full papers

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Cells

Controlling Cellular Uptake by Surface Chemistry, Size,and Surface Topology at the NanoscaleMarzia Massignani, Caterina LoPresti, Adam Blanazs, Jeppe Madsen,Steven P. Armes, Andrew L. Lewis, and Giuseppe Battaglia*

Keywords:� biomimetics

� cells

� intracellular delivery

� nanovectors

� polymersomes

Cell cytosol and the different subcellular organelles house the most

important biochemical processes that control cell functions. Effective

delivery of bioactive agents within cells is expected to have an enormous

impact on both gene therapy and the future development of new therapeutic

and/or diagnostic strategies based on single-cell–bioactive-agent inter-

actions. Herein a biomimetic nanovector is reported that is able to enter

cells, escape from the complex endocytic pathway, and efficiently deliver

actives within clinically relevant cells without perturbing their metabolic

activity. This nanovector is based on the pH-controlled self-assembly of

amphiphilic copolymers into nanometer-sized vesicles (or polymersomes).

The cellular-uptake kinetics can be regulated by controlling the surface

chemistry, the polymersome size, and the polymersome surface topology.

The latter is controlled by the extent of polymer–polymer phase separation

within the external envelope of the polymersome.

1. Introduction

It is now widely accepted that the delivery of appropriate

agents to specific biological sites is one of the most important

factors in the design of an effective therapy. Efficient

administration minimizes side effects, reduces toxicity, and

permits better dosage control. Several formulations based on

both natural and synthetic systems have been developed in the

last years and some of these have been evaluated in clinical

[�] Dr. G. Battaglia, M. Massignani, Dr. C. LoPresti

Biomaterials and Tissue Engineering, The Kroto Research Institute

Department of Engineering Materials, University of Sheffield

Broad Lane, Sheffield, S3 7HQ (UK)

E-mail: g.battaglia@sheffield.ac.uk

A. Blanazs, J. Madsen, Prof. S. P. Armes

Department of Chemistry, University of Sheffield

Sheffield (UK)

Prof. A. L. Lewis

Biocompatibles UK Ltd.

Farnham, Surrey (UK)

: Supporting Information is available on the WWW under http://www.small-journal.com or from the author.

DOI: 10.1002/smll.200900578

� 2009 Wiley-VCH Verl

trials.[1] To date, one of themost challenging biological barriers

to overcome is the cell membrane. The cell cytosol hosts more

than 90% of the biochemical processes that control the

complexity of life;[2] the safe delivery of appropriate actives

that offer control over such processes is promising for any new

therapy. Drugs can be designed to passively penetrate the cell

membrane or to trigger more active transportation.[2] How-

ever, largehydrophilic actives suchasnucleic acids andproteins

cannot cross the cellmembrane. Instead, their internalization is

controlled by a complex process termed endocytosis.[2] There

are several known endocytic pathways.[3] Although mechan-

istically very different, they share two fundamental steps:

i) physical engulfment of the molecule, particles, or pathogen

via invagination of the plasma membrane, and ii) rapid

digestion of the internalized materials through acidification.

The local acidity within the subcellular-organelle lumen

responsible for the digestion of endocytized materials can be

as low as pH 4 in some cases.[3]

Effective cytosolic delivery enhances the potency of

anticancer drugs[4] and stimulates adaptive immune

responses,[5] as well as being crucial for gene therapies[6]

involvingeitherDNAorRNAdelivery.[7] Several formulations

have been reported to be effective for cytosolic delivery.[8]

These either exploit natural cell endocytosis[3] such as

liposomes,[9] cationic polymers,[10] and several nanoparticle

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Controlling Cellular Uptake at the Nanoscale

formulations[11] or involve controlled membrane disruption by

means of cell-penetrating peptides,[12] electroporation,[13]

controlled inorganic-particle precipitation,[14] carbon nano-

tubes,[15] or ultrasonic treatment.[16] However, these methods

are often compromised by high levels of cytotoxicity and hence

translation into preclinical studies has been very difficult to

achieve in practice.[17]

