Serum Albumin-Alginate Microparticles Prepared by Transacylation

Subscriber access provided by NEW YORK UNIV

Article

Serum Albumin-Alginate Microparticles Prepared by Transacylation: Relationship between Physicochemical, Structural and Functional Properties Imane Hadef, Barbara ROGE, and Florence Edwards-Lévy Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.5b00536 • Publication Date (Web): 29 Jun 2015 Downloaded from http://pubs.acs.org on July 5, 2015

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Biomacromolecules is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Serum Albumin-Alginate Microparticles Prepared by Transacylation: Relationship between Physicochemical, Structural and Functional Properties AUTHOR NAMES Imane Hadef (,), Barbara Rogé (), Florence Edwards-Lévy ()* AUTHOR ADDRESS Institut de Chimie Moléculaire de Reims, CNRS UMR 7312. () U.F.R. Pharmacie, 51, rue Cognacq-Jay, 51096 Reims, France. () U.F.R. Sciences Exactes et Naturelles, Campus du Moulin de la Housse 51687 Reims, France.

ABSTRACT:

Our laboratory develops a method of microencapsulation using a transacylation reaction in a W/O emulsion. The method is based on the creation of amide bonds between free amine functions of a protein (human serum albumin (HSA)) and ester groups of propylene glycol

ACS Paragon Plus Environment

1

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 42

alginate (PGA) in the inner aqueous phase after alkalization. The aim of this work is to study the influence of physicochemical properties of HSA-PGA mixtures on microparticle characteristics. Microparticles were prepared varying the concentrations of PGA and HSA, then characterized (inner structure, size, swelling rate, release kinetics). PGA and each polymer mixture used in the microencapsulation procedure were examined in order to elucidate the mechanism of microstructure formation. It was found that the morphology and functional properties of HSAalginate microparticles were related to the two polymer concentrations in the aqueous solution. Actually, the polymer concentration variations led to physicochemical changes, which affected the microparticle structure and functional properties.

KEYWORDS: Microparticles, transacylation, serum albumin, alginate, internal structure, swelling rate, release kinetics. INTRODUCTION Microencapsulation is a process by which tiny particles (solids, liquids or gases) are surrounded by a coating or embedded in a homogeneous or heterogeneous matrix to give small microspheres/microcapsules of 1 µm to 1 mm average diameter. This technology has been used in several fields (including pharmaceutics, agriculture, food, printing, cosmetics and textile, etc.) in order to isolate and protect the encapsulated substances from the external environment as well as to control their release profile 1,2,3. Among the numerous available microencapsulation techniques, the selection of the appropriate process is governed by the physical and chemical properties of core and coating materials and by the intended application 3,4,5. A chemical encapsulation method was developed in our laboratory, based on the use of a transacylation


2

Page 3 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

reaction, creating covalent bonds between proteins and polysaccharides 6,7 whilst avoiding bifunctional reagents generally used for protein-polysaccharide cross-linking8,9. Indeed, this method involves the gel formation ability of propylene glycol alginate (PGA), in alkaline medium, by reaction with polyamine compounds (e.g. proteins) 10 (Figure 1). The use of such natural substances (proteins and polysaccharides) leads to biocompatible and biodegradable microparticles, making them appropriate for biomedical applications. Moreover, the covalent bonds created between these polymers enhance the microparticle stability.

Figure 1. Transacylation reaction between protein and PGA The process was first implemented to form stable membranes around alginate spheres. In this procedure, the PGA and the protein were added to a Na-alginate solution: gel spheres were formed by dropwise addition into a calcium solution (macrospheres) 6 or by calcium gelation in W/O emulsion (microspheres) 7,11 . The transacylation reaction was then started by alkalization of the suspension, giving rise to a membrane formed around the spheres, made of a protein directly bound to the polysaccharide through amide bonds. The multi-stage protocol was simplified afterwards by getting rid of the calcium alginate spheres and keeping only the PGA and the protein in the aqueous solution. Microparticles, formed of a covalent network, could be prepared using transacylation between the two aqueous polymers in a W/O emulsion after alkalization (Figure 2) 12,13. The preliminary microscopic observations carried out on the cross


3

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 42

sections of HSA-alginate microparticles prepared via the latter procedure showed differences in their inner structure depending on polymer content in the aqueous solution.

Figure 2. Microparticle preparation using a transacylation reaction between HSA and PGA in W/O emulsion Hence, the aim of this work is to elucidate the influence of HSA and PGA concentrations as well as their physicochemical properties on microparticle characteristics, especially on their internal structure, swelling capacity and drug release kinetics. For that purpose, microparticles were prepared using the transacylation reaction in W/O emulsion at various concentrations of HSA and PGA. Their internal structures were observed on cross sections, and their swelling rates were evaluated after numerous cycles of water washings. PGA and each polymer mixture used in the microencapsulation procedure were characterized in order to understand the mechanism of microstructure formation and to identify a relationship between polymer physicochemical properties (esterification degree, pH, viscosity and thermodynamic incompatibility) and microparticle characteristics. Moreover, a comparative study of lysozyme release kinetics from microparticles of distinct inner structures was also realized with the aim of proving the advantage


4

Page 5 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

of using HSA-alginate microparticles for drug sustained release and demonstrating the effect of microparticle inner structure on their functional properties. MATERIALS AND METHODS 1. Microparticle preparation Microparticles were prepared by transacylation between HSA and PGA in W/O emulsion. The aqueous phase consisted of a HSA-PGA aqueous solution while the oily phase was composed of isopropyl myristate added with a surfactant. 1.1.Preparation of HSA-PGA aqueous solutions Human serum albumin (HSA) was provided by LFB biomédicaments (Vialebex, 200mg/mL injection solution) and was freeze-dried before use. Propylene glycol alginate (PGA) (Profoam, 84% esterification degree) was kindly offered by FMC BioPolymer. The aqueous solutions were prepared at various mass concentrations of PGA (1% to 4% (w/w)) and HSA (2.5% to 20% (w/w)) in water (water for irrigation, Versol, Aguettant). The solutions were kept 24 h at 4°C, then characterized (viscosity, microscopic observation and pH measurement) and used in the preparation procedure. 1.2.Preparation of the microparticles Standard procedure. After elimination of air bubbles by centrifugation, 6 mL of HSA-PGA aqueous solution was emulsified at 1500 rpm in 40 mL of isopropyl myristate (SDF) containing 5% (w/v) sorbitan trioleate (Span 85, Sigma-Aldrich), with a mechanical stirrer (Heidolph RGL500 stirring motor, Prolabo) equipped with an anchor-shaped blade.


