Influence of the Spatial Distribution of Cationic Functional Groups at Nanoparticle Surfaces on Bacterial Viability and Membrane Interactions

Article abstract of DOI:10.1021/jacs.0c02737

While positively charged nanomaterials induce cytotoxicity in many organisms, much less is known about how the spatial distribution and presentation of molecular surface charge impact nanoparticle-biological interactions. We systematically functionalized diamond nanoparticle surfaces with five different cationic surface molecules having different molecular structures and conformations, including four small ligands and one polymer, and we then probed the molecular-level interaction between these nanoparticles and bacterial cells. Shewanella oneidensis MR-1 was used as a model bacterial cell system to investigate how the molecular length and conformation of cationic surface charges influence their interactions with the Gram-negative bacterial membranes. Nuclear magnetic resonance (NMR) and X-ray photoelectron spectroscopy (XPS) demonstrate the covalent modification of the nanoparticle surface with the desired cationic organic monolayers. Surprisingly, bacterial growth-based viability (GBV) and membrane damage assays both show only minimal biological impact by the NPs functionalized with short cationic ligands within the concentration range tested, yet NPs covalently linked to a cationic polymer induce strong cytotoxicity, including reduced cellular viability and significant membrane damage at the same concentration of cationic groups. Transmission electron microscopy (TEM) images of these NP-exposed bacterial cells show that NPs functionalized with cationic polymers induce significant membrane distortion and the production of outer membrane vesicle-like features, while NPs bearing short cationic ligands only exhibit weak membrane association. Our results demonstrate that the spatial distribution of molecular charge plays a key role in controlling the interaction of cationic nanoparticles with bacterial cell membranes and the subsequent biological impact. Nanoparticles functionalized with ligands having different lengths and conformations can have large differences in interactions even while having nearly identical zeta potentials. While the zeta potential is a convenient and commonly used measure of nanoparticle charge, it does not capture essential differences in molecular-level nanoparticle properties that control their biological impact.

Full text of DOI:10.1021/jacs.0c02737

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Article
Influence of spatial distribution of cationic functional groups at
nanoparticle surfaces on bacterial viability and membrane interactions
Yongqian Zhang, Natalie V. Hudson-Smith, Seth D. Frand, Meghan S Cahill,
Larissa S. Davis, Z. Vivian Feng, Christy L. Haynes, and Robert J Hamers
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.0c02737 • Publication Date (Web): 13 May 2020
Downloaded from pubs.acs.org on May 17, 2020
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Influence of spatial distribution of cationic functional groups at
nanoparticle surfaces on bacterial viability and membrane interactions
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Yongqian Zhang†, Natalie V. Hudson-Smith‡, Seth D. Frand⊥, Meghan S. Cahill‡, Larissa S. Davis†§, Z.
Vivian Feng⊥, Christy L. Haynes‡, Robert J. Hamers†*
†University of Wisconsin-Madison, Department of Chemistry, Madison, WI 53706, USA
‡University of Minnesota Twin Cities, Department of Chemistry, Minneapolis, MN 55455, USA
⊥Augsburg University, Department of Chemistry, Minneapolis, MN 55454, USA
ABSTRACT: While positively charged nanomaterials induce cytotoxicity in many organisms, much less is known about how
the spatial distribution and presentation of molecular surface charge impacts nanoparticle-biological interactions. We
systematically functionalized diamond nanoparticle surfaces with five different cationic surface molecules having different
molecular structures and conformations, including four small ligands and one polymer, and we then probed the molecular-
level interaction between these nanoparticles and bacterial cells. Shewanella oneidensis MR-1 was used as a model bacterial
cell system to investigate how molecular length and conformation of cationic surface charges influence their interactions with
the Gram-negative bacterial membranes. Nuclear magnetic resonance (NMR) and X-ray photoelectron spectroscopy (XPS)
demonstrate the covalent modification of nanoparticle surface with the desired cationic organic monolayers. Surprisingly,
bacterial growth-based viability (GBV) and membrane damage assays both show only minimal biological impact by the NPs
functionalized with short cationic ligands within the concentration range tested. Yet NPs covalently linked to a cationic
polymer induce strong cytotoxicity, including reduced cellular viability and significant membrane damage at the same
concentration of cationic groups. Transmission electron microscopy (TEM) images of these NP-exposed bacterial cells show
that NPs functionalized with cationic polymers induce significant membrane distortion and production of outer membrane
vesicles, while NPs bearing short cationic ligands exhibit only weak membrane association. Our results demonstrate that the
spatial distribution of molecular charge plays a key role in controlling the interaction of cationic nanoparticles with bacterial
cell membranes and subsequent biological impact. Nanoparticles functionalized with ligands having different lengths and
conformations can have large differences in interactions even while having nearly identical zeta potentials. While zeta
potential is a convenient and commonly used measure of nanoparticle charge, it does not capture essential differences in
molecular-level nanoparticle properties that control their biological impact.
12-15
and neutral surfaces,2,
and NPs with higher surface
INTRODUCTION
charge density lead to higher bacterial toxicity and
The surface chemistry of nanoparticles (NPs) plays a key
role in controlling the interactions of nanoparticles with
each other and with their surroundings.1-11 The intentional
functionalization of nanoparticles with molecular coatings
such as short-chain ligands or polymeric coatings provides
one approach to controlling their environmental and
biological impacts.3, 5-10
membrane permeability.16-17
While the impact of cationic surface functional groups on
nanoparticle-cell interactions is widely recognized as
important, much less is known about how the molecular
conformation of surface ligands and the resulting spatial
distribution of charge affect their resulting biological
impact. Prior studies have shown that differences in the
arrangement of surface ligands can impact the ability of
nanoparticles to penetrate cell membranes,8, 16-18 suggesting
the importance of NP surface ligand conformation to
membrane interactions. One challenge with understanding
the influence of surface molecular layers is that self-
assembled monolayers on gold and most other
nanoparticles of interest can be easily removed or displaced
under biological conditions.19-21 Nanodiamond is an
excellent platform for understand the influence of surface
Prior studies have frequently reported that nanoparticles
with cationic functional groups interact more strongly with
bacterial cells compared with those functionalized with
neutral or negatively charged groups,3,
These
5, 7, 9, 12
differences have frequently been attributed to the favorable
electrostatic interactions between positively charged NP
surfaces and negatively charged cell membranes.2-3, 10, 12 NPs
with cationic surfaces generally show higher cell-
membrane affinity and penetration compared to anionic
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Page 2 of 12
molecular groups because functionalization via an “all- Materials. All reagents were purchased from Sigma-
carbon” scaffold provides outstanding stability.22-25 In prior
work using nuclear magnetic resonance (NMR) T2
measurements, we found that when a model cationic
polymer was covalently linked to diamond nanoparticles,
approximately 45 % of the segments are highly mobile,
while the rest (~55 %) remain tightly bound on the NP
surfaces.26 In contrast, nanoparticles functionalized with
short molecular linkers must have the charged groups
closer to the surface.
