Concentration of polymer nanoparticles through dialysis:
Efficacy and comparison with lyophilization for PEGylated
and zwitterionic systems
Matteo Maraldi1, Raffaele Ferrari2, Renato Auriemma1, Mattia Sponchioni1,*, and Davide Moscatelli1
1Department of Chemistry, Materials and Chemical Engineering “Giulio Natta”,
Politecnico di Milano, Via Mancinelli 7 - 20131 Milano, Italy.
2Department of Chemistry and Applied Biosciences, Institute for Chemical and
Bioengineering, ETH Zurich, Vladimir-Prelog-Weg 1, 8093 Zurich, Switzerland.
* Corresponding author: Mattia Sponchioni; E-mail: mattia.sponchioni@polimi.it
Abstract
Biodegradable polymeric nanoparticles (NPs) are attracting increasing attention as carriers for drug delivery. However, one of the main factors limiting their transition to the market is their premature degradation and release of the payload during the storage. Therefore, for increasing the formulation shelf-life, the removal of water is of paramount importance. In this work, we synthesized both polyethylene glycol (PEG)-stabilized and zwitterionic NPs via Reversible Addition Fragmentation Chain Transfer (RAFT) Polymerization. We demonstrated that lyophilization leads the PEGylated NPs to irreversible aggregation, while the stability of the zwitterionic NPs was preserved only using a cryoprotectant. Therefore, we developed an alternative method for the NP concentration, based on the dialysis against a concentrated PEG solution. This method was optimized in terms of concentration factor (Fc), the ratio between the final and initial NP concentration, by acting on the PEG concentration in the dialysis medium, on its volume and on the initial NP concentration. With this approach, Fc up to 40 can be achieved in less than 10 h, preserving the possibility of redispersing the NPs to their original particle size distribution. Therefore, the dialysis
proposed herein is a valuable alternative to lyophilization for the concentration of polymer NPs preserving their stability.
Keywords: Polymer; Nanoparticles; Dialysis; Lyophilization; Storage; Concentration; PEG; Phosphoryl choline; Zwitterions
1
Introduction
Polymeric nanoparticles (NPs) are a class of colloids in the submicron size scale that are finding growing interest for application in medicine1. In the recent years, a huge effort
has been performed in order to develop biodegradable polymer NPs to be used as carriers for the controlled and targeted delivery of different active principles, including lipophilic drugs, proteins and genetic material2–4. The ability of these colloids to degrade
under physiological conditions and the easiness of elimination of their biocompatible degradation products make them particularly suitable for this application, ensuring that no polymer accumulates in the body5–7. In particular, the recent advances in the field
proposed NPs structurally composed of amphiphilic block-copolymers as a versatile tool for the encapsulation and controlled release of therapeutics8–10. In this
configuration, the lipophilic portion of the copolymer, usually a polyester, forms the biodegradable core of the NPs, whereas the hydrophilic one acts as shell and serves as stabilizer10–12. The versatility of such system arises from the possibility of tuning
independently both parts of the copolymer, each with a specific outcome on the NP physicochemical properties13. In this context, the golden standard in the realization of
the hydrophilic part of the copolymer is represented by poly(ethylene glycol) (PEG) derivatives, able to reduce the protein adsorption on the particle through the hydration layer formed around it14,15. However, to face the so called “accelerated blood clearance”
recently discovered for PEGylated products16–19, valuable alternatives to PEG are
nowadays under investigation1,20. Among all the types of polymers studied,
poly(zwitterions) represent a promising class of materials21–23. Indeed, they permit the
avoidance of the protein opsonization without causing any polymer-specific antibody production thanks to their high hydrophilicity24. Specifically, since its earliest reported
2-methacryloyloxyethyl phosphorylcholine (MPC) has been the mainstay component used in the synthesis of phosphorylcholine-bearing polymers and NPs25–27.
Despite this evolution in the carrier composition and architecture, one major concern in the translation of polymer NPs from the bench to the clinic is still ensuring a sufficiently long shelf-life to the formulation. This requires first of all the preservation of the colloidal stability of the system28,29. Indeed, the storage of the NPs in aqueous
suspensions for a prolonged period causes a physical destabilization of the system which consists in the particle fusion or aggregation30. Furthermore, an additional
drawback that occurs to the system in an aqueous environment is represented by the chemical instability, consisting in the hydrolysis of the polymer that composes the particle. This is particularly relevant for biodegradable polyesters, among which the most typical for the synthesis of polymer NPs include e.g. polycaprolactone, polylactic acid, and poly(lactic acid-co-glycolic acid))31. Finally, an additional issue is the
premature drug leakage from the NPs, which compromises the therapeutic index of the formulation32.
