Freezing or wrapping, the role of particle size behind the
mechanism of nanoparticle − biomembrane interaction
Shengwen Zhang, Andrew Nelson,* Paul A. Beales*
Centre for Molecular Nanoscience, School of Chemistry, University of Leeds, Leeds, UK.
A.L.Nelson@leeds.ac.uk; P.A.Beales@leeds.ac.uk
KEYWORDS
nanoparticle, giant unilamellar vesicles, supported monolayer, membrane interactions, lipid lateral
mobility, electrochemical impedance spectroscopy
ABSTRACT
Understanding the interactions between nanoparticles (NPs) and biological matter is a high priority
research area due to the importance of elucidating the physical mechanisms underlying the interactions
leading to NP potential toxicity as well as NP viability as therapeutic vectors in nanomedicine. Here, we
use two model membrane systems, giant unilamellar vesicles (GUVs) and supported monolayers, to
demonstrate the competition between adhesion and elastic energy at the nano-bio interface, leading to
different mechanisms of NPs − membrane interaction relating to NP size. Small NPs (18 nm) cause a
‘freeze-effect’ of otherwise fluid phospholipids, significantly decreasing phospholipid lateral mobility.
The release of tension through stress-induced fracture mechanics results in a single microsize hole in the
1
GUVs after interaction. Large particles (>78 nm) promote membrane-wrapping, which leads to an
increased lipid lateral mobility and the eventual collapse of the vesicles. Electrochemical impedance
spectroscopy on the supported monolayer model confirms that different sized NPs interact differently
with the phospholipids within close vicinity of the electrode during the lipid desorption process. The
timescale of these processes are in accordance with the proposed NP/GUV interaction mechanism.
2
1. Introduction
With the advance of nanoparticle (NP) engineering, intensive investigations into their potential
industrial and biomedical applications have been initiated.1-3 Various studies using NPs to assist the
intracellular delivery and controlled release of drug molecules have shown encouraging results.4,5
However, significant safety concerns have arisen, with supporting findings from cytotoxicity studies.2, 6,
7
Thus, the issue addressing how NPs interact with the biomembrane has become a pressing concern.1, 2
Whether NPs are seen as the therapeutic solution to enhanced drug delivery or a potential health threat,
the mechanisms behind the NP-biomembrane interactions are yet to be fully established.
The complexities of cell membranes and the diverse properties of NPs make it extremely challenging
to study the interactions between the plasma membrane and NPs. This work takes a reductionist
approach to study the interactions between spherical SiO2 NPs with narrow size distributions and
dioleoyl phosphatidylcholine (DOPC) model membranes at physiological pH. It has been reported that
SiO2 NPs adsorb on the biomembrane surface8 and SiO2 NPs across a broad range of particle sizes
perturb the permeability barrier of biomembranes, even at very low concentrations.9 Two
complementary model membrane systems, giant unilamellar vesicles (GUVs) and supported
monolayers, are used in this work. Free-standing bilayer GUVs are of a similar size and morphology to
native cells and facilitate microscale imaging and analysis,10-12 while supported monolayers allow
detailed electrochemical investigations at the molecular level.13 DOPC is used as the sole component for
the biomembranes, eliminating possible complexities arising from compositionally-distinct domains in
multi-component mixtures. DOPC has a melting temperature (Tm) ≈ −20 °C, hence is in a fluid,
physiologically relevant state at room temperature. Moreover, Hg supported DOPC monolayers have
been reported to undergo a series of well-defined and reversible electric field induced reorientations
(Figure SI-1).13-16 Monitoring how these structural transitions are influenced by extraneous stimuli gives
insight into the interactions at this nano-bio interface.
3
2. Experimental Section
2.1. Materials
DOPC and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl)
(18:1 Liss Rhod PE) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL). Dextran, Alexa
Fluor 647, 10,000 MW was obtained from Invitrogen Molecular Probes. Colloidal SiO 2 NPs (LUDOX
SM-30) with 18 nm (dia) were purchased from Sigma Aldrich Ltd. 30 wt% suspension in water.