2. Results and Discussion

We have recently demonstrated successful cytosolic

delivery of plasmid DNA using pH-sensitive polymer vesicles

(that is, polymersomes)without affecting the cellularmetabolic

activity or even inducing inflammation.[18] Polymersomes are

produced by the spontaneous self-assembly of appropriate

synthetic amphiphilic copolymers to formmembrane-enclosed

sacs,[19] which mimic natural membrane structures such as

myelins, tissue-like structures, multilamellar aggregates, and

vesicles.[20] In particular, polymersomes are much more stable

than their low-molecular-weight counterparts[20] and in vivo

experiments have confirmed much longer circulation times

than PEG-based liposomes.[21] In the present study polymer-

somes were formed using poly(2-(methacryloyloxy)ethyl

phosphorylcholine)-co-poly(2-(diisopropylamino)ethyl

methacrylate) (PMPC25–PDPA70) and poly(ethylene glycol)-

co-poly(2-(diisopropylamino)ethyl methacrylate) (PEG23–

PDPA15) diblock copolymers comprising a highly biocompa-

tible[22] PMPC or PEG block and a pH-sensitive PDPA block,

where the subscripts denote the respective mean degrees of

polymerization. PDPA-based copolymers form stable vesicles

at physiological pH[18] but dissociate on millisecond time-

scales[23] to give individual copolymer chains (unimers) on

lowering the solution pH. This morphological transition occurs

at around thepKa (6.4)of thePDPAblock,which is comparable

to the critical local pH encountered during the endocytic

pathway. Thus, once internalized by cells, this reduction in

local pH triggers dissociation of the PMPC–PDPA polymer-

somes.The resultingdramatic increase in thenumber of species

present (each polymersome comprises many individual

copolymer chains) generates an osmotic shock that can lyse

the endosome membrane, permitting release of the encapsu-

lated active. The solution pH gradually returns to its original

valueof 7.4, leading todeprotonationof thePDPAchains.Thus

the amphiphilic character of the copolymer chains is restored

and polymersome self-assembly and/or interaction with the

endosomal membrane occurs. As illustrated in Figure 1,

such pH sensitivity is fundamental for escape from the

endosomes. The delivery efficiency was studied in detail using

primary human-dermal-fibroblast (HDF) cells donated by

patients; such unmodified primary cells are well-suited for

understanding cell–polymersome interactions. Each experi-

ment was performed in triplicate using various batches ofHDF

cells originating from three different patients. This is an

important point, because the typical patient-to-patient varia-

bility encountered in human therapy has been at least partially

explored.

Figure 1a and b shows confocal laser scanning microscopy

(CLSM) images of HDF cells after 24 h incubation with dye-

small 2009, 5, No. 21, 2424–2432 � 2009 Wiley-VCH Verlag Gmb

loaded pH-sensitive PMPC–PDPA and pH-insensitive poly-

(ethylene glycol)–poly(butylene glycol) (PEG–PBG) poly-

mersomes, respectively. In both cases the dyewas rhodamineB

octadecyl ester perchlorate. CLSM allows precise imaging of

cells at different focus levels, which allows volumetric analysis

of the localization of the different fluid-phase markers within

the cells. In the case of pH-sensitive polymersomes, the

rhodamine B dye is released throughout the cell cytosol. As

shown by costaining of the cell with both DNA-binding green

SYTO9andwitha lysosomeprobe(yellow lysotracker), thedye

clearly escapes the endolysomal compartments and stains

almost all subcellular compartments, including the nucleus

(Figure 1a). In contrast, the dye is confined within the

endolysomal compartments for the pH-insensitive polymer-

some (Figure 1b). In control experiments, cells were also

treated with free rhodamine B octadecyl ester perchlorate and

no significant fluorescence was detected.

Successful escape from the endosome has also been

demonstrated by incubating HDF cells with polymersomes

loadedwithpropidium iodide (PI) dye (Figure 1c). PIhas ahigh

affinity for DNA and, once bound to it, its dye fluorescence is

enhanced by several orders of magnitude. PI is also imperme-

able to cell membranes and is commonly used to assess cell

viability.[21] Dead cells normally have ‘‘leaky’’membranes that

allow passive diffusion of PI. Once encapsulated within the

PMPC–PDPA polymersomes, the PI is delivered within live

primary HDF cells and stains their nuclei. After PI delivery,

the cell viability was 80% as judged by an MTT-ESTA assay

(Figure 1d) compared to untreated cells. This result also

suggests that the PMPC–PDPA polymersomes can trigger the

endocytic pathway in order to deliver their cargowithin the cell

cytosol. It is emphasized that, in control experiments, primary

HDF cells were incubated with free PI and no fluorescence was

detected after 24 h. Thus the PMPC–PDPA copolymer

nanovector is essential for efficient intracellular delivery of

such dyes.

In order to better understand the subcellular fate of the

copolymer, primary HDF cells were also incubated with

copolymer chains that had been covalently labeled with

rhodamine 6G (Rh–PMPC–PDPA). The HDF cells were

counter-stained with both green SYTO9 and yellow lyso-

tracker. The z-stack micrographs in Figure 1e confirm the

homogeneous distribution of Rh–PMPC–PDPA copolymer

within the cell cytosol.Both the top viewand the side view show

the distribution of the copolymer within the cells, indicating

efficient uptake. In addition, counter-staining of the cells with

DNA and lysosome fluid-phase marker allows the relative

levels of uptake of the copolymer within the cell to be

compared. This can be understood more in detail by

colocalization studies[24] involvingRh–PMPC–PDPA, SYTO9

(Figure 1f), and Lysotracker (Figure 1g), respectively. As

shown in Figure 1f, some of the copolymer colocalizes with

DNA, indicating that it is able to reach the cell nuclei. In

Figure 1e, copolymer–lysosome colocalization indicates that

most of the copolymer chains escape the endosomes and enter

the cell cytosol.

Although the complex cell internalization pathways for

PMPC–PDPApolymersomes clearlywarrant further study, the

most importantfinding is thatPMPC–PDPApolymersomesare

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full papers G. Battaglia et al.

