j.biomaterials.2003.08.069 (1)

8/13/2019 j.biomaterials.2003.08.069 (1)

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Biomaterials 25 (2004) 17711777

Preparation and characterization of cationic PLGA nanospheres

as DNA carriers

M.N.V. Ravi Kumar*,1, U. Bakowsky, C.M. Lehr

Department of Biopharmaceutics and Pharmaceutical Technology, Saarland University, Saarbr .ucken, D 66123, Germany

Received 11 May 2003; accepted 11 August 2003

Abstract

Nanoparticles formulated from biodegradable polymers such as poly(lactic acid) (PLA) and poly(lactide-co-glycolide) (PLGA)

are being extensively investigated as non-viral gene delivery systems due to their controlled release characteristics and

biocompatibility. PLGA nanoparticles for DNA delivery are mainly formulated by an emulsion-solvent evaporation technique

using PVA as a stabilizer generating negatively charged particles and heterogeneous size distribution. The objective of the present

study was to formulate cationically modified PLGA nanoparticles with defined size and shape that can efficiently bind DNA. An

Emulsion-diffusion-evaporation technique to make cationic nanospheres composed of biodegradable and biocompatible co-

polyester PLGA has been developed. PVA-chitosan blend was used to stabilize the PLGA nanospheres. The nanospheres were

characterized by atomic force microscopy (AFM), photon-correlation spectroscopy (PCS), and Fourier transform infrared

spectroscopy (FTIR). Zeta potential and gel electrophoresis studies were also performed to understand the surface properties of

nanospheres and their ability to condense negatively charged DNA. The designed nanospheres have a zeta potential of 10 mV at pH

7.4 and size under 200 nm. From the gel electrophoresis studies we found that the charge on the nanospheres is sufficient to

efficiently bind the negatively charged DNA electrostatically. These cationic PLGA nanospheres could serve as potential alternatives

of the existing negatively charged nanoparticles.

r 2003 Elsevier Ltd. All rights reserved.

Keywords: Biodegradable; Chitosan; Gene therapy; Nanoparticles; PLGA

1. Introduction

Biodegradable colloidal particles have received con-

siderable attention as a possible means of delivering

drugs and genes by several routes of administration.

Special interest has been focused on the use of particles

prepared from polyesters like PLGA, due to their

biocompatibility and to their resorbability through

natural pathways [1]. Various methods have been

reported for making nanoparticles viz., emulsion-eva-

poration[2], salting-out technique[3], nanoprecipitation

[4], cross-flow filtration [5] or emulsion-diffusion tech-

nique [6,7]. Indeed PLGA particles are extensively

investigated for drug [810] and gene delivery [11,12],

but still improvements in the existing methods are

needed to overcome the difficulties in terms of reprodu-

cibility, size, and shape. The size and shape of the

colloidal particles are influenced by the stabilizer and the

solvent used. Most investigated stabilizers for PLGA

lead to negatively charged particles and the plasmid

incorporation is achieved via double emulsion technique

during particle preparation. This could pose problems in

the stability and biological activity of the plasmid due to

the involvement of organic solvents during the prepara-

tion process. This can be overcome by using cationically

modified particles that can bind and condense negatively

charged plasmids by simply adding nanoparticles to

plasmid or vice versa. Literature suggests PVA as most

popular stabilizer for the production of PLGA nano-

particles leading to negatively charged particles, never-

theless, investigations have been carried out using

other stabilizers as well [13]. Vila et al. investigated

double emulsion technique for making PLGA-lecithin

nanoparticles for protein delivery using PVA-chitosan

blend as coating material [14]. The particle size and

charge reported were 500729nm and 21.871.1 mV

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*Corresponding author. Tel.: +91-172-214683 ext 2057; fax: +91-

172-214692.

E-mail address: [email protected] (M.N.V. Ravi Kumar).1Present address: Department of Pharmaceutics, National Institute

of Pharmaceutical Education and Research (NIPER), SAS Nagar,

Sector 67, Punjab 160062, India.

0142-9612/$- see front matterr 2003 Elsevier Ltd. All rights reserved.

doi:10.1016/j.biomaterials.2003.08.069

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respectively [14]. However, sole cationic PLGA nano-

spheres with chitosan as a modifier for gene delivery can

hardly be seen in the literature. Therefore, attempts were

made to develop a technique that produces uniform and

much smaller nanospheres with cationic surface modifica-

tion, which can readily bind and condense DNA.