Colloidal SiO2 microspheres of 78 nm were purchased from Polyscience, Inc. as 5 wt% aqueous
dispersion. 180 nm AngstromSphere SiO2 particles were purchased from Fiber Optic Center Inc. as dry
form and dispersed in water according to the technique provided by the supplier. The particle size was
measured in both 100 mM KCl + 10 mM PBS and 165 mM NaCl + 10 mM HEPES electrolyte at a
particle concentration of 500 g mL-1 (Zetasizer Nano S, Malvern Instruments Ltd.). The SiO2 colloidal
systems are stable for at least 3 days.
2.2. Methods
Confocal laser scanning microscopy and fluorescence recovery after photobleaching (FRAP)
measurements were conducted on DOPC GUVs. GUVs were prepared by the electroformation
technique in a 300 mM sucrose solution described previously from DOPC with 0.5 mol % fluorophore
(Liss Rhod PE).12 The experiment was conducted at room temperature. Confocal laser scanning inverted
microscope Zeiss LSM T-PMT/LSM700 and Zeiss ZEN software were used for the experiments. Glass
bottom culture dishes (MatTek), pretreated with 10 % BSA solution to prevent GUVs adhering to the
glass,
were
used
for
microscopy.
Microscope
buffer
(10
mM
4-(2-hydroxyethyl)-1-
piperazineethanesulfonic acid + 165 mM NaCl) of pH 7.4 was added such that GUVs sedimented to the
bottom of sample and sat on cover slip. The FRAP data fitting was performed with IGOR Pro
(Wavemetrics, Inc).
4
Electrochemical studies were performed on DOPC monolayers supported on mercury film electrodes
(MFEs). The fabrication of the wafer based MFEs had been described previously.14, 17, 18 All potentials
in this paper were quoted versus the potential of a Ag/AgCl:3.5 mol dm–3 KCl reference electrode
separated from the electrolyte by a porous glass frit and a Pt counter electrode (Metrohm UK Ltd.).
Supported DOPC monolayers on MFEs were developed by initially spreading 13 μL of a 2.54 mmol
dm−3 solution of DOPC in pentane (HPLC grade, Fisher Scientific Chemicals Ltd.) at the
argon|electrolyte interface in the electrochemical cell, followed by slowly lowering the MFE through the
phospholipid on the electrolyte interface.14,
19, 20
The integrity of the monolayer was confirmed with
rapid cyclic voltammetry between −0.2 and −1.6 at 40 V s–1.14 The KCl (0.1 mol dm–3) electrolyte was
prepared from Analar KCl (Fisher Chemicals Ltd.) calcined at 600 °C for 5 h and dissolved in 18.2 MΩ
Milli-Q water with added 0.01 mol dm–3 phosphate buffer (pH 7.4).