5

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 42

After 5 min stirring, the transacylation reaction was triggered by adding 4 mL of 2% (w/v) NaOH /EtOH (95% (v/v)) solution to the emulsion, and agitation was maintained for a further 15 min. The reaction was stopped by adding 150 mL of imidazole buffer (0,05M, pH 7) and agitation was continued for 2 min. Microparticles were separated by centrifugation and washed successively with a 2% (w/v) polysorbate aqueous solution (Tween20, Acros-Organics) and with distilled water. Modified procedures. Some variations were carried out separately on the standard procedure; all other parameters previously detailed were kept unchanged. Polymer concentrations were fixed at 10% HSA and 2% PGA (w/w). First, the effect of the amount of NaOH solution was studied by adding 8 mL of 2% (w/v) NaOH /EtOH (95% (v/v)) solution to the emulsion instead of 4 mL used in the standard procedure. Two further experiments were also performed by preparing the HSA-PGA solution in phosphate buffer of pH 5 and 8, in order to study the influence of the pH of the initial aqueous solution on microparticle properties. Another assay was achieved using PGA of 55 % esterification degree (FMC BioPolymer) with the aim of identifying the effect of ester amount on microparticle characteristics. 2. Polymer characterization 2.1.Determination of PGA esterification degree


6

Page 7 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

The degree of esterification (DE) is defined as the amount of ester groups (COOR) compared to the total amount of carboxylic acid and ester groups (COOH+COOR). The degree of esterification of PGA was confirmed by two methods: titrimetry and 1H-NMR analysis. Titrimetry is a universal method generally used for the determination of polysaccharide DE, nevertheless it requires a large amount of ester. 1H-NMR analysis could solve the last inconvenience but it needs to be validated, by a standard method, for each polysaccharide ester. Titrimetry. 5 g of PGA was previously acidified in a mixture of 5 mL HCl (2.7 M) and 100 mL ethanol (60% (v/v)) for 10 min 14 and then precipitated in 150 mL of ethanol (95 %(v/v)). The suspension was centrifuged at 5000 rpm for 5 min (Beckman GS.15 centrifuge), filtered, washed several times with ethanol (95 % (v/v)) and finally air-dried. The acidification step was required to protonate the free carboxylate functions in order to take into account the whole sum of carboxylic groups in DE calculation. 500 mg of acidified PGA were dissolved in 100 mL water and 1 mL of indicator solution was added (mixture of bromothymol blue, phenol red and cresol red). Free carboxylic acid functions were titrated by 0.1N NaOH aqueous solution (volume VA). 25 mL of 0.1 N NaOH solution were added afterwards in order to hydrolyze the ester functions. After 30 min, the excess of NaOH was titrated by 0.1 N HCl solution (volume VB). The analysis was performed in triplicate and the DE (titrimetry) was calculated according to the equation:

DE titrimetry= 1

25-VB x100% 25-VB+VA

H-NMR analysis. PGA was dissolved in D2O and freeze dried prior to NMR spectrum

acquisition in order to exchange alginate hydroxyl protons by deuterium. The NMR spectra (600 MHz, Bruker Avance III spectrometer) were measured at 318°K with dilute sample of PGA in


7

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 42

D2O in order to diminish the viscosity of the sample. Measuring the spectra at 318°K had the additional advantages to improve the signal resolution and to shift the signal of residual water away from alginate protons signals. Perdeuterated 3-(trimethylsilyl) propionate sodium salt (TSP) was added as chemical shift reference. NMR data were acquired by Topspin software and treated by Mestrenova software. The DE(NMR) was calculated from the integral ratio of esterified propylene glycol protons ( I PG ester

at 1.22 and 1.28 ppm) to the alginate backbone protons ( I alginate between 3 to 5.5 ppm). The

integral sum of esterified and residual propylene glycol protons, which could interfere with the sugar region (IPG ester (3 to 5.5 ppm) and I PG residual (3 to 5.5 ppm) respectively), was subtracted.

DE NMR=

I PG ester (1.22, 1.28 ppm) /3] x100 I alginate/5

Where:

I alginate = ∑ I (3 to 5.5 ppm) – I PG residual (3 to 5.5 ppm) – I PG ester (3to 5.5 ppm) I PG residual (3 to 5.5 ppm) = I (1.15 ppm) (Corresponding to 3 protons) I PG ester 3 to 5.5 ppm = I 1.22 ppm + I 1.28 ppm (Corresponding to 3 protons) 2.2.Determination of PGA molecular weight by size exclusion chromatography Size exclusion chromatography (SEC) was performed using a HPLC pump (Knauer HPLC pump 364). The flow rate was 0.3 mL/min and the injection volume was 20 µl. The instrument set-up consisted of SEC-columns serially connected (SHODEX OH pack sb-805 HQ and sb-804 HQ) and set in an oven at 25°C (Igloo-cil; Interchim) which was followed by RI detector (IOTA2). The system was also equipped with a guard column. The mobile phase was 50 mM


8

Page 9 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

NaNO3 solution filtered through a 0.45 µm filter (Merck Millipore). After 24h of equilibration of the SEC system with the mobile phase, a series of dextran calibration standards was injected. The samples were prepared by dissolving 10 mg of polysaccharide in 2 mL of mobile phase (5mg/mL) followed by filtering through 0.45 µm filter (PTFE filters, Millex samplicity). Data acquisition was performed using HOBOware Software. 2.3.Viscosity measurement The viscosities of PGA solutions and of HSA-PGA mixtures prepared with various concentrations in water (0.5 % to 4% PGA with or without 10% HSA (w/w)) were measured. Dependence of shear stress and viscosity on shear rate was obtained by means of Rheomat RM300 rheometer (Lamy rheology) using a co-axial cylinder system (diameters of cylinders: 14 mm and 15.2 mm). Shear rate was raised from 100 to 1000 s−1 within 50 sec at 25°C. Viscosities were recorded at 1000 s−1 and analyses were performed in triplicate. 2.4.Phase diagram Three centrifuges were tried (Beckman GS-15, Beckman coulter Avanti J-E and Beckman optima 120.000 rpm), under various conditions (2000 rpm to 40 000 rpm, for 10 min to 3h and at 25°C or 4°C), in order to visualize the phase separation of HSA-PGA mixtures, however no mixture showed a visible phase separation. The phase diagram was then established based on the microscopic aspect of the polymer aqueous solutions. A drop of HSA-PGA mixture was placed on a Neubauer cell (Counting chamber with 0,01 mm special depth, Petroff, Marienfeld) and observed with a 60x microscopic objective (UplanApo 60x/0.90, BX51 microscope, Olympus). The two polymers were considered thermodynamically incompatible when a water-in-water


