Journal of Colloid and Interface Science Preparation of uniform

Journal of Colloid and Interface Science 323 (2008) 267–273
Contents lists available at ScienceDirect
Journal of Colloid and Interface Science
www.elsevier.com/locate/jcis
Preparation of uniform-sized PELA microspheres with high encapsulation
efficiency of antigen by premix membrane emulsification
Qiang Wei a,b , Wei Wei a,b , Rui Tian a,b , Lian-yan Wang a , Zhi-Guo Su a , Guang-Hui Ma a,∗
a
b
State Key Lab of Biochemical Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, PR China
Graduate School of Chinese Academy of Sciences, Beijing, PR China
a r t i c l e
i n f o
a b s t r a c t
Article history:
Received 3 February 2008
Accepted 19 April 2008
Available online 30 April 2008
Relatively uniform-sized poly(lactide-co-ethylene glycol) (PELA) microspheres with high encapsulation
efficiency were prepared rapidly by a novel method combining emulsion-solvent extraction and premix
membrane emulsification. Briefly, preparation of coarse double emulsions was followed by additional
premix membrane emulsification, and antigen-loaded microspheres were obtained by further solidification. Under the optimum condition, the particle size was about 1 μm and the coefficient of variation
(CV) value was 18.9%. Confocal laser scanning microscope and flow cytometer analysis showed that the
inner droplets were small and evenly dispersed and the antigen was loaded uniformly in each microsphere when sonication technique was occupied to prepare primary emulsion. Distribution pattern of
PEG segment played important role on the properties of microspheres. Compared with triblock copolymer
PLA–PEG–PLA, the diblock copolymer PLA–mPEG yielded a more stable interfacial layer at the interface of
oil and water phase, and thus was more suitable to stabilize primary emulsion and protect coalescence of
inner droplets and external water phase, resulting in high encapsulation efficiency (90.4%). On the other
hand, solidification rate determined the time for coalescence during microspheres fabrication, and thus
affected encapsulation efficiency. Taken together, improving the polymer properties and solidification rate
are considered as two effective strategies to yield high encapsulation.
© 2008 Elsevier Inc. All rights reserved.
Keywords:
Premix membrane emulsification
PELA microspheres
Uniform-sized
Coalescence
Encapsulation efficiency
1. Introduction
Biodegradable microspheres have received more and more attentions for therapeutic application such as controlled release
and drug targeting [1–4]. Among several techniques to fabricate
biodegradable microspheres [5], the water-in-oil-in-water (W/O/W)
double emulsion method is generally considered as one of the
most convenient ways to encapsulate water soluble protein [6–8].
Although the process is conceptually simple, many factors, either
material- or process-related, are still critical to the obtained microspheres [7].
To date, polylactide (PLA) is the most commonly used polymer because of its proven biodegradable and biocompatible. However, PLA has some drawbacks, resulting from hydrophobic nature,
which is related to their rapid sequestration by the mononuclear
phagocyte system [9,10]. Furthermore, the affinity between hydrophobic polymer and hydrophilic protein is not satisfied, and
leads to a low encapsulation efficiency of protein drug [11]. High
encapsulation efficiency has been considered to be an important
requirement for the successful fabrication of microspheres con-
*
Corresponding author. Fax: +86 10 82627072.
E-mail address: [email protected] (G.-H. Ma).
0021-9797/$ – see front matter
doi:10.1016/j.jcis.2008.04.058
©
2008 Elsevier Inc. All rights reserved.
taining protein. As an alternative to PLA, copolymer poly(lactideco-ethylene glycol) (PELA) has showed much potentials because
it improves the properties of the microspheres products [8,12,13].
Herein, two PELA polymers, diblock copolymers (PLA–mPEG) based
on monomethoxypoly(ethylene glycol) (mPEG) and triblock copolymer (PLA–PEG–PLA) based on poly(ethylene glycol) (PEG) were
synthesized and used as polymer materials, which was expected
to yield high encapsulation efficiency during microspheres fabrication.