The impedance spectroscopy measurements were carried out by applying the logarithmically
distributed frequencies between 65000 and 0.1 Hz at −0.35 V, the position of zero charge (PZC) for Hg,
and −1.32 V (the DOPC desorption potential) using Autolab (Ecochemie, Utrecht, Netherlands).14, 19, 20
An AC amplitude of 0.002 V was applied during the measurements. The impedance data obtained were
then transformed with EXCEL (Microsoft) to the complex capacitance plane, plotting C’ vs –C” in the
unit of F cm-2. All the electrochemical measurements were conducted in a Faraday cage. At the
desorption potential, the RC time constants were obtained from the impedance data: RC = 1/(2f1), with
f1 as the characteristic frequency for this process where the imaginary capacitance reaches a maximum
value. The electrode capacitance, C, can then be calculated from C = RC / Ru, of which Ru is the
uncompensated resistance. The second relaxation process of DOPC desorption with and without the
presence of NPs can be analyzed in a similar manner, except for the adsorption of 182 nm particles, of
which  is calculated from an impedance model developed for this system. 14, 19, 20
3. Results and Discussion
5
3.1. NP-biomembrane interactions on GUVs
Morphology of the GUVs is studied by confocal microscopy. The morphology changes of GUVs
after interacting with SiO2 NPs are remarkably dependent on the size of NPs. Significantly, 18 nm NPs
create unusual crinkles and permanent holes in GUVs (Figure 1a-d), transforming the previously
smooth, spherical GUVs into crumpled ‘paper bags’ with microscale openings. These unusual holes and
curvatures on the GUVs are also shown in 2D (Figure 1d). 10 kDa fluorescent dextran was added to
these samples and was observed to immediately enter deformed GUVs, confirming that these are indeed
holes, not dye-excluding lipid domains. In rare events, the spherical GUV buckles into a ‘helmet’
morphology without forming a micropore (Figure 1c). Nonetheless, crumpled, ‘pot’-shaped vesicles
with a single micropore are more commonly observed. In direct contrast, the outcome of the interaction
between GUVs and 182 nm particles is dramatically different. During the initial interaction, wrapping
events can be clearly observed on the surface of GUVs (Figure 1e). These wrapping events deplete the
lipids, and eventually lead to GUV breakdown.
This striking size-dependence of the NP-biomembrane interaction can also be observed in their
distinctly different effects on lipid lateral mobility. Fluorescence recovery after photobleaching (FRAP)
is employed to investigate the modifications in lipid lateral mobility.21 Diffusion coefficients, D, are
calculated based on the mobility of lipid-based fluorescent probes within the bilayer of GUVs.
Normalized fluorescence recovery curves are presented in Figure 2. The FRAP data are fitted to a
classic fluorescence recovery model:22, 23
f(t) = A × {exp(−2τD/t) × [I0(2τD/t) + I1(2τD/t)]}
(1)
in which t is time, A is the recovery level, τD is characteristic recovery time, I0 and I1 are modified
Bessel functions of the first kind. Diffusion is calculated from D = w2/4τD (w is the radius of the circular
bleached area).22
6
Figure 1. Confocal laser scanning microscopy of DOPC GUVs after NPs interactions: (a-d) GUVs after
20 min interaction with 18 nm SiO2: (a, b) reconstituted 3D images of GUVs with unusual curvature and
stabilized holes; (c) a helmet shaped GUV; (d) 2D confocal image of deformed GUVs and dextran
fluorescence leakage after interaction with 18 nm 25 μg mL-1 SiO2 NPs; (e) 3D reconstruction of a GUV
7
interacting with 182 nm SiO2 NPs; (f) schematic view of our interpretation of the effect of size on the
NP-membrane interactions: small NPs adsorb on to the membrane (top) and the membrane wraps the
large NPs (bottom). Depth profile is shown in Figure a, b, c, and e. Insets to c: 2D and 3D view of the
same GUV; insets to d, additional deformed vesicles from the same experiment; inset to e: 2D view of a
GUV undergoing a wrapping event.
Figure 2. Representative FRAP recovery curves of DOPC in the presence of 18 nm SiO2, 78 nm SiO2,
and 182 nm SiO2 with particle concentration of 500 μg mL-1. Recovery curves are fit to equation (1) for
each data set presented (red lines). Inset presents the initial recoveries and the fittings.
The good fits of this model to our data in Figure 2 suggest that the intramembrane molecular
dynamics of DOPC with and without the presence of NPs are well described by a single characteristic
diffusion coefficient. Attempts to fit the fluorescence recovery using two coexisting diffusion
coefficients within the membrane did not improve the fits. This confirms that, on microscopic length
scales, the GUV bilayers display uniform lipid dynamics, suggesting that the membrane consists of a
single homogeneous phase on these length scales. Mean values of diffusion coefficient are listed in
Table 1. Typical recovery level is between 90 – 95%, which we attribute to bleaching of the
fluorophores.
8
Table 1. Mean diffusion coefficients of DOPC in GUVs before and after the interactions with SiO2 NPs
(datasets ranging between 6 to 12 GUVs per measurement).
