SEC profiles of (A) PAsp(DTPA)2, (B) PAsp(DTPA:In)-2, and (C) PAsp(DTPA:Eu)-2 in phosphate buffer (10 mM, pH 7.4) containing 150 mM NaCl at 37 °C. Poly(sodium methacrylate) was used as an internal standard.

Figure 5. Time dependence of the SEC profiles of (A) PAsp(DTPA)1, (B) PAsp(DTPA:In)-1, and (C) PAsp(DTPA:Eu)-1 in phosphate buffer (10 mM, pH 7.4) containing 150 mM NaCl at 37 °C. Poly(sodium methacrylate) (P(NaMA) (Rv = 11.2 mL) was used as an internal standard in each sample.

polymers, as seen in Figure 6B for PAsp(DTPA:In)-2 and in Figure 6C for PAsp(DTPA:Eu)-2. Degradation of PAsp(DTPA)-2 and its metal-containing derivatives was more significant than the degradation of PAsp(DTPA)-1 and its metal-loaded counterparts. One can attribute the faster degradation in this set of polymers to the presence of the N[N-(2-aminoethyl)-2-aminoethyl] pendant groups. We analyze the SEC traces in Figures 5 and 6 in two ways to obtain semiquantitative data on the degradation rates. In Figure 7A, we plot the shift in peak position as a function of incubation

methacrylate) internal standard at retention volume (Rv) = 11.2 mL. With time, there was a slow but steady decrease in the area under the peak accompanied by a shift of the peak maximum to longer retention volumes. These are symptoms of polymer backbone degradation. The changes over the first two days were small but became more pronounced after day 4. Results for the metal-containing polymers are presented in Figure 5B and C. When these polymers were incubated at 37 °C, aliquots taken over time showed pronounced shifts in the SEC peak maxima to longer retention times, accompanied by a decrease in peak area. The changes over the first two days were small, but they became increasingly pronounced after day 4. Moreover, for the metal-loaded polyaspartamides, a new peak appeared at longer Rv values (Rv = 16). This peak can be attributed to lower molecular weight fragments from backbone degradation. It is interesting to note that, in the case of the polyaspartamides, the severity (and rate) of degradation was greater for the metal-containing polymers than for the metal free polymers and more for the Eu3+-containing polymers than for the In3+-containing polymers. To determine the effects of varying the number of DTPA groups per polymer on the polymer degradation rate, we examined the hydrolytic stability of PAsp(DTPA)-2, in which 53% of the DET pendant groups were conjugated to the DTPA functionalities. The remaining 47% of the pendant groups have an −NH−CH2CH2−NH2 functionality, which is expected to be monoprotonated at pH 7.4.20 Samples of this polymer were also loaded with Eu3+ ions (PAsp(DTPA:Eu)-2) and with In3+ ions (PAsp(DTPA:In)-2). The degradation of these polyaspartamides was also analyzed by SEC. For PAsp(DTPA)-2 (Figure 6A), there is a noticeable peak shift in the SEC trace and a small decrease in peak area even after 2 days. After 1 week, more profound changes were observed, including new peaks at 15.8 mL and at 16.4 mL, with the latter corresponding to the position of the solvent peak. The changes in the SEC traces over time were similar for both metal-containing

Figure 7. Time dependence of the changes in the SEC traces of PAsp(DTPA)-1 and PAsp(DTPA)-2 and their metal-loaded derivatives in terms of (A) the shift in the retention volume of the original peak maximum and (B) the percentage decrease of the area of the original peak maximum.

time as a measure of the change in polymer length. The shift in peak position to longer Rv suggests that these polymers became shorter due to chain degradation. In Figure 7B, we attempt to estimate the fraction of the initial PAsp(DTPA)-1 polymer that survives the degradation process. To make this estimate, we examine the areas of the polymer peaks in Figure 5 and assume that the fraction of the peak that overlaps the peak of the original polymers is due to polymer 1303