To overcome these problems, the best option, when feasible, is the removal of water from the system. In fact, this would avoid the hydrolysis of the polymer matrix as well as the drug diffusion outside the carrier. There are many methods in literature to concentrate the NP suspensions or to remove completely the water. Among these methods it is worth mentioning evaporation, dialysis and lyophilization33–35. The former
is the most industrially used due to its high efficiency in the removal of both organic solvents and water. However, it is not suitable for drug delivery applications, because of the presence of thermo-labile compounds that may undergo degradation at high temperatures28,36. Another appealing process for drying polymer NPs is the
lyophilization, or freeze-drying. In fact, it enables drying the NPs in an industrially relevant timescale and with reasonable consumption of energy37. However, the main
drawback of this process is the dispersibility of the NPs after drying and the preservation of the colloidal stability once the formulation is re-suspended, due to the several stresses that are produced during the process38. These include the mechanical
stresses that the crystallization of ice exerts on the NPs and the phase separation during freezing leading to a concentrated particulate phase that may induce the NP fusion39. To
protect the colloids during the freezing and desiccation steps40–43. However, even in this
scenario, certain products do not survive coalescence during the process.
Therefore, a strategy that could be industrially appealing for drying polymer NPs preserving their stability after resuspension is urgently needed to improve their shelf-life. This is in fact a fundamental requisite for a new formulation in order to enter the clinical trials.
In this work, we study the possibility of removing water without inducing aggregation from two types of biodegradable NPs already tested as drug delivery carriers24,44,45. Both
of them are obtained from the self-assembly of amphiphilic block copolymers synthesized via reversible addition-fragmentation chain transfer (RAFT) polymerization and sharing the same biodegradable hydrophobic block. This is obtained from macromonomers produced via the ring opening polymerization (ROP) of ε-caprolactone. On the other hand, the hydrophilic part of the copolymer is made of PEG methacrylate-based units in one case and polyMPC in the other, with the aim of comparing the most diffuse NPs for drug delivery with the more recent zwitterionic NPs. Since the former are not able to preserve their stability after lyophilization, we propose and optimize a concentration method based on the dialysis against a concentrated solution of PEG. The performance of this method, expressed in terms of concentration factor (Fc), is discussed with respect to the different operational conditions of the process (i.e. buffer volume, PEG concentration and initial NP concentration). In particular, we show that with this method we can obtain the removal of up to 97% of water in an industrially relevant timescale.
2
Materials and Methods
2.1
Materials
ε-Caprolactone (ε-CL, 97%, MW=114.14 g/mol), 2-hydroxyethyl methacrylate (HEMA, 97%, MW=130.14 g/mol), Ethanol absolute (EtOH, ≥99.8%, MW=46 g/mol), 4,4’-Azobis(4-cyanovaleric acid) (ACVA, ≥98%, MW=280.28 g/mol), 4-Cyano-4-(phenyl-carbothioylthio) pentanoic acid (CPA, ≥97%, MW=279.38 g/mol), toluene (≥99.5%, MW=92.14 g/mol), N,N-Dimethylformamide (DMF, 99%, MW=73.09 g/mol), Potassium peroxodisulfate (KPS, ≥99%, MW=270,33g/mol), Sodium sulfate,
poly(ethylene glycol) methyl ether methacrylate solution (PEGMA2000, 50% w/w
Mn=2000 Da), Tin(II) 2-ethylhexanoate (>92.5-100.0%) and polyethylene glycol (PEG8000, Mn=8000 Da) were purchased from Sigma-Aldrich, Steinheim, Germany.
Cellulose ester membrane (Spectra/Por, Molecular weight cut-off (MWCO) 100-500
Da, wet 0.05% NaN3), SnakeSkin Dialysis Tubing (3.5 kDa MWCO, 35 mm dry) and
Slide-A-Lyzer Dialysis cassettes (3.5 kDa MWCO, 0.5-3mL capacity) were purchased from Fisher-Scientific, Rodano, Italy.
2.2
Synthesis of the hydrophilic Macro-RAFT agent
Two different hydrophilic macromolecular chain transfer agents (macro CTA), which constitute the hydrophilic block of the final copolymer were synthesized via RAFT polymerization. One was obtained from PEGMA and one from MPC. They were synthesized using CPA as RAFT agent, ACVA as initiator and ethanol or a mixture of ethanol/Acetic buffer as solvent. The degree of polymerization of the macro-RAFT agent (n) was set as the ratio between the monomer and the RAFT agent. For the PEG-based macro CTA, 1 g of PEGMA2000 (0.5 mmol), 0.028 g of CPA (0.1 mmol) and 7 mg of ACVA (0.025 mmol) were dissolved in 4 ml of ethanol in a round bottom flask equipped with a magnetic stirrer. The solution was purged by bubbling nitrogen for 15 minutes at room temperature and then heated up to 65 °C in an oil bath under magnetic stirring. After 24 h, 22 mg of ACVA in 100 µl of Ethanol were added to the system to re-initiate the polymerization. After further 24 h, the product obtained was precipitated twice in a 10-fold excess of diethyl ether and centrifuged at 3500 rpm in order to remove the unreacted monomer. The precipitate was dried in a vacuum oven at 35 °C overnight and recovered as a pink powder.
For the MPC-based macro-RAFT agent, 2.24 g of MPC (0.0256mmol), 85 mg of CPA (0.302 mmol) and 0.0283 g of ACVA (0.1 mmol) were dissolved in 10 ml of a solution of ethanol/acetic buffer (10 mM, pH = 4.5) 50/50 v/v. The solution was purged by bubbling nitrogen for 15 minutes at room temperature and then heated up to 65 °C in an oil bath under magnetic stirring. After 24 h, the product obtained was precipitated twice in a 10-fold excess of acetone and centrifuged at 5000 rpm to remove the unreacted
monomer. The precipitate was dried in a vacuum oven at 35 °C overnight and recovered as a pink powder.