Chitosan was selected in these studies because, other thanits cationic charge it has been recognized for its

mucoadhesivity, biodegradability and ability to enhance

the penetration of large molecules across mucosal surfaces

[15].To our knowledge, not many studies were reported

describing such a well defined shape and size (o200nm)

of the nanospheres, particularly when PLGA and high

molecular weight polymers like chitosan were used.

2. Materials and methods

2.1. Materials

Poly(l-lactide-co-glycolide) (PLGA) (70:30 lactide:

glycolide) and Poly(vinyl alcohol) were obtained from

Polysciences, Inc. and MoWiol, Germany, respectively.

Chitosan Hydrochloride (Seacure 210, 83% deacety-

lated) was obtained from Pronova Biopolymer, Norway.

The b-galactosidase expression plasmid pCMVb was

purchased from ATCC (Manassas, VA, USA) and

transformed intoE.coli DH5a: Gigaprep from 2500 ml

of over-night culture was performed according to the

manufacturers instructions (QIAGEN, Hilden, Ger-

many). The DNA was precipitated in 70% ethanol and

reconstituted in water to 1 mg/ml. All other chemicals

and reagents used in this study were from Aldrich-

Sigma, Germany.

2.2. Preparation of PLGA nanospheres

Nanospheres were prepared by a new emulsion-

diffusion-evaporation technique as shown in the

Fig. 1. The methodology in brief goes as follows:

200 mg of PLGA is dissolved in 10 ml ethyl acetate at

room temperature. The organic phase is then added to

an aqueous stabilizer mixture containing 100 mg of PVA

and 30 mg of chitosan in 10 ml water under stirring. The

emulsion is stirred at room temperature for 3 h before

homogenizing at 13,500 rpm for 10 min using an Ultra-

Turrax T25 homogenizer (Janke and Kunkel GmbH

KG, Staufen, Germany). To this emulsion water is

added under stirring resulting in nano-precipitation.

Stirring is continued on a water bath maintained at

40C to remove organic solvent.

2.3. FTIR spectroscopy

To assess the modification of the polymer surfaces an

FT-IR (ATR) spectrometer (Perkin Elmer system 2000)

was used. For the measurements, the particles in

solution were spread directly onto the ATR crystal

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Ethyl acetate+

PLGA

Stirring 1000 rpm2 hours

Passed through0.2 m filter

AqueousPVA-Chitosan

2 hours

Passed through

0.2 m filter

Stirring 1000 rpm

Mixing

3 hoursStirring 1000 rpm

Homogenize10 min 13,500 rpm

Water

Stirring 2 hoursWater bath, 40 oC

Add organic to aqueous NH2

NH2

NH2

NH2

NH2

NH2

NH2

NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2 NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2 NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2 NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2 NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2NH2NH2

NH2

NH2

NH2NH2

NH2

NH2NH2

NH2

NH2

NH2

NH2NH2

NH2

NH2NH2

Fig. 1. Schematic representation of PLGA nanospheres preparation process.

M.N.V. Ravi Kumar et al. / Biomaterials 25 (2004) 177117771772

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(ATMS 45, 7 cm in longitude). The water was

evaporated by a nitrogen stream. The spectrum was

collected in a range between 4500 and 850 cm1 with a

resolution of 1 cm1 (100 scans per sample).

2.4. Photon correlation spectroscopy

Particle size was determined by photon correlation

spectroscopy (PCS) on an ALV 5000 (Laser Vertriebs-

gesellschaft mbH, Langen, Germany) at a scattering

angle of 90 (sampling time 200 s). Autocorrelation was

performed using the contin method. For PCS

measurements, all samples were diluted 50 fold in

demineralized water, resulting in comparable viscosities.

Hence, no corrections for the effect of the additives were

necessary.