Aldrich, unless noted otherwise. Nanopure water
(resistivity ≥18 MΩ ・cm, Thermo Scientific Nano-pure
system GenPure UV-TOC/UF xCAD plus) was used for all
experiments. The terminal alkenes used to functionalize
diamond nanoparticles (DNPs) in this paper are allyl
trimethylammonium bromide, tert-Butyl N-allylcarbamate
(98%), 11-trimethylammonium-1-undecene bromide and
5-hexenoic acid (98%). Allyl trimethylammonium bromide
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3
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7
8
was synthesized by
a
reported method;25 and 11-
9
trimethylammonium-1-undecene bromide was synthesized
by a modified procedure.30 See Supporting Information for
detailed synthesis procedures and 1H NMR and 13C NMR
Here, we demonstrate how the spatial distribution of
charged molecular functional groups at the surface of
diamond nanoparticles impacts their interactions with a
Gram-negative model bacterium. Figure 1 shows the five
cationic diamond NP surfaces that we prepared, using an
approach we described previously.25 To identify the
influence of primary vs. quaternary amino groups, we
prepared diamond nanoparticles (DNPs) linked via 3-
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(Supporting
trimethylammonium
trimethylammonium-1-undecene
Information,
S1).
Both
and
allyl
11-
absorb
bromide
bromide
moisture over time to form insoluble complex, thus need to
be stored in a moisture-free environment.
+
carbon chains to terminal primary amino groups (C3-NH3 -
Preparation of different positively charged diamond
nanoparticle surfaces. Detonation diamond nanoparticles
(5 nm average primary particle size, Nanostructured &
Amorphous Material Inc.) (DNPs) were used for studies
reported here. These DNPs were functionalized with five
different positively charged ligands using a radical-based
method we reported previously.25 Figure 1 shows an
overview of the functionalization process and the chemical
structures of the five different positively charged DNP
surfaces. The DNPs were heated in a tube furnace with a
flow of pure hydrogen gas (1 atm, 50 standard cm3 min-1) at
600 °C for 6 h to hydrogen-terminate the surface atoms.25, 31
The TEM micrographs of the hydrogenated- DNPs from our
previous study show an average core particle size of 5 nm
DNPs) and quaternary (tetramethylammonium) amino
+
groups (C3-N(CH3)3 -DNPs. To explore the influence of
chain length we also prepared DNPs with a C11 chain
terminated with tetramethylammonium group (C11-
+
N(CH3)3 -DNPs). To determine the influence of charges
distributed along the length of the molecules we prepared
DNPS functionalized with tetraethylenepentamine (TEPA-
DNPs). Finally, to determine the influence of ligand
conformational flexibility, we synthesized DNPS with a
covalently bonded poly(allylamine) (PAH-DNPs). We use
Shewanella oniedensis MR-1 (S. oneidensis) as a model
biological organism due to its ubiquity and importance in
the environment.27-28 Previous studies have shown S.
oneidensis to be more hardy than other Gram-negative
bacteria models, signifying that any impacts measured here
are likely to be amplified in other strains. In addition,
working with S. onidensis MR-1 allows comparison to other
studies, including those with ligands similar to those
studies here.29
Using the above surface modifications, we employed a
bacterial Live/Dead (L/D) assay, a growth-based viability
(GBV) assay, and transmission electron microscopy (TEM)
to characterize how nanoparticles functionalized with these
molecules interacted with Shewanella. Our results show
that PAH-DNPs induce high cytotoxicity by disrupting the
integrity of bacterial cell membranes. Surprisingly,
however, the nanoparticles functionalized with shorter
linear molecules all exhibited only weak association with
bacterial cell membranes and little toxicity, even though
their measured zeta potentials are similar to that of the
PAH-modified nanoparticles. While zeta potential is a
convenient and commonly used measure of nanoparticle
charge, it does not capture essential differences in
molecular-level nanoparticle properties that control their
biological impact. Our results suggest that the presence of
conformationally flexible molecular groups extending away
from the nanoparticle core and therefore able to intercalate
into the outer molecular layers of bacterial cell membranes
is a key molecular factor controlling biological impact of
cationic functionalized nanoparticles.
Figure 1. Functionalization scheme of five cationic
amine-containing monolayer binding to 5 nm-core
hydrogen-functionalized diamond nanoparticles
(DNPs). Molecules and DNPs are not drawn to
Experimental Section
scale.