9

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 42

emulsion was observed. The distance between the slide and the coverslip in a Neubauer cell is 0.01 mm, which was sufficient to save the aspect of emulsion globules. 3. Microparticle characterization 3.1.Optical microscopic observation Microparticles were colored with methylene blue and observed by means of a light microscope (BX51, Olympus) equipped with a digital camera (DP26, Olympus) for image recording. The observations were performed on intact particles in order to check their morphologies. The internal structures of the particles were observed on 20 µm thick cross sections obtained with a cryotome (Leica CM1850, Leica Microsystems). The image acquisition and treatment were performed by Stream Motion software (Olympus). 3.2.Fractal analysis of microparticle cross sections In order to assess the homogeneity of particle inner structure, fractal dimension (DB) was calculated on binary images by means of FracLac V. 2.5 software (ImageJ plugin) using the count of boxes containing pixels. For each microparticle batch, the calculation was carried out on five microphotographs taken with the same magnification factor (x10). The ROIs were selected manually by taking into account only the covalent network of the particles. DB ranges from 1 for regular particle structures to 2 for irregular ones. 3.3.Elementary analysis The mass composition of lyophilized microparticles was determined by elementary analysis (Flash EA 1112 series, Thermo Electron). Protein content was calculated considering that


10

Page 11 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

nitrogen was its specific marker in the HSA-alginate network. The 6.25 conversion factor used for the calculation of protein content was based on the 16 % nitrogen weight average found in serum albumin15. PGA content was obtained by subtraction. 3.4.Microparticle size measurement Particles were sized by a laser diffraction technique (Mastersizer 2000 analyzer equipped with a Hydro SM sample dispersion unit, Malvern) and particle size distribution measurements were conducted on three replicates. 3.5.Microparticle swelling rate Microparticle batches were prepared from PGA-HSA aqueous solutions with increasing concentrations [(HSA%; PGA%) (w/w): (7.5%; 2%), (10%; 2%), (10%; 3%), (10%; 4%), (12%; 4%)]. Agitation was stopped after 15 min of transacylation and before the neutralization step. The emulsion was allowed to stand for 15 min and microparticle sediment volume was marked (Vi for initial volume). The liquid phase was removed and 150 mL of imidazole buffer (0,05M, pH 7) was added. The agitation was resumed for 10 min at 300 rpm, then stopped. The suspension was allowed to stand for 15 min and the sediment volume was noted. In the same way, imidazole buffer was removed and the microparticle sediment was washed successively with a 2% (w/v) polysorbate aqueous solution, then with distilled water for several times. The sediment volume was marked after each washing-sedimentation cycle until constancy (final volume Vf). The swelling rate τ, was calculated according to the equation:

τ =

Vf - Vi x100 Vi


11

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 42

3.6.Lysozyme release kinetics In order to bring out the effect of microparticle architecture on the release kinetics, 3 batches of distinct structures were studied [(HSA%; PGA%) (w/w): (7.5%; 2%), (10%; 2%), (10%; 4%)]. Lysozyme from chicken egg white (Sigma) was picked as a model protein for the release study. 10 mg of freeze-dried microparticles were impregnated with 6 mg of lysozyme dissolved in water. Lysozyme was assumed to enter the particles with the solvent, during particle rehydration11. The initial loading volumes and concentrations of lysozyme solutions were adapted according to the swelling capacity of each structure type so that the amount of lysozyme (6 mg) was kept constant and was entirely entrapped in the particles, no residual liquid remained between the rehydrated particles at the end of the loading. After 24h, the loaded particles were then immersed in 10 mL of phosphate-buffered saline medium (PBS, pH 7.4). Lysozyme release kinetics was determined at 37 °C under magnetic agitation (600 rpm). 0.4 mL of the release medium was taken at constant time intervals and the amount of released lysozyme was determined by measuring the absorbance of the supernatant at 280 nm (UV spectrophotometer Beckman Coulter DU-720) after dilution and centrifugation. Every time the release medium was taken away, 0.4 mL of PBS solution was added to compensate the volume taken for the measurements. A blank test was carried out for each batch to check the residual HSA interference and the background noise was subtracted afterwards. It is important to note that the absorbance of the supernatant at the first seconds following the addition of the release medium was measured, during preliminary experiments, to prove that no significant amount of lysozyme was still free. The experiments were conducted in triplicate.


12

Page 13 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

3.7.Statistical analysis Student's t-test was used for statistical analysis. RESULTS AND DISCUSSION 1. Polymer Properties 1.1.

PGA esterification degree

PGA is a partially esterified commercial derivative of alginate, a polyanionic polysaccharide originally extracted from brown seaweed algae 16 (structure shown on Figure 3). Its degree of esterification was calculated by 1H-NMR analysis and by titrimetry. 1H-NMR analysis confirms that the used PGA is highly esterified (~ 80%) which is in good agreement with titrimetric results and with supplier data (Table 1). 1H-NMR spectrum also showed that the PGA sample contained traces of propylene glycol which probably remained after the industrial process of PGA production. The latter is usually based on the reaction of alginic acid with gaseous propylene oxide under mild conditions of pressure, temperature and pH 17,18. Moreover, two ester types are detected where 70 % of PGA is in the form of the primary 2-hydroxyprop-1-yl alginate, the remaining 30% consists of the secondary 1-hydroxyprop-2-yl ester (Figure 4). Similar results were presented by J. F. Kennedy et al., for which the two types of esters were found in an equilibrium with the more thermodynamically favorable primary ester predominating in the ratio 4:1. In fact, the two PGA forms are obtained after nucleophilic attack of alginate carboxylic function on an unsymmetrical epoxide and their ratio depends upon the employed conditions (acidic or basic) 19.


13

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 42

HO HO O O

O

O

OH

O

O

O OH

O

O

OH

O

HO O

HO

OH

O

β,D- mannuronic ester

OH O O OH

O

OH

α,L-guluronic ester

HO

β,D-mannuronic ester

O

α,L-guluronic acid

Figure 3. Propylene glycol alginate (PGA) Table 1. PGA physicochemical characterization PGA properties

DE (NMR)

R1: R2(NMR)a

value

80%

70:30

DE (titrimetry)

84%

DE (supplier data)

84%

MW b

1 095 KD

a

R1: 2-hydroxyprop-1-yl and R2: 1-hydroxyprop-2-yl. b Peak top molecular weight measured by SEC.


14

Page 15 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 4.1H-NMR spectrum of PGA (DE 84%) in D2O (600MHz, 318°K) 1.2.

PGA molecular weight and viscosity

The effects of polysaccharide structure, molecular weight and concentration on their viscosity in solution were widely studied 20,21,22,23. The high molecular weight of used PGA (Table 1) led to high viscosities which in turn increased with PGA concentration in the aqueous solution. It seems that PGA content also controls the mixture viscosities even for 10% HSA while the protein contribution is insignificant. Thus, the higher the concentration of PGA in the blend, the higher the viscosity of HSA-PGA mixture (Figure 5).