Figure 1. CLSM micrographs and colocalization study examining endosomal escape by the

rhodamine-labeled PMPC–PDPA chains. a) CLSM z-stack micrographs showing live HDF cells

containing PMPC–PDPA polymersomes (200nm, 24h exposure) loaded with rhodamine

octadecyl ester perchlorate B (red channel) and then treated with lysotracker (yellow channel)

and DNA staining SYTO9 (green channel). b) CLSM z-stack micrographs showing live HDF cells

containingPEG–PBGpolymersomes(200 nm,24hexposure) loadedwithrhodamineoctadecyl

esterperchlorateB (redchannel)andDNAstainingSYTO9(greenchannel). c)HDFcellsexposed

to polymersomes (200nm,24hexposure) loadedwith PI and thend) analyzedbyMTTassay to

examine cell viability. e) The first micrograph shows HDF cells treated with polymersomes

(200 nm, 24h exposure) (red channel), the second shows the nucleus-staining SYTO 9 (green

channel), the thirdshows the lysotracker (yellowchannel) and the forth shows thecombination

of all three channels merged together (composed). f) The micrograph shows colocalization

(white)betweenrhodamine-labeledpolymersomes(red)andSYTO9(green).g)Themicrograph

indicates colocalization (white) between rhodamine-labeled polymersomes (red) and

lysosomes (yellow). Error bars¼� standard deviation (n¼3).

2426 www.small-journal.com � 2009 Wiley-VCH Verlag GmbH & Co. KGaA, Weinhe

not only takenup by cells but are also able to

deliver material into the cytosol, suggesting

that theconventionalendocyticpathwaycan

be avoided, as anticipated.

As shown in Figure 2a, PMPC–PDPA

polymersomes have been employed to

deliver payloads into 23 different types of

cells including primary animal cells, primary

human cells, cancer cells, and various cell

lines. The graphical data shown in Figure 2a

summarize the results obtained by testing

the cell viability and the delivery efficiency

(calculated as the percentage of cells that

show significant fluorescence after incuba-

tion with polymersomes) of the whole panel

after 24 h incubation with rhodamine B-

octadecyl-ester-loaded PMPC–PDPApoly-

mersomes. With the exception of red blood

cells, all cell types exhibit high uptake levels

for these polymersomes. This apparent

anomaly is readily explained: red blood

cells are one of the few types of eukaryotic

cells that do not exhibit active endocyto-

sis.[3,7]

The connection between polymersome

internalization and active endocytosis was

confirmed by inhibition studies on HDF

cells. The bar chart in Figure 2b shows the

extent of cellular uptake normalized with

respect to the positive control (i.e., 1 h

incubation at 37 8C) of rhodamine-conju-

gatedPMPC–PDPAandrhodamine-loaded

PEG–PDPA polymersomes and PMPC–

PDPA polymersomes. Incubation of cells

at 4 8C should minimize most of the move-

ment within the cellular membrane,[25]

thereby switching off all the endocytic

pathways. As shown, both PEG- and

PMPC-based polymersomes show either

very limited or no cellular uptake under

these conditions, confirming that both these

polymersome formulations require an

active endocytic pathway to enter cells.

Incubation with chlorquine, another endo-

cytosis blocking agent, almost completely

prevented internalization. HDF cells were

also incubated with nocodazole. This drug

inhibits microtubule polymerization[26] and

hence interferes with the various biochem-

ical processes that require microtubule

assembly for mobility. These include trans-

portation of the trafficking vesicles coming

from/to the cellular membrane and from/to

endosomes, endoplasmic reticulum, and the

Golgi apparatus. CLSM images shown in

Figure 2c (PMPC–PDPA) and d (PEG–

PDPA) indicate that the polymersomes,

although internalized, are blocked in the

vicinity of the cellular membrane when

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Controlling Cellular Uptake at the Nanoscale

Figure 2. Viability, internalization, and inhibition studies performed by means of MTT-ESTA

assay, flow cytometry, and CLSM. a)MTT-ESTA assay performedon 23 different cell types, both

cell lines and primary cells, compared to their intracellular uptake (delivery efficiency) of

polymersomes. b) Inhibition study performed on HDF cells treated with polymersomes at 4 8Cafter treatment with nocodazole and chlorquine, respectively, relative to control samples

incubated with polymersomes for 1 h at 37 8C. c) CLSM z-stack micrographs showing HDF cells

treated with nocodazole and then loaded with either PMPC–PDPA polymersomes (c) or PEG–

PDPA polymersomes (d). Error bar¼� standard deviation (n¼ 3).

HDF cells are pretreated with nocodazole. This can be

visualized more clearly on the z–x slide section: hardly any

material reaches the cell interior. Cells were also treated with

lysotracker, showing that the microtubule disruption also

affects internalizationof thefluid-phasemarker, confirming the

observations made for the polymersomes.

Inorder toquantify thedegreeof internalization,HDFcells

were digested using a nonionic detergent (tritonX-100) and the

resulting dispersions were analyzed by UV–Vis absorption

spectroscopy. Calibration and standard curves were con-

structed by adding known concentrations of dye-labeled

polymersomes to the digested untreated cell dispersions. This

protocol allows the amount of internalized material per single

cell to be monitored as a function of the incubation time. In

small 2009, 5, No. 21, 2424–2432 � 2009 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

addition to the influence of the surface

chemistry, the effect of varying the poly-

mersome dimensions on their kinetics of

internalization has also been studied.