Particle size is also a crucial parameter in drug release system [14–16]. However, the method for controlling particle size and
size distribution is limited. In conventional processes, the double
emulsions are prepared by mechanical stirring or homogenization,
and thereby the size distribution of microspheres obtained is very
broad. We previously reported that preparation of uniform-sized
PLA microspheres by direct SPG (Shirasu Porous Glass) membrane
emulsification while methylene chloride (MC) was used as organic
solvent [17,18]. Typically, the primary emulsion was permeated
through the uniform pores of the membrane into the external water phase by the pressure of nitrogen gas to form uniform-sized
multiple droplets. However, MC suffers from the drawback of high
toxicity. In contrast, ethyl acetate (EA) has much low toxicity and
could thus be good alternative [8,19]. Previous studies showed that
a high interfacial tension between dispersed phase and membrane
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Q. Wei et al. / Journal of Colloid and Interface Science 323 (2008) 267–273
surface played important role on the uniform droplet formation
[20,21]. Unfortunately, the water solubility of EA (8.7%, w/v) is
higher than that of MC (2.0%, w/v) [8], it cannot meet the requirement of direct SPG membrane technique for preparation of uniform emulsion with relatively hydrophilic dispersed phase. Therefore, it is still highly expected to develop a method to prepare
uniform-sized microspheres with good applicability to relatively
hydrophilic materials.
Premix membrane emulsification would be a promising technique due to the high transmembrane flux [22]. The oil phase
containing polymers does not have sufficient time to spread on the
membrane surface. Thus, in present study, a novel method combining double emulsion-extraction and premix membrane emulsification was proposed. Briefly, preparation of coarse double emulsions
was followed by additional premix membrane emulsification with
proper pressure, and protein-loaded microspheres were obtained
by further solidification. EA was employed as organic solvent and
PLA was also chosen as polymer for comparison with PELA. The
recombinant hepatitis B surface antigen (HBsAg) was selected as
a prototype to testify the encapsulation efficiency [23]. The effect of volume ratio of oil phase and external water phase on size
distribution of the microspheres was investigated. Flow cytometer
analysis and confocal laser scanning microscope were occupied to
compare the preparation method of primary emulsion. The effects
of polymer composition and solidification technique on the encapsulation efficiency of microspheres were also investigated and
explained in details.
2. Materials and methods
2.1. Materials
PLA with a molecular weight (Mw ∼ 40,000 kDa) was purchased from the Institute of Medical instrument (Shandong, China).
Monomethoxypoly(ethylene glycol) (mPEG 2000, Mw = 2000 Da)
was obtained form Fluka (Switzerland). Poly(ethylene glycol) (PEG
2000, Mw = 2000 Da) was purchased from Bio Basic Inc. (Canada).
Stannous octoate was ordered from Sigma (Germany). Poly(vinyl
alcohol) (PVA-217, degree of polymerization 1700, degree of hydrolysis 88.5%) was provided by Kuraray (Japan). Hepatitis B surface antigen (Mw ∼ 24 kDa) was kindly supplied by Hualan Biological Engineering Incorporation (Xinxiang, China) and concentrated
to 1.5 mg/ml. Fluorescein isothiocyanate (FITC) was purchased from
Sigma–Aldrich Inc. (Germany). SPG membrane (pore size of the
membrane was 5.2 μm) was provided by SPG Technology Co. Ltd.
(Japan). All other reagents were of analytical grade.
PELA copolymers (PLA–mPEG and PLA–PEG–PLA) were synthesized according to the procedure reported by Lucke et al. [24],
in which the mPEG block and PEG block have a same molecular weight of 2000 Da. The Mw, Mn and Mw/Mn of copolymers
were evaluated by gel permeation chromatography. Mean number molecular weight (Mw) of copolymers PLA–mPEG and PLA–
PEG–PLA were 43,000 Da and 39,000 Da and the polydispersity
(Mw/Mn) were 1.63 and 1.49, respectively.