The products were analyzed by gel permeation chromatography (GPC) and proton nuclear magnetic resonance (1H NMR, in CDCl
3 for the PEG-based macro CTA and in
deuterium oxide for the polyMPC macro CTA) to assess the molecular weight distribution, degree of polymerization and monomer conversion (see Section 2.8).
2.3
Synthesis of the oligo-ester macromonomer
HEMA-CL macromonomers constitute the hydrophobic block of the final copolymer. HEMA-CL was synthesized via ROP using HEMA as initiator and tin octanoate as catalyst. The molar ratio between the monomer and the initiator provides the degree of polymerization of the macromonomer (q). In order to obtain a macromonomer with
q=5, hereinafter HEMA-CL5,25.4 g of CL (0.222 mmol) and 15 mg of Na2SO4 were
added to a round bottom flask equipped with a magnetic stirrer. The flask was closed with a rubber stopper and heated to 125 °C in an oil bath. After that, a solution of 5.85 g of HEMA (45 μmol) and 90 mg of Tin(II) 2-ethylhexanoate (0.225 μmol) was added to the flask under stirring. The polymerization proceeded for 2.5 h. The resulting HEMA-CL5 was characterized by GPC and NMR in CDCl3 to assess the average molecular
weight and degree of polymerization (see Section 2.8).
2.4
Block copolymer synthesis
The hydrophilic macro CTAs were chain extended with HEMA-CL5 via RAFT
emulsion polymerization to produce amphiphilic block copolymers self-assembled into NPs. The degree of polymerization of the final block is given by the molar ratio between the macromonomer and the macro CTA (p). Briefly, to obtain 5PEGMA200060CL5, where 5 and 60 represent n and p, respectively (see Scheme 1), 0.2
g of 5PEGMA2000, 0.8 g of HEMA-CL5 and 2 mg of ACVA were dissolved in a mixture
of ethanol/acetic buffer (10 mM, pH=4.5) 30/70 v/v in a round bottom flask equipped with a magnetic stirrer. The solution was purged by bubbling nitrogen for 15 minutes at room temperature and then heated to 65 °C in an oil bath under magnetic stirring. After 24 h, 2 mg of ACVA were added to the system to re-initiate the polymerization. After further 24 h, the reaction was quenched by exposure to air and the product dialyzed
against 1 L of deionized water using regenerated cellulose (RC) membranes with a MWCO of 3.5 kDa (Spectra/Por) for 24 h. Size and polydispersity were analyzed through dynamic light scattering (DLS) analysis using a Malvern Zetasizer Nano ZS. In order to obtain the MPC-based NPs (25MPC30CL5), 52 mg of 25MPC, 0.144 g of HEMA-CL5 and 0.6 mg of ACVA were dissolved in a mixture of ethanol/acetic buffer
(10 mM, pH=4.5) 30/70 v/v. The solution was purged by bubbling nitrogen for 15 minutes at room temperature and then heated to 65 °C in an oil bath under magnetic stirring. After 24 h, the NPs obtained were dialyzed against deionized water as reported previously. Size and polydispersity were analyzed through DLS analysis using a Malvern Zetasizer Nano ZS.
2.5
Concentration by Dialysis
A weighted amount of the nanoparticle suspension (10 g) was introduced in a pre-weighted dialysis bag SnakeSkin Dialysis Tubing (3.5 kDa MWCO) and dialyzed against a solution of PEG8000 at concentrations ranging from 20 to 180 g/L in water. The
volume of the counter-dialysis medium ranged from 10 to 200 mL and the dialysis was performed at room temperature for 24 h. At different time points, the dialysis tube was collected, carefully dripped and weighted to determine the loss of water from the NP suspension. Finally, it was placed again in the dialysis medium.
2.6
Lyophilization
The NP suspension was diluted 3 mg/ml. In order to perform the freeze-drying process, a certain amount of glucose was added to 2 ml of NP suspension, as cryoprotectant. Specifically, different concentrations of glucose were used: 0.1; 0.5; 1.0; 2.0 % w/w. Once the glucose was completely solubilized, the suspension was quickly frozen with liquid nitrogen and then lyophilized for 24 h. The freeze-dryer (Telstar Lyoquest-Eco55) was set to -56 °C at a pressure of 0.1 mBar. After this procedure, the dried NPs were re-dispersed in 2 mL of distilled water. The suspension obtained was analyzed via DLS to determine the particle size distribution and average hydrodynamic diameter.