According to Figure S1 in the Supporting

Information, the hydrodynamic polymer-

some diameter can be effectively controlled

by extruding the polymersome dispersions

through differently sized polycarbonate

membranes, as judged by dynamic light

scattering (DLS).[27]

In Figure 3 the number of internalized

vesicles per single cell was plotted as a

functionof incubation time. It is evident that

both the surface chemistry and vesicle size

markedly affect the internalization kinetics.

The rate of internalization of the PMPC–

PDPA polymersomes is an order of magni-

tude faster than that of the PEG–PDPA

polymersomes. This substantial difference

in uptake kinetics between the PMPC and

PEG polymersomes seems to be more or

less constant over the 100–400-nm size

range investigated. The cellular uptake

behavior observed for the 50:50 (mol/mol)

PMPC–PDPA/PEG–PDPA binary mix-

ture appears to be primarily dictated by

the PMPC chains at short incubation times,

with intermediate behavior being observed

at longer incubation times. This is particu-

larly evident for the 400-nm vesicles.

However, a 75:25 PMPC–PDPA/PEG–

PDPAbinarymixtureexhibits fasteruptake

in the first few hours regardless of the

polymersome dimensions. The 25:75

PMPC–PDPA/PEG–PDPA formulation is

taken up more slowly than the 75:25

formulation, but the mean vesicle diameter

alsoplays a role indetermining theseuptake

kinetics. In particular, the kinetic data

obtained for the 400-nm polymersomes is

unexpected: these hybrid species are inter-

nalized much faster than even those poly-

mersomes comprising solely PMPC–PDPA

copolymer.

Several studies have demonstrated that the particle size has

a major effect on the internalization kinetics.[28] Maximum

rates of uptake were obtained for particle diameters of

30–40 nm, with progressively slower kinetics being observed

for larger particles. On this basis, it is reasonable to assume that

100-nm polymersomes should be internalized faster than

400-nmpolymersomes. This is indeed the case for both strongly

cell-binding PMPC and weakly cell-binding PEG polymer-

somes. However, the hybrid polymersomes formed from the

binary copolymer mixtures do not follow this trend. Recently,

we have demonstrated that simply mixing two membrane-

forming copolymers allows the formation of nanometer-sized

domainson thepolymersome surface.[29] Thesedomains aredue

to microphase segregation between the thermodynamically

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full papers G. Battaglia et al.

Figure 3. The effect of polymersomesurface chemistry anddimensions on cellular internalization. Thenumber of internalizedpolymersomesper cell

was plotted as a function of the incubation time for three different polymersome diameters. Error bars¼� standard deviation (n¼3).

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immiscible PMPC and PEG chains. The size and the

morphology of such domains are strongly controlled by the

copolymer molar ratio. As shown in Figure 4, we have

developed a method to visualize the domains by selectively

staining the PMPC chains using phosphotungstenic acid (PTA)

(see supplementary information for more details).[29] This

electron-dense salt is known to react with the carboxylic ester

groups on the PMPC chains.[30] Transmission electron micro-

scopy (TEM) images both shown as raw and fast Fourier

transform(FFT)-filtered (shown inbothgrayscaleand inacolor

gradient) indicate the presence of nanometer-sized PMPC

domains, confirming that the PMPC and PEG polymersomes

do not mix homogeneously. The average size of these domains

increases linearly[29] as the polymersome dimensions are

increased. Thus the two copolymer chains phase-separate to

form �10-nm domains within a 100-nm polymersome, while

larger (�40 nm) domains are formed within a 400-nm

polymersome. The ‘‘patchy’’ nature of the external surface

of the binary mixture formulations allows the internalization

kinetics to be rationalized as a function of the polymersome

size. InFigure 4, alongside theTEMmorphologies observed for

different polymersome formulations, the number of polymer-

somes internalized after incubation for 24 h is plotted against

themean polymersome diameter. In the case of polymersomes

comprising purely PMPC surfaces, the mean diameter

significantly affects their rate of cellular uptake. Thus the

100-nm polymersomes are internalized up to two orders of

magnitude faster than the 400-nm polymersomes (Figure 4a

and b). The purely PEG-based polymersomes exhibit a similar

trend (Figure 4i–j), albeit with much lower efficiency.

Incorporation of just 25mol% PEG–PDPA within a

PMPC–PDPA polymersome changes its topology and PEG-

rich domains become visible, forming a pattern typical of

bimodal separation (Figure 4c).Generation of suchPEG-rich

domains forces the PMPC chains to adopt a smaller surface

area compared to a pristine PMPC-based polymersome of

similar size. This change in topology also affects the

internalization kinetics, since fewer 100-nm ‘‘hybrid’’ poly-

mersomes are internalized faster than the pristine PMPC

polymersomes. However, the mean hybrid polymersome

diameter affects cellular uptake much less than that for the

pristine PMPC polymersomes. The polymersome obtained

from the 50:50 binary mixture displays much bigger domains

(Figure 4e), which seems to suggest enhanced spinodal

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decomposition. In this case, the rate of uptake is intermediate