2.2. Preparation of microspheres
Microspheres loaded with antigen were prepared by two-step
procedure. Briefly, the coarse double emulsions were first prepared,
0.4 ml HBsAg aqueous solution was mixed with 4 ml ethyl acetate
containing polymers (200 mg) by sonication (Branson Sonicator)
for 15 s in an ice bath to form primary emulsion, and homogenization with 6000 rpm for 15 s (IKA homogenizer) was used as
comparison. The W/O was further emulsified into external aqueous
phase containing 1% (w/v) PVA and 1% (w/v) NaCl by magnetic stirring for 60 s at 300 rpm to prepare coarse double emulsions. The
Fig. 1. Schematic diagram of a miniature kit for premix membrane emulsification.
coarse double emulsions were then poured into the premix reservoir. As schematized in Fig. 1, double emulsions with smaller and
relatively uniform size were achieved by extruding the coarse double emulsions through the SPG membrane with a high pressure of
300 kPa. The microspheres were obtained by different solidification
technique. In direct solidification technique, the obtained double
emulsion droplets were poured quickly into 800 ml solution containing 1% (w/v) NaCl (solidification solution) under magnetic stirring for 5 min to solidify the microspheres. In step-solidification
technique, the obtained double emulsion droplets were preliminarily solidified by a little amount volume (20 or 40 ml) of presolidification solution containing 1% (w/v) PVA and 1% (w/v) NaCl
for 1 min, and then were completely solidified by 800 ml solution
containing 1% (w/v) NaCl (solidification solution) under magnetic
stirring for 5 min. The obtained microspheres were collected by
centrifugation and washed with distilled water for three times and
then lyophilized for 2 days. Unless specified, the formulation conditions were as follows: polymer was PLA–mPEG, primary emulsion was prepared by sonication technique and microspheres were
solidified by direct solidification technique.
2.3. Characterization of microspheres
The surface morphology of microspheres was observed by a
JSM-6700F (JEOL, Japan) scanning electron microscope (SEM). The
volume-mean diameter of microspheres was measured by laser
diffraction using Mastersizer 2000 (Malvern, UK). The particle size
distribution was expressed as a coefficient of variation (CV) value,
which is defined as:
CV =
n
(di − d̄)2
i =1
N
1/ 2
d̄,
where di is the diameter of the ith particle, d̄ is the numberaverage diameter and N is the total number of particles counted.
2.4. Flow cytometer analysis of antigen distribution in microspheres
FITC was covalently linked to HBsAg in a way as described
previously [25]. Briefly, a solution of FITC (5 mg/ml) was slowly
added into HBsAg solution. After an incubation period of 12 h
at 4 ◦ C, unbound FITC was separated by ultrafiltration. The FITCHBsAg loaded microspheres and blank microspheres were added
into phosphate buffered saline (PBS) to form a suspension with
a concentration of 2.0 × 106 microspheres/ml. The samples were
Q. Wei et al. / Journal of Colloid and Interface Science 323 (2008) 267–273
Fig. 2. Effects of volume ratio of oil phase to external water phase on size distribution.
analyzed on FL1-H channel by a LSR flow cytometer (Becton Dickinson).
2.5. CLSM imaging of microspheres
The microspheres samples prepared for flow cytometer analysis
were also observed by CLSM (Confocal Laser Scanning Microscope).
Suspensions containing microspheres in Petri dish were observed
by a TCS SP2 CLSM (Leica). The transmitted image was taken.
2.6. Turbidity determination of primary emulsion
The stability of primary emulsion was indicated by its turbidity. The turbidity of primary emulsion was determined according
to a method previously described [26]. Briefly, the primary emulsions were prepared by different polymers and added into a quartz
cell and absorbance was then measured at 620 nm with a spectrophotometer as a function of time. The turbidity was calculated
from standard absorbance–turbidity curve with formazin suspension as standards. Formazin was prepared by reacting hydrazine
sulfate with hexamethylenetetrammonium, and standards of formazin turbidity units (FTU) was prepared by appropriate dilution.
2.7. Measurement of protein encapsulation efficiency
The total antigen loaded efficiency in PELA microspheres was
determined by dissolving twenty milligrams of the freeze-dried
microspheres in 1 ml of 5% (w/v) sodium dodecyl sulfate (SDS)
containing 0.1 M sodium hydroxide solution. The amount of antigen was determined by micro bicinchoninic acid (micro-BCA) assay
(Pierce, USA). Blank microspheres were used as control. The encapsulation efficiency of antigen was calculated by the following
formula:
EE =
total amount of antigen loaded
total amount of antigen added
× 100%.