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

different from those of PAsp(DTPA:In)-1 in Figure 5. In addition, the flow scintillation analyzer used to detect the presence of 111In has a much larger bore than that of other detectors, leading to artificial peak broadening. This effect and possible polymer adsorption to the column packing material led to a broad tail in the elution profile, extending to retention times corresponding to the low molecular weight cutoff (M < 10 000) of the column. By comparison, aqueous 111In3+ in citrate buffer gives a symmetric peak centered at 14.8 min extending out to 15.7 min. This means that the loss of small amounts of low molar mass material containing 111In might not be discriminated due to their elution under the tail of the polymer trace. The main result, however, is that the peak profile is almost unchanged over the 4 days of the experiment. This result allows us to conclude that loss of the pendant group via deamidation is not a major degradation pathway for PAsp(DTPA:In)-1 on the time scale of 4 days. Mechanism of the Polymer Degradation. As mentioned in the Introduction, there is a broad interest in the spontaneous deamidation and cleavage reactions of Asn-containing polypeptides.12 The general conclusion from these studies is that peptide cleavage (backbone degradation) is slower than deamidation and that deamidation rates have half-times commonly on the order of tens of days. The mechanism of this reaction was investigated in detail by Capasso et al.,21 and we summarize this mechanism, specialized to the case of polyaspartamides, in Scheme 2. For pH < 9, the first step of the reaction involves equilibrium deprotonation of an amide nitrogen, followed by intramolecular attack by the nitrogen on the carbonyl carbon, leading to formation of a tetrahedral intermediate (not shown here). Subsequent proton transfer from a general acid to the leaving group leads to cleavage of the peptide bond with formation of a succinimide ring. At pH < 9, formation of the tetrahedral intermediate is rate limiting. The succinimide, then hydrolyzes to a ring-opened carboxyamide. Side-chain cleavage (deamidation) involves deprotonation of the backbone amide nitrogen on the C-terminal side of the reaction center. The pendant group is expelled following ring closure. Backbone cleavage begins with deprotonation of the amide nitrogen of the pendant group. Capasso et al.21 comment that although the deamidation and backbone cleavage reactions proceed through similar pathways, they have very different rate constants. For Asn-containing polypeptides, the deamidation rate is dominant. The cleavage reaction occurs to a significant extent only when deamidation is precluded. For the polyaspartamide samples that we examined, the reverse appears to be the case. The cleavage reaction is dominant, and for PAsp(DTPA:111In)-1, deamidation, if it occurs at all on a time scale of 4 days, does not contribute to a measurable extent to the HPLC-SEC traces in Figure 8. When we compare the results for PAsp(DTPA), PAsp(DTPA:In)-1, and PAsp(DTPA:In)-2, we see that the cleavage rates are sensitive to the nature of the pendant group and its charge. In all three examples, it is likely that the cleavage pathway is favored as a consequence of the positively charged nitrogen of the pendant group, which makes the adjacent amide nitrogen more acidic. This effect is offset to some extent when the pendant group contains a DTPA-In (−1 charge) or DTPA itself (−2 or −3 at pH 7.4). Thus, PAsp(DTPA:In)-2, in which 45% of the DET groups have a terminal −NH2, undergoes more rapid degradation than the polymers with DTPA on each pendant group.

that has not yet undergone degradation. We call this fraction “Δarea”. The y-axis in Figure 7B is 100(1 − Δarea/area), where “area” refers to the area of the peak for the initial polymer. An example showing how Δarea/area is calculated from the SEC traces before and after degradation of one PAsp(DET)-1 sample is presented in Figure S12 in the Supporting Information. It is possible that this analysis may overestimate the fraction of surviving polymer if partial degradation of one of the longer components of the original polymer leaves a fragment that itself is comparable in length to one of the shorter components of the original sample. We expect that this contribution to the data is small. These plots reveal that the polymer backbone degradation rate for these polyaspartamides is very sensitive to the nature of the pendant groups. The polyaspartamides with pendant groups partially converted to DTPA degraded faster than the polyaspartamides with pendant groups fully converted to DTPA. For example, we found that, after 14 days of incubation, about 65% of PAsp(DTPA)-1 remained intact, whereas only 15% of PAsp(DTPA)-2 was undegraded. Possible Pendant Group Loss via Deamidation. Polymers intended to carry 111In as radioimmunoconjugates need to be stable during the delivery not only against backbone degradation but also against pendant group cleavage. Deamidation leading to loss of the DET-DTPA-In pendant group would release radioactive indium in an uncontrolled manner in the form of a low molecular weight DET-DTPA conjugate. To assess the importance of this reaction under the model physiological conditions described above, we take advantage of the fact that an HPLC-SEC instrument with a radioactivity (γ-ray) detector can monitor the loss of radioactive low molecular weight substances from polymer with a much shorter elution volume. To carry out these experiments, a sample of PAsp(DTPA)-1 was radiolabeled with 4 MBq 111In (half-life 2.8 days) for 1 h at room temperature and then the remaining DTPA groups were saturated with cold In3+ as described in the Experimental Section. We refer to this polymer as PAsp(DTPA:111In)-1. This polymer was incubated at 37 °C in 10 mM phosphate buffer (pH 7.4) containing 150 mM NaCl, and samples were taken periodically over 4 days and analyzed by HPLC-SEC. Because of the radioactive decay of 111In, we present normalized (corrected for radioactive decay) elution curves in Figure 8. These experiments used a different HPLC instrument, a different aqueous SEC column, and a different detector, than those used for the degradation experiments described above. Thus, the peak shapes and retention volumes for the PAsp(DTPA:111In)-1 polymer for the traces in Figure 8 are