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and no evidence for graphitic shells.25 The reactant solution
incubated at 30C for 4-6 h or until midlog phase. These
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8
containing benzoyl peroxide (Luperox A98, >98%), the
reactant molecule of interest, and an appropriate
anhydrous solvent were very briefly (~10 min) dried over
3 Å molecular sieves to help sequester any water present in
the benzoyl peroxide. The supernatant was removed, mixed
with hydrogen-terminated DNPs, and heated at 80 ⁰C for 3.5
h while stirring under an argon atmosphere. The solution
was cooled slowly to room temperature overnight. Details
of the conditions used for functionalizing these five surfaces
cultures were centrifuged at 750 g for 10 min and
resuspended with 1x Dulbecco’s phosphate buffered saline
(DPBS) (Corning) for 10 min. This DPBS solution was
centrifuged again at 750 g for 10 minutes and resuspended
in HEPES buffer (2 mM, pH 7.4) to obtain a concentration
with an optical density of 0.1 at 600 nm. In this study, we
normalize nanoparticle doses based on the total amine
concentration of the surrounding ligands, as described
below. In a 96-well plate, aliquots of Shewanella were
introduced, followed by addition of functionalized DNPs to
achieve amine concentrations (after dilution) of 1.24, 2.5,
5, 10, 20, 40 and 80 nM. After 1 h of exposure, 5-L aliquot
was removed from each of the 96 wells and added to 195 L
of fresh LB broth in triplicate and optical density readings
were collected with a BioTek Synergy Mx Microplate Reader
at 600 nm every 20 min over the course of 19 h at 30 C and
with medium intensity agitation for 1 min prior to each
reading. Growth curves were analyzed as described
previously.33
9
are included in Supporting information (S2).
The
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functionalized DNPs were purified by rinsing in the
corresponding reaction solvent (50 mL per 50 mg DNPs)
with sonication to ensure resuspension, followed by
centrifugation at 4480 g (gravitational constant) until the
samples fully pellet (typically 5-50 min). The washing and
centrifugation steps were repeated with water (50 mL per
50 mg DNPs, 1x). The final samples were resuspended in 1
mM HCl solution and centrifuged at 14400 g for 30 mins.
Aggregated NPs were removed, leaving behind individual
NPs and small clusters for biological studies.
Bacterial Live/Dead Assay (L/D assay). The
LIVE/DEAD® BacLightTM Viability Kit (ThermoFisher
Scientific) was used to quantify bacterial membrane
damage by different cationic DNPs. S. oneidensis
suspensions were prepared and cultured. Functionalized
DNPs were introduced into the suspensions at
concentrations identical to those used in GBV
measurements. Samples were then distributed in a 96-well
plate in at least triplicate and were exposed to a stain
consisting of a mixture of SYTO 9 dye and propidium iodide
for 15 min following the manufacturer’s recommendations.
Briefly, fluorescence measurements were performed using
a plate reader (BioTek Synergy Mx Microplate Reader) with
excitation wavelength of 485 nm and emission at 528 nm
(for SYTO9 stain) and 638 nm (for propidium iodide stain).
The ratio of fluorescence intensities at 528 nm and 638 nm,
I528/I638, was compared with that of control samples of live
and dead bacteria, yielding an average fraction of bacteria
with intact membranes (live bacteria). The bacterial
viability assay is based on growth rates of nanoparticle-
exposed bacterial populations compared with growth of
control populations. Experimental variations between
bacterial populations can yield individual measurements
with calculated viability slightly greater than 100%;
however, replicate measurements show that none of the
measurements exceed 100% viability in a statistically
significant manner.
Characterization of diamond NPs surfaces. We
measured the hydrodynamic diameters and zeta potentials
of the functionalized DNPs using dynamic light scattering
(DLS) and electrophoretic light scattering (Marvern
Zetasizer ZS). The results from these measurements is
provide in the Supporting Information (S3). In summary,
the average hydrodynamic diameter of all five types of
nanoparticles after functionalization is 41.0 nm, and the
average zeta potential is + 42.7 mV. Chemical compositions
and surface structures of all functionalized diamond
nanoparticles were characterized by both X-ray
photoelectron spectroscopy (XPS) and nuclear magnetic
resonance (NMR). X-ray photoelectron spectroscopy (XPS)
measurements were performed using Thermo Scientific K-
alpha XPS system with a micro-focused monochromated Al
K X-ray source and 180° hemispherical analyzer with a
128-channel detector using and analyzer pass energy of 50
eV. XPS data were analyzed using Casa XPS software32 and
were energy-referenced to the C(1s) peak at 284.8 eV
binding energy. Samples for XPS were prepared by drop-
casting 20 L of a dilute sample onto doped silicon wafers
(B-doped, 0.1-1 ohm·cm resistivity, Electro-Optic
Materials). NMR measurements were performed using a
Bruker Avance-600 spectrometer with TCI-F cryoprobe. To
ensure quantitative results, the recycling time (d1) was set
to 10 s, which is more than 5 times longer than the T1 of the
protons of interest. Large solvents peaks were reduced in
intensity using solvent suppression to achieve high-quality
data. DNPs samples for NMR analysis were further
centrifuged down (14100 g) to form pellets and re-
dissolved in D2O. This process replaced H2O with D2O as
much as possible and was repeated at least 3 times. The
samples were then further disaggregated with sonication.
Transmission
Electron
Microscopy
(TEM).
Transmission electron microscopy (TEM) images of S.
oneidensis MR-1 exposed to nanodiamonds were obtained
with a FEI Tecnai T12 with a 120-kV operating voltage after
resin embedding. S. oneidensis MR-1 that were exposed to
functionalized DNPs were washed in D-PBS and suspended
in HEPES buffer. The exposed bacteria suspension was
centrifuged down to a pellet, washed 3x with 0.1 M
cacodylate buffer, and fixed for 50 minutes in 2.5%
glutaraldehyde in 0.1 M sodium cacodylate buffer. Fixed cell
pellets were washed with sodium cacodylate buffer,
dehydrated with increasing concentrations of ethanol
solutions (30, 50, 70, 80, 90, 95, 100% EtOH in water), and
rinsed with propylene oxide three times. Then, the pellets
were soaked in a 2:1 propylene oxide/epoxy resin for two
Evaluation of toxicity toward bacteria: To evaluate the
toxicity of five different cationic functionalized DNPs, we
used a growth-based viability (GBV) assay.33 Shewanella
oneidensis MR-1 (S. oneidensis) were grown from stock
solution that was stored in 30% glycerol at -80 C; the
bacteria were inoculated into Luria-Bertani broth (LB) agar
plates and incubated at 30 C for 18-24 h. The resulting
bacterial colonies were inoculated into 10 mL LB broth and
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+
+
hours, 1:1 propylene oxide/epoxy resin overnight, fresh 1:1
propylene/epoxy resin for 5 hours, and then finally fixed in
pure resin for in a vacuum oven. Slices of the pellet were
microtomed with a Leica uC6 microtome for imaging.