3. Results and discussion
In premix membrane emulsification, fine double emulsions
could be obtained by disrupting the coarse double emulsions with
SPG membrane. The effects of volume ratio of oil phase and external water phase on particle size and size distribution were investigated in present study. As shown in Fig. 2, with the increase of
external water phase volume, the obtained microspheres showed
a bigger size and broader size distribution (SEM was not shown).
The viscosity of the coarse double emulsions could be a good explanation for these results. When external water phase volume
269
Fig. 3. Comparison of the preparation method of primary emulsion: (a) sonication technique, (b) homogenization technique and (c) corresponding flow cytometer
analysis.
increased, a larger amount of EA diffused from the oil phase into
the external water due to its good solubility in water (8.7%, w/v).
Consequently, the coarse double emulsions became more viscous
and difficult to be disrupted into fine double emulsions, which
affected the uniformity of the obtained double emulsions. The optimum volume ratio of oil phase and external water phase (1:6)
was performed in the following study. The corresponding particle
size of the obtained microspheres was about 1 μm and CV value
was 18.9%.
When the size is in particular ranges, the microspheres possess passive targeting property. What’s more, microspheres with
a diameter under 5 μm would be ideal for passive targeting of
antigen presenting cells [14–16], and induce high humoral immunization response. However, preparation of microspheres with
such small particle requires an efficient method to prepare stable primary emulsion. It is well known that reducing droplets size
of the primary emulsion is known as one promising approach to
improve the stability of double emulsion [27]. Herein, homogenization and sonication were occupied to testify their effects on
the primary emulsion. The antigen was labeled with FITC and then
encapsulated into the microspheres. During the solvent-extraction
process, the microspheres were solidified so rapidly that inner
droplets did not have sufficient time to coalesce. Thus, the interior structure of microspheres almost could represent the original distribution of inner droplets within oil droplets. As shown
in Figs. 3a and 3b, the inner droplets of primary emulsion prepared by sonication were small and evenly dispersed in the microsphere, whereas the size of inner droplets was very large and
could not be effectively encapsulated while using homogenization technique. Correspondingly, the signal of FITC labeled antigen
showed a sharp peak for the microspheres fabricated by sonication during flow cytometer analysis, whereas a broad distribution
for the microspheres prepared by homogenization and the obtained signal was partly overlapped with the signal of the blank
microspheres (Fig. 3c), indicating that some of the microspheres
prepared by homogenization did not encapsulate any FITC labeled
antigen and that the antigen was loaded uniformly in each microsphere under the sonication process. From these two aspects,
sonication was an adequate technique and adopted in following
study.
3.1. Effects of polymer composition
The morphologies of microspheres fabricated by premix membrane emulsification were visualized by SEM and shown in Fig. 4.
Relatively uniform-sized microspheres were successfully prepared
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Q. Wei et al. / Journal of Colloid and Interface Science 323 (2008) 267–273
Fig. 5. The stability of primary emulsion of different polymers.
Fig. 4. SEM photographs of microspheres prepared by different polymers: (a) PLA,
(b) PLA–PEG–PLA and (c) PLA–mPEG.
Table 1
Preparative parameters and properties of corresponding microspheres
Batch
Polymer
Pre-solidification
solution volume
(ml)
Particle size
(μm)
Encapsulation
efficiency
(%)
A
B
C
D
E
PLA
PLA–PEG–PLA
PLA–mPEG
PLA–mPEG
PLA–mPEG
0
0
0
40
20
1.39 ± 0.05
1.21 ± 0.07
1.13 ± 0.03
1.22 ± 0.05
1.33 ± 0.07
61.3 ± 3.7
75.1 ± 5.1
90.4 ± 4.3
83.9 ± 3.9
77.3 ± 4.1
by different polymers. The CV values of microspheres were 21.5,
24.1 and 18.9% for PLA, PLA–PEG–PLA and PLA–mPEG, respectively.