2.7
Characterization techniques
The structure and the effective number of repeating units composing the polymer chains were determined by characterizing all the macromonomers and the polymers synthesized via 1H NMR analysis, performed on a Bruker 400 MHz spectrometer. The
average molecular weight (MW) and the MW distribution were characterized using either aqueous or organic size exclusion chromatography (SEC) analysis. The organic SEC was performed using THF as eluent at 1 ml/min flow rate and 35 °C. The instrument (Jasco 2000 series) was equipped with differential refractive index (RI) three PL gel columns (Polymer laboratories Ltd., UK; two columns had pore sizes of the Mixed-C type and one was an oligopore; 300 mm length and 7.5 mm ID) and a precolumn. A universal calibration was applied based on polystyrene (PS) standards from 580 Da to 3,250,000 Da (Polymer Laboratories). Whereas the aqueous SEC was performed using a mixture of 0.05 M Na2SO4 water solution and acetonitrile (ACN)
80/20 v/v and a 0.5 ml/min flow rate at a temperature of 35 °C. The instrument (Jasco 2000 series) was equipped with differential refractive index (RI) and three Suprema columns (Polymer Standards Service; particle size 10 mm, pore sizes of 100, 1000, and 3000 Å) and a precolumn. A universal calibration was applied based on PEG standards (Polymer Laboratories). DLS measurements were performed with a Malvern Zetasizer Nano ZS at a scattering angle of 173°. The sample was filtered with a 0.45 μm syringe filter to remove dust and diluted to 0.5% w/w with pre-filtered deionized water before the measurement. Three independent measurements, 11 runs each, were taken. Both PEGylated and MPC-based NPs were analyzed before and after the concentration, either through lyophilization or dialysis. In particular, after concentration, the NP suspension was diluted to 0.5% w/w with pre-filtered deionized water and vortex-mixed for 30 seconds before the analysis.
3
Results and discussion
3.1
NP synthesis
First, we synthesized two types of NPs composed of amphiphilic block-copolymers with the same lipophilic part but different hydrophilic blocks. The synthesis is achieved by a combination of ROP and RAFT polymerization, as shown in Scheme 1.
In the first step, the oligoester macromonomer adopted for the synthesis of the lipophilic core of the NPs was produced through ROP of CL using HEMA as initiator. The good control offered by the ROP enabled us to obtain an oligoester (HEMA-CL5) with an
average degree of polymerization (q) close to the target of 5 (see Figure S1 and Table
1) and able to further polymerize via free radical chemistry due to the vinyl bond of
HEMA.
In the second step, the hydrophilic macro CTAs (25MPC and 5PEGMA2000) were
synthesized via RAFT solution polymerization. Also in this case, it was possible to tune the average degree of polymerization (n) to the target values of 5 for the PEGMA and 25 for the polyMPC (full NMR characterization in Figures S2 and S3, respectively, and details in Table 1) by simply modulating the mole ratio between the monomer and CTA. The good control of the RAFT over the polymerization is then testified by the narrow molecular weight distribution, with polydispersity (Ð) lower than 1.3, as expected for a pseudo-living polymerization46.
Finally, we chain-extended the macro CTAs produced in this step with HEMA-CL5 via
RAFT emulsion polymerization to obtain the final block-copolymers and their simultaneous self-assembly into core-shell NPs. The hydrophobic polyester is in fact expected to be confined in the NP core, with the hydrophilic portion exposed to the surface. High monomer conversions and good control over the degree of polymerization (p) is confirmed by 1H NMR (see Table 1 and Figures S4 and S5). Also, the produced
NPs proved to be stable and narrowly dispersed, with a size suitable for drug delivery applications (Table 2).
Overall, both the lipophilic block and the NP size for the two samples under investigation were comparable, thus allowing the direct analysis of the effect of the hydrophilic block composition over the most suitable storage protocol, choosing between dialysis and lyophilization.
3.2
PEG-based NPs
With narrowly distributed biodegradable NPs in our hands, we first optimized the storage protocol to preserve the physicochemical properties of PEG-based NPs, which represent the golden standard for drug delivery.
Specifically, the first method tested was the lyophilization. This is the technique of choice in the clinical practice, mainly due to the complete removal of water as well as the avoidance of heating, which could lead to degradation of the payload13. However,
the PEG-based NPs synthesized in this work did not survive the freeze-drying also when up to 10% w/w of cryoprotectant was added. Indeed, after rehydration, only large particles, not suitable for systemic administration, were obtained (see Figure S6). This may be due to the poor ability of PEG of crystalizing at low temperature. Indeed, since the polymer is characterized by a low glass transition temperature, once the water is removed, the stabilizer block may be embedded in the polymer, thus causing coalescence. We then demonstrated that the freeze-drying is not a suitable storage process for this kind of PEGylated NPs and alternative techniques are required.
In particular, we tested the possibility of concentrating these NPs by dialysis against a solution of PEG8000. This is a cheap and widely available polymer, differently from other
compounds proposed in the literature for similar purposes, such as dextran47. According
to the work of Stanley et al.48, the osmotic pressure (π) of PEG
8000 solutions can be
represented as a polynomial function of the PEG to water ratio (w/w) (G), as reported in
Equations 1 and 2. π (atm)=−1.38 G2 ∗T +134.3 G2+3.0G for 0.11<G<1.18(10<wt %<54) ¿10<T <40 °C (1) π (atm)=−1.29 G2 ∗T +127.6 G2+3.0G for 0.03<G<0.11(3 <wt %<10) ¿10<T <40 °C (2)
A NP suspension contained in a dialysis membrane immersed in a high osmotic pressure solution receives an osmotic stress that produces a flow of water from inside to outside the membrane. This natural movement of water serves to counterbalance the difference in osmotic pressure between the two sides of the dialysis membrane. Consequently, this process causes the suspension of NPs to be concentrated until there is an osmotic balance on both sides of the membrane. To quantify this process, we took
into account the concentration factor, Fc. This parameter is defined as the ratio between the final and the initial NP concentration (Cfin and C0, respectively), as described in
Equation 3.
