Protein decorated membranes by specific molecular interactions

PAPER www.rsc.org/softmatter | Soft Matter

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

Protein decorated membranes by specific molecular interactions†

Rainer Nehring,a Cornelia G. Palivan,a Susana Moreno-Flores,b Alexandre Mantion,c Pascal Tanner,a

Jose Luis Toca-Herrera,b Andreas Th€unemannc and Wolfgang Meier*a

Received 10th February 2010, Accepted 15th April 2010

First published as an Advance Article on the web 18th May 2010

DOI: 10.1039/c002838j

Here we characterize new metal-functionalized amphiphilic diblock copolymers, developed for both

surface and solution molecular recognition applications. Polybutadiene-block-poly(ethylene oxide)

copolymers functionalized with nitrilotriacetic acid and tris(nitrilotriacetic acid) were complexed with

nickel(II) to obtain coordination sites for oligohistidine residues of model proteins. Mixtures of

functionalized polymers with the respective non-functionalized block copolymers self-assemble in

aqueous solution into vesicular structures with a controlled density of the metal end-groups on their

surface. In solution, binding of His6-tagged green fluorescent protein (EGFP) and red fluorescent

protein (RFP) to the vesicle surface was quantified by fluorescence correlation spectroscopy. Small-

angle X-ray scattering indicates an increase of the membrane thickness by 2–3 nm upon protein

binding. Block copolymer monolayers at the air–water interface and on solid support served as a model

system to characterize the protein-decorated membranes by Brewster angle microscopy and AFM.

High resolution AFM of solid-supported, hydrated monolayers indicates that the proteins form densely

packed and partially ordered arrays with the cylindrically shaped EGFP molecules lying flat on the

surface of the films.

Introduction

Many experimental approaches in nanotechnology, biochemistry

and medicine for diagnostic or drug delivery systems require

the immobilization of proteins onto substrates.1–3 Therefore,

the control of surface properties of biomaterials as well as the

modulation of binding capacity and specific recognition are

essential for the development of functional surfaces to be used in

molecular recognition and medical diagnostics,4–6 heterogeneous

biocatalysis,7,8 cellular patterning9,10 and high effective purifica-

tion of specific proteins.11,12

For example, in order to overcome some of the limitations in

protein purification, the use of ‘‘tag’’ concepts has found wide

application, whereby a generic peptide (e.g. oligohistidines) is

aDepartment of Chemistry, University of Basel, Klingelbergstrasse 80,CH-4056 Basel, Switzerland. E-mail: [email protected]; Fax:+41 (0)61 267 3850; Tel: +41 (0)61 267 3802bCIC BiomaGUNE—Biosurfaces unit, Paseo Miram�on 182, E-20009 SanSebasti�an, SpaincBAM—Federal Institute for Materials Sciences and Testing, RichardWillst€atterstraße 11, 12489 Berlin, Germany

† Electronic supplementary information (ESI) available: Fig. S1 showsa transmission light microscope image of a drop ofPB39-PEO36-SA-OH/PB39-PEO36-SA-TrisNTA.d-Ni2+ 10 : 1, in bidi-stilled water as a function of time. Table S1 presents the results of DLSfor copolymers and copolymers/metal-functionalized analogues.Table S2 shows the results of SAXS data analysis after protein bind-ing to the metal centers of vesicle surfaces. Fig. S2 shows cryo-TEMimages of PB60-PEO34-SA-OH/PB60-PEO34-SA-NTA.d-Ni2+solutions.Fig. S3 shows SAXS data from PB60-PEO34-SA-OH/PB60-PEO34-SA-NTA.d-Ni2+solutions in PBS. Fig. S4 shows the FCS data forprotein-bound fraction as function of pH and protein incubation time.Fig. S5 shows the Langmuir compression isotherm of the diblockcopolymer PB60-PEO34-OH on double distilled water with thecorresponding BAM images. See DOI: 10.1039/c002838j

This journal is ª The Royal Society of Chemistry 2010

introduced at the N- or C-terminal of the target protein to form

a ‘‘binding’’ structure.13–16 In addition, the application of

immobilized metal ion affinity chromatography for the isolation

of recombinant proteins has attracted great interest, due to its

selectivity and mild elution conditions.17–19 Hearn et al. even

proposed a new class of immobilized metal ion chelate complexes

to selectively bind proteins as a function of pH or ionic

strength.20 In this context, the combination of polymers with

NTA ligands offers a wide range of possibilities for selective

protein immobilization.21 Bruening et al. described the use of

thick poly(acrylic acid) (PAA) brushes derivatized with NTA–

Cu2+ where more than 10 monolayers of protein can be immo-

bilized in PAANTA–Cu2+ brushes.22 Block copolymer

membranes are considerably thicker and chemically and

mechanically more stable than conventional lipid bilayers, while

mimicking natural biomechanical properties.23 In addition, their

chemical constitution, the relative lengths and structure of the

different blocks, or even the architecture of the whole polymer

can be designed with respect to the desired application, which

make them more versatile than lipidic membranes.24

Likewise, it is shown that amphiphilic block copolymers can

be functionalized with metal chelating moieties without affecting

their ability to self-assemble into well-defined membrane struc-

tures as vesicles.25 This vesicular structure was proposed as an

ideal model system for biological compartments in aqueous

media26–29 and for particular copolymer types the system can

possess pH-induced sensitivity30 or metal binding can be used to

induce formation of domains.31

The attachment of proteins to surfaces is crucial for drug

targeting approach using polymer particles as well as in many

biotechnological processes and applications.32–34 Additionally,

the oriented immobilization of densely packed proteins to model

surfaces is a key step for 2D protein crystallization.35 For these

Soft Matter, 2010, 6, 2815–2824 | 2815

http://dx.doi.org/10.1039/C002838J

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

purposes, one needs to control the adsorption strength and

protein orientation, which can be achieved only for specific

binding of the protein to a homogeneous and stable interface.36

Model amphiphilic block copolymers with narrow molecular

weight distributions, controllable block lengths, and high purity

can be used for building well-defined monolayer and membrane

surfaces.25 These requirements are met by poly(butadiene)-block-

poly(ethylene oxide) diblock copolymers where the hydrophobic

blocks (PBs) are crosslinkable under UV light.23 We previously

modified such polymers with NTA–Me2+ end groups and showed

that these groups are accessible for molecular recognition at

vesicle surfaces.25

Here we were interested in gaining more insight into the

selective binding between functional polymer membranes based

on Ni2+–NTA functionalized poly(butadiene)-block-poly-

(ethylene oxide) diblock copolymers and His-tagged proteins,

both in vesicles dispersions and on surfaces based on polymer

monolayers. We characterized the structure of the metal deco-

rated vesicles and their binding to selected His-tagged model

proteins by varying the density of the Ni2+–NTA groups at the

membrane surfaces or the experimental conditions such as the

composition of the buffer. Vesicular structures were investigated

by a combination of induced coupled plasma atomic emission

spectroscopy, fluorescence correlation spectroscopy, small angle

X-ray scattering, and x-potential measurement experiments.