Compared with smooth surface of PLA microspheres, the PLA–PEG–
PLA microspheres exhibited pitted and porous surface, whereas
the PLA–mPEG yielded microspheres with rough appearances and
numerous saliences. Irregular surface structures were observed
similarly in the case of microspheres prepared with another amphiphilic copolymer, such as PLA–mPEO [28]. The heterogeneous
surface of microspheres might be formed due to different distribution pattern of PEG segment in copolymers [7], which could be
seen from Fig. 1 in Supporting materials.
As shown in Table 1, the PELA microspheres possessed higher
encapsulation efficiency as compared to PLA counterpart. The result can be ascribed to the improvement of affinity between polymer and antigen. Interestingly, there is still an evident difference
between the PELA copolymers. Although PLA–mPEG and PLA–PEG–
PLA possessed the same content of PEG segment, the PLA–mPEG
microspheres yielded higher encapsulation efficiency (90.4%) than
that of PLA–PEG–PLA microspheres (75.1%). To our knowledge, limited study has performed on detailed mechanism about the results.
We supposed that these might be affected by the different distribution pattern of PEG segments in polymers and these would be
discussed in present study.
It was known that the encapsulation efficiency of microspheres
was strongly affected by the stability of the primary emulsion during the double emulsion process [29,30]. Coalescence between the
inner droplets within primary emulsion leads to separation of oil
phase and water phase. The stability of the primary emulsion was
measured by the time required for phase separation [31]. However, the time was not easy to be determined. In present study,
we proposed a more quantitative method to evaluate the stability by measuring turbidity of the primary emulsion. The turbidity
decreased with the growth of inner droplets by coalescence and
was easily monitored by spectrophotometer. The turbidity changes
of primary emulsion fabricated by different polymers as a function of time were investigated and are shown in Fig. 5. All primary
emulsions had a same initial turbidity, but the turbidity of the primary emulsion prepared by PLA decreased dramatically from 250
to 81 FTU in the initial 10 h, whereas the turbidity of the PLA–
PEG–PLA copolymer decreased slightly to 219 FTU in the initial
10 h and the turbidity of the PLA–mPEG copolymer was kept constant during 24 h, indicating that PELA could improve the primary
emulsion stability. These results were in good agreements with encapsulation efficiency of different polymers.
When PELA was employed in double emulsion process, the hydrophilic PEG segments would extend into the water phase and
hydrophobic PLA segments would spread into the oil phase. As illustrated in Fig. 6, in the present case of PELA, the polymer itself
would form an interfacial layer and act as surfactant because of
its amphiphilic character, thereby stabilizing the primary emulsion.
Surfactants could improve the stability of the primary emulsion
[30,32], but should be limited to the minimum level to avoid possible toxic [1,33]. On the other hand, distribution pattern of PEG
segment would also play important role on the properties of the
interface between oil phase and water phase. As shown in Fig. 6,
the PLA–mPEG would form a more stable interfacial layer than
that of PLA–PEG–PLA probably due to spatial hindrance of triblock
copolymer [29], thus led to a more stable primary emulsion.
Q. Wei et al. / Journal of Colloid and Interface Science 323 (2008) 267–273
271
Fig. 6. Schematic illustration of polymer distribution on the interface between internal phase (W 1 ) and oil phase.
Fig. 7. Schematic illustration of the coalescence process between inner
droplets (W 1 ) and external aqueous phase (W 2 ) (oil droplet was dispersed in external aqueous phase W 2 ).