15

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 42

Figure 5. Viscosities of PGA aqueous solutions (triangles joined by continuous line) and (10% HSA)+PGA aqueous mixtures (squares joined by dashed lines) as a function of PGA concentration, at 25°C (values recorded at 1000 s−1 shear rate, n=3). 1.3.

pH of HSA-PGA mixtures

The pH of HSA-PGA mixtures depends on the concentration ratio of the two polymers in the aqueous phase. Due to the presence of free carboxylic functions, concentrated solutions of PGA exhibit acidic pH (~5). However, the increase of HSA concentration in the mixture induces a buffering capacity and stabilizes the pH at a neutral value (6.5 to 7) for [HSA] / [PGA] ratios greater than 4 (Figure 6).


16

Page 17 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 6. pH of HSA-PGA aqueous mixtures as a function of polymer concentration ratio. 1.4.

HSA-PGA phase diagram

The phase behavior of mixed biopolymer solutions is quantitatively characterized by the phase diagram (mainly represented by the binodal curve), describing the boundary conditions of phase separation and the partition of components between the phases 24,25,26,27. In this work, the phase diagram is studied in order to establish a relationship between the disparities observed among microparticle inner structures and the polymer thermodynamic incompatibility in the aqueous solution. Even though the phase diagram is usually drawn at a given pH and ionic strength, we chose not to set these parameters to constant values in order to mimick the conditions of microparticle preparation. In most cases, the pH values of HSA-PGA mixtures were higher than HSA isoelectric point (PI 4.7 28) so that no ionic complexation might occur. To draw the phase diagram and the binodal curve, several methods are commonly used for which a prior centrifugation step is required to separate the two incompatible phases (e.g. phase volume ratio 25 and node determination 29 methods). In the present case, it was not possible to visualize a phase separation of HSA-PGA mixtures even after ultracentrifugation. The close densities of the two phases and/or the high viscosities of PGA-rich phases could be the causes of this non-separation 30. Therefore, the microscopic aspects of the mixtures were used to distinguish the monophasic from the biphasic domains but without being able to draw a binodal curve or tie lines. When water-in-water emulsion was observed, the blend was considered as biphasic (example shown on Figure 7). By this approach, the HSA-PGA phase diagram was established (Figure 8).


17

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 42

Figure 7. Optical microphotograph of 20% HSA-2% PGA (w/w) aqueous solution placed on Neubauer cell. At low polymer concentrations, HSA and PGA are co-soluble in the aqueous phase (domain M, Figure 8). Their concentrations are below the threshold of phase separation and repulsive interactions are negligible 24,31,32. By increasing the polymer concentrations, the two macromolecules quickly show a thermodynamic incompatibility, giving rise to two distinct phases (domain B, Figure 8). This incompatibility could be observed as the two polymers are of the same charge (HSA and PGA negatively charged when pH > pI HSA). Under these conditions, important repulsive interactions could occur and the interactions between the polymers of the same nature are then privileged 26,33–35,36. The thermodynamic incompatibility could also be induced when HSA-solvent interactions and PGA-solvent interactions are favored compared to the solvent-solvent and the protein-polysaccharide ones24,36. Moreover, the incompatibility domain might be enlarged due to the high molecular weight of the used PGA24. On the other hand, HSA exhibits a limited solubility when it is highly concentrated in the blend (> 15 % w/w), leading to microscopic lumps and heterogeneous solutions (domain I, Figure 8). This third domain could be biphasic (gel/liquid) or triphasic (gel/liquid/liquid) depending on PGA concentration.


18

Page 19 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 8. HSA-PGA phase diagram based on microscopic aspect of polymer aqueous mixtures. (M: for monophasic domain, B: for biphasic domain and I: for HSA insolubility domain). 2. Effect of polymer mixture properties on microparticle structure and composition 2.1.

Microparticle feasibility

It was found that under the given preparation conditions, no microparticle could be obtained from HSA-PGA mixtures in the monophasic area of the phase diagram because of the low polymer concentrations in this domain (domain M, Figure 8 and Table 2): no microparticle could be observed in the reaction medium, and no sediment could be separated by centrifugation. It was also the case for the HSA-PGA mixtures with the lowest concentrations in the biphasic domain (domain B, Figure 8 and Table 2). When the polymer concentrations are higher in the biphasic domain, a stable network can be created between the two polymers despite the phase separation, and the standard procedure gives rise to microparticles presenting a spherical shape (example shown on Figure 9). Microparticles can also be prepared from mixtures containing HSA concentrations higher than 15% (w/w) (domain I, Figure 8 and Table 2). It is important to note that when microparticle preparation is possible,


19

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 42

no complexation phenomenon could occur in the initial aqueous phase between HSA and PGA (pH > pI).

Figure 9. Optical microphotograph of microparticles prepared with 10% HSA and 2% PGA (w/w) and colored with methylene blue. 2.2.

Microparticle inner structure

The inner structures of microparticles prepared with increasing concentrations of HSA and PGA, were studied via microscopic observation of cross sections. When microparticle preparation is possible in the biphasic domain (domain B, Figure 8), their internal structure is dependent of HSA and PGA concentrations in the initial aqueous solution (Table 2). At lower polymer concentrations in the biphasic area, microspheres presenting a lacework-like structure are obtained. By increasing the polymer concentrations, a gradual transition towards a vesicular structure enclosing a liquid content is observed.


20

Page 21 of 42

Table 2. Optical microphotographs of HSA-alginate microparticle cross sections as a function of

4%

polymer concentrations (% w/w) and thermodynamic incompatibility.

Cross sections

1.5%

2%

3%

Domain a

B

B

B

B

B

B+I

Domain a

B

B

B

B

B

B+I

Cross sections

No microparticle

Domain a

B

B

B

B

B

B+I

B

B

B

B

B+I

No microparticle

No microparticle

M

M

B

B

B

B+I

2.5%

5%

7.5%

10%

12%

20%

Cross sections

Cross sections

Domain a

1%

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

No microparticle

B

Cross sections

Domain a

↑PGA / HSA

a: Phase diagram domains shown on Figure 8; M: monophasic, B: Biphasic and I: HSA insolubility, domains. In response to tree-like patterns observed on many of these cross sections (example on Figure 10), a fractal analysis was carried out on each microphotograph to quantify the differences between microparticle internal structures 37,38.