Block copolymer monolayers were prepared by Langmuir–

Schaefer and incubated with a His-tagged protein to investigate

protein binding to metal-functionalised polymer surfaces.

Atomic force microscopy revealed the first structural details of

the protein-decorated membranes and indicated that polymer

membranes or monolayers might induce the formation of highly

ordered protein arrays.

Materials and methods

Materials

All reagents and solvents were purchased from Aldrich or Fluka

with the highest purity grade and used as received unless other-

wise noticed.

Milli-Q water (Nanopure diamond, Barnstead) with 18.2 MU

cm resistivity was used to prepare the protein solutions and

samples for the atomic force microscopy (AFM) measurements.

His6-enhanced green fluorescent protein (His6-EGFP) and His6-

red fluorescent protein (His6-RFP) (BioVision, California, USA)

were used without further purification. His-EGFP was used as

delivered, while His6-RFP was reconstituted with bidistilled

water to a concentration of 1 mg mL�1, prior to use.

Vesicle preparation. The synthesis of the block copolymers

used is described in Nehring et al.25 In order to form vesicles by

the film rehydration method, the polymer (or a mixture of

different polymers) was dissolved in chloroform in a round-

bottomed flask. The solvent was evaporated under reduced

pressure, until a thin polymer film was formed on the inner glass

surface of the flask. Each film was either hydrated with Milli-Q

water, bidistilled water or buffer, respectively by overnight

rotation of the flask (25 rotations per minute) at room

2816 | Soft Matter, 2010, 6, 2815–2824

temperature under atmospheric pressure. Vesicle concentrations

ranged from 5 mM up to 2 mM.

Methods

Optical microscopy. Vesicle emulsions in bidistilled water or

PBS-buffer were examined with a transmission light microscope

(DMIRE2, Leica Camera AG, Solms, Germany) at a magnifi-

cation of 100 � 10 (oil immersion objective: HCX PL APO CS

100�, aperture 1.4) in bright field or with a polarisation filter.

For fluorescence images, the microscope was used in fluorescence

mode (fluorescence lamp, ebq. 100 isolated, Jena Germany).

Prior to visualization, the protein-decorated vesicle solutions

were dialyzed to remove excessive and unbound protein.

Inductively coupled plasma atomic emission spectroscopy.

Nickel(II) content in the metal-functionalised vesicles was

determined by ICP-AES. After filtration through a PC

membrane (10 mm pore diameter) the vesicle solutions were

injected from a cyclone spray chamber into the argon plasma at

a temperature of 10 000 K.

Dynamic light scattering. DLS measurements were carried out

using a Malvern Instruments particle sizer (Zetasizer Nano ZS,

Malvern Instruments, UK) equipped with a He–Ne laser (l/nm

632.8) working in backscattering mode at a scattering angle of

2q ¼ 173�. The solutions were placed into 10 � 10 mm plastic or

quartz cuvettes. The Stokes–Einstein relation was used to

calculate the vesicle hydrodynamic diameter.

x-Potential and size measurements. The x-potential and diam-

eter of the polymer vesicle solutions were measured at room

temperature with a Zetasizer Nano-ZS (Malvern Instruments,

UK). The x-potential was calculated using Henry’s equation and

the Smoluchowski approximation, as implemented in the Zeta-

sizer Control Software.

Fluorescence correlation spectroscopy. FCS measurements

were performed with a Zeiss LSM 510-META/Confocor2 laser-

scanning microscope (Zeiss AG, Germany) equipped with an

argon laser (l/nm 488) or a helium/neon laser (l/nm 594) and

a 40� water-immersion objective (Zeiss C/Apochromat 40�, NA

1.2). Vesicle solutions extruded through a PC membrane (200 nm

pore diameter) with concentrations between 800 mM and 0.375

mM were incubated with solutions (50 nM) of the His-tagged

proteins at room temperature. Protein/polymer vesicle mixtures

were measured immediately and after 3 hours at room temper-

ature in special chambered quartz-glass holders (Laboratory-

Tek; 8-well, NUNC A/S), which provide optimal measurement

conditions by reducing evaporation. Fluorescence intensity

fluctuations were analyzed in terms of an autocorrelation func-

tion with the LSM 510/Confocor software package (Zeiss AG,

Germany). Each measurement of 30 s was repeated 10 times;

results are reported as the average of three independent experi-

ments. Adsorption and bleaching effects were reduced by

exchanging the sample droplet after 5 minutes of measurement.

The excitation power of the Ar laser was 15 mW, and the exci-

tation transmission at l/nm 488 was 5%. The excitation power of

the He/Ne laser was 10 mW, and the excitation transmission at



Scheme 1 Structure of a Me2+–NTA modified poly(butadiene)–poly-

(ethylene oxide) (PB–PEO) block copolymer. The anionic polymerization

was quenched at the PEO’s end to yield a semiester of succinic anhydride

(SA). The carboxy end of the semiester of the succinic anhydride was

connected to N,N-bis[(tert-butyloxycarbonyl-tert-butyl)methyl]-L-lysine

via an amide bond. tert-Butyl groups protected the carboxylic ends of the

nitrilic-tri acetic acid (NTA.p). After deprotection of the tert-butyl

groups we achieved three active carboxylic end groups: N,N-bis[(tert-

butyloxycarbonyl)methyl]-L-lysine (NTA.d), ready to complex nickel(II).

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

l/nm 594 was 3%. To reduce the number of free fitting

parameters, the diffusion times of free proteins (His6-EGFP and

His6-RFP) as well as the structure factor were determined inde-

pendently and fixed in the fitting procedure of the autocorrela-

tion function. Fluorescence autocorrelation functions were

normalized to an equal number of molecules in the confocal

volume.

Small angle X-ray scattering. SAXS measurements of polymer

solutions were performed with a SAXSess camera (Anton Paar,

Austria), using a vacuum tight quartz capillary (1 mm in diam-

eter). This camera was attached to a laboratory X-ray generator

(PW3830, PANanalytical) and was operated with a fine focus

glass X-ray tube at 40 kV and 50 mA (CuKa, l/nm 0.1542). The

scattering vector is defined in terms of the scattering angle q, and

the wavelength of the radiation thus q ¼ 4p/lsin(q). SAXS data

were recorded for 60 min in a q-range of 0.085 nm�1 to 5.0 nm�1

with a CCD detection system (Anton Paar). The two-dimen-

sional intensity data were converted to one-dimensional data and

deconvoluted (desmeared) with the software SAXSQuant 2.0

(Anton Paar). The measured intensity was corrected by sub-

tracting the intensity from the capillary filled with water or PBS

buffer. Data were fitted using a SANS Package with Igor Pro

6.0.4 (Wavemetrics), using a polydisperse core–shell and diluted

lamellar model were implemented.37–40

Langmuir isotherms. A Langmuir trough was used to obtain

a homogeneous monolayer of the amphiphilic diblock copoly-

mers at the air/water interface. For surface pressure–area (P–A)

isotherms, either a Langmuir–Blodgett minitrough (surface area/

cm2 273) or a Brewster Angle Microscope (BAM) trough (surface

area/cm2 432) were used (both from KSV Instruments Ltd.,

Helsinki, Finland; solid PTFE/Teflon troughs equipped with two

symmetrically moving hydrophilic Delrin barriers and a Wil-

helmy plate film balance).