2.5. Zeta potential measurements

Surface charge of nanoparticles was determined by

zeta potential measurement on a Malvern Zetasizer 2000

HS (Malvern, UK) with a flow measurement cell

connected to a Mettler DL 25 (Mettler-Toledo, Giessen,

Germany) auto-titrator via a circulating system. Within

the 250 ml sample container at the titrator, 510 ml of

nanoparticle samples were diluted with demineralized

water to a final volume of 200 ml. The pH was adjusted

to 3 by using HCl (1n) before titration to pH 10 with

NaOH (0.1n). Measurements of the zeta-potential were

carried out at 0.5 pH increments at 25C. The

instrument was calibrated routinely with a 50mV

latex standard.

2.6. Gel electrophoresis and determination of unbound

DNA

NanoparticleDNA complexes were prepared by

mixing the nanoparticles with plasmid at a concentra-

tion of 10 mg/ml in 25 mm Hepes (pH 7.4) as well as in

deionized water (pH 6.0). The complex formation

studies were performed at room temperature and

allowed to stand for 15 min to attain complexes. The

nanoparticleDNA complexes were electrophored on an

agarose gel (1% ethidium bromide included for visua-

lization) for 90 min at 5 V/cm. Images were acquired

using a Geldoc 2000 gel documentation system (Bio-Rad,

Munich, Germany) equipped with a UV transluminator.

Molecular Analyst, version 1.1 software (Bio-Rad) was

used for band integration and background correction.

2.7. Atomic force microscopy

The size and surface morphology of the PLGA

particles was analyzed by atomic force microscopy

(AFM) Nanoscope IV Bioscopet (Digital Instruments,

Veeco) in tapping mode using a Si3N4cantilever with a

spring constant of about 34N/m and a resonance

frequency of about 200 kHz. Scanning was performed

at a scan speed of 0.5 Hz with a resolution of

512 512 pixels. The tip loading force was minimized

to avoid structural changes of the sample.

3. Results and discussions

3.1. Nanospheres formation

The routine emulsion-solvent evaporation technique

being used for formulating PLGA nanoparticles is

believed to produce heterogeneous size distribution

[16]. Various formulation factors and characteristics of

the nanoparticles have a key role to play in biological

applications. The foremost factor that could have an

influence on the transfection and cellular uptake is the

size of the nanoparticles. Prabha et al. [16]have studied

the size-dependency of nanoparticle-mediated gene

transfection with fractionated nanoparticles.

Recent reports suggests that a fraction of the

stabilizer PVA always remains associated with the

nanoparticles despite repeated washings because PVA

forms an interconnected network with the polymer at

the interface[17]. We came across similar factors while

formulating nanoparticles using PVA as a stabilizer and

discussed in the following sections. Above all, the

stability and biological activity of the plasmid have

been major concerns due to the involvement of organic

solvents during the preparation process[16,17]. Keeping

the above factors in mind we developed a method forformulation of cationically modified PLGA nanoparti-

cles. An emulsion-diffusion-evaporation technique using

ethylacetate as organic solvent and PVA-chitosan blend

as a stabilizer yielded uniform spherical cationic nano-

spheres. We have screened several solvents and found

that the particle size is at lower end when ethylacetate

was employed (data not shown). Stabilizers (PVA and

Chitosan) concentration has been optimized for the

smallest particle size for this method (data not shown).

We believe that the nanospheres formation involves the

mechanism as described: Stirring causes the dispersion

of the solvent as irregular sized globules in equilibrium

with the continuous phase and the stabilizer is then

absorbed on the larger interface created; homogeniza-

tion further results in the smaller globules; the addition

of water and the heating step destabilizes the equili-

brium and causes to diffuse the organic solvent to the

external surface. During the transport of solute, new

smaller globules less than 200 nm are produced; the

heating step also helps to have a final colloidal

suspension free of organic solvent and more uniform

in size. Nanospheres were also prepared by eliminating

one or two steps of the complete method and the results

obtained are presented asTable 1. Eliminating either of

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the steps resulted in increase in the particle size, which is

in agreement to our discussion.

3.2. FTIR characterization

The nanospheres were characterized by FTIR. The

characteristic peaks obtained from PVA, chitosan and

PLGA were compared with the peaks resulted from

nanospheres. The characteristic peaks at 1511 and

3015 cm1 due to amino groups from chitosan were

also found in the nanospheres prepared from PVA-

chitosan blend, suggesting the cationic modification.