Sections were stained with uranyl acetate and lead citrate
and placed on copper TEM grids (Ted Pella Inc) for imaging.
(Fig. 2c), C3- N(CH3)3 -DNPs (Fig. 2d) and C11- N(CH3)3 -
DNPs (Fig. 2e) show two peaks that are centered at 399.6
eV and 402.2 eV, and are consistent with the C-N nitrogen
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3
4
5
6
7
8
(399.6 eV),37 -NH3 and -N(CH3)3 nitrogen (402.2 eV,
protonated amines)34-35 moieties that are present at these
surfaces. Figures 2f-j show C(1s) spectra for all five cationic
surfaces. The peak at 284.8 eV is present in all carbon 1s
spectra and corresponds to C-C and C-H from diamond
cores and hydrocarbon chains at DNP surfaces.23 Peaks
centered at 286 eV are also observed in all C(1s) spectra and
are consistent with the C-N moieties that are present in all
these surfaces. Nitrogen moieties such as C-N and C=N at
carbon nanoparticle surfaces have been reported to appear
at approximately 285.9 ± 0.1 eV.25, 35 Additional peaks at
288.1 eV are observed in PAH-DNPs (Fig. 2f) and TEPA-
DNPs (Fig. 2g). C(1s) spectra, which we attribute to C(1s)
atoms in amide linkages38 (N-C=O) resulting from the EDC
coupling. The carbon 1s spectra show consistent results
with the nitrogen spectra for each cationic DNP surfaces,
which further verified the presence of C-N, CO-NH and -
NH2/NH3+ moieties in our samples. XPS measurements were
done and quantified independently on both Si and Au
substrates (drop-cast substrates) to verify data consistency.
The line shapes of N(1s) and C(1s) were very consistent
among measurements using different drop-case substrates
and among replicates (Supporting Information S4). The XPS
results in Figure 2 confirm that the DNP surfaces have been
functionalized with different amine moieties through
covalent bonds, yielding the structures shown in Fig. 1.
Quantitative analysis of the XPS data (Supporting
Information S5 for details) yields an average surface
coverage of 0.7 ± 0.2 molecules per nm2 for the DNPs
functionalized with linear ligands, consistent with one
+
+
Normalization of S. oneidensis responses to amine
concentration.
While
biological
responses
to
nanomaterials are frequently plotted using nanoparticle
concentrations as an experimental variable, our work is
aimed at understanding the effect of the charge amino
functional groups. The number of amino groups per
nanoparticle varies with the identity of the ligand. For
example, nanoparticles functionalized with small molecule
ligands bearing a single amino group present a smaller
number of amino groups compared with DNPs
functionalized with cationic polymers. Therefore, to
understand if this inherently higher amine content in PAH-
DNPs contributes to the significantly higher membrane
damage and cytotoxicity, we analyzed the responses of GBV
and L/D assay using the total concentration of exposed
amine groups in each sample. We used quantitative 1H NMR
to determine the total concentration of amine moieties in
each sample and normalized the GBV toxicity and L/D
membrane damage to yield the resulting biological impact
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for
a given concentration of amino groups. This
normalization allows us to compare the bio- responses to
per amine among different DNP surfaces. For quantitative
NMR analysis, three known amounts of each propylamine
(with D2O), propyl trimethylamine (with D2O), 11-
trimethylamine (with D2O), tetraethylenepentamine (with
D2O) and poly(allylamine) (with D2O) were used as external
standards, and peak integrations were compared to these
standards. Relaxation delay was set to at least 5 times the
T1 of the standards and integrated peak areas are
normalized with the number of scans and the receiver gain.
RESULTS
Characterization of functionalized DNPs. In order to
verify that the DNPs are functionalized as expected, we
characterized the functionalized DNDs using x-ray
photoelectron spectroscopy (XPS). Figures 2a-e show the
N(1s) spectra for all five different cationic surfaces:
polyallylamine hydrochloride-DNPs (PAH-DNPs, Fig. 2a),
tetraethylenepentamine-DNPs (TEPA-DNPs, Fig. 2b), C3
primary amine-DNPs (C3-NH3 -DNPs, Fig. 2c), C3
quaternary amine-DNPs (C3-N(CH3)3 -DNPs, Fig. 2d) and
C11 quaternary amine-DNPs (C11-N(CH3)3 -DNPs, Fig. 2e).
The nitrogen region for PAH-DNPs (Fig. 2f) shows two
peaks at 399 eV and 401 eV. TEPA- DNPs (Fig. 2g) has two
peaks at 399.6 eV and 401 eV. Nitrogen 1s peaks that are
centered at 399-400 eV are often assigned as non-
protonated amine moieties such as C-N, -NH2;34-35 while
nitrogen peaks that are at higher binding energies (400 eV-
402 eV) have been reported as amide bonds (-NH-CO) and
+
+
+
Figure 2. XPS spectra of N(1s) regions (a-e) and C(1s)
regions (f-j) of PAH-DNPs, TEPA-DNPs, C3-NH3+-
DNPs, C3-N(CH3)3+-DNPs and C11-N(CH3)3+-DNPs.