Prior to experiments, the trough was thoroughly cleaned with

chloroform and ethanol, rinsed with water (double-distilled or

ultrapure from ELGA, resistivity 18 MU cm, pH 5.5), and filled

with the aqueous subphase based on pure water or a phosphate

buffer saline, 50 mM Na2HPO4, 10 mM KCl (Fluka AG, Buchs,

CH, puriss.), pH 7.4. The barriers were cleaned with ethanol and

rinsed with bidistilled water. The Wilhelmy plate (chromatog-

raphy paper, ashless Whatman Chr 1, perimeter 20 mm) was

equilibrated for at least 20 min. The surface was cleaned

repeatedly through compression–surface aspiration–expansion

cycles and checked for impurities. The polymer was spread

dropwise from chloroform solution (c ¼ 1 mg mL�1) on the air/

water interface. The solvent was allowed to evaporate for 10 min,

and the monolayers were compressed at the rate of 10 mm min�1.

The surface was cleaned and checked for impurities after each

measurement. All experiments were performed at 20 �C in a dust-

free room; for additional protection from impurities, the trough

was housed in a Plexiglas cabinet.

Brewster angle microscopy. BAM was performed in order to

visualize the formation of the film at the interface. A BAM2plus

Brewster angle microscope (Nanofilm Technologie GmbH,

G€ottingen, Germany) with a Nd:YAG laser operated at a wave-

length of l/nm 532, Nikon 10� Plan Epi SLWD objective (N.A.


0.30), and monochrome CCD camera attached to real-time

frame grabber was used, mounted over the Langmuir trough.

The images were captured in line scan mode and corrected for

geometry and contrast.

Langmuir–Schaeffer films on graphite. Highly oriented pyro-

litic graphite (HOPG, grade-2, Structure Probe, Inc., USA) was

used as hydrophobic substrate for film transfer and atomic force

microscopy measurements.

Langmuir–Blodgett films of PB60-PEO34-SA-NTA.d and

PB60-PEO34-SA-OH: a standard-sized LB trough (R&K, Berlin,

Germany) of 160 cm2 horizontal area and 3 mm depth was used

to prepare the LB films. The trough was equipped with a Wil-

helmy plate that allowed measuring the surface pressure of the

air–liquid interface. 40–50 mL of polymer solution (1 mg mL�1)

were deposited onto the water surface and let to equilibrate for

5 minutes. The copolymer film was compressed until a surface

pressure of 35.5 mN m�1 was reached, at which the film is in the

condensed phase (see P–A isotherms in the ESI†). Horizontal

transfer from the air–water interface to the HOPG substrate was

performed under these conditions by means of a motorized lifter

(R&K, Berlin, Germany). The substrates were detached from the

interface after 2 hours and kept immediately in water or PBS

before experiments.

Atomic force microscopy. The surface morphology of polymer

films before and after protein incubation was studied in

tapping� mode using a Multimode atomic force microscope

equipped with a Nanoscope V controller (Veeco Instruments,

Santa Barbara, CA, USA). SiN probes with oxide sharpened tips

and cantilever constants of 0.32 N m�1 (NP-S and DNP-S, Veeco

Instruments) were mounted in an enclosed liquid cell that made it

possible to perform the measurements in liquids. All substrates

were imaged in phosphate buffer before and after incubation

with protein (incubation time 1.5 h, protein concentration 0.1 mg

mL�1, 311 nM).

Results and discussion

Characterisation of metal-functionalised polymer vesicles

Polybutadiene-block-poly(ethylene oxide) copolymers function-

alized with terminal nitrilotriacetic acid (NTA) (Scheme 1) or

Soft Matter, 2010, 6, 2815–2824 | 2817


Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

tris(nitrilotriacetic acid) (trisNTA) were synthesized and com-

plexed with metals as recently described.25

Previous experiments already indicated that mixtures of NTA-

functionalized or Me–NTA-functionalized polymers and their

non-functionalized counterparts self-assemble in aqueous solu-

tion into vesicular structures that allow a selective binding of His-

tagged proteins to their surface. However, there have been no

investigations on how protein binding is influenced by variations

in the density of the metal-functionalized polymers at the surface

of the vesicles and varying environmental conditions, such as the

nature of the buffer, incubation time, etc. In order to form

vesicles in various media we used polybutadiene-block-poly-

(ethylene oxide)-succinic acid semiesters, PB-PEO-SA-OH,25

with low polydispersity (PDI 1.1), as shown in Table 1.

For vesicle preparation we mixed a solution of polybutadiene-

block-poly(ethylene oxide)-succinic acid semiester (PB39-PEO36-

SA-OH and PB60-PEO34-SA-OH, respectively) with the

corresponding Ni-NTA functionalized polymer (PB39-PEO36-

SA-trisNTA.d-Ni2+ and PB60-PEO34-SA-NTA.d-Ni2+,

respectively), at a defined mixing ratio (i.e., 10 : 1 if not stated

otherwise). As typical for the film rehydration method, the

samples initially show a broad size distribution with vesicle sizes

ranging from several micrometres down to ca. 100 nm. The size

distribution was reduced by repeated extrusion through filters of

defined pore diameter, which also provides some control to the

vesicle diameters (Fig. S1, ESI†). It was reported that PB–PEO

copolymers with wPEO > 0.35 would yield both vesicles and

worm-like micelles.41 However, in our system also for the poly-

mer with w¼ 0.42 we exclusively observed vesicles. This could be

due to the relatively low pH at which the vesicles are formed (pH

4), so that the terminal carboxylic groups are protonated.31 We

observed in cryo-TEM a minor population of vesicles attached to

worm-like potentially helically intertwined structures (Fig. S2†).

After extrusion the vesicles were characterized by DLS (Table

S1, ESI†). Interestingly, the observed vesicle hydrodynamic

radius does not depend on polymer chemical constitution (i.e.