Fc=Cfin C0
(3)
In order to optimize this process for the concentration of a suspension of polymeric NPs, we first conducted low-volume tests. Specifically, 1.5 ml of 1% w/w NP suspensions were dialyzed against 5 mL of PEG8000 solutions at different concentrations.
Kinetic analysis were then carried out to track the concentration of both the NP suspension and the polymer solution over time as shown in Figure 1 and Figure S7, respectively.
As it can be retrieved from Figure 1a, independently from the PEG8000 concentration,
the presence of the polymer is able to produce an osmotic pressure that causes the migration of water out of the dialysis membrane and in turn the NP concentration. This concentration led to a final weight of the suspension up to 50% of the initial weight. In turn, we excluded that this reduction in weight could be due to external phenomena (i.e. evaporation) by demonstrating that the weight lost by the NP suspension is gained by the dialysis medium (see Table S1). Moreover, we demonstrated that by increasing the concentration of PEG in the medium, we were able to increase both the concentration rate as well as the Fc reached at the equilibrium, as shown in Figure 1b. However, comparing the different concentrations of PEG8000 tested, it can be noticed that for small
volumes of NP suspensions, only a partial amount of water is removed, limiting the potential of the technique and thus the ability to concentrate NPs. Furthermore, the tests performed increasing the concentration of the polymer show that it is not possible to achieve concentration factors higher than 2, for small volumes.
Therefore, in order to investigate more in details the effect of the osmotic pressure on the concentration of NP suspensions, we designed a second set of experiments for larger volumes of NP suspensions and buffers. Five PEG8000 concentrations were investigated,
namely 20 mg/ml, 43 mg/ml, 87 mg/ml, 130 mg/ml and 180 mg/ml, and the volumes of buffer were scaled-up to 100 ml and 200 ml, as described in Table S2. Indeed, we show in Figure 2 how in this case we were able to remove a significantly larger amount of
water from the NP suspension. Compared to smaller volumes, it is possible to observe in Figure 2a that more than 95% of the suspension weight is lost at the equilibrium. Furthermore, the concentration of PEG8000 in the dialysis medium significantly
influences the water removal rate, which represents a fundamental aspect for the scale-up of this method and its applicability in the clinic. It is worth mentioning that before reaching the equilibrium, the system is characterized by a linear decrease of the suspension weight during time. Hence, the concentration factor follows a zero-order kinetic with time with the slope depending on the PEG8000 concentration. Furthermore,
with PEG8000 concentrations higher than 87 mg/ml, the equilibrium is reached within 24
h, which is not much higher than the time required for the complete freeze-drying of a NP suspension. However, if we further increase the PEG8000 concentration up to 180
mg/ml we were able to remove more than 97% of water in less than 10 h, a suitable timescale for a large scale application. Indeed, as shown in Figure 2b, the Fc at equilibrium seems to only depend on the PEG concentration and not on the buffer volume.
In fact, we found that Fc shows an exponential increase with the osmotic pressure π, as highlighted in Figure 3a. In this Figure, it is possible to retrieve that Fc is also affected by the initial concentration of the NP suspension. Interestingly, higher Fc can be obtained at increasing NP initial concentration. Specifically, this dependence follows the relation expressed in Equation 4.
Fc=1.7 eBπ (4)
Where π is a polynomial function of the PEG concentration as defined in Equation 2 and B is a linear function of the initial concentration of the NP suspension (C0) as
described in Equation 5.
B=0.02 C0+0.71 (5)
The mathematical model expressed by Equations 4-5 and plotted as continuous lines in
Figure 3a is in good agreement with the set of experimental data (symbols) also
showed in Figure 3a. Moreover, the model allows to predict the trend of Fc by taking into account both the contribution of π and C0 .In fact, it can be used as guidelines in
the tuning of the concentration factor at the equilibrium by varying the initial concentration of NPs and the polymer concentration as depicted in Figure 3b.
Finally, after having demonstrated that high Fc can be obtained in an industrially-relevant time scale, we also confirmed that through this method, the NP stability after concentration and dilution is preserved. In fact, independently from the initial concentration of the NP suspension, no aggregate formation was observed. Furthermore, the NPs showed almost unchanged particle size distribution before and after dialysis (Figure S8), with polydispersity indexes always lower than 0.1, a clear indication that the NP aggregation was avoided49,50.