Sample NP concentration
D (μm2 s-1)
S.D. ±
GUV
0
3.10
0.34
18 nm
5 g mL-1
3.12
0.64
25 g mL-1
0.61
0.72
50 g mL-1
0.18
0.12
500 g mL-1
0.25
0.14
50 g mL-1
3.01
0.37
500 g mL-1
5.64
1.80
50 g mL-1
2.66
0.36
500 g mL-1
7.14
2.40
78 nm
182 nm
We find that small NPs substantially decrease lipid mobility, while larger NPs lead to a discernible
increase in lateral diffusion in the membrane. The average lateral diffusion coefficient (D) in DOPC
GUVs is 3.1 ± 0.34 μm2 s-1 (Table 1) and is consistent with literature values.24,
25
We observe a
negligible change in D when the GUVs are exposed to SiO2 NPs at low concentrations, e.g. 5 μg mL-1
for 18 nm NPs, 50 μg mL-1 for 78 nm and 182 nm particles. Significantly, D decreases when increasing
the concentration of 18 nm SiO2 NPs. This decrease is also dependent on incubation time, suggesting a
progressive interaction, and is ~45 times lower after 60 minutes’ interaction with 18 nm NPs (Figure SI2). This progressive decrease in lipid mobility suggests that the ‘freeze-effect’ does not proceed by a
sharp first order phase transition as would be expected for a thermodynamic fluid-to-gel phase
transformation in a membrane crossing its Tm. Notably, D increases after exposing DOPC membranes to
high doses of 182 nm and 78 nm NPs. The increased mobility is attributed to defects in the GUV
9
membrane created by the wrapping mechanism. Moreover, the substantially larger SD suggests that
these disorganized lipids are far from equilibrium.
3.2. NP-biomembrane interactions on supported monolayers
Electrochemical impedance spectroscopy (EIS) is a non-invasive, yet extremely sensitive method to
study interactions at the lipid interface. In this work, EIS is applied to corroborate the interactions of the
DOPC assemblies with SiO2 particles and to gain further insight into the mechanisms at the molecular
level. In this classical model of a DOPC monolayer supported on a Hg electrode, the configuration of
the assembled phospholipids can be varied by altering the applied electric field (Figure SI-1). Close to
the PZC of Hg13 an intact DOPC monolayer exists with its apolar tail groups physically associated with
the apolar Hg surface14,
15, 17, 20
resembling the outer leaflet of phospholipid bilayer vesicles. Such a
supported monolayer can be electrically represented by a simple RC circuit of solution resistance R and
monolayer capacitance, C, in series. Following an EIS measurement, this circuit gives rise to a single
RC semi-circle in the complex capacitance plot19 with a specific capacitance of ~1.8 μF cm–2 (Figure
3a.1). Some impedance terminologies are illustrated in Figure SI-3.
Primary EIS results suggest that SiO2 particles of all sizes adsorb on supported monolayers but do not
alter the dielectric constant or thickness of the DOPC monolayer. The monolayer capacitance is not
significantly modified after interaction with each of the three SiO2 NP samples (Figure 3.a.1), as is also
confirmed by AC voltammetry results (Figure SI-4). However, additional and identical capacitive
elements arise at low frequencies after DOPC interaction with all sizes of NPs (Figure 3a.2), showing a
similar structural interference of adsorbed NPs with a well-oriented phospholipid monolayer.8, 19, 20 The
high affinity of DOPC for Hg compared to SiO2 constrains the lipids to the Hg electrode and therefore
allows us to separate the initial NP adsorption process from subsequent structural changes in the lipid
layer induced by NP – DOPC interactions. This can be clearly demonstrated by quantitative comparison
of relative adhesion energies: at the PZC of Hg, DOPC has a spreading pressure, or adhesion energy, of
10
52  10-3 J m-2,13 much greater than the adhesion energy between SiO2 and phosphatidylcholine, which
has been measured as 0.5 − 1 10-3 J m-2.26 Thus particles of all sizes adsorb on the DOPC layer on Hg
at this potential.