21

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 42

Figure 10. Optical microphotograph of a microparticle cross section colored with methylene blue (initial aqueous phase composition: 5% HSA and 4% PGA (w/w)). Unlike vesicular microcapsules prepared with high polymer concentration, for which DB is between 1.5 and 1.7, microparticles having a lacework structure present the highest fractal dimension values (DB > 1.8). This means that for the latter, cross-linked regions are less homogeneous than those of vesicular capsules (Figure 11, Table 2). Therefore, microparticles having transitional structures show intermediate DB values ranging between 1.7 and 1.8. Accordingly, these analyses could be a good tool to evaluate microparticle heterogeneity and structure type.


22

Page 23 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 11. Fractal dimension « DB » variations calculated by frac_lac (ImageJ) on HSA-alginate microparticle cross sections. The sizes of the internal cavities (Table 2) are larger than the sizes of the globules present in the biphasic aqueous phases before emulsification (Figure 7). However, the phase separation might play a role in the development of the internal structure of microparticles by organizing in part the spatial distribution of the two polymers inside the droplets. Even if it was not possible to draw tie lines, it is supposed that the upper left part of region B on the phase diagram consists of mixtures with a continuous PGA-rich phase, whereas the lower right part consists of mixtures with a continuous HSA-rich phase. This phase inversion might be one of the causes explaining morphological changes observed when varying the PGA:HSA ratio. However, the phase separation in the aqueous droplets is not the only phenomenon responsible for the variation in internal structure of the network. In this work section, a relationship between polymer properties, previously introduced, and microparticle structure is assessed. In fact, the observed morphological disparities are the results of three main physicochemical variations induced by increasing polymer concentrations in the initial aqueous solution, as detailed hereinafter: (NaOH/polymers) concentration ratio. It should be reminded that microparticles are formed in W/O emulsion by a transacylation reaction between ester groups of PGA and amine functions of HSA, after diffusion of NaOH in the aqueous droplets (Figure 1 and Figure 2). The formation of the covalent network implies the simultaneous presence of alkaline reagent, HSA and PGA. Based on this concept, the concentration ratio of NaOH to polymers in the aqueous phase ([NaOH]/ [HSA+PGA]) seemed to be the first factor influencing microparticle structure. In fact,


23

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 42

the higher the (NaOH/ polymers) concentration ratio, the thicker the microparticle membrane and/or the tougher the polymer network. This hypothesis was confirmed by varying separately the added volume of NaOH solution used for triggering the transacylation reaction on the one hand, and PGA esterification degree in order to vary the number of available ester groups on the other hand. Significant morphological modifications were observed in the two cases (Table 3(A), (B)). Increasing the amount of NaOH results in a more compact network up to the center, which means that NaOH, when available, could diffuse farther in the aqueous phase where amine and ester functions are still unreacted (Table 3(A)). Similar results were previously obtained on HSA-alginate coated beads where a progressive increase of membrane thickness and rigidity modulus was observed with increasing NaOH concentration

39, 40

. This observation can also be

compared to the conclusions of a study concerning gelation of alginate with calcium ions 41, and showing that the degree of inhomogeneity observed in these gels was linked to the relative rate of diffusion of polymer and gel-inducing ions. It is important to note that even when added in excess compared to the two aqueous polymers, NaOH is partly consumed in the hydrophobic external phase due to the saponification of isopropyl myristate and sorbitan trioleate. On the other side, the replacement of the initial PGA (DE 84%) by another PGA of lower DE (55%) leads to microparticles with thicker membranes (Table 3(B)). In this case, the amount of available esters is not sufficient to consume completely the alkaline reagent at the same depth as with the initial PGA, and NaOH is still available to diffuse deeper into the aqueous droplets. The 55% DE PGA and the initial PGA show close viscosities and similar pH values in aqueous solution.


24

Page 25 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

HSA buffering capacity. As the transacylation reaction is pH-dependent (ideal pH is about 9.3 to 10.5

42

), the structure of HSA-alginate microparticles could be affected by the buffering

capacity exerted by the protein when concentrated in the aqueous solution. This buffering power would consume more NaOH and make it less available for the transacylation reaction. This would result in more vesicular structures at high concentrations of HSA. In fact, this suggestion is upheld after two further experiments in which HSA and PGA were dissolved in phosphate buffer solutions of pH 5 and 8 instead of pure water. The resulting mixtures had a final pH value of 5.8 and 7.1, respectively. Observations of the cross sections show that microparticle structure is highly dependent on the initial pH of the aqueous solution. For acidic initial conditions (pH 5.8), where the pH value is steadied far below the optimal reaction pH, microparticles are vesicular and less ramified, unlike those prepared from a pH 7.1 solution (Table 3(C)). A hypothesis for the mechanism of network formation can be drawn from these results confirming that the presence of HSA hinders the diffusion of hydroxide ions through the center. The relative locations of PGA and HSA at the interface might determine some starting points for the entry of hydroxide ions, maybe through PGA-rich domains, or through sorbitan trioleate molecules used to stabilize the emulsion and crossing the interface. The colocalization of PGA ester groups and HSA amino groups consumes hydroxide ions during the transacylation reaction, and after reaction this location offers no more resistance to a further hydroxide ion, able to enter deeper inside the droplet. This could explain the fractal patterns observed on many of the cross sections, looking like small brooks giving rise to larger rivers from the outside towards the center of the particles.


25

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 42

PGA viscosity. Another factor that could also control HSA-alginate microparticle structure is the mixture viscosity. The high viscosity of PGA, which in turn increased with its content in the blend, could impede NaOH diffusion and lead to vesicular particles at high concentration of PGA (Figure 5, Table 2).


26

Page 27 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Table 3. Effect of the variations in microencapsulation protocol on the structures and sizes of HSA-Alginate microparticles: Effect of (A) NaOH (2% in EtOH 95% (v/v)) added volume; (B) PGA esterification degree; (C) pH of the aqueous solution. (Initial aqueous phase composition: 10% HSA and 2% PGA (w/w)). (A) Volume of NaOH solution Molar ratio (NaOH/ester) Molar ratio (NaOH/(ester + amine)) Mean diameter

4 mL

8 mL

4.7

9.5

1.7

3.4

908 µm

620 µm

DE of PGA Molar ratio (NaOH/ester)

84%

55%

4.7

6.6

Mean diameter

908 µm

890 µm

262 µm

326 µm

Cross sections

(B)

Cross sections membrane thickness

(C) pH

Phosphate buffer pH 5.0

water pH 6.5

Phosphate buffer pH 8.0

Mean diameter

983 µm

908 µm

708 µm

Cross sections


27

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

2.3.