between the pure PMPC and the pure PEG polymersomes

with a discernible size effect, although much less pronounced

than that observed for the pure PMPC polymersomes. The

most interesting formulation is the 25:75 PMPC–PDPA/

PEG–PDPA binary mixture: here discrete PMPC domains

are formed within a PEG matrix with a typical bimodal

decomposition pattern (Figure 4g). Although only 25mol%

of the polymersome surface comprises PMPC chains, its

uptake kinetics are almost as rapid as that observed for the

pure PMPC polymersomes. In addition, this hybrid topology

results in faster uptake kinetics for larger polymersomes

(Figure 4h). According to our previous studies,[29] such

100-nm-diameter polymersomes contain approximately

10-nm domains, while 400-nmpolymersomes comprise larger

domains of around 40 nm. If the rate of uptake reaches a

maximum at a particle size of 30–40 nm,[28] this suggests that

the domain size (rather than the overall polymersome

diameter) actually controls cellular uptake. Clearly, the

PMPC chains have a much higher affinity for cell membranes

than the PEG chains. However, binary mixtures of these two

copolymers produce either PMPC- or PEG-rich domains and

it is the size of these domains that seems to control the rate of

internalization. This effect is enhanced for the 400-nm

polymersomes since their 40-nm-domain sizes seem to be

optimal for cell internalization.[28]

3. Conclusions

In conclusion, we report a cytosolic delivery system that can

deliver a wide range of molecular payloads into cells without

compromising their viability. Detailed kinetic studies indicate

that not only are the polymersome dimensions and surface

chemistry vital for efficient intracellular delivery but that

the precise surface topology of the polymersome also plays an

important role. This has allowed us to identify two comple-

mentary pH-sensitive polymersomes based on PMPC and

PEG: judicious binary mixtures of these two diblock copoly-

mers leads to the formation of polymersomes with specific

domains within their exterior envelopes. Such microphase

separation in turn allows the rate of cellular uptake to be fine-

tuned.

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Controlling Cellular Uptake at the Nanoscale

Figure 4. TEMimagesshowingbothselectivestainingof thePMPCchainsatdifferentPMPC/PEGmolar ratiosandalsocelluptakeafter24 hincubation

asa functionof thepolymersomediameter.BinarymixturesofPMPC–PDPAandPEG–PDPAcopolymers formhybridpolymersomesthatdisplaysurface

domains due to microphase segregation. TEM images were analyzed using FFT filtering to minimize background noise. The resulting images are

displayedingrayscaleandusingacolorgradientsoastohighlight thenanoscaledomainsonthepolymersomesurface.a)TEManalysisofPMPC–PDPA

polymersomes.b)Numberofpolymersomes internalizedpercell after incubationofHDFcells for24 hwithPMPC–PDPApolymersomesasa functionof

polymersome dimensions. c) TEM analysis of 75:25 PMPC–PDPA/PEG–PDPA polymersomes. d) Number of polymersomes internalized per cell after

incubationofHDFcells for24 hwith75:25 PMPC–PDPA/PEG–PDPApolymersomesasa functionofpolymersomesize.e) TEManalysisof50:50PMPC–

PDPA/PEG–PDPApolymersomes. f)Numberofpolymersomes internalizedper cell after incubationofHDFcells for24 hwith50:50 PMPC–PDPA/PEG–

PDPApolymersomesasafunctionofpolymersomesize.g)TEManalysisof25:75PMPC–PDPA/PEG–PDPApolymersomes.h)Numberofpolymersomes

internalizedper cell after incubationofHDFcells for 24 hwith25:75 PMPC–PDPA/PEG–PDPApolymersomesasa functionofpolymersomesize. i) TEM

analysisof PEG–PDPApolymersomes. j)NumberofPEG–PDPApolymersomes internalizedpercell after incubationofHDFcells for24 hasa functionof

polymersome size. Scale bar¼ 50nm; error bar¼� standard deviation (n¼ 3).

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full papers G. Battaglia et al.

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4. Experimental Section

Materials: MPC (>99%) was kindly donated by Biocompa-

tibles UK Ltd. DPA was purchased from Scientific Polymer Products

(USA). CuBr (99.999%), CuCl (99.995%), 2,2-bipyridine (bpy),

methanol, isopropanol, 2-bromoisobutyl bromide, and triethylamine

were purchased from Aldrich and were used as received. The silica

used for removal of the atom transfer radical polymerization

(ATRP) copper catalyst was column chromatography grade silica

gel 60 (0.063–0.200mm) purchased from E. Merck (Darmstadt,

Germany). 2-(N-Morpholino)ethyl 2-bromo-2-methylpropanoate

(ME-Br) initiator was synthesized according to a previously

reported procedure.[31] Chloroform was purchased from Fisher

Scientific, phosphate buffer saline (PBS) tablets from Oxoid Ltd,

and sepharose 4B and rhodamine B octadecyl ester perchlorate

from Sigma-Aldrich. Ham’s F12 media, Dulbecco’s modified

Eagle’s medium (DMEM), and fetal calf serum were bought from

Biosera (UK) and L-glutamine, penicillin streptomycin, and

amphotericin B were bought from Sigma (UK). Collagenase A

was purchased from Boehringer-Mannheim (Lewes, UK). For the

MTT-ESTA assay, 3-(4,5-dimethylthiazol-2-yl)-2,5 diphenyl tetra-

zolium bromide (MTT) was purchased from Sigma-Aldrich (UK) and

hydrochloric acid from BDH AnalaR. DPA (Scientific Polymer

Products, USA) was passed through the column supplied by the

manufacturer to remove inhibitor. Monohydroxy-capped poly-

(ethylene oxide) was purchased from Fluka and freeze-dried

before use to remove water. Regenerated cellulose (RC) dialysis

tubing was purchased from Spectrum Labs (Spectra Por 6,

molecular weight cut-off 3.5 kD).