Crucially, the size-dependence of the NP-biomembrane interaction becomes evident near the lipid
desorption potential, where the lipids can leave the Hg electrode and freely interact with the NPs in
solution. Close to the lipid desorption potential, the surface pressure, or adhesion energy, of DOPC on
Hg approaches zero.13 This allows the lipids to leave the Hg surface and to interact freely with the NPs,
yet still physically remain in the vicinity of the electrode.15 At high frequencies, the lipids (and NPs) are
outside the double-layer of the electrode, giving an extrapolated characteristic capacitance of ~20 μF
cm-2, typical for an uncoated Hg electrode (Figure 3b.4).13,
14
At low frequencies, adsorbed NPs
substantially alter the secondary low frequency relaxation process and this modification is dependent on
the NP diameter (Figure 3b). The secondary relaxations of the impedance profile represent the
adsorption-desorption of charged species into and out of the electrode double-layer13, 20 coupled with the
fluctuating electric field. Time constants, τ, for this movement are presented in Figure 3b.4. In the
absence of NPs, τ is associated with the movement of desorbed DOPC assemblies, which are formed
when the DOPC bilayer structures leave the electrode surface. Notably, τ is 11 times smaller in the
presence of 18 nm NPs, suggesting that small NPs adsorbed on DOPC stabilize the SiO2-lipid bilayer
transition state within the Debye length of the electrode, hence facilitating adsorption-desorption of
DOPC. In contrast, τ is two orders of magnitude larger in the presence of 182 nm particles, indicating a
hindered adsorption-desorption process, where desorbed phospholipids wrap around large particles
moving away from the electrode surface. These results are consistent with our findings from the GUV
experiments indicating that desorbed DOPC bilayer structures wrap around the larger particles and form
hybrid organic-inorganic structures away from the electrode surface. The extended mechanism of
bilayer wrapping hinders the lipid adsorption-desorption process in the presence of the electric field.
11
Figure 3. Impedance spectroscopy presented in complex-capacitance plane of supported DOPC
monolayer on Hg in 0.1 mol dm–3 KCl with 0.01 mol dm–3 PBS buffer pH 7.4 before and after 50
minutes’ interaction with SiO2 NPs of 500 μg mL-1 at (a) -0.35 V with schematic view presented in a.2;
12
(b) -1.32 V with a schematic view of the interactions of b.1: DOPC; b.2: DOPC with 18 nm SiO 2; b.3:
DOPC with 182 nm SiO2; b.4: summary of time constant (black dot) and extrapolated zero frequency
capacitance (blue triangle) with error bar marking the SD from a set of 3 experiment results.
In general, particle size, shape, charge density, surface chemistry and crystallinity determine how
particles interact with biomembranes.3 In this current system, we interpret our observations to arise from
the balance between adhesion and elastic energies at the membrane surface.27 Quantitative analysis
shows that the bending energy barrier for DOPC membranes around 18 nm NPs is two orders of
magnitude larger than that for 182 nm NPs (Supporting Information – 5). Thus, the energy required for
membrane bending is the dominant barrier when particles are small,27,
28
suppressing membrane
wrapping.29 Adhesion arises from Van der Waals forces and electrostatic interaction between the
negative charges (circa 2 µC cm-2)30 on the SiO2 surface and the positively charged region of the DOPC
P−–N+ dipole, which is believed to alter the tilt angle of the head group, condensing the surface area per
lipid.31 This increased lipid packing density reduces lateral mobility, resulting in a rigid membrane with
high lateral tension, i.e. a ‘freeze-effect’. This tension is eventually released through stress-induced
fracture mechanics, which creates a single microsize hole in the membrane. In a recent study with
binary GUVs, solidification of membranes has been shown to cause pores in the presence of surfactants
that minimize the line tension created at the rim of the pore.32 Therefore we suggest that small SiO2 NPs
may also function as ‘line-actant’ to stabilize holes within biomembranes. Since the adhesion energy
between SiO2 and phosphatidylcholine membranes is in the range of 0.5 − 1 10-3 J m-2,26 we can
estimate that bending and adhesive energies equate for a particle size in the range 28 − 40 nm
(Supporting Information – 5). For 182 nm particles (clearly outside this range), wrapping is
energetically more favorable, as observed by fluorescence microscopy and supported by electrochemical
results. NPs of 78 nm diameter interact with DOPC GUVs in a similar, but less aggressive manner. A
13
moderate increase of lipid lateral mobility gradually followed by GUV breakdown is observed, which is
also attributed to the defects in the GUVs created by the wrapping mechanism.