Page 28 of 42

Microparticle mass composition

The determination of HSA content allowed to compare between the initial concentrations of polymers in the aqueous solution and their concentrations in freeze-dried particles. It also could explain microparticle behaviors during drug release. As PGA does not contain any nitrogenous function, the mass composition of freeze-dried HSA-alginate microparticles was achieved by elementary analysis based on the estimation of nitrogen percentage in the dry matter. It is found that all microparticles are highly rich in HSA (> 60 % w/w). Moreover and for a given PGA concentration, HSA microparticle content is in constant increase when increasing HSA concentration in the initial aqueous solution (Figure 12, Table 2). These results indicate that PGA ester groups are in excess compared to accessible free amine functions and are still available to react with more HSA molecules. Figure 12(B) compares the measured HSA content (full line) to the theoretical maximal HSA content (dotted line) of dried microparticles for a fixed PGA concentration. This figure shows that for a PGA concentration of 3%, a low HSA concentration in the initial solution leads to a leakage of PGA during the washing steps. Increasing the HSA concentration allows to trap growing amounts of PGA.


28

Page 29 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 12. HSA content (% w/w), estimated by elementary analysis, in freeze-dried HSAalginate microparticles: (A) as a function of initial polymer concentrations in the aqueous solution, (B) as a function of initial HSA concentration in the aqueous solution where [PGA] = 3% (w/w) (triangles joined by dotted line). The squares joined by the full line in (B) present the theoretical maximal content of HSA in dried microparticles. Data corresponding to the dotted line in (A) are presented in (B). 3. Effect of the inner structure on microparticle functional properties 3.1.Microparticle swelling rate and size


29

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 30 of 42

Microparticle swelling rate. The swelling rate of HSA-alginate microparticles is directly related to their internal structure (Figure 13). Unlike tough microcapsules obtained at higher polymer concentrations, the lacework-like microspheres are more resilient and able to absorb water up to nine times their volume. Moreover, microparticle swelling rate is related to the fractal dimension. Figure 14 presents the fractal dimensions corresponding to the microparticles of Figure 13. Two slopes can be distinguished on this graph, corresponding to vesicular particles on the one hand and to lacework-like ones on the other hand, which makes possible the prediction of the swelling rate from DB values for each structure type.

Figure 13. HSA-alginate microparticle swelling rate (%) as a function of total polymer concentration and particle internal structure (Full squares for lacework-like microparticles, circles for vesicular ones and empty square for intermediate ones). (HSA; PGA) aqueous concentrations % (w/w) used for microparticle preparation are indicated under each structure.


30

Page 31 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 14. HSA-alginate microparticle swelling rate (%) vs fractal dimension (Full squares for lacework-like microparticles, circles for vesicular ones and empty square for intermediate ones). Microparticle size. It is known that the granulometric distribution of an emulsion is directly related to the generated viscous shear forces

43,44

. Basically, and for a given hydrophobic phase,

the lower the viscosity of the internal aqueous phase in W/O emulsion, the smaller the droplets, which subsequently give rise to smaller microparticles after alkalization. Based on these notions, it was expected that HSA-alginate microparticle mean size would rise with PGA concentration in the aqueous blend because of the begotten increase in viscosity (Figure 5). This rule might be true if HSA-alginate microparticles were sized before the washing steps. In fact, the difference in swelling capacities of microparticles in water, depending on their inner structure, disrupted the rising relationship between the aqueous phase viscosity, the droplet size and microparticle diameter. Therefore, HSA-alginate microparticle diameter increased initially with viscosity and reached a maximum where water-swellable microspheres were obtained and then decreased again for tough vesicular particles with a lower swelling capacity (Figure 15, Table 2).


31

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 32 of 42

Figure 15. (A) HSA-alginate microparticle mean diameters (µm) as a function of HSA and PGA concentrations in the aqueous solution, (B) HSA-alginate microparticle mean diameters (squares), swelling rates (triangles) and aqueous solution viscosities at 25°C (diamonds) as a function of PGA concentration in the aqueous solution where [HSA] =10% (w/w). Data corresponding to the dotted line in (A) are presented in (B). 3.2.Lysozyme release kinetics It is known that the release kinetics of a drug from a particulate delivery system is governed by multiple factors, such as drug-matrix interactions, pH and ionic strength of release medium


32

Page 33 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

and microparticle size and porosity 45,46. Therefore and for a better understanding of the protein behavior, lysozyme is often selected as a model for release studies in vitro 47,48. The aim of this section is to prove HSA-alginate microparticle advantage in protein sustained release and to elucidate the effect of their microstructure on the release kinetics. The chosen microparticle batches are of distinct structures but of close diameters before freeze-drying and have a similar mass composition in the aim of studying the effect of the microstructure on lysozyme release kinetics, after minimizing the influences of microparticle size and polymer interaction disparities. Drug release induced by microparticle degradation is not discussed in this work by referring to the relative microparticle stability over the short duration of the experiments (8h). Lysozyme is a globular basic protein characterized by a molecular weight of 14.4 kDa 49 and an isoelectric point of 10.7 50. At neutral pH, lysozyme is positively charged while HSA and PGA bear negative charges (pI of HSA is 4.7 28). Under these conditions, ionic attractive interactions, occurring between positive charged lysozyme and negative polymers51, could promote lysozyme loading efficiency in addition to drug diffusion towards the interior of the particles. After adding the release medium (PBS buffer which contains salts), the ionic interactions were supposed to be disrupted and lysozyme release is enhanced 52. On the other hand, HSA-lysozyme hydrophobic interactions 53 moderately uphold the adsorption 47 of the lysozyme on the matrix and control their release. Noting that the use of hydrophilic polymers (HSA and PGA) could enhance the release rate by avoiding the excessive hydrophobic adsorption of lysozyme, unlike hydrophobic PLGA (Poly (lactic-co-glycolic acid)) microparticles commonly used for protein encapsulation 46. This combination of factors makes HSA-alginate microparticles suitable for protein encapsulation with a sustained release profile (Figure 16).