+
amines that are in their protonate states ( -NH3 ,
-
+
N(CH3)3 ).23, 34, 36 From our results shown in Fig. 2a-e, the
amide bond moiety at 401 eV binding energy is only
observed for PAH-DNPs and TEPA-DNPs, and is consistent
with these surface structures because the PAH and TEPA
molecules are linked to the carboxylate DNP surfaces
monolayer coverage as previously reported.25
Surface structural analysis using NMR. While XPS
analysis shows that the above procedure forms molecular
+
through amide linkage. The N(1s) spectra of C3-NH3 -DNPs
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layers with exposed amine groups as depicted in Fig. 1,
1
2
further insight into the specific chemical structures of the
1
molecular ligands upon attachment is needed. We used H
NMR to characterize the functionalized NP surfaces, as
shown in Fig. 3 and Table 1. While a detailed analysis of the
NMR data is included in the Supporting Information, several
important observations are noted here. We first note that
vinyl protons, which would appear in the 5-6 ppm region,
are not observed in any of the five spectra shown in Fig. 3.
This result demonstrates that the terminal vinyl groups of
the starting ligands have reacted in the course of above
procedure and that the ligands are attached to the DNP
surface through the vinyl terminus. Further confirmation of
the chemical structure of the surface-attached molecules is
obtained from a more detailed analysis of the spectra
(Supporting Information S6). Table 1 summarizes the
assignments and chemical species types for all peaks in the
1H NMR spectra of the five amine-functionalized DNP
surfaces. The 1H-NMR data in Fig. 3 provide detailed
structure confirmation that the five cationic surfaces have
the chemical structures depicted in Figure 1. As described
in the methods section, in order to help understand the
roles of amino groups in controlling the biological impact
and toxicity of cationic nanoparticles, we quantify the total
concentration of amino groups for each DNP surfaces by
integrating the areas of peaks characteristic of amine
moieties (labeled in c, f, g, i, j, l1-8, m, n and o). The measured
concentrations of the amino groups in each stock samples
are included in the Supporting Information, S6, Table S3. All
functionalized DNP solutions were then diluted to the same
amino group concentration for the bacterial exposure.
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Figure 3. 1H NMR spectra of a) C3-NH3+-DNPs, b) C3-
N(CH3)3+-DNPs c) C11-N(CH3)3+-DNPs, d) TEPA-DNPs
and e) PAH-DNPs. Asterisk denotes for solvent peaks.
Table 1. Peak assignment for 1H-NMR spectra of
amine-functionalized DNP surfaces.
Biological impact of cationic DNPs on S. oneidensis.
Growth-based viability and Live/Dead assay. We examined
the influence of the five different positively charged amine
functionalized- DNPs on the Gram-negative bacterium
Shewanella oneidensis MR-1 (S. oneidensis) using a growth-
based viability assay.33 Figure 4a shows viability of the S.
oneidensis after introduction of functionalized DNPs for 1 h,
reported for different concentration of surface-tethered
amino groups. None of the DNPs functionalized with small
1H 
(ppm)
0.8
DNP surfaces
Assignment
Type
-(CH2)n
a
b
+
1.2
2.03
0.07-0.15
3.1
-(CH2)n
C3-NH3
+
c
d-e
g
-CH2NH3
-(CH2)n
+
+
molecule ligands (e.g., C3-N(CH3)3 -DNPs, C3-NH3 -DNPs,
+
C11-N(CH3)3 -DNPs and TEPA-DNPs) have a significant
+
+
-N(CH3)3
C3-N(CH3)3
negative influence on the viability of S. oneidensis over the
concentration range studied; In contrast, the PAH-DNPs
show significant toxicity even at low amine concentration of
10 nM (equivalent DNPs concentration: 0.4 mg/L), and
induce 100% cell death at 20 nM amine concentration
(DNPs: 0.8 mg/L) and above. To confirm that the toxicity is
a nanoparticle surface-specific effect, we also conducted
control studies exposing S. oneidensis to the free ligands. As
shown in Fig. 4b, none of the ligands, including PAH, show
any toxicity across the same amine concentration range.
Thus, we conclude that the toxicity of PAH is only observed
when it is attached to the DNPs and is therefore a
“nanoparticle-surface-specific” effect. To understand the
specific interactions that led to the high cytotoxicity, we
employed the Live/Dead (L/D) assay to quantify the
membrane integrity of the cells.39-41 The L/D assay is a
fluorescence-based method that uses two different
+
+
-CH2N(CH3)3
3.9
1.33
f
h1-10
-(CH2)n
+
+
3.11
j
i
-N(CH3)3
C11-N(CH3)3
3.35
-CH2N(CH3)3
-(CH2)n
1.02-1.21
3.46-3.66
k1-5
l1-8
TEPA
PAH
-CH2NH
-CH2NH2
-CH2
1.17
1.59
2.67
n
o
-CH-CH2
+
m
-CH2NH3
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fluorescent dyes that bind to nucleic acid: green fluorescent (including PAH) induce significant membrane damage
SYTO® 9 and red fluorescent propidum iodide (PI) dyes.42
These two stains have different abilities to penetrate
healthy bacterial cells: the SYTO9 stain (excitation at 485
nm, emission at 528 nm) is membrane-permeant, whereas
the PI stain (excitation at 485 nm, emission at 638 nm) only
crosses damaged cell membranes. Figure 4c shows the
influence of the five cationic-DNPs on the integrity of cell
membranes of S. oneidensis, presented as ratios of
across the same concentration range, while the covalently
linked PAH polymer induces membrane damage when
attached to DNP surface. The reduced viability revealed by
the GBV assay and the membrane damage revealed by the
L/D assays demonstrate that amino groups present in an
amino-containing polymer (such as PAH) induce much
strong biological interactions compared with the same
concentration of primary or quaternary amino groups that
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Figure 4. Growth-based viability results of S. oneidensis upon exposure to all cationic diamond nanoparticles (a)
and free cationic ligands (b) (n = 4), the bacterial Live/Dead assay results assessing membrane integrity of S.
oneidensis by all cationic DNPs (c), and free cationic ligands (d) (n = 6), and (e) -potential values of all cationic
DNPs in HEPES buffer (bacterial medium) (n > 5 from technical replicates). Asterisk denotes statistical differences
among measurements using one-way ANOVA Tukey’s multiple comparison test (p < 0.05). All error bars represent
the standard deviation.