PMPC25–PDPA70 copolymer synthesis: PMPC25–PDPA70 was

synthesized by ATRP as reported elsewhere.[32] Briefly, a Schlenk

flask with a magnetic stir bar and a rubber septum was charged

with CuBr (25.6mg, 0.178mmol) and MPC (1.32 g, 4.46mmol).

ME-Br initiator (50.0mg, 0.178mmol) and bpy ligand (55.8mg,

0.358mmol) were dissolved in methanol (2mL), and this solution

was deoxygenated by bubbling N2 for 30min before injection into

the flask using a syringe. The [MPC]/[ME-Br]/[CuBr]/[bpy] relative

molar ratios were 25:1:1:2. The polymerization was conducted

under a nitrogen atmosphere at 20 8C. After 65min, a deoxyge-

nated DPA (6.09 g, 28.6mmol) and methanol (7mL) mixture was

injected into the flask. After 48 h, the reaction solution was diluted

by addition of isopropanol (about 200mL) and then passed

through a silica column to remove the spent Cu catalyst.

Vesicle preparation and propidium iodide incapsulation: In a

typical experiment, PMPC25–PDPA70 copolymer (20mg) was

added to a glass vial and dissolved in a solution of 2:1 chloro-

form/methanol at a concentration of 3mgmL�1. The solvent was

evaporated under vacuum in sterile conditions, resulting in a

copolymer film deposited on the walls of the vial. The copolymer

film was rehydrated under sterile conditions using 100mM PBS

at pH 2 to form a 0.5% w/w copolymer suspension. The solution

was then sterilized by filtration (200-nm pore size). The

solution pH was then increased to pH 6 and propidium iodide

was added to the vesicle dispersion, which was then sonicated for

30min. Vesicles encapsulating propidium iodide were purified via

gel permeation chromatography (GPC) using a size exclusion

column containing sepharose 4B and PBS at pH 7.3 to elute the

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vesicles. Rh–PMPC30–PDPA60 copolymer was synthesized by an

ATRP procedure as reported elsewhere.[32] Briefly, a Schlenk flask

with a magnetic stir bar and a rubber septum was charged with

MPC (1.20 g, 4.05mmol). Rh initiator (83.8mg, 0.135mmol) was

dissolved in anhydrous methanol (0.75mL) and added to the

MPC. The flask was washed with 0.75mL anhydorus methanol,

which was added to the MPC solution. The solution was then

deoxygenated by bubbling N2 for 30min. After this period, a

mixture of CuBr (19.37 mg, 0.135mmol) and bpy ligand

(42.17mg, 0.171mmol) was added to the reaction mixture after

mixing the solids with a spatula. The [MPC]/[Rh]/[CuBr]/[bpy]

relative molar ratios were 30:1:1:2. The reaction was carried out

under a nitrogen atmosphere at 20 8C. After 40min, a deoxyge-

nated DPA (1.73 g, 8.10mmol) and methanol (2mL) mixture was

injected into the flask. After 48 h, the reaction solution was diluted

by addition of methanol (about 70mL) and opened to the

atmosphere. When the resulting suspension had turned green,

chloroform (about 200mL) was added to thoroughly dissolve the

copolymer and the resulting solution was passed through a silica

column to remove the catalyst. After removal of the solvent, the

solid was taken up into chloroform/methanol 3:1 and dialyzed for

3 days against this solvent mixture with daily changes of solvent

in order to remove residual bpy ligand. After evaporation, the solid

was dispersed in water, freeze-dried, and dried in a vacuum oven

at 80 8C for 48 h.

PEO23–PDPA15 copolymer synthesis: DPA monomer, PEO23–Br

macroinitiator, and bpy were weighed into a round-bottomed

flask. Methanol was separately degassed under nitrogen for

20min, added to the reaction solution (overall monomer

concentration¼50% v/v) and further degassed for 10min. This

mixture was then heated to 50 8C and CuCl catalyst was added

under constant nitrogen flow. The reaction solution turned dark

brown and became more viscous, indicating the onset of

polymerization. After polymerization for 16 h, an aliquot was

removed for 1H NMR analysis. The reaction was terminated by

exposure to air after 1H NMR analysis indicated no remaining

monomer (>99.9% conversion). The crude copolymer was

dissolved in tetrahydrofuran (THF) and the spent catalyst was

removed by column chromatography (silica stationary phase).

Evaporation of the THF produced a viscous yellow copolymer. To

remove unreacted PEO initiator and bpy from the reaction product,

the copolymer was dissolved in minimal THF and dialyzed against

water for 1week. The solution was then exchanged with methanol

for 2 days and the methanol then evaporated to yield a clear

viscous polymer. The copolymer was then dried under vacuum and

characterized by 1H NMR spectroscopy and THF GPC.