Figure 8. Time dependence at 37 °C of the HPLC profiles of PAsp(DTPA:111In)-1 in phosphate buffer solution (10 mM, pH 7.4) containing 150 mM NaCl. 1304

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

Scheme 2. Mechanism of the Backbone Cleavage and Deamidation Reactions of Polyaspartamide in Buffer at pH 7.4a

a The rate-limiting step20 is the formation of a tetrahedral intermediate (not shown) formed upon attack of the deprotonated amide nitrogen on the carbonyl carbon, leading to formation of a succinimide intermediate.

the kinetics of polymer degradation. For PAsp(DTPA)-2, in which 53% of the pendant groups had a DTPA substituent and 47% of the pendant groups had an unsubstituted diethylenetriamine group (with a net positive charge), the degradation reaction was significantly faster. The slower degradation of PAsp(DTPA)-1 and its In3+ and Eu3+ complexes compared to PAsp(DTPA)-2 can be attributed to the bulky nature of the DTPA pendant groups as well as the net negative charge associated with the DTPA−metal complex. The most important conclusion that we draw from these experiments is that PAsp(DTPA)-1 is sufficiently stable under model physiological conditions to be used as a carrier for 111In3+ in radioimmunotherapy assays in which one monitors biodistribution over a time period of 24 to 72 h.

The slower degradation rates of PAsp(DTPA)-1 and its metal loaded polymers, compared to the PAsp(DET) in the Kataoka experiments,9,10 may also be attributed to the bulky nature of the DTPA pendent groups and the positively charged nitrogen of DET pendant groups. In their experiments, which used PAsp(DET) for DNA or protein delivery to cells, rapid polymer degradation following cellular uptake is important to avoid the toxicity commonly associated with cationic polymers. In the applications we envision, in which PAsp(DTPA) will be used as a carrier for 111In in the form of radioimmunoconjugates, the pharmacokinetic time scale for tumor uptake is on the order of 24 to 48 h. It appears that the polyaspartamide derivatives fully functionalized with DTPA groups remain stable for that time scale at physiological pH.

■

■

CONCLUSIONS We described the synthesis of two types of metal-chelating polymers, one based on a polyaspartamide backbone and the other based on a polyglutamide backbone. The polymers were characterized by 1H NMR, to determine the composition and the number average degree of polymerization, and by SEC to determine the polydispersity. Polydispersities were narrow, and within experimental error, all pendant groups in PAsp(DTPA)1 and PGlu(DTPA) contained a DTPA chelating group. Under model physiological conditions (37 °C, pH 7.4, 10 mM phosphate buffer, containing 150 mM NaCl), the aspartamide polymers underwent spontaneous degradation over a period of several days to weeks, as monitored by SEC. For PAsp(DTPA)-1, the polymer fully loaded with Eu3+ or In3+ ions degraded somewhat more quickly than the metal-free polymer. Backbone hydrolysis appears to involve anchimeric participation of the pendant amide groups, an idea supported by the observation that the corresponding polygluatamide polymer remained hydrolytically stable over 14 days. The nature of the pendant group plays an important role in affecting

ASSOCIATED CONTENT

S Supporting Information *

Materials and methods used for the synthesis of β-benzyl-Laspartate-N-carboxyanhydride (BLA-NCA) and γ-benzyl-Lglutamate-N-carboxyanhydride (BLG-NCA), polybenzylaspartate, polybenzylglutamate, PAsp(DET)-1, PAsp(DET)-2, and PGlu(DET), and their 1H NMR spectra; table of 1H NMR endgroup analyses and SEC data for all polymer samples; SEC chromatographs of polybenzylaspartate and polybenzylglutamate; thermogravimetric analysis (TGA) measurements on PAsp(DTPA)-1 and PGlu(DTPA) samples; and definition of the change of peak area [100(1 − Δarea/area)] in Figure 7. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]; raymond.reilly@ utoronto.ca. 1305

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306

Biomacromolecules

Article

Present Addresses

(20) Itaka, K.; Kanayama, N.; Nishiyama, N.; Jang, W.-D.; Yamasaki, Y.; Nakamura, K.; Kawaguchi, H.; Kataoka, K. J. Am. Chem. Soc. 2004, 126, 13612. (21) Capasso, S.; Mazzarella, L.; Sorrentino, G.; Balboni, G.; Kirby, A. J. Peptides 1996, 17, 1075.