PB39-PEO36 vs. PB60-PEO34) but rather on the extrusion

process. Indeed, in PBS buffer extrusion through 400 nm pores

yields vesicles with a hydrodynamic radius of 380 nm while

extrusion through 200 nm pores results in a hydrodynamic

radius of 118 nm. Interestingly, extrusion through smaller pore

sizes (e.g. 80 nm) leads to bimodal particle size distributions

indicating a lower limit for this process. While extrusion

through 200 nm pores seems to be insensitive towards solvent

effects, extrusion through 400 nm pores in bidistilled water

yields considerably smaller vesicles than in PBS buffer with

a hydrodynamic radius of 240 nm. This phenomenon might be

Table 1 Polybutadiene-block-poly(ethylene oxide) diblock copolymers used

Diblock copolymerGPC

1H-NMR

Nna (PB) Nn

a (PEO)

PB60-PEO34-SA-OH 60 34PB60-PEO34-SA-NTA.d 60 34PB39-PEO36-SA-OH 39 32PB39-PEO36-SA-trisNTA.d 39 36

a Nn is the number of monomer units. b wPEO is the molecular weight fractio

2818 | Soft Matter, 2010, 6, 2815–2824

due to a smaller size of the vesicles prior to extrusion in bidis-

tilled water, where no charge screening effects are to be expec-

ted. If not stated otherwise, all the following experiments were

performed with vesicles containing 10 mol% of the corre-

sponding NTA (trisNTA) functionalized polymer and extruded

through 200 nm pores.

Complementary information about the morphology and

structural details of the vesicles were obtained from SAXS

measurements, as this technique enables the determination of the

membrane thickness and inter-particle interactions. Moreover,

this technique is accurate enough to monitor changes in

membrane thickness induced by protein binding.

Fig. 1a shows the scattering curve of a solution of PB60-PEO34-

SA-OH/PB60-PEO34-SA-NTA.d-Ni2+ obtained after extrusion

through a 400 nm pore PC membrane in PBS buffer, at pH 7.4.

In the q > 0.3 nm�1 region data were fitted using a core–shell

model, assuming a constant shell thickness but a varying vesicle

radius (set to be very large). Clearly, the curve fits with q�2 and

q�4, indicative of a vesicular system, and, at the same time, the

lack of a Guinier plateau suggests an overall object size diameter

larger than 100 nm, in good agreement with DLS analysis indi-

cating a hydrodynamic radius of 380 nm. The deviations between

experimental data and the fit in the low q region can be explained

by the presence of a minor fraction of worm like structures

attached to the vesicles. A more detailed description of the

scattering curve can be found in the ESI† (Fig. S3 and Table S2).

In contrast, the scattering curve of a solution of PB39-PEO36-

SA-OH/PB39-PEO36-SA-NTA-Ni2+ vesicles (extruded through

a 400 nm membrane in bidistilled water) can be fitted using

a lamellar model in the q > 0.3 nm�1 region (Fig. 1b), as the

radius of curvature is small enough to use this approximation.

Even if fitting with a multi-lamellar model remains possible,

experimental data do not directly support it. The analysis of the

lower q region is impossible, as inter-particle interactions lead to

strongly distorted experimental curves.

SAXS analysis shows that the membrane thickness correlates

with the molar mass of the block copolymer used to form the

vesicle. Values of 8.9 � 0.2 nm and 10.2 � 0.2 nm for PB39 and

PB60-polymer membrane thickness are in good agreement with

already published data for similar PB–PEO block copolymers.42

To test the influence of the density of the Ni2+–NTA groups at

the surface of vesicles on protein binding, we prepared mixtures

of non-functionalized diblock copolymers with increasing frac-

tions of metal-NTA functionalized diblock copolymers (from 0%

to 60%, see Table 2). The overall Ni2+ content after extrusion

through PC membranes with pores of 200 nm diameter was

determined by ICP-AES spectroscopy.

for self-assembling in dilute aqueous solutions25

GPC GPC/1H-NMRwPEO

b Mw/Mnc (diblock) Mn (diblock)/g mol�1

0.32 1.1 43000.29 1.1 47500.42 1.1 35000.29 1.1 3700

n of PEO. c Mw/Mn is the polydispersity.



Fig. 2 x-Potential (a) and diameter (b) of PB60-PB34-SA-OH/xPB60-

PB34-SA-NTA.d-Ni2+ dispersions in PBS buffer, pH 7.4, extruded

through PC membranes with pores of 200 nm diameter.

Fig. 1 (a) SAXS data from PB60-PEO34-SA-OH/PB60-PEO34-SA-

NTA.d-Ni2+solutions in PBS, extruded through a 400 nm pore PC

membrane. (b) SAXS data from PB39-PEO36-SA-OH/PB39-PEO36-SA-

NTA-Ni2+ extruded through a 400 nm membrane in bidistilled water.

Plain line indicates the fit curve. The vertical line denotes the limit where

the inter-particle interactions are negligible (q > 0.3 nm�1).

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

Although the metal concentration in the vesicle solution

should increase with the concentration of Ni–NTA–polymer

(1.16 mg L�1 to 6.97 mg L�1) in the mixture, our experiments

indicate virtually no difference in the Ni2+-concentration of the

prepared vesicles. Data show that already for the samples with

10 mol% of functionalized block copolymer the membranes are

‘saturated’ with metal. We consider this to be a result of

a reduced solubility of the metal-functionalized block copolymer

compared to the corresponding non-functionalized ones in both

the organic solvent used for sample preparation and the buffers

used during film rehydration. This is supported by the fact that

samples with higher Ni–NTA polymer concentrations frequently

contained non-dissolved polymer particles prior to extrusion.

Control experiments with the ‘soluble’ part indicated that the

loss of Ni-ions during extrusion is negligible, as expected due to

the strong coordination character of NTA/trisNTA moieties.

Table 2 Metal content in different metal functionalized vesicles of PB60-PE

Sample x (%)

PB60-PEO34-SA-OH 0PB60-PEO34-SA/xPB60-PEO34-SA-NTA.d-Ni2+ 10PB60-PEO34-SA/xPB60-PEO34-SA-NTA.d-Ni2+ 15PB60-PEO34-SA/xPB60-PEO34-SA-NTA.d-Ni2+ 40PB60-PEO34-SA/xPB60-PEO34-SA-NTA.d-Ni2+ 60Blank (PBS-buffer) 0


The almost constant surface concentration of Ni–NTA is

confirmed by x-potential measurements, which show that the

extruded polymer dispersions (vesicles of PB60-PB34-SA-OH/

PB60-PB34-SA-NTA.d-Ni2+, 10 : 1), have an average negative

surface potential between �25 and �30 mV in PBS, which does

not significantly depend of the mole fraction of Ni–NTA func-

tionalized polymer in the initial block copolymer mixture.

(Fig. 2). At the same time the Ni–NTA groups do not signifi-

cantly affect the vesicle hydrodynamic radii within the experi-

mental errors, as already shown previously.25 This in turn

indicates that the vesicle coating does not modify the colloidal

properties of the vesicles.