fluorescence intensity of SYTO9 to PI stain that were
exposed with NP-cells matrices. Because the signals from
SYTO9 stain represents all cells and signals from PI stain
represents only cells with damaged membranes, the
fluorescence intensity ratio of SYTO9 to PI is related to the
fraction of cells with intact membranes. The L/D assay
exhibits the same trend as the GBV assay where the four
DNPs functionalized with small-molecule ligands (C3-
N(CH3)3 -DNPs, C3-NH3 -DNPs, C11-N(CH3)3 -DNPs and
TEPA-DNPs) had almost no effect on the integrity of S.
oneidensis membrane, while PAH-DNPs induce significant
membrane damage (20% of all cells having damaged
membranes) at 10 nM amine concentration (DNPs: 0.4
mg/L) and up to 60 % membrane damage at higher amine
concentration (Fig. 4c).
are tethered more closely to the DNP surfaces (such as
linear ligands with terminal amino groups). We further note
that if we had instead presented these data as a function of
nanoparticle concentration instead of amino concentration,
the strong interactions of PAH would be even more
apparent because of the higher number of ligand density of
PAH-NDPs per nanoparticle (Supporting Information S7).
Prior studies have often correlated toxicity of nanoparticles
with the presence of a net positive charge.2, 13-15 However,
recent studies using both experiments and MD simulations
have suggested that, at high amine densities, both the total
number of charged groups and the fraction of amine in a
charged state can be less than those at intermediate
coverages, due to Coulombic interactions between
neighboring sites.43-44 We measured the zeta potential (ζ
potential) for all five cationic surfaces in both nanopure
water and HEPES buffer as this is the biological exposure
medium. Figure 4e shows that the ζ potential for the DNPs
+
+
+
The results from the L/D control studies using free
ligands (Fig. 4d) are consistent with the results of GBV
measurements, showing that none of the free ligands
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that are functionalized with linear small molecule ligands PAH-DNPs. Figure 5a-b shows TEM images of untreated S.
+
+
+
1
2
3
(C3-NH3 -DNPs, C3-N(CH3)3 -DNPs, C11-N(CH3)3 - DNPs
and TEPA-DNPs) and the DNPs that are functionalized with
oneidensis cells in bright- field imaging mode. Figure 5c-d
shows two representative TEM images of the bacterial cells
+
polymer (PAH-DNPs) are statistically identical in
ζ
after exposure to C3-N(CH3)3 -DNPs; no major membrane
potential, except for TEPA-DNPs with slightly lower ζ
potential. The previous molecular dynamics studies have
shown that when the net surface charge density is
sufficiently high, the zeta potential is only weakly
dependent on the density of surface charges, due to charge
compensation by the counter-ions. 45-47 In the present work,
charge compensation by counter-ions likely plays a role in
controlling the zeta potentials, such that the the multi-
cationic PAH shows almost identical zeta potential as the
mono-cationic molecular ligand at DNP surfaces. An
important conclusion from this work is that the ζ potential
alone is not a good parameter to predicting biological
impact of functionalized nanoparticles. Overall, the GBV
toxicity and L/D assay results show that the DNPs
functionalized with the small linear molecules do not have
any negative effect on the S. oneidensis viability or on
membrane integrity, while the PAH-functionalized DNPs
induce significant cell death and membrane damage even at
lower concentrations. These results show that neither
amine concentration nor the surface charge (measured by ζ
potential) is the key factor in this difference; rather, the
ligand conformation and spatial distribution of amino
groups must be playing important roles in controlling the
biological interactions.
distortion or disruption is visible, and the DNPs appear to
be adhered to the outer layer of the cell membranes
(indicated in red circle). In contrast, Figure 5e-f shows TEM
image of the bacterial cells after exposure to PAH- DNPs; in
this case, significant membrane distortion with spherical
protrusions are observed on the outside of the bacterial
membranes (indicated in red squares). These structures
appear similar to previously reported images of outer
membrane vesicles (OMVs).48-50 Prior work has shown that
OMVs can be formed as a stress response.51 Similarly, the
structures observed in our studies may be an indication of
stress induced by PAH-DNPs. We have included additional
TEM images at larger size in the Supporting Information S8.
In summary, the TEM images in Fig. 5 demonstrate that the
S. oneidensis cell surfaces respond much more strongly to
the NPs functionalized with the polymer PAH compared
with NPs functionalized with the linear small molecule
ligand, even when the total concentration of amine groups
is the same.
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Role of molecular structure. Although the strong
correlation between bacteria viability (Fig. 4a-b) and
membrane damage (Fig. 4c-d and 5) was expected, our
overall results are surprising in two ways. First, control
experiments using the free PAH cationic polymer show that
free PAH polymer does not induce membrane damage or
decrease in cell viability across the tested amine
concentrations. The cytotoxicity of PAH-DNPs only results
from surface-attached PAH polymers. Second, both GBV and
L/D assays show that on a “per amine” basis, each amino
group of PAH-DNPs induces significantly higher toxicity
than an equal number of amino groups on a linear small
molecule ligand. Viability measurements normalized to
Cell membrane deformation induced by polyamine
functionalized DNPs. To determine the influence of
functionalized DNPs on S. oneidensis cell membranes, we
analyzed the structure of S. oneidensis cell membranes after
exposure to functionalized DNPs using TEM. Since all four
small-molecule ligand-functionalized DNPs show similar
+
responses in GBV and L/D assay, we chose C3-N(CH3)3 -
DNPs as a representative linear ligand for comparison with
Figure 5. TEM images of (a-b) healthy S. oneidensis without exposure to cationic DNPs, (c-d) S.
oneidensis after exposure to linear molecular ligand functionalized DNPs (C3-N(CH3)3+-DNPs) and (e-f)
S. oneidensis after exposure to polymer functionalized DNPs (PAH-DNPs). TEM images were taken using
7
S. oneidensis that was treated with 80 nM amine concentration at DNP surfaces.
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amino concentration (Fig. 4a-d) show that the NMR T2 measurements showed that the PAH molecules
1
2
3
4
5
6
7
8
concentration of surface-exposed amino groups in solution
is not the key factor in this difference, nor is the zeta
potential. Dielectrophoretic light-scattering estimates the
zeta potential by measuring dielectrophoretic mobility and
exhibited two distinct populations, corresponding to sites
bonded to the surface and other sites consisting of highly
flexible polymer loops and tails. Similar conformations have
been reported for other polymers at planar and
nanoparticle surfaces.56-60 The presence of loops and tails
leads to an average charge distributed over a finite
thickness adjacent to the DNP surfaces, extending the
double-layer and increasing the hydrodynamic thickness.