A phospholipid matrix forms the major part of the plasma membrane, implying NP-phospholipid
interactions alone can compromise its barrier function.9 Indeed, recent work with lung alveolar
carcinoma cells suggests non-endocytotic uptake of the very same 18 nm SiO2 NPs at 4 °C, a
temperature at which active membrane trafficking through ATP driven endocytosis is inhibited.33
Therefore, the fundamental NP-biomembrane interaction mechanisms we report here are still applicable
to complex biomembrane systems and biological cells, and will facilitate further work in both
nanomedicine and nanotoxicology.
Conclusions
With highly consistent observations at microscopic and molecular length scales, we demonstrate the
size-dependent interaction mechanisms between NPs and biomembrane models. Small NPs cause a
‘freeze-effect’ of otherwise fluid phospholipids, significantly decreasing phospholipid lateral mobility.
The tension created is released through stress-induced fracture mechanics resulting in micro-sized
opening in GUVs. Large particles promote membrane-wrapping, which leads to moderate increases of
lipid lateral mobility and the eventual collapse of the vesicles. The size-dependent interaction
mechanisms are confirmed electrochemically using the same phospholipids within close vicinity of the
electrode during the lipid desorption process. The size-dependent mechanisms of NPs − membrane
interaction are attributed to the competition between adhesion and elastic energy at the nano-bio
interface.
ASSOCIATED CONTENT
SUPPORTING INFORMATION
14
Details of electrochemical data analysis and supplementary information on the model membrane
systems are available in Supporting Information. This material is available free of charge via the
Internet at http://pubs.acs.org
AUTHOR INFORMATION
Corresponding Authors:
* Tel: +44 113 343 6409. Fax: +44 113 343 6452. E-mail: A.L.Nelson@leeds.ac.uk
* Tel: +44 113 343 9101. Fax: +44 113 343 6452. E-mail: P.A.Beales@leeds.ac.uk
ACKNOWLEDGMENT
A.N. and S.Z. thank the Brian Mercer Award for Innovation (2008) from the Royal Society (U.K.) for
funding this work. P.B. thanks the Biomedical and Health Research Centre (BHRC) in Leeds for
funding and support. This work is also supported by the ENNSATOX programme funded by EU FP7
under grant agreement no. NMP-229244.
ABBREVIATIONS
DOPC, dioleoyl phosphatidylcholine; EIS, electrochemical I mpedance spectroscopy; FRAP,
fluorescence recovery after photobleaching; GUV, giant unilamellar vesicles; MFE, mercury film
electrode; NP, nanoparticles; PZC, position of zero charge.
REFERENCES
1.
Nel, A.; Xia, T.; Madler, L.; Li, N., Toxic Potential of Materials at the Nanolevel. Science 2006,
311, (5761), 622-627.
15
2.
Yamashita, K.; Yoshioka, Y.; Higashisaka, K.; Mimura, K.; Morishita, Y.; Nozaki, M.; Yoshida,
T.; Ogura, T.; Nabeshi, H.; Nagano, K.; Abe, Y.; Kamada, H.; Monobe, Y.; Imazawa, T.; Aoshima, H.;
Shishido, K.; Kawai, Y.; Mayumi, T.; Tsunoda, S.-i.; Itoh, N.; Yoshikawa, T.; Yanagihara, I.; Saito, S.;
Tsutsumi, Y., Silica and titanium dioxide nanoparticles cause pregnancy complications in mice. Nat
Nano 2011, 6, (5), 321-328.