Overall, for PEG-based NPs, the lyophilization is not a valuable option for the removal of water, due to the impossibility of reconstituting the NPs to their original particle size distribution. However, the dialysis demonstrated to be an efficient process to remove up to 97% of the water present in the formulation without inducing destabilization of the NPs and in an industrially-relevant timescale. This phenomenon is particularly interesting considering also the exponential dependence of the Fc on the osmotic pressure and thus on the PEG8000 concentration that can further enhance significantly the
removal of water from the system. This can be taken in consideration for reducing the costs for transportation as well as to limit the premature drug diffusion from the NP core.
3.3
MPC-based NPs
First, we wanted to demonstrate that the dialysis is an effective method for concentrating MPC-based NPs as well as for the PEGylated ones. Therefore, we tracked the Fc of the MPC-based biodegradable NPs during time at a concentration of PEG8000 in
the dialysis medium of 130 mg/ml. As shown in Figure 4, the dialysis proved to be effective for both the types of NPs. In fact, the colloidal system stabilized by the MPC block achieved the same Fc as that of PEGylated NPs. Furthermore, the rate of water removal was similar for the two systems, confirming that the ability of concentrating the suspension depends mainly on the process itself and not on the composition of the NPs. Once the dialysis method was proved to be efficient for this type of NPs, the lyophilization was performed in order to achieve a complete removal of water from the suspension. MPC-based NPs were concentrated by lyophilization using 1% w/w of
glucose as cryoprotectant. The NPs were then re-suspended in water and their particle size distribution before and after freeze-drying compared in Figure 5.
The result shows a small increase in the NP size after the lyophilization and re-dispersion. However, the average size is still in an acceptable range for drug delivery (30-300 nm). In addition, the NPs preserve their narrow distribution, thus confirming an efficient re-dispersability of the formulation differently from the PEG-based NPs. This may be due to the ability of the MPC to crystallize at low temperature. This phenomenon stabilizes the formulation avoiding the coalescence of the polymeric particles. However, even in this case, a low percentage of cryoprotectant was required to enable the re-dispersion of the NPs preserving the original size. Therefore, the dialysis method is still a valuable alternative when the addition of exogenous compounds is unacceptable.
4
Conclusions
In this work, we optimized an alternative method to the traditional freeze-drying for the removal of water from biodegradable polymer NP suspensions. In fact, we demonstrated that lyophilization leads to irreversible aggregation in the case of PEGylated NPs, the current golden standard as drug carriers. Therefore, we proposed and optimized the dialysis of NP suspensions against a concentrated PEG8000 solution as
an efficient process for the NP concentration. Through this method, at a PEG concentration of 180 mg/ml it was possible to obtain Fc up to 40 in less than 10 h, without inducing the NP destabilization. We proposed an exponential dependency of the Fc reached at the equilibrium on the osmotic pressure, being the exponent a linear function of the initial NP concentration. The same protocol demonstrated to be efficient also for the concentration of MPC-based NPs. Indeed, for these NPs also the lyophilization enabled to obtain a redispersible product, provided that 1% w/w of cryoprotectant is added. Then, the dialysis can be a valuable solution when cryoprotectants are not tolerated.
Supporting Material: Electronic supporting information are available on the publisher
detailed parameters of the dialysis experiments and the proof of preserved NP stability after dialysis.
Acknowledgments: The authors are grateful to Luca Garghentini for his help with the
freeze-drying experiments.
Conflict of Interests: The authors declare no conflict of interests for this work.
References
1. Ferrari, R., Sponchioni, M., Morbidelli, M. & Moscatelli, D. Polymer nanoparticles for the intravenous delivery of anticancer drugs: the checkpoints on the road from the synthesis to clinical translation. Nanoscale 10, 22701–22719 (2018).
2. Salata, O. V. Applications of nanoparticles in biology and medicine. J.
Nanobiotechnology 2, 3 (2004).
3. Zhang, L. et al. Nanoparticles in Medicine: Therapeutic Applications and Developments. Clin. Pharmacol. Ther. 83, 761–769 (2008).
4. Panyam, J. & Labhasetwar, V. Biodegradable nanoparticles for drug and gene delivery to cells and tissue. Adv. Drug Deliv. Rev. 55, 329–347 (2003).
5. Moscatelli, D. & Sponchioni, M. Bioresorbable polymer nanoparticles in the medical and pharmaceutical fields. in Bioresorbable Polymers for Biomedical
Applications 265–283 (Elsevier, 2017).
doi:10.1016/B978-0-08-100262-9.00012-4.
6. Hans, M. L. & Lowman, A. M. Biodegradable nanoparticles for drug delivery
and targeting. Curr. Opin. Solid State Mater. Sci. 6, 319–327 (2002).
7. Kamaly, N., Yameen, B., Wu, J. & Farokhzad, O. C. Degradable
Controlled-Release Polymers and Polymeric Nanoparticles: Mechanisms of Controlling Drug Release. Chem. Rev. 116, 2602–2663 (2016).
Their Use in Stabilizing Fully Biodegradable Polymer Nanoparticles. Macromol.
Chem. Phys. 218, 1700380 (2017).
9. Wang, F., Bronich, T. K., Kabanov, A. V, Rauh, R. D. & Roovers, J. Synthesis and Evaluation of a Star Amphiphilic Block Copolymer from Poly(ε-caprolactone) and Poly(ethylene glycol) as a Potential Drug Delivery Carrier.