Polymersome formation: Rh–PMPC30–PDPA60 was dissolved in

a 2:1 chloroform/methanol mixture to form a 0.05mM solution.

0.05�10�3moles of this solution were added to three different

vials each containing 1�10�3moles of PMPC25–PDPA70 copoly-

mer, PEO23–PDPA15 copolymer, a 50:50 mixture of these, a 25:75

mixture of these, and a 75:25 mixture of these. The polymers were

dissolved in 2:1 chloroform/methanol mixture, and the solvent

was evaporated under vacuum, resulting in a copolymeric film

deposited on the walls of the vial. The copolymer film was

rehydrated under sterile conditions using 100mM PBS at pH 2 to

form a 0.250mM copolymer solution. The solution was filter

sterilized using a 0.2-mm pore filter before the pH was increased

bH & Co. KGaA, Weinheim small 2009, 5, No. 21, 2424–2432

Controlling Cellular Uptake at the Nanoscale

to 6.4. Bioactive molecules (propidium iodide, rhodamine B

octadecyl ester perclorate) to be encapsulated were added at this

stage to the polymer dispersion. The pH of the solution was

increased to 7.3 before being sonicated for 5min. The solution

was passed back and forth 31 times through a membrane with

pore sizes of 100, 200, and 400 nm using two gas-tight glass

syringes. Finally vesicles were purified via GPC using a size-

exclusion column containing Sepharose 4B and using PBS at

pH 7.3 to elute the vesicles.

Cell culture: Primary HDFs were isolated from skin obtained

from abdominoplasty or breast reduction operations (according to

local ethically approved guidelines, NHS Trust, Sheffield, UK).

Primary cultures of fibroblasts were established as previously

described.[33] Briefly, the epidermal layer of the skin was removed

by trypsinization and the remaining dermal layer was washed in

PBS. The dermis was then minced using surgical blades and

incubated in 0.5% (w/v) collagenase A at 37 8C overnight in a

humidified CO2 incubator. A cellular pellet was collected from the

digest and cultured in DMEM supplemented with 10% (v/v) fetal

calf serum, 2mM L-glutamine, 100 IUmL�1 penicillin, 100mgmL�1

streptomycin, and 0.625mgmL�1 amphotericin B. Cells were

subcultured routinely using 0.02% (w/v) ethylenediaminetetraa-

cetic acid (EDTA) and used for experimentation between passages

4 and 9.

MTT-ESTA assay: All cells (HDF cells, epidermal cheratinocites,

MLO-A5, limbal epithilial cells, cho cells, Sc25, Scc4, Fadu, KB

cells, melanoma C8161, melanoma A357, MCF7, lung carcinoma

H1299, endothelial cells, oesophagos fibroblasts and keratino-

cites) were seeded in standard 24-well plates at a density of

3�104 cells per well and grown for two days in culture medium.

Blood cells (macrophages, monocites, and red blood cells) were

left and treated in suspension. Cell monolayers/suspensions were

treated for 24 h with fresh medium containing 1mgmL�1 of

polymersomes. Viable cell density was then assessed using the

MTT assay. Briefly, treated/untreated cells were washed thor-

oughly twice with PBS and then incubated in MTT solution

(0.5mgmL�1 MTT in PBS, 1mL per well of 24-well plate or per cm2

of cultured tissue) for 45min at 37 8C and 95% air/5% CO2.

Intracellular metabolic activity reduces MTT to a purple formazan

salt. Subsequently, the solution was aspirated and the insoluble

formazan product was solubilized by adding isopropanol (0.5mL

per well of 24-well plate or 1mL cm�2 cultured tissue) and

incubating for 10min. The optical density at 540 nm was then

measured using a plate-reading spectrophotometer (Dynex

Technologies, MRX II).

Dynamic light scattering: DLS measurements were performed

with a Brookhaven Instruments 200 SM laser light scattering

goniometer using a He�Ne 125mW laser operating at 633 nm.

The vesicle dispersions were placed into glass vials. Single

analyses of 10min exposure were performed on each sample.

Particle sizes were estimated using the CONTIN multiple pass

method of data analysis.

Epifluorescence microscopy and confocal laser scanning

microscopy: Standard 6-well plates were seeded at a density of

5�104 cells per well for HDF cells, and grown for two days in

culture medium (see above). Cells were replenished with fresh

medium (1.5mL) and polymersomes were loaded at 50 mM. Cells

were incubated for 24 h in a humidified CO2 incubator. Thereafter,

small 2009, 5, No. 21, 2424–2432 � 2009 Wiley-VCH Verlag Gmb

the cell monolayers were washed three times with PBS and left in

fresh PBS for visualization. Samples were visualized by epifluor-

escence microscopy CLSM. Micrographs of living cells were taken

at lex¼543 nm/lem¼560 nm for the detection of rhodamine. This

was performed using a ZEISS LSM 510M instrument.

Flow cytometry measurements/delivery efficiency: Cells were

trypsinized using trypsin-EDTA, a cellular pellet was collected, and

the cells were resuspended in PBS. Cells were then analyzed using

flow cytometry.