Protein binding to the metal centers of vesicles

As a first, qualitative analysis of protein binding to Ni–NTA-

functionalized vesicles we used fluorescence microscopy on giant,

micrometre-sized vesicles as a model. As shown in Fig. 3a and b,

His-tagged red fluorescence protein (His6-RFP) binds in

bidistilled water to the surface of both PB39-PEO36-SA-/PB39-

PEO36-SA-TrisNTA.d-Ni2+, and PB39-PEO36-SA-/PB39-PEO36-

SA-TrisNTA.d-Cu2+ vesicles, respectively. Similar results were

obtained for PB60-PEO34-SA-OH/PB60-PEO34-SA-NTA.d-Ni2+

vesicles in PBS buffer at pH 7.4 upon incubation with His-tagged

enhanced green fluorescence protein (His6-EGFP). The latter is

also in good agreement with previous studies that indicated

a dissociation constant of 12.3 mM for His6-EGFP.25 It should be

noted that His6-EGFP required a significantly longer incubation

O34-SA-/xPB60-PEO34-SA-NTA.d-Ni2+ diblock copolymers

ccalc/mg L�1 cmeasured/mg L�1

0 0.011.16 0.291.74 0.304.64 0.346.97 0.310 0.04

Soft Matter, 2010, 6, 2815–2824 | 2819


Fig. 3 Fluorescence microscopy images of fluorescent protein labeled

vesicles: (a) PB39-PEO36-SA-/PB39-PEO36-SA-TrisNTA.d-Ni2+ (c/mM

800) incubated with His6-RFP in bidistilled water, (b) PB39-PEO36-SA-/

PB39-PEO36-SA-trisNTA.d-Cu2+ (c/mM 800) incubated with His6-RFP in

bidistilled water, (c) PB60-PEO34-SA-/PB60-PEO34-SA-NTA.d-Ni2+

(c/mM 200) incubated with His6-EGFP. Here bleaching of His6-EGFP

fluorescence in the course of the incubation was observed. Scale bar

is 10 mm.

Fig. 4 (a) FCS autocorrelation curves: solution of His6-RFP (50 nM) in

bidistilled water (A), solution of His6-RFP incubated with polymeric

vesicles of PB39-PEO36-SA-OH/PB39-PEO36-SA-TrisNTA-Ni2+ (B).

Curves normalized to 2 to facilitate comparison. (b) Fraction of bound

His6-RFP as function of the Ni2+ content exposed at PB39-PEO36-SA-

OH/PB39-PEO36-SA-TrisNTA-Ni2+ vesicles surface, in bidistilled water.

It should be noted that incubating His6-RFP with vesicles without Ni2+-

trisNTA groups leads to less than 3% of unspecific protein binding to the

PEO brushes at the vesicle surface; i.e., the unspecific binding of this

protein is very limited and can be safely neglected.

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

time to observe the protein binding than for His6-RFP (around

one hour, compared to 5 minutes in the case of His6-RFP).

Control experiments with vesicles without Ni-NTA groups

clearly and unambiguously show that no binding takes place in

these experiments, strongly indicating that non-specific protein

adsorption on the PEO vesicles corona is negligible under these

conditions. Moreover, these experiments clearly demonstrate

that protein binding is really due to specific interactions between

the metal centers and the oligohistidine segments of the proteins.

In order to quantify the protein binding and to evaluate the

influence of experimental parameters (e.g. incubation time, pH,

nature of the buffer), we performed fluorescence correlation

spectroscopy (FCS) experiments. FCS is known for its sensitivity

with respect to the interaction between proteins and their ligands,

which can under specific conditions achieve single-molecule

detection.43 In FCS, the laser-induced fluorescence of fluorescent

molecules in a very small probe volume is auto-correlated in time

to give information about their diffusion time. The diffusion time

is related to the hydrodynamic radius of the fluorescent mole-

cules via the Stokes–Einstein relation, and changes in diffusion

time provide information about the binding of the fluorescent

molecules to larger target molecules.44,45 In our experiment the

size difference between the free protein (His6-EGFP or His6-

RFP) and the protein bound to the surface of metal-function-

alized vesicles with diameters >100 nm allowed us to differentiate

between the two species unambiguously.

FCS measurements of PB60-PEO34-SA-/PB60-PEO34-SA-

NTA.d-Ni2+ vesicles incubated with His6-EGFP (50 nM), in

PBS-buffer, indicate the presence of two populations of objects

with diffusion times s1 ¼ 80 ms and s2 ¼ 10.5 ms. The small

diffusion time is related to the free protein still present in solu-

tion, while the high diffusion time accounts for the protein bound

to the vesicles. In order to further enhance protein binding to the

vesicles we increased the incubation time, and changed the pH in

the range between 7.0 and 8.5 (Fig. S4, ESI†). An increased

incubation time of two hours increased the fraction of His6-

EGFP protein bound to the metal centers at the surface of the

vesicles, for the whole range of pH. In addition, a pH value of 7.0

is optimum for His6-EGFP protein binding. However, with

increasing incubation time bleaching of His6-EGFP fluores-

cence46 competes with the increasing fraction of the protein

bound to vesicles, thus preventing us to obtain quantitative data

on the total amount of bound protein. As a result, the binding

2820 | Soft Matter, 2010, 6, 2815–2824

affinity of His6-EGFP we reported in PBS (a value of KD ¼ 12 �0.8 mM)25 was not significantly changed by varying the pH or the

incubation time. In contrast, the binding behavior of His6-RFP

to the PB39-PEO36-SA-/PB39-PEO36-SA-trisNTA.d-Ni2+ vesicles

in bidistilled water was completely different: most of the protein

bound immediately to the vesicles (incubation time < 5 minutes).

The best fit of the autocorrelation function of the time-dependent

fluorescence signal indicates the presence of two populations: one

with a diffusion time sd ¼ 120 ms (less than 20%), and a second

one with s2 ¼ 5.8 ms (around 80%) (Fig. 4a). The population

with the small diffusion time represents the free His6-RFP, and

the major population represents the vesicles-bound protein, as

attributed by its much longer diffusion time. The presence of the

major population representing the bound protein explains also

the intense fluorescent corona of the vesicles observed in fluo-

rescence microscopy (see Fig. 3a and b). In order to calculate the

dissociation constant for His6-RFP bound to PB39-PEO36-SA/

PB39-PEO36-SA-TrisNTA-Ni2+ vesicles in bidistilled water, we



Fig. 5 SAXS curves of PB60-PEO34-SA-OH/PB60-PEO34-SA-NTA.d-

Ni2+ vesicles in PBS buffer with (a) Rh¼ 118 nm, (b) Rh¼ 380 nm and (c)

PB39-PEO36-SA-OH/PB39-PEO36-SA-NTA-Ni2+ with Rh ¼ 118 nm in

bidistilled water. Arrows indicates scattering curves determined upon

addition of His6-EGFP for (a) and (b) and of His6-RFP for (c) to the

vesicle solution. Vertical lines point to q ¼ 0.3 nm�1, limit upon which

downward interparticles interactions preponderate. Data were modelled

using the data present above q ¼ 0.3 nm�1 and the fitting models

described in the ESI†.