3.
Nel, A. E.; Madler, L.; Velegol, D.; Xia, T.; Hoek, E. M. V.; Somasundaran, P.; Klaessig, F.;
Castranova, V.; Thompson, M., Understanding biophysicochemical interactions at the nano-bio
interface. Nat Mater 2009, 8, (7), 543-557.
4.
Vivero-Escoto, J. L.; Slowing, I. I.; Wu, C.-W.; Lin, V. S. Y., Photoinduced Intracellular
Controlled Release Drug Delivery in Human Cells by Gold-Capped Mesoporous Silica Nanosphere.
Journal of the American Chemical Society 2009, 131, (10), 3462-3463.
5.
Bernardos, A.; Mondragon, L.; Aznar, E.; Marcos, M. D.; Martinez-Manez, R.; Sancenon, F.;
Soto, J.; Barat, J. M.; Perez-Paya, E.; Guillem, C.; Amoros, P., Enzyme-Responsive Intracellular
Controlled Release Using Nanometric Silica Mesoporous Supports Capped with “Saccharides”. ACS
Nano 2010, 4, (11), 6353-6368.
6.
Napierska, D.; Thomassen, L. C. J.; Rabolli, V.; Lison, D.; Gonzalez, L.; Kirsch-Volders, M.;
Martens, J. A.; Hoet, P. H., Size-Dependent Cytotoxicity of Monodisperse Silica Nanoparticles in
Human Endothelial Cells. Small 2009, 5, (7), 846-853.
7.
Tao, Z.; Toms, B. B.; Goodisman, J.; Asefa, T., Mesoporosity and Functional Group Dependent
Endocytosis and Cytotoxicity of Silica Nanomaterials. Chemical Research in Toxicology 2009, 22, (11),
1869-1880.
8.
Vakourov, A.; Brydson, R.; Nelson, A., Electrochemical modeling of silica nanoparticle-
biomembrane interaction. Langmuir 2011, accepted manuscript.
16
9.
de Planque, M. R. R.; Aghdaei, S.; Roose, T.; Morgan, H., Electrophysiological Characterization
of Membrane Disruption by Nanoparticles. ACS Nano 2011, 5, (5), 3599-3606.
10. Beales, P. A.; Bergstrom, C. L.; Geerts, N.; Groves, J. T.; Vanderlick, T. K., Single Vesicle
Observations of the Cardiolipin-Cytochrome c Interaction: Induction of Membrane Morphology
Changes. Langmuir 2011, 27, (10), 6107-6115.
11. Beales, P. A.; Vanderlick, T. K., Specific Binding of Different Vesicle Populations by the
Hybridization of Membrane-Anchored DNA. The Journal of Physical Chemistry A 2007, 111, (49),
12372-12380.
12. Beales, P. A.; Vanderlick, T. K., Partitioning of Membrane-Anchored DNA between Coexisting
Lipid Phases. The Journal of Physical Chemistry B 2009, 113, (42), 13678-13686.
13. Bizzotto, D.; Nelson, A., Continuing Electrochemical Studies of Phospholipid Monolayers of
Dioleoyl Phosphatidylcholine at the Mercury‚àíElectrolyte Interface. Langmuir 1998, 14, (21), 62696273.
14. Zhang, S.; Nelson, A.; Coldrick, Z.; Chen, R., The Effects of Substituent Grafting on the
Interaction of pH-Responsive Polymers with Phospholipid Monolayers. Langmuir 2011, 27, (13), 85308539.
15. Nelson, A., Electrochemical analysis of a phospholipid phase transition. Journal of
Electroanalytical Chemistry 2007, 601, (1-2), 83-93.
16. Brukhno, A. V.; Akinshina, A.; Coldrick, Z.; Nelson, A.; Auer, S., Phase phenomena in
supported lipid films under varying electric potential. Soft Matter 2011, 7, (3), 1006-1017.