3.3. PCS measurements

The nanospheres when analyzed by dynamic light

scattering demonstrated a unimodal size distribution for

PVA alone and PVA-Chitosan blend formulated by

emulsion-diffusion-evaporation technique (Fig. 2).

However, there is no indication of nanosphere forma-

tion when chitosan was used alone, which is in

agreement with the reported studies that PVA is

necessary to stabilize PLGA particles[18]. Prabha et al.[16] in their recent report compared the difference

between the PCS vs. TEM measurements in terms of

particle size and found huge difference. The PVA is

known to form layers of aggregates (B5 layers) around

the surface of nanoparticles contributing towards the

hydrodynamic diameter of nanoparticles [19,20]. The

discrepancy in the size of nanoparticles could be that the

dynamic light scattering method gives the hydrodynamic

diameter rather than the actual diameter of nanoparti-

cles, therefore a comparison of the particle size with

other techniques as well is worth it. The mean

hydrodynamic particle diameter was found to be

111.774.2 nm when PVA was alone used as stabilizer,

whereas, 181.573 nm when a combination of PVA and

chitosan were used as a blend. The increase in size is

expected and attributed to the high molecular weight

chitosan. We have compared the size of the particles as

analyzed by PCS and AFM techniques and presented as

Table 2.

3.4. Zeta potential measurements

The zeta potential value is an important particle

characteristic as it can influence both particle stability as

well as particle mucoadhesion. In theory, more pro-

nounced zeta potential values, being positive or nega-

tive, tend to stabilize particle suspension. The

electrostatic repulsion between particles with the same

electric charge prevents the aggregation of the spheres

[21]. Mucoadhesion, on the other hand, can be

promoted by a positive zeta potential value. The mucus

layer itself is at a neutral pH value an anionic

polyelectrolyte [22]. Consequently, the presence of the

positively charged groups on the particles could lead to

electrical charge interactions between the mucus and the

particles. In the present studies nanospheres were made

with PVA alone and a blend of PVA-Chitosan to attain

surface modification. The particles made of PVA (1%

w/v) alone were negatively charged (8 mV at pH 7.4).

Zeta potential titration provided proof of successful

cationic surface modification when a blend of PVA-

chitosan (1.3% w/v) was used. The final nanoparticle

suspension using PVA-chitosan blend has a pH of 4 and

a zeta potential of 36mV, which suggests that the

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0

20

40

60

80

100

0.1 10 100 1000

PVA alonePVA-Chitosan BlendChitosan alone

SizeDistribution(%)

Particle Size

Fig. 2. Particle size distributions of the nanospheres as measured by

PCS.

Table 1

Eliminating one or two steps of the method and the resultant particle size

No. O/W emulsion stirring 1000rpm Homogenization 13,500rpm Add. water & evaporation Particle size by PCS (nm)

1 Yes No No 884717

2 Yes Yes No 40378

3 Yes Yes Yes 18173

Results are presented as mean (n 3)7standard deviation.

Table 2

Nanospheres as measured by PCS and AFM

No. Stabilizer PCS measurement

(nm)

AFM measurment

(nm)

1 PVA 111.774.2n 3 100.276.2n 47

2 Chitosan Not detectable 24.972.7n 117

3 PVA-chitosan 181.573 n 3 187714.4 n 112

PCS: Number in parenthesis represents number of replicates; AFM:

number in parenthesis represents number of particles measured.

Results are presented as mean7standard deviation.


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suspension would be stable. The zeta potential at pH 3.0

was 46 mV; however, it decreased with increase in pH

and reached to 10 mV at pH 7.4 (Fig. 3). AFM images

show uniform cationic modification, which is evident

through uniform DNA coating onto the nanospheres

due to the electrostatic interaction between phosphate

groups of DNA and the NH2groups of chitosan on thesurface (Fig. 5E). This has been confirmed by gel

electrophoresis studies in the later sections.

3.5. Gel electrophoresis

The binding of the cationic PLGA nanospheres to the

polyanionic DNA was studied using analysis of the

electrophoretic mobility of the DNA within an agarose

gel. Efficient complexation of pCMVbeta by cationic

PLGA nanospheres leads to immobilisation. These new

PLGA nanospheres were able to immobilise pCMVbeta

plasmid (Fig. 4). Negligible amounts of free DNA in thelane of 100 particles:1 DNA and no free DNA there

after is the proof of good complex at the ratio 100:1 and

beyond.