The encapsulation efficiency of microspheres was determined
by the protein lose in microspheres fabrication. How the protein
lost during double emulsion-solvent extraction method has not
been studied thoroughly. A reasonable explanation was that the
loss of encapsulated protein during the fabrication process essentially from the fact that the protein in the inner droplets tend to
merge with external water phase [7]. Herein, we speculated that
an oil film prevented the coalescence of the inner droplets and external aqueous phase. As illustrated in Fig. 7, inner droplets moved
to the interface of oil phase and external water phase and separated from external water phase by a thin oil film. Once the oil
film was evacuated, the inner droplets would merge into the external water phase rapidly and disappear, resulting in the loss of
protein. It would be an effective strategy to yield high encapsulation by restraining the evacuation of the oil film separating the
inner droplets and external water phase. When PELA was used, the
PLA segment was anchored at the interface due to interfacial layer
(Fig. 6), thus the evacuation of oil film therein would be restrained,
and higher encapsulation efficiency of antigen is an expected consequence. As mentioned above, the interfacial layer of PLA–mPEG
was more stable than that of PLA–PEG–PLA. The properties of oil
film within PLA–mPEG copolymer was more suitable for protecting
the coalescence of inner droplets and external water phase, led to
a higher encapsulation efficiency of antigen.
3.2. Effects of solidification technique
In order to control the solidification rate, the process of solidification was modified with a procedure of additional presolidification. The double emulsions obtained by premix membrane emulsification were preliminarily partially-solidified by a lit-
Fig. 8. SEM photographs of microspheres prepared by different solidification techniques: (a) batch C, (b) batch D and (c) batch E.
tle amount aqueous solution and then completely solidified. PLA–
mPEG was selected as polymer, and the detailed preparative parameters and properties of the microspheres are listed in Table 1.
The solidification rate of microspheres fabricated by direct solidification technique is faster than those of step-solidification
techniques. Among the step-solidification techniques, the solidification rate of microspheres would increase with the increase
of pre-solidification aqueous solution volume. Thus, batch C >
batch D > batch E in terms of solidification rate of obtained microspheres.
The surface morphologies of microspheres were observed by
SEM and shown in Fig. 8. All the microspheres revealed rough
appearances, which was more developed with the increase of solidification rate. As shown in Table 1, the particle size of microspheres fabricated by direct solidification technique had a lightly
smaller size compared with those of step-solidification techniques.
Solidification rate is known to directly influence the internal structure of microspheres [34]. The coalescence of inner droplets within
oil droplets might form interconnecting channels inside of the
obtained microspheres [8]. In present study, rapid solidification
would reduce the coalescence time, and thus led to a more compact structure. To represent their effects on internal structure of
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Q. Wei et al. / Journal of Colloid and Interface Science 323 (2008) 267–273
4. Summary
Relatively uniform-sized amphiphilic PELA microspheres were
prepared rapidly and conveniently by a novel method combining
emulsion-solvent extraction and premix membrane emulsification
while EA was used as organic solvent. Under the optimum condition, the particle size was about 1 μm and the CV value was 18.9%.
Confocal laser scanning and flow cytometer analysis showed that
the inner droplets were small and evenly dispersed and the antigen was loaded uniformly in each microsphere when sonication
technique was occupied to prepare primary emulsion.
Effects of polymer composition and solidification technique on
encapsulation efficiency were also investigated in this study. Improving the properties of polymers and shorting the time required for solidification of microspheres were found as two effective strategies to yield high encapsulation of antigen. Distribution
pattern of PEG segment of polymers played important role on
the properties of microspheres. Compared with triblock copolymer
PLA–PEG–PLA, the diblock copolymer PLA–mPEG yielded a more
stable interfacial layer at the interface of oil and water phase, and
thus was more suitable to stabilize primary emulsion and protect
coalescence of inner droplets and external water phase, resulting in
high encapsulation efficiency (90.4%). On the other hand, solidification technique determined solidification time during microsphere
fabrication, and thus affected coalescence time. The encapsulation
efficiency increased with the increase of solidification rate of microspheres. These uniform-sized PELA microspheres prepared by
this novel method might have great potential as protein drug carrier.
Acknowledgments
We thank the financial support of the National Nature Science
Foundation of China (20536050, 20221603 and 20636010) and Chinese Academy of Sciences (KJCX2.YW.M02).
Supplementary material
Supplementary material for this article may be found on ScienceDirect, in the online version.
Please visit DOI: 10.1016/j.jcis.2008.04.058.
Fig. 9. SEM photographs of microspheres after incubation in PBS for 70 days:
(a) batch C, (b) batch D and (c) batch E.
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