17
17. Coldrick, Z.; Steenson, P.; Millner, P.; Davies, M.; Nelson, A., Phospholipid monolayer coated
microfabricated electrodes to model the interaction of molecules with biomembranes. Electrochimica
Acta 2009, 54, (22), 4954-4962.
18. Coldrick, Z.; Penezic, A.; Gasparovic, B.; Steenson, P.; Merrifield, J.; Nelson, A., High
throughput systems for screening biomembrane interactions on fabricated mercury film electrodes.
Journal of Applied Electrochemistry 2011, 41, (8), 939-949.
19. Whitehouse, C.; O'Flanagan, R.; Lindholm-Sethson, B.; Movaghar, B.; Nelson, A., Application
of Electrochemical Impedance Spectroscopy to the Study of Dioleoyl Phosphatidylcholine Monolayers
on Mercury. Langmuir 2003, 20, (1), 136-144.
20. Protopapa, E.; Maude, S.; Aggeli, A.; Nelson, A., Interaction of Self-Assembling B-Sheet
Peptides with Phospholipid Monolayers: The Role of Aggregation State, Polarity, Charge and Applied
Field. Langmuir 2009, 25, (5), 3289-3296.
21. Nam, J.; Beales, P. A.; Vanderlick, T. K., Giant Phospholipid/Block Copolymer Hybrid
Vesicles: Mixing Behavior and Domain Formation. Langmuir 2010, 27, (1), 1-6.
22. Soumpasis, D. M., Theoretical analysis of fluorescence photobleaching recovery experiments.
Journal of Biophysical Society 1983, 41, (1), 95-97.
23. Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. W., Mobility measurement by
analysis of fluorescence photobleaching recovery kinetics. Biophysical Journal 1976, 16, (9), 10551069.
24. Fahey, P. F.; Webb, W. W., Lateral diffusion in phospholipid bilayer membranes and
multilamellar liquid crystals. Biochemistry 1978, 17, (15), 3046-3053.
18
25. Baksh, M. M.; Jaros, M.; Groves, J. T., Detection of molecular interactions at membrane
surfaces through colloid phase transitions. Nature 2004, 427, (6970), 139-141.
26. Anderson, T. H.; Min, Y.; Weirich, K. L.; Zeng, H.; Fygenson, D.; Israelachvili, J. N.,
Formation of Supported Bilayers on Silica Substrates. Langmuir 2009, 25, (12), 6997-7005.
27. Deserno, M., Elastic deformation of a fluid membrane upon colloid binding. Physical Review E
2004, 69, (3), 031903.
28. Ginzburg, V. V.; Balijepalli, S., Modeling the Thermodynamics of the Interaction of
Nanoparticles with Cell Membranes. Nano Letters 2007, 7, (12), 3716-3722.
29. Roiter, Y.; Ornatska, M.; Rammohan, A. R.; Balakrishnan, J.; Heine, D. R.; Minko, S.,
Interaction of Nanoparticles with Lipid Membrane. Nano Letters 2008, 8, (3), 941-944.
30. Sonnefeld, J., Determination of Surface Charge Density Constants for Spherical Silica Particles
Using a Linear Transformation. J. Colloid Interface Sci. 1996, 183, (2), 932-938.
31. Wang, B.; Zhang, L.; Bae, S. C.; Granick, S., Nanoparticle-induced surface reconstruction of
phospholipid membranes. Proceedings of the National Academy of Sciences 2008, 105, (47), 1817118175.
32. Sakuma, Y.; Taniguchi, T.; Imai, M., Pore Formation in a Binary Giant Vesicle Induced by
Cone-Shaped Lipids. Biophysical Journal 2010, 99, (2), 472-479.
33. Mu, Q.; Hondow, N.; Krzeminski, L.; Brown, A.; Jeuken, L.; Routledge, M., Mechanism of
cellular uptake of genotoxic silica nanoparticles. Manuscript in preparation.
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