33

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 34 of 42

Figure 16. Lysozyme release at 37°C from HSA-alginate microparticles of distinct inner structures. Microparticles were prepared at: (a) 7.5% HSA, 2% PGA (diamonds); (b): 10% HSA, 2% PGA (squares) and (c) 10% HSA, 4% PGA (triangles) (w/w). All differences were significant (p< 0.05) except for dashed line encircled points. As regards the microstructure effect, the exploration of HSA-alginate microparticles of distinct structures showed significant differences on lysozyme release kinetics (p < 0.05, except for dashed line encircled points on Figure 16). Even if the use of microcapsules instead of microspheres is supposed to reduce interactions between the drug and the matrix by minimizing the drug-polymers surface area 54, HSA-alginate vesicular microcapsules, prepared at high polymer concentrations (10% HSA and 4% PGA (w/w)), show the slowest drug release with the lowest curve (Figure 16 (c)). This result could be due to the compact structure and the low porosity of the covalent network forming the capsule membrane, which mainly control the drug diffusion. Otherwise, drug uniformly dispersed in the sphere matrix could increase the initial burst effect 55

which probably occurs in lacework-like microparticles prepared at lower concentrations of

HSA and PGA (7.5% HSA and 2% PGA (w/w)) (Figure 16 (a)). Sphere porosity could also


34

Page 35 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

control the initial burst effect owing to the leaching which takes place at the outer wall of the sphere as it becomes hydrated 56–58. But the main factor governing lysozyme release rate from HSA-alginate microparticles seems to be their swelling capacity. The loading was performed by rehydrating the freeze-dried samples with a limited volume of aqueous solution, which did not lead to an extensive swelling of the particles. Moreover, the presence of lysozyme on anionic binding sites of the microparticles further limits their swelling. During the release, on the contrary, loaded particles were placed in a large volume of release medium, which is in favor of swelling. Furthermore, the release of lysozyme with time is likely to be accompanied by a swelling of the particles. The swelling bringing additional release medium and ions inside the particles is supposed to lead to a faster release of the encapsulated protein. The swelling study conducted on empty particles showed that lacework type (a) particles (fractal dimension: 1.86) had a more intense swelling capacity than vesicular type (c) particles (fractal dimension: 1.65), with an intermediate result for type (b) particles (fractal dimension: 1.75) (Figure 14). The swelling capacities of the particles, related to the fractal dimensions of the internal networks, could therefore partially explain the differences between release curves. The release curves show that particles (a) and (c) did not reach a plateau at the end of the experiment. Concerning particles of type (b), a 100% release could not be reached because the interactions between the matrix and the lysozyme still seemed to be prevailing. The bound fraction could probably be displaced by adding some more ions to the release medium.

CONCLUSIONS


35

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 36 of 42

This work showed that the properties of HSA-alginate microparticles prepared by transacylation in W/O emulsion were directly related to the two polymer concentrations in the aqueous solution. At lower polymer concentrations, lacework-like microspheres were obtained, while a gradual transition into vesicular microcapsules was observed with increasing polymer concentrations. The differences between the structures could be assessed by fractal analysis carried out on cross section microphotographs. These morphological disparities were the result of several physicochemical variations: the (NaOH/ polymers) concentration ratio, the PGA esterification degree and viscosity as well as the HSA buffering capacity. In fact, the higher the (NaOH/ polymers) concentration ratio, the more homogeneous was the microparticle internal structure. On the other hand, the high viscosity of PGA solutions and the pH buffering capacity of HSA impeded the alkaline reagent diffusion and led to vesicular particles. Interfacial phenomena, which could also influence the disposition of the two biopolymers in the aqueous droplets before the reaction, are currently under investigation in our laboratory. Moreover, the striking differences in microparticle inner structures influenced their functional properties. Unlike tough microcapsules obtained at higher polymer concentrations, the laceworklike microspheres were extremely water-swellable. This mechanical property could be exploited in dermatological applications to absorb burn or wound exudates for example. Furthermore, noticeable gaps were observed on lysozyme release kinetics from the different structure types (microsphere and microcapsule), with a sustained release profile which makes HSA-alginate microparticles appropriate for protein encapsulation. Corresponding Author


36

Page 37 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

* Address: Institut de Chimie Moléculaire de Reims, U.F.R. Pharmacie ; 51, rue Cognacq-Jay 51095 Reims Cedex, France. E-mail: [email protected] ACKNOWLEDGMENTS “La Ville de Reims” is highly acknowledged for funding this research. We thank Dr. Christophe BLIARD for SEC system furnishing, Anthony ROBERT for NMR spectrum acquirement and Sylvie LANTHONY for elementary analyses. REFERENCES (1)

Dubey, R.; Shami, T. C.; Bhasker Rao, K. U. Def. Sci. J. 2009, 59, 82–95.

(2)

Gupta, A.; Dey, B. Sunsari Tech. Coll. J. 2013, 1.

(3)

Poshadri, A.; Kuna, A. JRes ANGRAU 2010, 38, 86–102.

(4)

Benoît, J.-P.; Richard, J.; Venier-Julienne, M.-C. Tech. Ing. 2013.

(5)

Gouin, S. Trends Food Sci. Technol. 2004, 15, 330–347.

(6)

Lévy, M.-C.; Edwards-Lévy, F. J. Microencapsul. 1996, 13, 169–183.

(7)

Callewaert, M.; Laurent-Maquin, D.; Edwards-Lévy, F. Int. J. Pharm. 2007, 344, 161–

164. (8)

Andry, M.-C.; Edwards-Lévy, F.; Lévy, M.-C. Int. J. Pharm. 1996, 128, 197–202.

(9)

Pariot, N.; Edwards-Lévy, F.; Andry, M.-C.; Lévy, M.-C. Int. J. Pharm. 2002, 232, 175–

181. (10) McDowell, R. H. J. Soc. Cosmet. Chem. 1970, 21, 441–457.


37

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 38 of 42

(11) Hurteaux, R.; Edwards-Lévy, F.; Laurent-Maquin, D.; Lévy, M.-C. Eur. J. Pharm. Sci. 2005, 24, 187–197. (12) Munin, A.; Callewaert, M.; Edwards-Lévy, F. Cahiers de Formulation n°15 de la Société Chimique de France. EDP sciences. 2011. (13) Edwards-Lévy, F. Bioencapsulation Innov. 2013, 22–25. (14) Liu, L.; Cao, J.; Huang, J.; Cai, Y.; Yao, J. Bioresour. Technol. 2010, 101, 3268–3273. (15) Jones, D. B. Factors for converting percentages of nitrogen in foods and feeds into percentages of proteins; Washington, D.C. : U.S. Dept. of Agriculture, 1931; Vol. 183. (16) Sinquin, A.; Hubert, P.; Marchal, P.; Choplin, L.; Dellacherie, E. Colloids Surf. Physicochem. Eng. Asp. 1996, 112, 193–200. (17) Steiner, A. B.; McNeely, W. H. Ind. Eng. Chem. 1951, 43, 2073–2077. (18) Noto, V.; Pettitt, D. Process for the preparation of propylene glycol alginate from partially neutralized alginic acid. US3772266 A, November 13, 1973. (19) Kennedy, J. F.; Griffiths, A. J.; Philp, K.; Stevenson, D. L.; Kambanis, O.; Gray, C. J. Carbohydr. Polym. 1989, 11, 1–22. (20) Rinaudo, M. In Food Hydrocolloids; Nishinari, K., Doi, E., Eds.; Springer US: Boston, MA, 1994; pp 21–34. (21) Berth, G. Carbohydr. Polym. 1992, 19, 1–9. (22) Vauchel, P.; Arhaliass, A.; Legrand, J.; Kaas, R.; Baron, R. J. Phycol. 2008, 44, 515–517.