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

titrated the protein (50 nM) with increasing amounts of polymer

vesicles (see details in ESI†) (Fig. 4b). The data were fitted by

a Langmuir isotherm.25,47 The dissociation constant of His6-RFP

in bidistilled water has a value of 1.99 � 0.42 mM, in agreement

with that obtained when RFP was proposed to be used as

a selective copper sensing system (KD¼ 3.6� 1.1 mM).48 This KD

value is lower due to the presence of Tris-NTA moieties, which

favor protein binding by an increased number of metal centers

exposed at the surface of vesicles, as proposed by Tampe et al. in

the case of trisNTA-functionalised lipidic membranes.49–52 In

contrast to NTA-functionalised liposomes, in our system the

coordination sites are attached to the flexible PEO blocks of

the copolymer and protein binding might be influenced by the

presence of the surrounding PEO brushes at the surface of the

polymer vesicles. Interestingly, the KD values determined for our

system were in good agreement with previous data for the NTA-

functionalised liposomes.46 This is indicating no significant effect

of the PEO brushes on the protein binding process. For the

binding of His-tagged RFP to trisNTA-modified vesicles, the

binding affinity is even higher, according to its low KD value.

Similar values have been obtained when tris-NTA was used to

bind to oligohistidine-tagged ifnar2 protein,49 indicating that

tris-NTA provides better conditions for stable metal coordina-

tion. Van Broekhoven et al. also pointed out that a protein

bound to one NTA head group is not sufficiently stable to form

the complex, again supporting the lower dissociation constant.53

Interestingly the dissociation constant for His6-RFP bound to

PB39-PEO36-SA/PB39-PEO36-SA-trisNTA-Ni2+ vesicles in water

is found to be significantly lower than previous data for His6-

EGFP bound to the same type of vesicles.25 This is due to the

intrinsic properties of His6-RFP, such as an improved fluores-

cence lifetime and increased conformational stability, which

provides better conditions for protein binding and detection.54,55

It should be noted that control measurements with vesicles

without trisNTA–Ni2+ groups showed less than 3% of protein

binding. This clearly shows that under our experimental condi-

tions non-specific protein adsorption to the PEO brushes at the

vesicle surface is negligible.

We estimated the number of proteins bound to one vesicle with

brightness measurements, during the FCS experiment. The

number of RFP bound per vesicle was determined by dividing the

value of the molecular brightness of protein-bonded vesicles

(expressed as CPM) to that of freely diffusing RFP (CPMRFP ¼2.05 kHz). A number of 24 � 8 molecules of RFP bound per

vesicle was obtained when the protein amount added and incu-

bated with the vesicles dispersion was of 50 nM (in the case of

a 97% bounded protein fraction).56

SAXS was used to monitor the binding of His6-EGFP and

His6-RFP to NTA-functionalised polymer vesicles (Table S2,

ESI†). Fig. 5 shows the scattering curve evolution upon the

addition of protein. The measured signal I(q) can in first

approximation be described as the product of a structure factor

S(q), depending on the colloidal properties, and a form factor

F(q), depending only on the object shape. Depending on the size

and solvent used to disperse the vesicles, the majority of the

objects behave differently upon protein addition, both in terms

of form factor and structure factor. This indicates that the

membrane structures became asymmetric or their respective

colloidal properties changed significantly. Indeed, for the smaller


vesicles with Rh ¼ 118 nm (Fig. 5a) in PBS buffer the membrane

thickness clearly increases from 10.2 � 0.2 nm to 12.9 � 0.2 nm

upon protein binding for both model proteins. This increase

corresponds well to the molecular dimensions of EGFP,57 even if

a precise determination of the protein orientation at the vesicle

surface is difficult, and data quality prevented a better data

analysis. For large vesicles (Fig. 5b) with Rh ¼ 380 nm the

scattering curve of the probe remains constant upon the addition

of protein as both curves virtually overlap, and the membrane

thickness appears constant within the experimental error. This is

the consequence of the weak electron density change caused by

the protein adsorption on the membrane, which in turn does not

enable seeing the membrane thickness change, as the membrane

electronic contrast sharpness that the solvent did not increase.

Although for both vesicle sizes the protein binding occurs with

identical binding affinity, their orientation and organisation at

the vesicles surface might be different thus leading to different

results. In addition, we cannot exclude the fact that larger vesicles

have a higher ‘flexibility’ and undergo stronger shape fluctua-

tions in solution. Obviously these fluctuations interfere with

protein binding and presumably also the contrast observed in the

Soft Matter, 2010, 6, 2815–2824 | 2821


Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

SAXS experiments. Further, more systematic experiments are

necessary to clarify this behaviour. Interestingly, the samples

prepared in bidistilled water (with a Rh of about 118 nm) showed

much larger changes (Fig. 5c). This is presumably because the

electrostatic interactions are suddenly screened upon protein

adsorption, as shown by the strong changes in the low q area. The

object shape did not change significantly, as shown by the limited

changes in the high q region upon protein adsorption. Never-

theless, a qualitative analysis of the SAXS data indicates that

protein adsorption at the vesicles surface takes place.

Fig. 6 Height AFM images of the copolymer (PB60-PEO34-SA-NTA.d-

Ni2) films transferred on HOPG. (a) Topology of the copolymer film; (b)

the same copolymer film after being scratched with the AFM stylus. The

thickness of the copolymer can be ascertained by computing the height

differences between the scratched and the non-scratched area. Inset:

height profile along the highlighted line 2 shown in b.

Proteins bound at surface of metal-functionalised copolymer

films

To get more structural details of the protein-decorated

membranes, we used planar block copolymer monolayers created

at the air–water interface and transferred to solid-supports as

a model. The behavior of monolayers of the amphiphilic diblock

copolymer PB60-PEO34-SA-NTA.d-Ni2+ was characterized by

surface pressure–area (p–A) isotherms. The isotherms are

reproducible and reversible (Fig. S5, ESI†). Additionally, poly-

mer organization at the air–water interface was investigated by

Brewster angle microscopy (BAM). At high surface pressures

(i.e. the range relevant for the transfer experiments) BAM indi-

cated the presence of homogeneous and smooth films. Before

Langmuir–Sch€afer transfer to a solid support (HOPG), the

polymer monolayers were compressed to the surface pressure

applied in the transfer experiments. The compressed monolayers

maintained the pressure for longer than 100 min, indicating high

film stability.

Film depositions were performed at a surface pressure of

40 mN m�1 corresponding to 80% of the collapse pressure. In this

phase, the polymer films are assumed to have the most densely

packed, brush-like order.34 Scheme 2a shows the copolymer

chelator structure as well as the strategy of monolayer prepara-

tion. A monolayer is attained if a number of block copolymer

molecules are deposited at the air–water interface and the area of

the latter is reduced via movable barriers, thus compressing the

molecules to each other. At a sufficient compression, the copoly-

mer molecules self-organize so that the PB blocks are exposed to

the air, while the PEO-NTA blocks face the aqueous medium

(Scheme 2b). The monolayer can then be transferred to

a substrate. To increase the transfer efficiency hydrophobic

substrates (in this case, highly oriented pyrolitic graphite,

HOPG) are preferentially used.

AFM on the transferred films showed a corrugated topology

that extends over micrometres, with a typical root mean square

Scheme 2 Strategy for: (a) monolayer preparation (PB60-PEO34-SA-

NTA.d-Ni2) and (b) transfer to a solid support (HOPG substrate). BDn

represents the PB block and EOm represents the PEO block.