§

Institute for Technical Chemistry, Braunschweig University of Technology, Hans-Sommer-Straße 10, 38106 Braunschweig, Germany ∥ Departamento de Quimica Organica, Universidad de Sevilla, 41012 Seville, Spain Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors gratefully acknowledge financial support from the Canadian Institutes of Health Research/Natural Sciences and Engineering Research Council Collaborative Health Research Program (Grant No. CHRPJ 365423-09; CPG-95268), as well as from DVS Sciences. YL thanks the Chinese Scholarship Council for a scholarship to come to the University of Toronto. We also thank Mr. M. Soleimani for assistance with the TGA measurements, as well as D. Majonis and I. Herrera for helpful discussions.

■

REFERENCES

(1) Boswell, C. A.; Brechbiel, M. W. J. Nucl. Med. 2005, 46, 1946. (2) Valabrega, G.; Montemurro, F.; Aglietta, M. Ann. Oncol. 2007, 18, 977. (3) Costantini, D. L.; McLarty, K.; Lee, H.; Done, S. J.; Vallis, K. A.; Reilly, R. M. J. Nucl. Med. 2010, 7, 1084. (4) Lou, X.; Zhang, G.; Herrera, I.; Kinach, R.; Ornatsky, O.; Baranov, V.; Nitz, M.; Winnik, M. A. Angew. Chem., Int. Ed. 2007, 46, 6111. (5) Majonis, D.; Herrera, I.; Ornatsky, O.; Schulze, M.; Lou, X.; Soleimani, M.; Nitz, M.; Winnik, M. A. Anal. Chem. 2010, 82, 8961. (6) Boyle, A.; Liu, P.; Lu, Y.; Weinrich, D.; Ngo Ndjock Mbong, G.; Winnik M. A.; Reilly, R. To be submitted. (7) Shoichet, M. S. Macromolecules 2010, 43, 581. (8) (a) Uhrich, K. E.; Cannizzaro, S. M.; Langer, R. S; Shakesheff, K. M. Chem. Rev. 1999, 99, 3181. (b) Freiberg, S.; Zhu, X. X. Int. J. Pharm. 2004, 282, 1. (9) Lee, Y.; Miyata, K.; Oba, M.; Ishii, T.; Fukushima, S.; Han, M.; Koyama, H.; Nishiyama, N.; Kataoka, K. Angew. Chem., Int. Ed. 2008, 47, 5163. (10) Uchida, S.; Itaka, K.; Chen, Q.; Osada, K.; Miyata, K.; Ishii, T.; Harada-Shiba, M.; Kataoka, K. J. Controlled Release 2011, 155, 296. (11) Kanayama, N.; Fukushima, S.; Nishiyama, N.; Itaka, K.; Jang, W.-D.; Miyata, K.; Yamasaki, Y.; Chung, U.-I.; Kataoka, K. Chem. Med. Chem. 2006, 1, 439. (12) Robinson, N. E.; Robinson, Z. W.; Robinson, B. R.; Robinson, A. L.; Robinson, J. A.; Robinson, M. L.; Robinson, A. B. J. Peptide Res. 2004, 63, 426. (13) Itaka, K.; Ishii, T.; Hasegawa, Y.; Kataoka, K. Biomaterials 2010, 31, 3707. (14) Robinson, N. E.; Robinson, A. B. Proc. Natl. Acad. Sci. USA 2001, 98, 944. (15) Hermanson, G. T. Bioconjugate Techniques, 2nd ed.; Academic Press, Inc.: 2008. (16) (a) Habraken, G. J. M.; Wilsens, K. H. R. M.; Koning, C. E.; Heise, A. Polym. Chem. 2011, 2, 1322. (b) Vayaboury, W.; Giani, O.; Cottet, H.; Deratani, A.; Schué, F. Macromol. Rapid Commun. 2004, 25, 1221. (c) Knoop, R. J. I.; Habraken, G. J. M.; Gogibus, N.; Steig, S.; Menzel, H.; Koning, C. E.; Heise, A. J. Polym. Sci., Part A: Polym. Chem. 2008, 46, 3068. (17) de Marre, A.; Soyez, H.; Schacht, E.; Pytela, J. Polymer 1994, 35, 2443. (18) Sherry, A. D.; Cacheris, W. P.; Kuan, K.-T. Magn. Reson. Med. 1988, 8, 180. (19) Subramanian, K. M.; Wolf, W. J. Nucl. Med. 1990, 31, 480. 1306

dx.doi.org/10.1021/bm2018239 | Biomacromolecules 2012, 13, 1296−1306