38

Page 39 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(23) Fu, S.; Thacker, A.; Sperger, D. M.; Boni, R. L.; Buckner, I. S.; Velankar, S.; Munson, E. J.; Block, L. H. AAPS PharmSciTech 2011, 12, 453–460. (24) V.B.Tolstoguzov. Food Hydrocoll. 1991, 4, 429–468. (25) Polyakov, V. I.; Grinberg, V. Y.; Tolstoguzov, V. B. Polym. Bull. 1980, 2, 757–760. (26) Grinberg, V. Y.; Tolstoguzov, V. B. Food Hydrocoll. 1997, 11, 145–158. (27) Antonov, Y. A.; Grinberg, V. Y.; Zhuravskaya, N. A.; Tolstoguzov, V. B. Carbohydr. Polym. 1982, 2, 81–90. (28) Vlasova, I. M.; Saletsky, A. M. J. Appl. Spectrosc. 2009, 76, 536–541. (29) Kaul, A. In Aqueous Two-Phase Systems: Methods and Protocols; Hatti-Kaul, R., Ed.; Methods in Biotechnology; Humana Press, 2000; pp 11–21. (30) Tolstoguzov, V. B. Food Hydrocoll. 1995, 9, 317–332. (31) In Aqueous Biphasic Separations; Albertsson, P. ., Ed.; Springer US: Boston, MA, 1995. (32) Syrbe, A.; Bauer, W. J.; Klostermeyer, H. Int. Dairy J. 1998, 8, 179–193. (33) Schmitt, C.; Sanchez, C.; Desobry-Banon, S.; Hardy, J. Crit. Rev. Food Sci. Nutr. 1998, 38, 689–753. (34) Benichou, A.; Aserin, A.; Garti, N. J. Dispers. Sci. Technol. 2002, 23, 93–123. (35) Tolstoguzov, V. Crit. Rev. Biotechnol. 2002, 22, 89–174. (36) Piculell, L.; Lindman, B. Adv. Colloid Interface Sci. 1992, 41, 149–178.


39

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 40 of 42

(37) Fini, A.; Fernández-Hervás, M. J.; Holgado, M. A.; Rodriguez, L.; Cavallari, C.; Passerini, N.; Caputo, O. J. Pharm. Sci. 1997, 86, 1303–1309. (38) Cavallari, C.; Rodriguez, L.; Albertini, B.; Passerini, N.; Rosetti, F.; Fini, A. J. Pharm. Sci. 2005, 94, 1124–1134. (39) Edwards-Lévy, F.; Lévy, M.-C. Biomaterials 1999, 20, 2069–2084. (40) Rachik, M.; Barthes-Biesel, D.; Carin, M.; Edwards-Lévy, F. J. Colloid Interface Sci. 2006, 301, 217–226. (41) Skjåk-Bræk, G.; Grasdalen, H.; Smidsrød, O. Carbohydr. Polym. 1989, 10, 31–54. (42) Lévy, M.-C. Utilization of a transacylation reaction between an esterified polysaccharide and a polyaminated or polyhydroxylated substance for fabricating microparticles, microparticles thus obtained, methods and compositions containing them. WO1994004261 A1, March 3, 1994. (43) Shinnar, R. J. Fluid Mech. 1961, 10, 259. (44) Boxall, J. A.; Koh, C. A.; Sloan, E. D.; Sum, A. K.; Wu, D. T. Langmuir 2012, 28, 104– 110. (45) Freiberg, S.; Zhu, X. X. Int. J. Pharm. 2004, 282, 1–18. (46) Giteau, A.; Venier-Julienne, M.-C.; Aubert-Pouëssel, A.; Benoit, J.-P. Int. J. Pharm. 2008, 350, 14–26. (47) Jiang, G.; Woo, B. H.; Kang, F.; Singh, J.; DeLuca, P. P. J. Controlled Release 2002, 79, 137–145.


40

Page 41 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(48) Qodratnama, R.; Serino, L. P.; Cox, H. C.; Qutachi, O.; White, L. J. Mater. Sci. Eng. C 2015, 47, 230–236. (49) Cegielska-Radziejewska, R.; Leśnierowski, G.; Kijowski, J. Pol. J. Food Nutr. Sci. 2008, 58, 5–10. (50) Abeyrathne, N. S.; Lee, H. Y.; Ahn, D. U. Korean J. Food Sci. Anim. Resour. 2013, 33, 501–507. (51) Blanco, D.; Alonso, M. J. Eur. J. Pharm. Biopharm. 1998, 45, 285–294. (52) Park, T. G.; Yong Lee, H.; Sung Nam, Y. J. Controlled Release 1998, 55, 181–191. (53) Rojas, E. E. G.; dos Reis Coimbra, J. S.; Minim, L. A.; Saraiva, S. H.; da Silva, C. A. S. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2006, 840, 85–93. (54) Park, J. H.; Ye, M.; Yeo, Y.; Lee, W.-K.; Paul, C.; Park, K. Mol. Pharm. 2006, 3, 135– 143. (55) Kakish, H. F.; Tashtoush, B.; Ibrahim, H. G.; Najib, N. M. Eur. J. Pharm. Biopharm. 2002, 54, 75–81. (56) Le Corre, P.; Le Guevello, P.; Gajan, V.; Chevanne, F.; Le Verge, R. Int. J. Pharm. 1994, 107, 41–49. (57) Hiroaki, O.; Masaki, Y.; Toshiro, H.; Yayoi, I.; Shigeru, K.; Yasuaki, O.; Hajime, T. J. Controlled Release 1994, 28, 121–129. (58) Ghaderi, R.; Sturesson, C.; Carlfors, J. Int. J. Pharm. 1996, 141, 205–216.


41

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 42 of 42

For Table of Contents Use Only

Serum Albumin-Alginate Microparticles Prepared by Transacylation: Relationship between Physicochemical, Structural and Functional Properties AUTHOR NAMES Imane Hadef (,), Barbara Rogé (), Florence Edwards-Lévy ()* AUTHOR ADDRESS Institut de Chimie Moléculaire de Reims, CNRS UMR 7312. () U.F.R. Pharmacie, 51, rue Cognacq-Jay, 51096 Reims, France. () U.F.R. Sciences Exactes et Naturelles, Campus du Moulin de la Housse 51687 Reims, France.


42

Serum Albumin-Alginate Microparticles Prepared by Transacylation

Recommend Documents