2822 | Soft Matter, 2010, 6, 2815–2824

roughness (Rq) of 0.60 � 0.01 nm (Fig. 6a). By scratching

a reduced area within the film with the AFM stylus, it is possible

to obtain the layer thickness as the height difference between the

non-scratched region (2, in Fig. 6b) and the region where mate-

rial depletion occurs (1, in Fig. 6b). The value, 4.6 � 0.2 nm, is

close to the SAXS value for half of the membrane thickness

(5.1 � 0.2 nm). This indicates that the polymer molecules within

the film form a monolayer and adopt a similar conformation as

in the vesicle membranes.

We selected the recombinant and His6-tagged enhanced green

fluorescent protein (His6-EGFP) to test protein binding. The

EGFP is a cylindrically shaped, low-molecular weight protein

(29 kDa) whose structure is well-known: the b-sheet barrel

contains the fluorophore and is 4 nm height and 2 nm wide.

High-resolution AFM images of the copolymer films before and

after protein incubation reveal a change of topology that can be

directly related to the presence of the protein molecules on top of

the polymer layer.

A careful look at Fig. 7b can be clarifying with regard to the

orientation of the adsorbed protein. Fig. 8a shows a magnified

view where individual molecules are easily discernible. A profile

analysis along a row of molecules (light dashed line) gives us

information about the lateral molecular dimension. The average

value, 5.7 � 0.3 nm, is noticeably higher than the expected size

along the long axis of a single molecule of EGFP (4 nm).

Fig. 7 Height AFM images of the copolymer film (a) before protein

incubation (b) after protein incubation. The difference in topology can be

attributed to the presence of His6-EGFP on top of the copolymer

monolayer.



Fig. 8 (a) Magnified view of Fig. 7b where individual His6-EGFP are

discernible; (b) height profiles of a single molecule (dark dotted line) and

a row of molecules (light dashed line); (c) protein orientation in accor-

dance to the AFM results.

Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

Nonetheless, AFM images tend to overestimate lateral dimen-

sions due to the tip size, which can be three times as large as the

size of a single molecule. Vertical dimensions are not affected by

tip size, though. The profile along a single molecule (orange

dotted line) gives a value very similar to the short axis of the

EGFP (2 nm). It is thus very probable that the adsorbed mole-

cules are lying with their long axis oriented parallel to the

substrate plane.

Conclusions

We have shown that we are able to create membrane-like

structures with improved molecular recognition abilities, both in

solution and on surfaces starting from polybutadiene-block-

poly(ethylene oxide) copolymers functionalized with nitrilotri-

acetic acid and tris(nitrilotriacetic acid) complexed with metal(II).

Mixing Ni2+–NTA copolymers with their non-functionalized

analogues does not affect the self-assembly behavior of the

mixtures, i.e., they form vesicular structures in dilute aqueous

solutions, or monolayers at the air/water interface. Vesicles with

uniform distribution of metal sites and a negative surface

potential were prepared and characterized by optical micros-

copy, dynamic light scattering, small angle X-ray scattering and

x-potential measurements. Metal-functionalized polymers

monolayers were transferred via Langmuir–Schaefer technique

onto HOPG and investigated with AFM.

Accessibility and functionality of the metal site at the vesicle

surface were established with model proteins His6-EGFP and

His6-RFP. Fluorescence microscopy and FCS indicate that

proteins are selectively bound to the Me2+–NTA groups exposed

at the surface of the vesicles. Protein binding improvement was

observed upon fine tuning of the incubation conditions like pH,

buffer and binding time. Compared to His6-EGFP which showed

a moderate binding affinity, His6-RFP is characterized by a low

dissociation constant, much lower than those obtained for other

fluorescent proteins by Nehring et al.25 and Dorn et al.47 Metal-

functionalized polymer films adsorb in an oriented manner on

highly oriented pyrolytic graphite surfaces and are able to induce


the formation of densely packed protein arrays via binding to the

Ni–NTA group, as shown by AFM. This might offer new

possibilities for inducing 2D crystallization of membrane

proteins or preparing densely packed sensor devices.

We believe that these metal-functionalized polymeric

membranes have a large potential for selective immobilization

and alignment of proteins at vesicle/planar membrane surfaces,

due to their high cohesion and robustness, which make them

rather insensitive toward mechanical shear or presence of

detergents. They are thus a valuable equivalent for phospho-

lipids. Further improvements can be realized, for example by

covalently cross-linking the pendant poly(butadiene) blocks

double bonds to freeze the self-assembled structures and thus

provide additional stabilization.

Acknowledgements

Financial support was provided by the National Center of

Competence in Nanoscale Science, the Swiss National Science

Foundation NEST Projects 029084 and 043431 and MRTN-CT-

2004-005516. The authors thank Prof. Robert Tamp�e,

(University of Frankfurt/Main) for providing protected NTAs

and tris-NTAs, and David Hughes (Basel University) for reading

the manuscript. RN thanks Vimalkumar Balasubramanianam

for his strong contribution in formatting the text. AM thanks the

Adolf-Martens e.V. for an Adolf-Martens Fellowship and BAM,

Berlin for financial support.

Notes and references

1 M. P. Lutolf and J. A. Hubbell, Nat. Biotechnol., 2005, 23, 47–55.2 J. Z. H. N. A. Peppas, A. Khademhosseini and R. Langer, Adv.

Mater. (Weinheim, Ger.), 2006, 18, 1345–1360.3 H. Zhu and M. Snyder, Curr. Opin. Chem. Biol., 2001, 5, 40–45.4 B. M. Adhikari and S. Majumdar, Prog. Polym. Sci., 2004, 29, 699–

766.5 J. Homola, Chem. Rev., 2008, 108, 462–493.6 D. R. Shankaran, K. V. Gobi and N. Miura, Sens. Actuators, B, 2007,

121, 158–177.7 M. Eckert, A. Brethon, Y. X. Li, R. A. Sheldon and

I. W. C. E. Arends, Adv. Synth. Catal., 2007, 349, 2603–2609.8 M. Reetz, A. Zonta and J. Simpelkamp, Angew. Chem., Int. Ed. Engl.,

1995, 34, 301–303.9 D. Falconnet, G. Csucs, H. Michelle Grandin and M. Textor,

Biomaterials, 2006, 27, 3044–3063.10 J. T. Fink, M. Thery, A. Azioune, R. Dupont, F. Chatelain,

M. Bornens and M. Piel, Lab Chip, 2007, 7, 672–680.11 R. L. Ersson, J. C. Janson, Introduction to protein purification, John

Wiley & Sons, Inc., New York, 1998, pp. 3–40.12 M. T. W. Hearn, Physiochemical factors in polypeptide and proteins

and analysis by high performance chromatographic techniques,Academic Press, San Diego, 2000, pp. 72–235.