These effects are reflected in the larger hydrodynamic
diameter (dh=61.6 ± 2.2 nm) of DNPs modified with PAH
compared with DNPs modified with linear ligands (dh~30
nm). Other prior studies of polymer-modified nanoparticles
have reported similar morphologies,26, 59, 61 with loops and
exposed tails extending into the adjacent liquid medium.56,
within using
a model (such as the widely used
Smoluchowski model48, 51-53 to relate the measured mobility
to a zeta potential. However, this approach assumes that
particles are hard spheres with uniform charge distribution
and does not account for lateral inhomogeneities across the
NP surfaces or for more extended distribution of charge in
the electric double layer.54-55 Consequently, measurement
of the zeta potentials of DNPs functionalized with molecules
having different morphologies (e.g., linear ligands vs.
polymer-linked) does not adequately represent the
differences in spatial distribution of charge. Our results
show that the spatial distribution of charge plays a key role
in nanoparticle-membrane interactions. The main
difference between DNPs functionalized with linear ligands
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58, 60
We propose that these loops and tails are largely
responsible for the interaction of the PAH-NPs with S.
oneidensis cell membranes. Specifically, we attribute the
differential toxicity of PAH-ND and linear small molecule-
functionalized DNPs to differences in the radial extent of the
charged groups. Prior work29 using molecular dynamics
simulations has demonstrated that PAH binds to the LPS
layer in Gram-negative bacteria through electrostatic
interactions and eventually migrates to the core region of
the LPS. Therefore, we propose that it is the larger radial
extent of the loops and tails of the surface-linked PAH
polymer enables penetration of the PAH (and its associated
amino groups) into the lipopolysaccharide (LPS) layer at
the surface of the Gram-negative S. oneidensis cell
membrane. The positive charges allows the cationic NPs to
bind to the surface of the bacterial membrane through
electrostatic interactions, and the conformationally flexible
PAH loops and tails assist physical insertion into the LPS
layer.
+
(such as C3-NH3 -DNPs) and those modified with covalent-
bonded polymers (PAH-DNPs) is the conformation of the
surface molecules. Figure
6 represents a schematic
illustration of functionalized DNPs interacting with a model
LPS layer. For DNPs that are functionalized with short linear
molecules, the charges are located in a thin shell at the
periphery of the molecular monolayer, as shown in Fig. 6a.
In contrast, DNPs that are functionalized with polymers
have charges that extend further from the nanoparticle
surface. In prior work, we characterized the molecular
conformations of DNPs covalently bonded to PAH.26 Our
CONCLUSIONS
These studies demonstrate that while cationic charged
groups can enhance nanoparticle interaction with cellular
membranes, the detailed molecular structure of the
molecules, which controls the spatial distribution and
presentation of charged groups, plays a key role. While
nanoparticle charges are frequently characterized using the
zeta potential, this single parameter does not fully capture
the essential molecular aspects that control the
nanoparticle interactions.
Our work indicates that
biological impact of cationic nanoparticles is greatest when
the molecular functional groups extend away from the
nanoparticle core in a flexible manner that can facilitate
penetration of the molecules into the outermost molecular
layers of the bacterial membrane. This work provides new
perspectives on why cationic surface layers induce
membrane damage in bacterial cells, and will facilitate
further understanding of the fundamental mechanisms of
cell membrane interactions of engineered NPs.
Figure 6. Conceptual illustration depicting spatial
distribution of charge of DNPs functionalized with a) linear
molecular ligands and b) covalently linked polymer
interacting with a model LPS layer. While short molecular
ligands interact only with the outermost regions of the LPS
layers of the cellular membrane, loops and tails of the
polymer can insert into the layers, leading to membrane
disruption. Gray spheres represent C atoms; red O atoms;
blue N atoms, purple P atoms of LPS. Positive charges
correspond to amino groups; negative charges to
phosphate groups of LPS.
ASSOCIATED CONTENT
Supporting Information
Procedures of 11-trimethylamine-1-undecene bromide
synthesis
and
characterization,
details
of
DNPs
functionalization conditions, hydrodynamic size and zeta
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Journal of the American Chemical Society
1.
Klaine, S. J.; Alvarez, P. J. J.; Batley, G. E.; Fernandes, T. F.;
potential analysis of functionalized DNPs, XPS quantification of
Handy, R. D.; Lyon, D. Y.; Mahendra, S.; McLaughlin, M. J.; Lead, J. R.,
Nanomaterials in the environment: Behavior, fate, bioavailability,
and effects. Environ. Toxicol. Chem. 2008, 27, 1825-1851.
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functionalized-DNPs, biological responses plot of S. oneidensis
versus NP concentration and additional TEM images of NP-
exposed S.oneidensis.
2.
nanoparticle–cell interactions. Small 2010, 6, 12-21.
3. Fleischer, C. C.; Payne, C. K., Nanoparticle Surface Charge
Verma, A.; Stellacci, F., Effect of surface properties on
The Supporting Information is available free of charge on the
ACS Publications website.
Mediates the Cellular Receptors Used by Protein-Nanoparticle
Complexes. J. Phys. Chem. B 2012, 116, 8901-8907.
4.
Albanese, A.; Tang, P. S.; Chan, W. C. W., The effect of
AUTHOR INFORMATION
nanoparticle size, shape, and surface chemistry on biological
systems. Ann. Rev. Biomed. Engr. 2012, 14, 1-16.
9
Corresponding Author
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60
5.