Bioconjug. Chem. 16, 397–405 (2005).
10. Adams, M. L., Lavasanifar, A. & Kwon, G. S. Amphiphilic block copolymers for drug delivery. J. Pharm. Sci. 92, 1343–1355 (2003).
11. Kabanov, A. V & Alakhov, V. Y. Micelles of amphiphilic block copolymers as vehicles for drug delivery. in (eds. Alexandridis, P. & Lindman, B. B. T.-A. B. C.) 347–376 (Elsevier Science B.V., 2000). doi:https://doi.org/10.1016/B978-044482441-7/50016-5.
12. Rösler, A., Vandermeulen, G. W. M. & Klok, H.-A. Advanced drug delivery
devices via self-assembly of amphiphilic block copolymers. Adv. Drug Deliv.
Rev. 64, 270–279 (2012).
13. Capasso Palmiero, U., Sponchioni, M., Manfredini, N., Maraldi, M. & Moscatelli, D. Strategies to combine ROP with ATRP or RAFT polymerization for the synthesis of biodegradable polymeric nanoparticles for biomedical applications. Polym. Chem. 9, 4084–4099 (2018).
14. Veronese, F. M. & Pasut, G. PEGylation, successful approach to drug delivery.
Drug Discov. Today 10, 1451–1458 (2005).
15. Jokerst, J. V, Lobovkina, T., Zare, R. N. & Gambhir, S. S. Nanoparticle PEGylation for imaging and therapy. Nanomedicine 6, 715–728 (2011).
16. Shimizu, T. et al. Anti-PEG IgM and complement system are required for the
association of second doses of PEGylated liposomes with splenic marginal zone B cells. Immunobiology 220, 1151–1160 (2015).
17. Shiraishi, K. et al. Exploring the relationship between anti-PEG IgM behaviors and PEGylated nanoparticles and its significance for accelerated blood clearance.
J. Control. Release 234, 59–67 (2016).
18. Ishihara, T. et al. Accelerated Blood Clearance Phenomenon Upon Repeated Injection of PEG-modified PLA-nanoparticles. Pharm. Res. 26, 2270–2279 (2009).
19. Ishida, T. & Kiwada, H. Accelerated blood clearance (ABC) phenomenon upon repeated injection of PEGylated liposomes. Int. J. Pharm. 354, 56–62 (2008).
20. Sponchioni, M., Morosi, L., Lupi, M. & Capasso Palmiero, U.
Poly(HPMA)-based copolymers with biodegradable side chains able to self assemble into nanoparticles. RSC Adv. 7, 50981–50992 (2017).
21. Sponchioni, M., Rodrigues Bassam, P., Moscatelli, D., Arosio, P. & Capasso Palmiero, U. Biodegradable zwitterionic nanoparticles with tunable UCST-type phase separation under physiological conditions. Nanoscale 11, 16582–16591 (2019).
22. Cao, Z., Yu, Q., Xue, H., Cheng, G. & Jiang, S. Nanoparticles for Drug Delivery Prepared from Amphiphilic PLGA Zwitterionic Block Copolymers with Sharp Contrast in Polarity between Two Blocks. Angew. Chemie Int. Ed. 49, 3771– 3776 (2010).
23. Ladd, J., Zhang, Z., Chen, S., Hower, J. C. & Jiang, S. Zwitterionic Polymers Exhibiting High Resistance to Nonspecific Protein Adsorption from Human Serum and Plasma. Biomacromolecules 9, 1357–1361 (2008).
24. Capasso Palmiero, U., Maraldi, M., Manfredini, N. & Moscatelli, D. Zwitterionic Polyester-Based Nanoparticles with Tunable Size, Polymer Molecular Weight, and Degradation Time. Biomacromolecules 19, 1314–1323 (2018).
25. KADOMA & Y. Synthesis and Hemolysis Test of the Polymer Containing Phosphorylcholine Groups. Koubunshi Ronbunshu 35, 423–427 (1978).
26. Konno, T., Kurita, K., Iwasaki, Y., Nakabayashi, N. & Ishihara, K. Preparation of nanoparticles composed with bioinspired 2-methacryloyloxyethyl phosphorylcholine polymer. Biomaterials 22, 1883–1889 (2001).
27. Lomas, H. et al. Biomimetic pH Sensitive Polymersomes for Efficient DNA Encapsulation and Delivery. Adv. Mater. 19, 4238–4243 (2007).
28. Chacón, M., Molpeceres, J., Berges, L., Guzmán, M. & Aberturas, M. R. Stability and freeze-drying of cyclosporine loaded poly(d,l lactide–glycolide) carriers. Eur. J. Pharm. Sci. 8, 99–107 (1999).
29. De Jaeghere, F. et al. Formulation and Lyoprotection of Poly(Lactic Acid-Co-Ethylene Oxide) Nanoparticles: Influence on Physical Stability and In Vitro Cell Uptake. Pharm. Res. 16, 859–866 (1999).
30. Lemoine, D. et al. Stability study of nanoparticles of poly(ɛ-caprolactone), poly(d,l-lactide) and poly(d,l-lactide-co-glycolide). Biomaterials 17, 2191–2197 (1996).