3.6. AFM measurements

The size and surface morphology was analyzed by

AFM. When PVA was alone used in the preparation theparticle size is about 100 nm (Fig. 5A) and the reasons

for the discrepancy of the size between the two

measurements was discussed under Section 3.3. It

appears that lot of PVA is adhered to the particle

surface (Fig. 5A), which is a similar finding to the

reported studies, irrespective of the method used [16].

When chitosan was used, AFM analysis did show the

particle size to be very small (24.972.7nm) (Fig. 5B),

which is unlikely with high molecular weight polymers

like chitosan. Moreover, the particle shape is not well

defined and fused. We could not detect any particle size

by PCS. It appears that the nanospheres were uniform

and spherical in shape with smooth surfaces when PVA-

chitosan blend was used in the preparation (Fig. 5C and

D). Also the AFM pictures show no free/unbound

material when PVA-chitosan blend was used (Fig. 5C

and D). DNA is uniformly coated onto the nanospheres

(DNA shell of 22.472.1, n 65nm) (Fig. 5 panel E)

due to the electrostatic interaction between phosphates

groups of DNA and the NH2 groups of chitosan on

the surface as shown in the Fig. 1. The size of the

nanospheresDNA complexes is smaller and more

uniform when compared to the reported DNA-polymer

(in particular when chitosan is used) self-assemblies

[2325]. To our knowledge such a high resolution AFMimage clearly showing the electrostatic interaction

between positively charged PLGA nanospheres and

negatively charged DNA has not been shown before.

Many reports on PLGA particles are entirely based on

PCS studies while discussing size and very few reports

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-10

0

10

20

30

40

50

3 3.5 4 4.5 5 5.5 6 6.5 7 7.5 8 8.5 9 9.5 10

pH

ZetaPotentialmV

Fig. 3. Zetapotential titration curve of PLGA nanospheres coated

with PVA-chitosan blend.

M= MARKER DNAB =BLANK SPACE

M B 0 1 10 20 25 50100120140160 B M

Particle to DNA ratio

FreeDNA

(%)

0

10

20

30

40

50

60

70

80

90

100

0 1 10 20 25 50 100 120 140 160

(B)(A)

Fig. 4. PLGA-DNA complexes with increasing amounts of PLGA nanospheres were prepared and analysed for DNA immobilisation ability. The

amounts of free DNA were related to un-complexed DNA (100% mobile) run on the same gel. To quantify the DNA-immobilisation ability, the

cationic PLGA:DNA ratios (w/w) required for 100% immobilisation are compared in this graph. (Solid bars=Percentage of free DNA; white

bars=100% immobilization).


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have shown visual images of the nanoparticles. Fromthe present studies its clear that one should not

base only on PCS analysis for the particle formation

or size.

4. Conclusion

From these investigations it is evident that this

method forms uniform cationic PLGA nanospheres

that can bind DNA readily by electrostatic interaction.

These cationic surface modified PLGA nanospheres

avoid the usage of the plasmid during the particle

preparation process, where it has to stay in contact with

organic solvents for quite a while. PVA alone could not

give the cationic charge needed and chitosan alone could

not stabilize the particles, therefore, a blend of these two

is needed. PVA-chitosan blend not only giving the net

positive surface charge, but also produced particles with

uniform size and spherical shape, as observed by AFM.

Investigations were performed using as low as 50 mg and

as high as 500 mg of polymer and found the technique is

reproducible irrespective of the polymer amount, which

is one of the key findings of the study. Gene transfection

and cellular uptake studies in cultured cells are under

way. Subsequently, investigations on scale-up processwill be performed.

Acknowledgements

MNVRK is grateful to Alexander von Humboldt

foundation, Germany for providing with a personal

fellowship. U. Bakowsky wishes to thank Stiftung

Deutscher Naturforscher Leopoldina (BMBF/LPD-

9901/8-6).

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Fig. 5. AFM images of nanospheres (A) nanospheres with PVA alone as stabilizer (B) with chitosan alone (C) PVA-chitosan blend; (D) surface

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