Cho, E. C.; Xie, J. W.; Wurm, P. A.; Xia, Y. N., Understanding
* Robert J. Hamers, rjhamers@wisc.edu
the role of surface charges in cellular adsorption versus
internalization by selectively removing gold nanoparticles on the
cell surface with a I2/KI etchant. Nano Lett. 2009, 9, 1080-1084.
ORCID
6.
Setyawati, M. I.; Mochalin, V. N.; Leong, D. T., Tuning
Yongqian Zhang: 0000-0001-9637-5314
Natalie V. Hudson-Smith: 0000-0002-2642-0711
Meghan Cahill: 0000-0002-1514-7625
Z. Vivian Feng: 0000-0002-3329-3781
Christy L. Haynes: 0000-0002-5420-5867
Robert J. Hamers: 0000-0003-3821-9625
endothelial permeability with functionalized nanodiamonds. ACS
Nano 2016, 10, 1170-1181.
7.
Silva, T.; Pokhrel, L. R.; Dubey, B.; Tolaymat, T. M.; Maier,
K. J.; Liu, X. F., Particle size, surface charge and concentration
dependent ecotoxicity of three organo-coated silver nanoparticles:
Comparison between general linear model-predicted and
observed toxicity. Sci. Total Environ. 2014, 468, 968-976.
8.
Verma, A.; Uzun, O.; Hu, Y.; Hu, Y.; Han, H.-S.; Watson, N.;
Chen, S.; Irvine, D. J.; Stellacci, F., Surface-structure-regulated cell-
membrane penetration by monolayer-protected nanoparticles.
Nat. Mater. 2008, 7, 588.
Present Addresses
§ Department of Chemistry, Lawrence University, Appleton,
Wisconsin, 54911, United States.
9.
Verma, A.; Arshad, F.; Ahmad, K.; Goswami, U.; Samanta,
S. K.; Sahoo, A. K.; Sk, M. P., Role of surface charge in enhancing
antibacterial activity of fluorescent carbon dots. Nanotechnol.
2020, 31, 9.
Author Contributions
The manuscript was written through contributions of all
authors.
10.
Huang, K.; Hu, Y.; Yu, C. J.; Boerhan, R.; Jiang, G. Q.,
Charged surface groups of nanoparticles and the adsorbed
proteins codetermine the fate of nanoparticles upon interacting
with cells. Rsc Advances 2016, 6, 58315-58324.
Funding Sources
National Science Foundation, CHE-1503408
National Science Foundation, 00039202
National Science Foundation, DMR-1720415
National Institutes of Health, S10OD012245
11.
Mensch, A. C.; Hernandez, R. T.; Kuether, J. E.; Torelli, M.
D.; Feng, Z. V.; Hamers, R. J.; Pedersen, J. A., Natural organic matter
concentration impacts the interaction of functionalized diamond
nanoparticles with model and actual bacterial membranes.
Environmental science & technology 2017, 51, 11075-11084.
ACKNOWLEDGMENT
12.
Feng, Z. V.; Gunsolus, I. L.; Qiu, T. A.; Hurley, K. R.; Nyberg,
This work was supported by the National Science Foundation
under the Center for Sustainable Nanotechnology, CHE-
1503408. The CSN is part of the Centers for Chemical
Innovation Program. N. Hudson-Smith acknowledges support
through the National Science Foundation Graduate Research
Fellowship Program (00039202). XPS studies used facilities
and instrumentation supported by NSF through the University
of Wisconsin Materials Research Science and Engineering
Center (DMR-1720415). The Bruker Avance 600 NMR
instrument used in this work was supported by the National
Institutes of Health grants S10OD012245. Parts of this work
were carried out in the Characterization Facility, University of
Minnesota, which receives partial support from NSF through
the MRSEC program.
L. H.; Frew, H.; Johnson, K. P.; Vartanian, A. M.; Jacob, L. M.; Lohse,
S. E., Impacts of gold nanoparticle charge and ligand type on
surface binding and toxicity to Gram-negative and Gram-positive
bacteria. Chem. Sci. 2015, 6, 5186-5196.
13.
Goodman, C. M.; McCusker, C. D.; Yilmaz, T.; Rotello, V. M.,
Toxicity of gold nanoparticles functionalized with cationic and
anionic side chains. Bioconjugate chemistry 2004, 15, 897-900.
14.
Arvizo, R. R.; Miranda, O. R.; Thompson, M. A.; Pabelick,
C. M.; Bhattacharya, R.; Robertson, J. D.; Rotello, V. M.; Prakash, Y.;
Mukherjee, P., Effect of nanoparticle surface charge at the plasma
membrane and beyond. Nano Lett. 2010, 10, 2543-2548.
15.
Fleischer, C. C.; Payne, C. K., Nanoparticle–cell
interactions: molecular structure of the protein corona and cellular
outcomes. Accounts of chemical research 2014, 47, 2651-2659.
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Hong, S.; Leroueil, P. R.; Janus, E. K.; Peters, J. L.; Kober,
M.-M.; Islam, M. T.; Orr, B. G.; Baker Jr, J. R.; Banaszak Holl, M. M.,
Interaction of polycationic polymers with supported lipid bilayers
and cells: nanoscale hole formation and enhanced membrane
permeability. Bioconjugate chemistry 2006, 17, 728-734.
ABBREVIATIONS
DNPs, diamond nanoparticles; XPS, x-ray photoelectron
spectroscopy; NMR, nuclear magnetic resonance; PAH,
poly(allylamine hydrochloride)
17.
Giljohann, D. A.; Seferos, D. S.; Patel, P. C.; Millstone, J. E.;
Rosi, N. L.; Mirkin, C. A., Oligonucleotide loading determines
cellular uptake of DNA-modified gold nanoparticles. Nano Lett.
2007, 7, 3818-3821.
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Herbig, M. E.; Assi, F.; Textor, M.; Merkle, H. P., The cell
penetrating peptides pVEC and W2-pVEC induce transformation of
gel phase domains in phospholipid bilayers without affecting their
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