31. Cingolani, A. et al. A Methodologic Approach for the Selection of Bio-Resorbable Polymers in the Development of Medical Devices: The Case of Poly(l-lactide-co-ε-caprolactone). Polymers (Basel). 10, 851 (2018).
32. Coffin, M. D. & McGinity, J. W. Biodegradable Pseudolatexes: The Chemical Stability of Poly(D,L-Lactide) and Poly (ε-Caprolactone) Nanoparticles in Aqueous Media. Pharm. Res. 9, 200–205 (1992).
33. Tagalakis, A. D. et al. A method for concentrating lipid peptide DNA and siRNA nanocomplexes that retains their structure and transfection efficiency. Int. J.
Nanomedicine 10, 2673–2683 (2015).
34. Dalwadi, G., Benson, H. A. E. & Chen, Y. Comparison of Diafiltration and Tangential Flow Filtration for Purification of Nanoparticle Suspensions. Pharm.
Res. 22, 2152–2162 (2005).
35. Abdelwahed, W., Degobert, G., Stainmesse, S. & Fessi, H. Freeze-drying of nanoparticles: Formulation, process and storage considerations. Adv. Drug Deliv.
Rev. 58, 1688–1713 (2006).
36. Harivardhan Reddy, L., Vivek, K., Bakshi, N. & Murthy, R. S. R. Tamoxifen Citrate Loaded Solid Lipid Nanoparticles (SLNTM): Preparation, Characterization,
In Vitro Drug Release, and Pharmacokinetic Evaluation. Pharm. Dev. Technol.
11, 167–177 (2006).
37. Franks, F. Freeze-drying of bioproducts: putting principles into practice. Eur. J.
Pharm. Biopharm. 45, 221–229 (1998).
38. Choi, M. J., Briançon, S., Andrieu, J., Min, S. G. & Fessi, H. Effect of Freeze-Drying Process Conditions on the Stability of Nanoparticles. Dry. Technol. 22, 335–346 (2004).
39. Abdelwahed, W., Degobert, G. & Fessi, H. A pilot study of freeze drying of poly(epsilon-caprolactone) nanocapsules stabilized by poly(vinyl alcohol): Formulation and process optimization. Int. J. Pharm. 309, 178–188 (2006). 40. Fonte, P. et al. Stability Study Perspective of the Effect of Freeze-Drying Using
Biomacromolecules 15, 3753–3765 (2014).
41. Date, P. V, Samad, A. & Devarajan, P. V. Freeze Thaw: A Simple Approach for Prediction of Optimal Cryoprotectant for Freeze Drying. AAPS PharmSciTech
11, 304–313 (2010).
42. Bozdag, S., Dillen, K., Vandervoort, J. & Ludwig, A. The effect of freeze-drying with different cryoprotectants and gamma-irradiation sterilization on the characteristics of ciprofloxacin HCl-loaded poly(D,L-lactide-glycolide) nanoparticles. J. Pharm. Pharmacol. 57, 699–707 (2005).
43. Anchordoquy, T. J., Armstrong, T. K. & Molina, M. d. C. Low Molecular Weight Dextrans Stabilize Nonviral Vectors During Lyophilization at Low Osmolalities: Concentrating Suspensions by Rehydration to Reduced Volumes.
J. Pharm. Sci. 94, 1226–1236 (2005).
44. Lu, Q., Yi, M., Zhang, M., Shi, Z. & Zhang, S. Folate-Conjugated Cell Membrane Mimetic Polymer Micelles for Tumor-Cell-Targeted Delivery of Doxorubicin. Langmuir 35, 504–512 (2019).
45. Ferrari, R. et al. Biocompatible Polymer Nanoformulation To Improve the Release and Safety of a Drug Mimic Molecule Detectable via ICP-MS. Mol.
Pharm. 14, 124–134 (2017).
46. Sponchioni, M., Capasso Palmiero, U., Manfredini, N. & Moscatelli, D. RAFT copolymerization of oppositely charged monomers and its use to tailor the composition of nonfouling polyampholytes with an UCST behaviour. React.
Chem. Eng. 4, 436–446 (2019).
47. Vauthier, C., Cabane, B. & Labarre, D. How to concentrate nanoparticles and avoid aggregation? Eur. J. Pharm. Biopharm. 69, 466–475 (2008).
48. Stanley, C. B. & Strey, H. H. Measuring Osmotic Pressure of Poly(ethylene glycol) Solutions by Sedimentation Equilibrium Ultracentrifugation.
Macromolecules 36, 6888–6893 (2003).
49. Huynh, K. A. & Chen, K. L. Aggregation Kinetics of Citrate and Polyvinylpyrrolidone Coated Silver Nanoparticles in Monovalent and Divalent Electrolyte Solutions. Environ. Sci. Technol. 45, 5564–5571 (2011).
50. Gillich, T. et al. PEG-Stabilized Core–Shell Nanoparticles: Impact of Linear versus Dendritic Polymer Shell Architecture on Colloidal Properties and the
Reversibility of Temperature-Induced Aggregation. ACS Nano 7, 316–329 (2013).