13 R. Gutierrez, E. M. Martin del Valle and M. A. Galan, Sep. Purif.Rev., 2007, 36, 71–111.

14 L. P. Pathange, D. R. Bevan and C. Zhang, Anal. Chem., 2008, 80,1628–1640.

15 K. R. Sticha, C. A. Sieg, C. P. Bergstrom, P. E. Hanna andC. R. Wagner, Protein Expression Purif., 1997, 10, 141–153.

16 J.-F. Chiu, X. Sun and Q.-Y. He, Methods Mol. Biol., 2008, 424, 205–212.

17 J. A. Bornhorst and J. J. Falke, Methods Enzymol., 2000, 326, 245–254.

18 Y.-C. Liu, S.-Y. Suen and C.-S. Chang, J Chromatogr., B, 2003, 797,305–313.

19 M. Zachariou, Methods Mol. Biol. (Clifton, NJ), 2004, 251, 89–102.20 W. Jiang, M. Prescott, R. J. Devenish, L. Spiccia and M. T. W. Hearn,

Biotechnol. Bioeng., 2009, 103, 747–756.

Soft Matter, 2010, 6, 2815–2824 | 2823


Dow

nloa

ded

by B

unde

sans

talt

fuer

Mat

eria

lfor

schu

ng u

nd -

prue

fung

on

02 N

ovem

ber

2010

Publ

ishe

d on

18

May

201

0 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/C

0028

38J

View Online

21 S. A. Lauer and J. P. Nolan, Cytometry, 2002, 48, 136–145.22 J. Dai, Z. Bao, L. Sun, S. U. Hong, G. L. Baker and M. L. Bruening,

Langmuir, 2006, 22, 4274–4281.23 A. Mecke, C. Dittrich and W. Meier, Soft Matter, 2006, 2, 751–

759.24 A. Taubert, A. Napoli and W. Meier, Curr. Opin. Chem. Biol., 2004, 8,

598–603.25 R. Nehring, C. G. Palivan, O. Casse, P. Tanner, J. Tuxen and

W. Meier, Langmuir, 2008, 25, 1122–1130.26 M. Antonietti and S. Foerster, Adv. Mater. (Weinheim, Ger.), 2003,

15, 1323–1333.27 M. Angelova and D. Dimitrov, Faraday Discuss. Chem. Soc., 1986,

81, 303–311.28 M. Angelova and D. Dimitrov, Prog. Colloid Polym. Sci., 1988, 76,

59–67.29 D. E. Discher and F. Ahmed, Annu. Rev. Biomed. Eng., 2006, 8, 323–

341.30 S. Yu, T. Azzam, I. Rouiller and A. Eisenberg, J. Am. Chem. Soc.,

2009, 131, 10557–10566.31 D. A. Christian, A. Tian, W. G. Ellenbroek, I. Levental,

K. Rajagopal, P. A. Janmey, A. J. Liu, T. Baumgart andD. E. Discher, Nat. Mater., 2009, 8, 843–849.

32 E. Katz and I. Willner, Angew. Chem., Int. Ed., 2004, 43, 6042–6108.33 G. MacBeath and S. L. Schreiber, Science, 2000, 289, 1760–1763.34 H. Zhu, J. F. Klemic, S. Chang, P. Bertone, A. Casamayor,

K. G. Klemic, D. Smith, M. Gerstein, M. A. Reed and M. Snyder,Nat. Genet., 2000, 26, 283–289.

35 R. D. Kornberg and S. A. Darst, Curr. Opin. Struct. Biol., 1991, 1,642–646.

36 T. Haefele, K. Kita-Tokarczyk and W. Meier, Langmuir, 2006, 22,1164–1172.

37 P. Bartlett and R. H. Ottewill, J. Chem. Phys., 1992, 96, 3306–3318.38 S. R. Kline, J. Appl. Crystallogr., 2006, 39, 895–900.39 F. Nallet, R. Laversanne and D. Roux, J. Phys. II, 1993, 3, 487–502.40 P. Mittelbach and G. Porod, Acta Phys. Austriaca, 1962, 15, 122–147.

2824 | Soft Matter, 2010, 6, 2815–2824

41 S. Jain and F. S. Bates, Science (Washington, D. C.), 2003, 300, 460–464.

42 B. M. Discher, H. Bermudez, D. A. Hammer, D. E. Discher,Y.-Y. Won and F. S. Bates, J. Phys. Chem. B, 2002, 106, 2848–2854.

43 G. Walter Nils, C.-Y. Huang, J. Manzo Anthony and A. SobhyMohamed, Nat. Methods, 2008, 5, 475–489.

44 W. Al-Soufi, B. Reija, M. Novo, S. Felekyan, R. Kuehnemuth andC. A. M. Seidel, J. Am. Chem. Soc., 2005, 127, 8775–8784.

45 P. Schwille, in Fluorescence Correlation Spectroscopy: Theory andApplications, ed. R. Rigler and E. S. Elson, Springer Ser. Chem.Phys., 2001, vol. 65.

46 D. Zhang, H. Lans, W. Vermeulen, A. Lenferink and C. Otto,Biophys. J., 2008, 95, 3439–3446.

47 I. T. Dorn, K. R. Neumaier and R. Tampe, J. Am. Chem. Soc., 1998,120, 2753–2763.

48 Y. Rahimi, S. Shrestha, T. Banerjee and S. K. Deo, Anal. Biochem.,2007, 370, 60–67.

49 T. Andre, A. Reichel, K.-H. Wiesmueller, R. Tampe, J. Piehler andR. Brock, ChemBioChem, 2009, 10, 1878–1887.

50 S. Lata, A. Reichel, R. Brock, R. Tampe and J. Piehler, J. Am. Chem.Soc., 2005, 127, 10205–10215.

51 A. Tinazli, J. Tang, R. Valiokas, S. Picuric, S. Lata, J. Piehler,B. Liedberg and R. Tampe, Chem.–Eur. J., 2005, 11, 5249–5259.

52 R. Valiokas, G. Klenkar, A. Tinazli, A. Reichel, R. Tampe, J. Piehlerand B. Liedberg, Langmuir, 2008, 24, 4959–4967.

53 C. L. van Broekhoven and J. G. Altin, Immunol. Cell Biol., 2001, 79,274–284.

54 D. Shcherbo, E. M. Merzlyak, T. V. Chepurnykh, A. F. Fradkov,G. V. Ermakova, E. A. Solovieva, K. A. Lukyanov,E. A. Bogdanova, A. G. Zaraisky, S. Lukyanov andD. M. Chudakov, Nat. Methods, 2007, 4, 741–746.

55 J. P. Sumner, N. M. Westerberg, A. K. Stoddard, C. A. Fierke andR. Kopelman, Sens. Actuators, B, 2006, 113, 760–767.

56 P. Rigler and W. Meier, J. Am. Chem. Soc., 2006, 128, 367–373.57 R. Y. Tsien, Annu. Rev. Biochem., 1998, 67, 509–544.



Protein decorated membranes by specific molecular interactions

Documents

Transcript of Protein decorated membranes by specific molecular interactions