Inhibition study: Cells were left at 4 8C for 1 h and then treated

with cold polymersomes for 30min before flow cytometry. Cells

were incubated with chlorquine (0.05mM) for 1 h and then treated

with polymersomes for 1 h in the presence of the inhibitor before

being analyzed by flow cytometry. HDF cells were incubated with

nocodazole (0.01mgmL�1) for 1 h, then treated with polymer-

somes in the presence of inhibitors for an additional 1 h at 37 8C.Finally, they were washed three times with PBS before treatment

with lysotracker and visualization with confocal microscopy.

This was done with both rhodamine-tagged PMPC–PDPA and

rhodamine-encapsulated PMPC–PDPA and PEG–PDPA polymer-

somes.

Lysis experiment: Standard 6-well plates were seeded at a

density of 5�104 cells per well for HDF cells and grown for two

days in culture medium (see above). Cells were replenished with

fresh medium (1.5mL) and polymersomes were loaded at 50mM.

Cells were incubated for 5min, 1 h, 3 h, 5 h, 8 h, 24 h, 48 h, and

72 h in a humidified CO2 incubator. Thereafter, the cell mono-

layers were washed three times with PBS. Triton X100 (100mL)

was added to each well and cells were left for 1 h at �80 8C. Thedigested solution was transferred in a 96-well plate. Absorbance

analysis was performed by UV plate reader at 540 nm and the

values were related to a calibration curve. Calibration curves were

generated by adding known amount of polymersomes in a

digested cells solution. The number of vesicles per cell was

calculated by

Nvesiclepercell

¼wcopolymer NaggregatePMPC�PDPAfPMPC�PDPAþNaggregatePEG�PDPAfPEG�PDPAð ÞMwPMPC�PDPAfPMPC�PDPAþMwPEG�PDPAfPEG�PDPAð ÞNcells

(1)

where Nvesicle per cell is the number of vesicles internalized per single

cell, wcopolymer is the measured weight of copolymer after cell lyses,

Mw copolymer is the molecular weight of the copolymer, fcopolymer is

the molar fraction of the copolymer, Ncells is the total number of

cells used in each experiment, and Naggregate copolymer is the number

of aggregate of a single vesicle calculated by

Naggregate ¼4p d

2

� �2þ d2 � t� �2

amolecule(2)

where d is the vesicle diameter as measured by DLS, t is the

thickness as measured by TEM, and amolecules is the area per

molecule calculated using the method reported previously.[34,35]

TEM sample preparation and imaging: PTA solution prepara-

tion: 37.5mg of PTA were dissolved in 5mL of boiling distilled

water upon stirring for 5min. The pH of the solution was adjusted

to 7.0 by adding small drops of 5 M NaOH under continuous

H & Co. KGaA, Weinheim www.small-journal.com 2431

full papers G. Battaglia et al.

2432

stirring. The PTA solution was then filtered through a 0.2-mm filter.

Polymersome/PBS dispersions (5mL) at 5mgmL�1 were depos-

ited onto glow-discharged copper grids. After 1min, the grids were

blotted with filter paper and then immersed into the PTA staining

solution. The grids were blotted a second time and dried under

vacuum for 1min. PTA has been used as a selective staining agent

for the PMPC–PDPA block copolymer. Different dwelling times of

the grids in the PTA solution were tested in order to find the best

conditions for obtaining both negative and positive staining. After

10 s dwelling time, the majority of the vesicles resulted negatively

stained, while after 5 s the resulting staining was mainly positive.

TEM imaging analysis: TEM micrographs were analyzed using

DigitalMicrograph Demo 3.9.5 for GMS 1.4.5 developed and

commercialized by the Gatan Software Team of Gatan Inc.

(Pleasanton, CA, USA). Raw data were collected and analyzed by

FFT filtering techniques.

Acknowledgements

We thank Dr. Irene Canton for useful discussions and

assistance with cell-culture studies and Prof. Sheila MacNeil,

Ms. Claudia Marques, Ms. Pallavi Deshpande, Ms. Vanessa

Hearden, and Ms. Anna Tellechea Lopez for providing access to

the dermal and oral fibroblasts, epidermal and oral keratino-

cytes, dermal endothelium, melanoma A8161, C8375, H357,

Limbal epithelium cells. We thank Dr. John Haycock and Ms.

Celia Morris-Smith for providing access to the Rat Schwannoma

Nemap22 cells and Dr. Gwen Reilly for the Preosteocytes MLO-

A5 and the primary MSC Stem cells. We also acknowledge

Dr. Jonathan Bury and Dr. Nicola Green for help in extracting

the oesophagus cells and Prof. Martin Thornhill and Dr. Craig

Murdoch for assistance with all oral cancer cells and the blood

cell types. This work was sponsored by Biocompatibles Ltd.

MM is recipient of an EPSRC/White Rose LSI -DTC studentship.

CL is sponsored by EPSRC (EP/E03103X/1). SPA is the recipient

of a five-year Royal Society-Wolfson Research Merit Award.

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bH & Co. KGaA, Weinheim

Received: April 5, 2009Revised: May 12, 2009Published online: July 27, 2009

small 2009, 5, No. 21, 2424–2432